<p>This AOP evaluates one of the major processes known to be involved in regulating efficient mucociliary clearance (MCC)—ciliary function. MCC is a key aspect of the innate immune defense against airborne pathogens and inhaled chemicals and is governed by the concerted action of its functional components, the cilia and the airway surface liquid (ASL), which is composed of mucus and periciliary layers (Bustamante-Marin and Ostrowski, 2017). Disturbances in any of the processes regulating ciliary function can cause MCC dysfunction. Impaired MCC is linked to airway diseases such as chronic obstructive pulmonary disease (COPD) or asthma, both of which are characterized by decreased lung function and bear a significant risk of increased morbidity and mortality. </p>
<p>We propose here an AOP that attempts to delineate how exposure to oxidative insults lead to decreased lung function (Luettich et al., 2021). This AOP evaluates one of the major processes known to be involved in regulating efficient mucociliary clearance (MCC)—ciliary function. MCC is a key aspect of the innate immune defense against airborne pathogens and inhaled chemicals and is governed by the concerted action of its functional components, the cilia and the airway surface liquid (ASL), which is composed of mucus and periciliary layers (Bustamante-Marin and Ostrowski, 2017). Disturbances in any of the processes regulating ciliary function can cause MCC dysfunction. Impaired MCC is linked to airway diseases such as chronic obstructive pulmonary disease (COPD) or asthma, both of which are characterized by decreased lung function and bear a significant risk of increased morbidity and mortality. Given the individual and public health burden of the consequences of lung function impairment, gaining a greater understanding of the underlying mechanisms is extremely important in the risk assessment of respiratory toxicants.</p>
<p>The KE proposed here are moderately to highly essential, and we judge the overall biological plausibility of this AOP as strong. The KER <em>Oxidative stress leading to decreased CBF</em> is supported by multiple studies across different species with ample empirical evidence reflecting both dose-response and time concordance. The KER <em>Decreased CBF leading to decreased MCC</em> lacks this expanse of empirical evidence, or the evidence does not fully support the causality between the KE even though the relationship is logical and plausible. Overall, our quantitative understanding of the AOP is moderate.</p>
</div>
<div id="background">
<h3>Background</h3>
<p>With a surface area of ~100 m<sup>2</sup> and ventilated by 10,000 to 20,000 liters of air per day (National Research Council, 1988; Frohlich et al., 2016), the lungs are a major barrier that protect the body from a host of external factors that enter the respiratory system and may cause lung pathologies. Mucociliary clearance (MCC) is a key aspect of the innate immune defense against airborne pathogens and inhaled particles and is governed by the concerted action of its functional components, the cilia and the airway surface liquid (ASL), which comprises mucus and the periciliary layer (Bustamante-Marin and Ostrowski, 2017). In healthy subjects, ≥10 mL airway secretions are continuously produced and transported daily by the mucociliary escalator. Disturbances in any of the processes regulating ASL volume, mucus production, mucus viscoelastic properties, or ciliary function can cause MCC dysfunction and are linked to airway diseases such as chronic obstructive pulmonary disease (COPD) or asthma, both of which bear a significant risk of increased morbidity and mortality. The mechanism by which exposure to inhaled toxicants might lead to mucus hypersecretion and thereby impact pulmonary function has already been mapped in AOP148 on decreased lung function. However, whether an exposure-related decline in lung function is solely related to excessive production of mucus is debatable, particularly in light of the close relationship between mucus, ciliary function, and efficient MCC. To date, no single event has been attributed to MCC impairment, and it is likely that events described in this AOP as well as in AOP148 have to culminate to lead to decreased lung function.</p>
<p>With a surface area of ~100 m<sup>2</sup> and ventilated by 10,000 to 20,000 liters of air per day (National Research Council, 1988; Frohlich et al., 2016), the lungs are a major barrier that protect the body from a host of external factors that enter the respiratory system and may cause lung pathologies. Mucociliary clearance (MCC) is a key aspect of the innate immune defense against airborne pathogens and inhaled particles and is governed by the concerted action of its functional components, the cilia and the airway surface liquid (ASL), which comprises mucus and the periciliary layer (Bustamante-Marin and Ostrowski, 2017). In healthy subjects, ≥10 mL airway secretions are continuously produced and transported daily by the mucociliary escalator. Disturbances in any of the processes regulating ASL volume, mucus production, mucus viscoelastic properties, or ciliary function can cause MCC dysfunction and are linked to airway diseases such as chronic obstructive pulmonary disease (COPD) or asthma, both of which bear a significant risk of increased morbidity and mortality. The mechanism by which exposure to inhaled toxicants might lead to mucus hypersecretion and thereby impact pulmonary function has already been mapped in AOP148 on decreased lung function. However, whether an exposure-related decline in lung function is solely related to excessive production of mucus is debatable, particularly in light of the close relationship between mucus, ciliary function, and efficient MCC. To date, no single event has been attributed to MCC impairment, and it is likely that events described in this AOP as well as in AOP148, AOP424, and AOP425 have to culminate to lead to decreased lung function.</p>
<p>Acrolein, a ubiquitous environmental pollutant, is a highly reactive unsaturated aldehyde that exerts toxicity through several mechanism, including oxidative stress (Moghe et al., 2015). Acrolein exposure decreased CFTR-mediated Cl− transport in primary murine nasal septal epithelia, in human bronchial epithelial cells grown in monolayers and in human Calu-3 lung cancer cells (Alexander et al., 2012; Raju et al., 2013), transiently reduced CBF at low concentrations (0.5‒1 mM) and induced ciliostasis at high concentrations (> 1 mM) in rabbit tracheal epithelial cells (Romet et al., 1990), and significantly increased mucin production in rats (Chen et al., 2013; Liu et al., 2009; Borchers et al., 1998; Wang et al., 2009). In addition, exposure of Fischer rats to acrolein caused a left shift in the quasi-static compliance curves and increased lung volumes, indicative of airway obstruction (Costa et al., 1986).</p>
<h4>Ozone</h4>
<p>Tracheas of Wistar rats exposed to 1.5 ppm ozone for 1 h/day for 3 days exhibited reduced CFTR protein expression. Similarly, at 4 hours following a 30-min exposure to ozone, CFTR mRNA and protein were down-regulated in 16HBE14o- cells. At 24 hours post-exposure, a reduction in forskolin-stimulated CFTR Cl− conductance was observed (Qu et al., 2009).</p>
<p>Continuous, exposure of human nasal epithelial cells to different concentrations of ozone at 37°C for up to 4 weeks slightly (but not significantly) reduced CBF in healthy mucosa (7.1% at 500 µg/m<sup>3</sup> and 10.3% at 1000 µg/m<sup>3</sup>), and significantly in chronically inflamed mucosa (20.5/16.4%) at 2 weeks. During the third and fourth week of exposure at these higher concentrations CBF was significantly reduced in both healthy (after 3 weeks: 18.7/37.5%; after 4 weeks: 11.1/33.3%) and chronically inflamed mucosa (after 3 weeks: 33.8/26.8%; after 4 weeks: 21.4/38.6%). Low ozone concentrations (100 µg/m<sup>3</sup>) appeared to not have an effect on CBF (Gosepath et al., 2000).</p>
<p>Acute exposure (2 h) of adult ewes to 1.0 ppm ozone significantly reduced tracheal mucus transport velocity (TMV) at 40 min and 2 h post-exposure. Repeated exposure to 1.0 ppm ozone for 5 hper day, for 4 consecutive days showed a progressively significant decrease in TMV on the first and second days, and stabilized over the third and fourth days, around values ranging from -42% to -55% of the initial baseline. TMV remained depressed even after the end of exposure, persisting up to 5 days post-exposure (Allegra et al., 1991).</p>
<p>Acute exposure of healthy young adult subjects (aged 19 to 35 years, non-smokers) to 0.06 ppm ozone for 6.6 h resulted in a 1.71 + 0.50% (mean + SEM) decrease in FEV1 and a 2.32 + 0.41% decrease in FVC compared with air exposure (Kim et al., 2011).</p>
<p>A US-based study found inverse associations between increasing lifetime exposure to ozone (estimated median: 36 ppm; interquartile range 29–45 ppm; range 19–64 ppm) and FEF75 and FEF25–75 in adolescents (aged 18–20 years) (Tager et al., 2005).<br />
</p>
<h4>Cigarette smoke</h4>
<p>CFTR transcript and protein levels were reduced in human Calu-3 lung cancer cells exposed to the gas phase of cigarette smoke (Cantin et al., 2006b), human immortalized bronchial epithelial 16HBE14o- cells treated with 10% cigarette smoke extract (Hassan et al., 2014; Rasmussen et al., 2014; Xu et al., 2015), differentiated primary human bronchial epithelial cells exposed to whole cigarette smoke (Sloane et al., 2012; Hassan et al., 2014), and in airways of smokers compared to non-smokers (Dransfield et al., 2013). Following exposure to cigarette smoke, Cl<sup>−</sup> conductance (i.e., CFTR-mediated Cl<sup>−</sup> transport) decreased in primary human bronchial epithelial cells grown in monolayers (Lambert et al., 2014), differentiated primary human bronchial epithelial cells (Schmid et al., 2015; Chinnapaiyan et al., 2018), and nasal respiratory and intestinal epithelia of A/J mice (Raju et al., 2013; Raju et al., 2017).<br />
In the lower airways, healthy smokers and smokers with chronic obstructive pulmonary disease (COPD) showed reduced CFTR-dependent Cl<sup>−</sup> transport, whereas COPD former smokers showed an intermediate response to chloride-free isoproterenol solution compared to non-smokers. Similarly, amiloride-sensitive lower airway potential difference was also lower in healthy smokers and COPD smokers than in healthy non-smokers. This was linked to reduced CFTR protein levels in the airways of smokers compared to non-smokers, although there were no significant differences between healthy and COPD subjects (Dransfield et al., 2013). CFTR-dependent Cl<sup>−</sup> conductance as measured by nasal potential difference was also significantly reduced in healthy and COPD smokers compared to healthy non-smokers or to former smokers with COPD (Sloane et al., 2012). In addition, healthy never-smokers had higher mean sweat chloride concentrations than COPD smokers and COPD former smokers (Raju et al., 2013; Courville et al., 2014).</p>
<p>Multiple studies showed that exposure of primary human bronchial epithelial cells, either undifferentiated or differentiated at the air-liquid interface, to cigarette smoke decreased ASL height (Hassan et al., 2014; Lambert et al., 2014; Raju et al., 2016; Rasmussen et al., 2014; Schmid et al., 2015). Treatment of immortalized bronchial epithelial 16HBE14o- cells with 10% cigarette smoke extract for 48 hours also resulted in a significant reduction in ASL height (Xu et al., 2015).</p>
<p>Treatment of human sinonasal epithelial cells with cigarette smoke condensate significantly reduced forskolin-stimulated CBF (Cohen et al., 2009). CBF was also decreased in differentiated normal human bronchial epithelial cells exposed to whole cigarette smoke (Schmid et al., 2015), in cilia-bearing explant adenoid tissues treated with 5 and 10% cigarette smoke extract (Wang et al., 2012), in hamster oviducts treated with various mainstream cigarette smoke fractions (Knoll et al., 1995), and in nasal epithelial cells fom smokers with moderate and severe chronic obstructive pulmonary disease (COPD) (Yaghi et al., 2012).</p>
<p>Whole cigarette smoke exposure or treatment with cigarette smoke extract of normal human bronchial epithelial cells significantly lowered FoxJ1 mRNA and protein levels (Milara et al., 2012; Brekman et al., 2014; Valencia-Gattas et al., 2016; Ishikawa and Ito, 2017). Cigarette smoke extract treatment of normal human bronchial epithelial cells also reduced the expression of cilia-related transcription factor genes, including FOXJ1, RFX2, and RFX3, as well as that of cilia motility and structural integrity genes regulated by FOXJ1, including DNAI1, DNAH5, DNAH9, DNAH10, DNAH11, and SPAG6 (Brekman et al., 2014).</p>
<p>Exposure of human bronchial epithelial cells cultured at the air-liquid interface to cigarette smoke extract during differentiation significantly shortened the average cilia length compared to untreated cultures, and treatment of differentiated cultures prevented elongation of cilia seen in untreated cultures<br />
(Brekman et al., 2014). Whole smoke exposure of mouse tracheal epithelial cells differentiated at the air-liquid interface resulted in cilia shortening and also complete loss of cilia at 24 h post-exposure (Lam et al., 2013). Cilia length was also reduced in mouse nasal septal epithelial cells treated with cigarette smoke condensate (Tamashiro et al., 2009). Cilia length was reduced in endobronchial biopsies and airway brushings of smokers compared to nonsmokers (Leopold et al., 2009) and in COPD smokers compared to healthy smokers and nonsmokers (Hessel et al., 2014). In adults with adults with chronic sputum production, current and former smokers had a higher frequency of axonemal ultrastructural abnormalities than non-smokers and controls (Verra et al., 1994).</p>
<p>Nasomuciliary clearance time (determined by saccharin transit test) was significantly higher in smokers than in non-smokers and correlated positively with cigarettes per day and packs/year index (Proença et al., 2011; Baby et al., 2014; Yadav et al., 2014; Habesoglu et al., 2012; Pagliuca et al., 2015; Xavier et al., 2013; Dülger et al., 2018; Solak et al., 2018; Polosa et al., 2021).</p>
<p>Smoking decreased pulmonary function including forced vital capacity (FVC), forced expiratory volume in one second (FEV1) and FEF25–75 (Kuperman and Riker, 1973; Ashley et al., 1975, Tantisuwat and Thaveeratitham, 2014, Gold et al., 1996; Broekema et al., 2009).</p>
<h4>Nitrogen dioxide</h4>
<p>Exposure of human bronchial epithelial cells to 100, 400, and 800 ppb NO<sub>2</sub> decreased CBF by 2.3±1.7% , 3.0±2.5%, and 6.8±1.7% respectively, which was not significant relative to incubator controls. Exposure of the cells to 2000 ppb NO<sub>2</sub> significantly decreased CBF by 14.2±2.5% in comparison with controls (Devalia et al., 1993).</p>
<h4>Diesel engine exhaust</h4>
<p>Incubation of human primary bronchial epithelial cells differentiated at the air-liquid interface with Diesel exhaust particles (DEP; 100 µg/mL = 16.26 ng/mL phenanthrene, 3.65 ng/mL fluoranthene, 2.53 ng/mL pyrene) attenuated CBF in a time- and dose-dependent manner. Exposure to 10 µg/mL DEP decreased CBF by 40% (Q1 = 19, Q3 = 46) from baseline after 24-h incubation. Similarly, exposure to 50 µg/mL DEP, filtered DEP solution, or 100 µg/mL DEP decreased CBF by 51% (Q1 = 49, Q3 = 56), 33% (Q1 = 26, Q3 = 36), and 73% (Q1 = 65, Q3 = 83), respectively, from baseline after 24-h incubation. Changes in CBF started to become significant at 4 h with 50 µg/mL DEP and at 2 h with 100 µg/mL DEP compared to untreated cultures (Bayram et al., 1998).</p>
</div>
<div id="overall_assessment">
<h2>Overall Assessment of the AOP</h2>
<p>The experimental evidence to support the biological plausibility of the KERs from MIE to AO is moderate to strong overall for the AOP presented here, while there is a moderate concordance of dose-response relationships. The weakest evidence is for the KER of decreased CFTR function leading to decreased ASL height (KERC), due to both insufficient experimental evidence of causality and a scarcity of quantitative data on dose-related responses and temporal concordances. In terms of essentiality, we have rated all of the KEs as either moderate or high.</p>
<p>The experimental evidence to support the biological plausibility of the KERs from MIE to AO is moderate to strong overall for the AOP presented here, while there is a moderate concordance of dose-response relationships. In terms of essentiality, we have rated all of the KEs as either moderate or high.</p>
<p>AOPs such as this one can play a central role in risk assessment strategies for a wide variety of regulatory purposes by providing mechanistic support to an integrated approach to testing and assessment (IATA; (Clippinger et al., 2018)). IATAs are flexible frameworks that can be adapted to best address the regulatory question or purpose at hand. More specifically, this AOP can be applied to the risk assessment of inhaled toxicants, by enabling the development of testing strategies through the assembly of existing information and the generation of new data where they are currently lacking. Targeted approaches to fill data gaps can be developed using new approach methodologies (NAMs) informed by this AOP.</p>
<p>All KE proposed in this AOP network occur and are measurable in several species, including frogs, mice, rats, guinea pigs, ferrets, cats, dogs, cows, monkeys, and humans. The majority of the supporting empirical evidence derives from studies in rodent and human systems, and experimental findings in animals appear to be highly translatable to humans.</p>
<p>All KE proposed in this AOP occur and are measurable in several species, including frogs, mice, rats, guinea pigs, ferrets, cats, dogs, cows, monkeys, and humans. The majority of the supporting empirical evidence derives from studies in rodent and human systems, and experimental findings in animals appear to be highly translatable to humans.</p>
<p>Data regarding the applicability of KE to all life-stages from birth to adulthood are available for the MIE (Oxidative Stress), KE2 (Cilia Beat Frequency, Decreased), KE3 (Mucociliary Clearance, Decreased), and AO (Decreased Lung Function), and indicate that they apply to all life stages. It is also worth noting here that age-dependent decreases in CBF, MCC, and lung function have been demonstrated in several species (e.g., guinea pigs, mice, and humans) and reflect normal physiological aging processes (Bailey et al., 2014; Grubb et al., 2016; Ho et al., 2001; Joki and Saano, 1997; Paul et al., 2013; Sharma and Goodwin, 2006).</p>
<p>Data regarding the applicability of KE to all life-stages from birth to adulthood are available for the MIE (Oxidative Stress), KE1908 (Cilia Beat Frequency, Decreased), KE1909 (Mucociliary Clearance, Decreased), and AO (Decreased Lung Function), and indicate that they apply to all life stages. It is also worth noting here that age-dependent decreases in CBF, MCC, and lung function have been demonstrated in several species (e.g., guinea pigs, mice, and humans) and reflect normal physiological aging processes (Bailey et al., 2014; Grubb et al., 2016; Ho et al., 2001; Joki and Saano, 1997; Paul et al., 2013; Sharma and Goodwin, 2006).</p>
<p>Gender-specific data relevant to the AOP network are not as widely available as species-specific data, and to our knowledge, the role of gender has not been systematically evaluated for all KE described here. Informative evidence on gender differences stems from patients with chronic pulmonary diseases, such as cystic fibrosis, asthma, COPD, and bronchiectasis, that are characterized by decreased lung function. Considering the importance of efficient MCC—brought about by the interactions of ciliary function, ASL homeostasis and mucus properties—for normal physiological function, we consider this AOP applicable to both genders.</p>
<h3>Essentiality of the Key Events</h3>
<p>The definition of essentiality implies that the modulation of upstream KEs impacts the downstream KEs in an expected fashion. If blocked or failing to occur, the KEs in the current AOP will not necessarily stop the progression to subsequent KEs. Due to the complex biology of motile cilia formation and function, ASL homeostasis, mucus properties and MCC, the KEs and AO may be triggered because of alternative pathways or biological redundancies. However, when exacerbated, the KEs promote the occurrence of downstream events eventually leading to the AO. The causal pathway starting from the exposure to oxidants and leading to decreased lung function involves parallel routes with KEs, each of which is sufficient to cause the downstream KE to occur. Based on the evidence we judge the key events MIE (Oxidative Stress), KE2 (Cilia Beat Frequency, Decreased), and KE3 (Mucociliary Clearance, Decreased) as highly essential. </p>
<p>The definition of essentiality implies that the modulation of upstream KEs impacts the downstream KEs in an expected fashion. If blocked or failing to occur, the KEs in the current AOP will not necessarily stop the progression to subsequent KEs. Due to the complex biology of motile cilia formation and function, ASL homeostasis, mucus properties and their concerted impact on MCC, the KEs and AO may be triggered because of alternative pathways or biological redundancies. However, when exacerbated, the KEs promote the occurrence of downstream events eventually leading to the AO. The causal pathway starting from the exposure to oxidants and leading to decreased lung function involves parallel routes with KEs, each of which is sufficient to cause the downstream KE to occur. Based on the evidence we judge the MIE (Oxidative Stress), KE1908 (Cilia Beat Frequency, Decreased), and KE1909 (Mucociliary Clearance, Decreased) as highly essential. </p>
<h3>Weight of Evidence Summary</h3>
<p>We judge the overall biological plausibility of this AOP as strong. The KER <em>Oxidative stress leading to decreased CBF</em> is supported by multiple studies across different species with ample empirical evidence reflecting both dose-response and time concordance. The KER <em>Decreased CBF leading to decreased MCC</em> lacks this expanse of empirical evidence, or the evidence does not fully support the causality between the KE even though the relationship is logical and plausible. </p>
<h3>Quantitative Consideration</h3>
<p>Overall, our quantitative understanding of the AOP network is moderate.</p>
<p>There is robust evidence that provides an insight into several KER presented here, and the dose response and temporal relationship between the two KE in question are well described and quantified for different stressors across different test systems (Oxidative stress leading to decreased CFTR function, Decreased CFTR function leading to decreased ASL height; Decreased FOXJ1 protein leading to decreased motile cilia length/number; Decreased motile cilia length/number leading to decreased cilia beating frequency; Decreased cilia beat frequency leading to decreased MCC; Increased mucus viscosity leading to decreased MCC; Oxidative stress leading to decreased cilia beat frequency). </p>
<p>In some instances, we are less confident in our quantitative understanding. For example, for the KER Decreased ASL height leading to decreased cilia beat frequency, empirical evidence supporting causality between the two KE is lacking as is quantitative evidence. Dose response data as well as data supportive of the KE causality are limited for the KER Decreased MCC leading to decreased lung function. Similarly, the KER Decreased ASL height leading to increased mucus viscosity is supported by only a small number of studies, most of which evaluated the KEs in parallel, without interrogating dose responses and/or time responses. </p>
<p>There is robust evidence that provides an insight into the KER <em>Oxidative stress leading to decreased cilia beat frequency</em> and <em>Decreased cilia beat frequency leading to decreased MCC</em>, and the dose response and temporal relationship between the two KE in question are well described and quantified for different stressors across different test systems. In some instances, we are less confident in our quantitative understanding. For example, dose response data as well as data supportive of the KE causality are limited for the KER <em>Decreased MCC leading to decreased lung function</em>. </p>
<h2>Considerations for Potential Applications of the AOP (optional)</h2>
<p>Given the individual and public health burden of the consequences of lung function impairment, gaining a greater understanding of the underlying mechanisms is extremely important in the risk assessment of respiratory toxicants. An integrated assessment of substances with the potential to be inhaled, either intentionally or unintentionally, could incorporate inhalation exposure and dosimetry modelling to inform an in vitro approach with appropriate exposure techniques and cell systems to assess KEs in this AOP (EPA’s Office of Chemical Safety and Pollution Prevention, 2019). Standardization and robustness testing of assays against explicit performance criteria using suitable reference materials can greatly increase the level of confidence in their use for KE assessment (Petersen et al., In Press). Much of the empirical evidence that supports the KERs in the qualitative AOP described here was obtained from in vitro studies using well-established methodologies for biological endpoint assessment. Being chemical agnostic, this AOP can be applied to a variety of substances that share the AO. For example, impaired MCC and decreased lung function have a long-known relationship with smoking, but little is known about the consequences of long-term use of alternative inhaled nicotine delivery products such as electronic cigarettes and heated tobacco products. This AOP can form the basis of an assessment strategy to evaluate the effects of exposure to aerosol from these products based on the KEs identified here. </p>
<p>Given the individual and public health burden of the consequences of lung function impairment, gaining a greater understanding of the underlying mechanisms is extremely important in the risk assessment of respiratory toxicants. An integrated assessment of substances with the potential to be inhaled, either intentionally or unintentionally, could incorporate inhalation exposure and dosimetry modelling to inform an in vitro approach with appropriate exposure techniques and cell systems to assess KEs in this AOP (EPA’s Office of Chemical Safety and Pollution Prevention, 2019). Standardization and robustness testing of assays against explicit performance criteria using suitable reference materials can greatly increase the level of confidence in their use for KE assessment (Petersen et al., 2021). Much of the empirical evidence that supports the KERs in the qualitative AOP described here was obtained from in vitro studies using well-established methodologies for biological endpoint assessment. Being chemical agnostic, this AOP can be applied to a variety of substances that share the AO. For example, impaired MCC and decreased lung function have a long-known relationship with smoking, but little is known about the consequences of long-term use of alternative inhaled nicotine delivery products such as electronic cigarettes and heated tobacco products. This AOP can form the basis of an assessment strategy to evaluate the effects of exposure to aerosol from these products based on the KEs identified here. </p>
</div>
<div id="references">
<h2>References</h2>
<p>Bailey, K.L., Bonasera, S.J., Wilderdyke, M., Hanisch, B.W., Pavlik, J.A., DeVasure, J., et al. (2014). Aging causes a slowing in ciliary beat frequency, mediated by PKCε. Am. J. Physiol. Lung Cell. Mol. Physiol. 306, L584-L589.</p>
<p>Bustamante-Marin, X.M., and Ostrowski, L.E. (2017a). Cilia and Mucociliary Clearance. Cold Spring Harb. Persp. Biol. 9, a028241. </p>
<p>Clippinger, A.J., Allen, D., Behrsing, H., BéruBé, K.A., Bolger, M.B., Casey, W., et al. (2018). Pathway-based predictive approaches for non-animal assessment of acute inhalation toxicity. Toxicol. In Vitro 52, 131-145.</p>
<p>EPA’s Office of Chemical Safety and Pollution Prevention (2019). "FIFRA Scientific Advisory Panel Meeting Minutes and Final Report No. 2019-01 Peer Review on Evaluation of a Proposed Approach to Refine the Inhalation Risk Assessment for Point of Contact Toxicity: A Case Study Using a New Approach Methodology (NAM) December 4 and 6, 2018 FIFRA Scientific Advisory Panel Meeting". U.S. Environmental Protection Agency).</p>
<p>Frohlich, E., Mercuri, A., Wu, S., and Salar-Behzadi, S. (2016). Measurements of Deposition, Lung Surface Area and Lung Fluid for Simulation of Inhaled Compounds. Front. Pharmacol. 7, 181. </p>
<p>Grubb, B.R., Livraghi-Butrico, A., Rogers, T.D., Yin, W., Button, B., and Ostrowski, L.E. (2016). Reduced mucociliary clearance in old mice is associated with a decrease in Muc5b mucin. Am. J. Physiol. Lung Cell. Mol. Physiol. 310, L860-L867.</p>
<p>Ho, J.C., Chan, K.N., Hu, W.H., Lam, W.K., Zheng, L., Tipoe, G.L., et al. (2001). The effect of aging on nasal mucociliary clearance, beat frequency, and ultrastructure of respiratory cilia. Am. J. Respir. Crit. Care Med. 163, 983-988.</p>
<p>Joki, S., and Saano, V. (1997). Influence of ageing on ciliary beat frequency and on ciliary response to leukotriene D4 in guinea-pig tracheal epithelium. Clin. Exp. Pharmacol. Physiol. 24, 166-169.</p>
<p>Luettich, K., Sharma, M., Yepiskoposyan, H., Breheny, D., and Lowe, F. J. (2021). An Adverse Outcome Pathway for Decreased Lung Function Focusing on Mechanisms of Impaired Mucociliary Clearance Following Inhalation Exposure. Frontiers in Toxicology, 55.</p>
<p>National Research Council (1988). Air Pollution, the Automobile, and Public Health. Washington, DC: The National Academies Press.</p>
<p>Paul, P., Johnson, P., Ramaswamy, P., Ramadoss, S., Geetha, B., and Subhashini, A. (2013). The effect of ageing on nasal mucociliary clearance in women: a pilot study. ISRN 2013, 598589.</p>
<p>Petersen, E.J., Sharma, M., Clippinger, A.J., Gordon, J., Katz, A., Laux, P., et al. (2021). Use of Cause-and-Effect Analysis to Optimize the Reliability of In Vitro Inhalation Toxicity Measurements Using an Air–Liquid Interface. Chem. Res. Toxicol. 34, 1370–1385.</p>
<p>Sharma, G., and Goodwin, J. (2006). Effect of aging on respiratory system physiology and immunology. Clin. Interv. Aging 1, 253-260. </p>
<td><a href="/aops/220">Aop:220 - Cyp2E1 Activation Leading to Liver Cancer</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/17">Aop:17 - Binding of electrophilic chemicals to SH(thiol)-group of proteins and /or to seleno-proteins involved in protection against oxidative stress during brain development leads to impairment of learning and memory</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/284">Aop:284 - Binding of electrophilic chemicals to SH(thiol)-group of proteins and /or to seleno-proteins involved in protection against oxidative stress leads to chronic kidney disease</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/377">Aop:377 - Dysregulated prolonged Toll Like Receptor 9 (TLR9) activation leading to Acute Respiratory Distress Syndrome (ARDS) and Multiple Organ Dysfunction (MOD)</a></td>
<td><a href="/aops/377">Aop:377 - Dysregulated prolonged Toll Like Receptor 9 (TLR9) activation leading to Multi Organ Failure involving Acute Respiratory Distress Syndrome (ARDS)</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/411">Aop:411 - Oxidative stress [MIE] Leading to Decreased Lung Function [AO]</a></td>
<td><a href="/aops/411">Aop:411 - Oxidative stress Leading to Decreased Lung Function </a></td>
<td>MolecularInitiatingEvent</td>
</tr>
<tr>
<td><a href="/aops/424">Aop:424 - Oxidative stress [MIE] Leading to Decreased Lung Function [AO] via CFTR dysfunction</a></td>
<td><a href="/aops/424">Aop:424 - Oxidative stress Leading to Decreased Lung Function via CFTR dysfunction</a></td>
<td>MolecularInitiatingEvent</td>
</tr>
<tr>
<td><a href="/aops/425">Aop:425 - Oxidative Stress Leading to Decreased Lung Function via Decreased FOXJ1</a></td>
<td>MolecularInitiatingEvent</td>
</tr>
<tr>
<td><a href="/aops/429">Aop:429 - A cholesterol/glucose dysmetabolism initiated Tau-driven AOP toward memory loss (AO) in sporadic Alzheimer's Disease with plausible MIE's plug-ins for environmental neurotoxicants</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/452">Aop:452 - Adverse outcome pathway of PM-induced respiratory toxicity</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/464">Aop:464 - Calcium overload in dopaminergic neurons of the substantia nigra leading to parkinsonian motor deficits</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/470">Aop:470 - Deposition of energy leads to abnormal vascular remodeling</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/478">Aop:478 - Deposition of energy leading to occurrence of cataracts</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/479">Aop:479 - Mitochondrial complexes inhibition leading to left ventricular function decrease via increased myocardial oxidative stress</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/481">Aop:481 - AOPs of amorphous silica nanoparticles: ROS-mediated oxidative stress increased respiratory dysfunction and diseases.</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/482">Aop:482 - Deposition of energy leading to occurrence of bone loss</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/483">Aop:483 - Deposition of Energy Leading to Learning and Memory Impairment</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/505">Aop:505 - Reactive Oxygen Species (ROS) formation leads to cancer via inflammation pathway</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/521">Aop:521 - Essential element imbalance leads to reproductive failure via oxidative stress</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/26">Aop:26 - Calcium-mediated neuronal ROS production and energy imbalance</a></td>
<td>AdverseOutcome</td>
</tr>
<tr>
<td><a href="/aops/488">Aop:488 - Increased reactive oxygen species production leading to decreased cognitive function</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/396">Aop:396 - Deposition of ionizing energy leads to population decline via impaired meiosis</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/437">Aop:437 - Inhibition of mitochondrial electron transport chain (ETC) complexes leading to kidney toxicity</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/535">Aop:535 - Binding and activation of GPER leading to learning and memory impairments</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/171">Aop:171 - Chronic cytotoxicity of the serous membrane leading to pleural/peritoneal mesotheliomas in the rat.</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/138">Aop:138 - Organic anion transporter (OAT1) inhibition leading to renal failure and mortality</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/177">Aop:177 - Cyclooxygenase 1 (COX1) inhibition leading to renal failure and mortality</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/186">Aop:186 - unknown MIE leading to renal failure and mortality</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/200">Aop:200 - Estrogen receptor activation leading to breast cancer </a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/444">Aop:444 - Ionizing radiation leads to reduced reproduction in Eisenia fetida via reduced spermatogenesis and cocoon hatchability</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/447">Aop:447 - Kidney failure induced by inhibition of mitochondrial electron transfer chain through apoptosis, inflammation and oxidative stress pathways</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/476">Aop:476 - Adverse Outcome Pathways diagram related to PBDEs associated male reproductive toxicity</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/497">Aop:497 - ERa inactivation alters mitochondrial functions and insulin signalling in skeletal muscle and leads to insulin resistance and metabolic syndrome</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/457">Aop:457 - Succinate dehydrogenase inhibition leading to increased insulin resistance through reduction in circulating thyroxine</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/459">Aop:459 - AhR activation in the thyroid leading to Subsequent Adverse Neurodevelopmental Outcomes in Mammals</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/507">Aop:507 - Nrf2 inhibition leading to vascular disrupting effects via inflammation pathway</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/509">Aop:509 - Nrf2 inhibition leading to vascular disrupting effects through activating apoptosis signal pathway and mitochondrial dysfunction</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/510">Aop:510 - Demethylation of PPAR promotor leading to vascular disrupting effects</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/511">Aop:511 - The AOP framework on ROS-mediated oxidative stress induced vascular disrupting effects </a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/538">Aop:538 - Adverse outcome pathway of PFAS-induced vascular disrupting effects via activating oxidative stress related pathways </a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/260">Aop:260 - CYP2E1 activation and formation of protein adducts leading to neurodegeneration</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/450">Aop:450 - Inhibition of AChE and activation of CYP2E1 leading to sensory axonal peripheral neuropathy and mortality</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/501">Aop:501 - Excessive iron accumulation leading to neurological disorders</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/540">Aop:540 - Oxidative Stress in the Fish Ovary Leads to Reproductive Impairment via Reduced Vitellogenin Production</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/471">Aop:471 - Neuron defect induced early behavioral change</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/31">Aop:31 - Oxidation of iron in hemoglobin leading to hematotoxicity</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/534">Aop:534 - Succinate dehydrogenase (SDH) inhibition leads to cancer through oxidative stress</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/462">Aop:462 - Activation of reactive oxygen species leading the atherosclerosis</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/324">Aop:324 - Excessive reactive oxygen species leading to growth inhibition via uncoupling of oxidative phosphorylation and cell death</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/325">Aop:325 - Excessive reactive oxygen species leading to growth inhibition via uncoupling of oxidative phosphorylation and reduced cell growth</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/326">Aop:326 - Excessive reactive oxygen species leading to growth inhibition via uncoupling of oxidative phosphorylation and reduced cell proliferation</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/331">Aop:331 - Excessive reactive oxygen species leading to growth inhibition via lipid peroxidation and cell death</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/332">Aop:332 - Excessive reactive oxygen species leading to growth inhibition via lipid peroxidation and reduced cell growth</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/333">Aop:333 - Excessive reactive oxygen species leading to growth inhibition via lipid peroxidation and reduced cell proliferation</a></td>
<td><a href="/aops/596">Aop:596 - Excessive reactive oxygen species leading to growth inhibition via protein oxidation and cell injury/death</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/598">Aop:598 - Excessive reactive oxygen species leading to growth inhibition via protein oxidation and reduced cell proliferation</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/599">Aop:599 - Excessive reactive oxygen species leading to growth inhibition via fatty acid oxidation and cell injury/death</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/600">Aop:600 - Excessive reactive oxygen species leading to growth inhibition via fatty acid oxidation and reduced cell growth</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/601">Aop:601 - Excessive reactive oxygen species leading to growth inhibition via fatty acid oxidation and reduced cell proliferation</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/602">Aop:602 - Excessive reactive oxygen species leading to growth inhibition via oxidative DNA damage</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/603">Aop:603 - Excessive reactive oxygen species leading to growth inhibition via protein oxidation and cell cycle disruption</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/608">Aop:608 - Thyroid Hormone Excess Leading to Reduced, Swimming Performance via Hypomyelination</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/616">Aop:616 - organic UV filter and its Photoproducts reproductive toxicity pathways </a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/622">Aop:622 - Calcineurin inhibitor induced nephrotoxicity leading to kidney failure</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/624">Aop:624 - Altered glucocorticoid receptor signaling leading to MASLD progression via insulin resistance-associated mitochondrial dysfunction</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/625">Aop:625 - Altered glucocorticoid receptor signaling leading to MASLD progression via reduced very low-density lipoprotein export-associated mitochondrial dysfunction</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/626">Aop:626 - Altered glucocorticoid receptor signaling leading to MASLD progression via reduced VLDL export-associated mitochondrial dysfunction</a></td>
<p>Oxidative stress is produced in, and can occur in, any species from bacteria through to humans.</p>
<p><span style="color:#27ae60"><strong>Taxonomic applicability: </strong>Occurrence of oxidative stress is not species specific. </span></p>
<p><span style="color:#27ae60"><strong>Life stage applicability:</strong> Occurrence of oxidative stress is not life stage specific. </span></p>
<p><span style="color:#27ae60"><strong>Sex applicability: </strong>Occurrence of oxidative stress is not sex specific. </span></p>
<p><span style="color:#27ae60"><strong>Evidence for perturbation by prototypic stressor:</strong> There is evidence of the increase of oxidative stress following perturbation from a variety of stressors including exposure to ionizing radiation and altered gravity (Bai et al., 2020; Ungvari et al., 2013; Zhang et al., 2009). </span></p>
<h4>Key Event Description</h4>
<p style="margin-left:18.0pt">Oxidative stress is defined as an imbalance in the production of reactive oxygen species (ROS) and antioxidant defenses. High levels of oxidizing free radicals can be very damaging to cells and molecules within the cell. As a result, the cell has important defense mechanisms to protect itself from ROS. For example, Nrf2 is a transcription factor and master regulator of the oxidative stress response. During periods of oxidative stress, Nrf2-dependent changes in gene expression are important in regaining cellular homeostasis (Nguyen, et al. 2009) and can be used as indicators of the presence of oxidative stress in the cell.</p>
<p>Oxidative stress is defined as an imbalance in the production of reactive oxygen species (ROS) and antioxidant defenses. High levels of oxidizing free radicals can be very damaging to cells and molecules within the cell. As a result, the cell has important defense mechanisms to protect itself from ROS. For example, Nrf2 is a transcription factor and master regulator of the oxidative stress response. During periods of oxidative stress, Nrf2-dependent changes in gene expression are important in regaining cellular homeostasis (Nguyen, et al., 2009) and can be used as indicators of the presence of oxidative stress in the cell. </p>
<p>In addition to the directly damaging actions of ROS, cellular oxidative stress also changes cellular activities on a molecular level. Redox sensitive proteins have altered physiology in the presence and absence of ROS, which is caused by the oxidation of sulfhydryls to disulfides on neighboring amino acids (Antelmann & Helmann 2011). Importantly Keap1, the negative regulator of Nrf2, is regulated in this manner (Itoh, et al. 2010). </p>
<p style="margin-left:18.0pt">In addition to the directly damaging actions of ROS, cellular oxidative stress also changes cellular activities on a molecular level. Redox sensitive proteins have altered physiology in the presence and absence of ROS, which is caused by theoxidation of sulfhydryls to disulfides (2SH àSS) on neighboring amino acids (Antelmann and Helmann 2011). Importantly Keap1, the negative regulator of Nrf2, is regulated in this manner (Itoh, et al. 2010).</p>
<p>ROS also undermine the mitochondrial defense system from oxidative damage. The antioxidant systems consist of superoxide dismutase, catalase, glutathione peroxidase and glutathione reductase, as well as antioxidants such as α-tocopherol and ubiquinol, or antioxidant vitamins and minerals including vitamin E, C, carotene, lutein, zeaxanthin, selenium, and zinc (Fletcher, 2010). The enzymes, vitamins and minerals catalyze the conversion of ROS to non-toxic molecules such as water and O2. However, these antioxidant systems are not perfect and endogenous metabolic processes and/or exogenous oxidative influences can trigger cumulative oxidative injuries to the mitochondria, causing a decline in their functionality and efficiency, which further promotes cellular oxidative stress (Balasubramanian, 2000; Ganea & Harding, 2006; Guo et al., 2013; Karimi et al., 2017). </p>
<p style="margin-left:18.0pt">Protection against oxidative stress is relevant for all tissues and organs, although some tissues may be more susceptible. For example, the brain possesses several key physiological features, such as high O2 utilization, high polyunsaturated fatty acids content, presence of autooxidable neurotransmitters, and low antioxidant defenses as compared to other organs, that make it highly susceptible to oxidative stress (Halliwell, 2006; Emerit and al., 2004; Frauenberger et al., 2016).</p>
<p>However, an emerging viewpoint suggests that ROS-induced modifications may not be as detrimental as previously thought, but rather contribute to signaling processes (Foyer et al., 2017). </p>
<p> </p>
<p><strong>Sources of ROS Production </strong></p>
<p><strong>Direct Sources: </strong>Direct sources involve the deposition of energy onto water molecules, breaking them into active radical species. When ionizing radiation hits water, it breaks it into hydrogen (H*) and hydroxyl (OH*) radicals by destroying its bonds. The hydrogen will create hydroxyperoxyl free radicals (HO2*) if oxygen is available, which can then react with another of itself to form hydrogen peroxide (H2O2) and more O2 (Elgazzar and Kazem, 2015). Antioxidant mechanisms are also affected by radiation, with catalase (CAT) and peroxidase (POD) levels rising as a result of exposure (Seen et al. 2018; Ahmad et al. 2021). </p>
<p><strong>Indirect Sources</strong>: An indirect source of ROS is the mitochondria, which is one of the primary producers in eukaryotic cells (Powers et al., 2008). As much as 2% of the electrons that should be going through the electron transport chain in the mitochondria escape, allowing them an opportunity to interact with surrounding structures. Electron-oxygen reactions result in free radical production, including the formation of hydrogen peroxide (H2O2) (Zhao et al., 2019). The electron transport chain, which also creates ROS, is activated by free adenosine diphosphate (ADP), O2, and inorganic phosphate (Pi) (Hargreaves et al. 2020; Raimondi et al. 2020; Vargas-Mendoza et al. 2021). The first and third complexes of the transport chain are the most relevant to mammalian ROS production (Raimondi et al., 2020). The mitochondria has its own set of DNA and it is a prime target of oxidative damage (Guo et al., 2013). ROS is also produced through nicotinamide adenine dinucleotide phosphate oxidase (Nox) stimulation, an event commenced by angiotensin II, a product/effector of the renin-angiotensin system (Nguyen Dinh Cat et al. 2013; Forrester et al. 2018). Other ROS producers include xanthine oxidase, immune cells (macrophage, neutrophils, monocytes, and eosinophils), phospholipase A2 (PLA2), monoamine oxidase (MAO), and carbon-based nanomaterials (Powers et al. 2008; Jacobsen et al. 2008; Vargas-Mendoza et al. 2021). </p>
<h4>How it is Measured or Detected</h4>
<p><strong>Oxidative Stress. Direct measurement of ROS is difficult because ROS are unstable. The presence of ROS can be assayed indirectly by measurement of cellular antioxidants, or by ROS-dependent cellular damage:</strong></p>
<p><strong>Oxidative Stress:</strong> Direct measurement of ROS is difficult because ROS are unstable. The presence of ROS can be assayed indirectly by measurement of cellular antioxidants, or by ROS-dependent cellular damage. Listed below are common methods for detecting the KE, however there may be other comparable methods that are not listed </p>
<p>- Detection of ROS by chemiluminescence <span style="font-size:12px">(<span style="font-family:arial,helvetica,sans-serif">https://www.sciencedirect.com/science/article/abs/pii/S0165993606001683)</span></span></p>
<ul>
<li>Detection of ROS by chemiluminescence (https://www.sciencedirect.com/science/article/abs/pii/S0165993606001683) </li>
<li>Detection of ROS by chemiluminescence is also described in OECD TG 495 to assess phototoxic potential. </li>
<li>Glutathione (GSH) depletion. GSH can be measured by assaying the ratio of reduced to oxidized glutathione (GSH:GSSG) using a commercially available kit (e.g., http://www.abcam.com/gshgssg-ratio-detection-assay-kit-fluorometric-green- ab138881.html). </li>
<li>TBARS. Oxidative damage to lipids can be measured by assaying for lipid peroxidation using TBARS (thiobarbituric acid reactive substances) using a commercially available kit. </li>
<li>8-oxo-dG. Oxidative damage to nucleic acids can be assayed by measuring 8-oxo-dG adducts (for which there are a number of ELISA based commercially available kits),or HPLC, described in Chepelev et al. (Chepelev, et al. 2015). </li>
</ul>
<p>- Detection of ROS by chemiluminescence is also described in OECD TG 495 to assess phototoxic potential.</p>
<p> </p>
<p><br />
- Glutathione (GSH) depletion. GSH can be measured by assaying the ratio of reduced to oxidized glutathione (GSH:GSSG) using a commercially available kit (e.g., http://www.abcam.com/gshgssg-ratio-detection-assay-kit-fluorometric-green-ab138881.html). </p>
<p><strong>Molecular Biology:</strong> Nrf2. Nrf2’s transcriptional activity is controlled post-translationally by oxidation of Keap1. Assay for Nrf2 activity include: </p>
<p><br />
- TBARS. Oxidative damage to lipids can be measured by assaying for lipid peroxidation using TBARS (thiobarbituric acid reactive substances) using a commercially available kit. </p>
<ul>
<li>Immunohistochemistry for increases in Nrf2 protein levels and translocation into the nucleus Western blot for increased Nrf2 protein levels </li>
<li>Western blot of cytoplasmic and nuclear fractions to observe translocation of Nrf2 protein from the cytoplasm to the nucleus qPCR of Nrf2 target genes (e.g., Nqo1, Hmox-1, Gcl, Gst, Prx, TrxR, Srxn), or by commercially available pathway-based qPCR array (e.g., oxidative stress array from SABiosciences) </li>
<li>Whole transcriptome profiling by microarray or RNA-seq followed by pathway analysis (in IPA, DAVID, metacore, etc.) for enrichment of the Nrf2 oxidative stress response pathway (e.g., Jackson et al. 2014) </li>
<li>OECD TG422D describes an ARE-Nrf2 Luciferase test method </li>
</ul>
<p><br />
- 8-oxo-dG. Oxidative damage to nucleic acids can be assayed by measuring 8-oxo-dG adducts (for which there are a number of ELISA based commercially available kits),or HPLC, described in Chepelev et al. (Chepelev, et al. 2015).</p>
<p>In general, there are a variety of commercially available colorimetric or fluorescent kits for detecting Nrf2 activation.</p>
<p><strong>Molecular Biology: Nrf2. Nrf2’s transcriptional activity is controlled post-translationally by oxidation of Keap1. Assay for Nrf2 activity include:</strong><br />
- Immunohistochemistry for increases in Nrf2 protein levels and translocation into the nucleus; </p>
<table border="1">
<tbody>
<tr>
<td>
<p><strong>Assay Type & Measured Content </strong></p>
</td>
<td>
<p><strong>Description </strong></p>
</td>
<td>
<p><strong>Dose Range Studied </strong></p>
</td>
<td>
<p><strong>Assay Characteristics (Length/Ease of use/Accuracy) </strong></p>
</td>
</tr>
<tr>
<td>
<p>ROS </p>
<p>Formation in the Mitochondria assay (Shaki et al., 2012) </p>
</td>
<td>
<p>“The mitochondrial ROS measurement was performed flow cytometry using DCFH-DA. Briefly, isolated kidney mitochondria were incubated with UA (0, 50, 100 and 200 µM) in respiration buffer containing (0.32 mM sucrose, 10mM Tris, 20 mM Mops, 50 µM EGTA, 0.5 mM MgCl2, 0.1 mM KH2PO4 and 5 mM sodium succinate) [32]. In the interval times of 5, 30 and 60 min following the UA addition, a sample was taken and DCFH-DA was added (final concentration, 10 µM) to mitochondria and was then incubated for 10 min.Uranyl acetate-induced ROS generation in isolated kidney mitochondria were determined through the flow cytometry (Partec, Deutschland) equipped with a 488-nm argon ion laser and supplied with the Flomax software and the signals were obtained using a 530-nm bandpass filter (FL-1 channel). Each determination is based on the mean fluorescence intensity of 15,000 counts.” </p>
<p> </p>
</td>
<td>
<p>0, 50,100 and 200 µM of Uranyl Acetate </p>
<p> </p>
</td>
<td>
<p> Long/ Easy High accuracy </p>
<p> </p>
</td>
</tr>
<tr>
<td>
<p>Mitochondrial Antioxidant Content Assay Measuring GSH content (Shaki et al., 2012) </p>
<p> </p>
</td>
<td>
<p>“GSH content was determined using DTNB as the indicator and spectrophotometer method for the isolated mitochondria. The mitochondrial fractions (0.5 mg protein/ml) were incubated with various concentrations of uranyl acetate for 1 h at 30 °C and then 0.1 ml of mitochondrial fractions was added into 0.1 mol/l of phosphate buffers and 0.04% DTNB in a total volume of 3.0 ml (pH 7.4). The developed yellow color was read at 412 nm on a spectrophotometer (UV-1601 PC, Shimadzu, Japan). GSH content was expressed as µg/mg protein.” </p>
</td>
<td>
<p>0, 50, </p>
<p>100, or </p>
<p>200 µM </p>
<p>Uranyl Acetate </p>
</td>
<td>
<p> </p>
</td>
</tr>
<tr>
<td>
<p>H2O2 Production Assay Measuring H2O2 Production in isolated mitochondria (Heyno et al., 2008) </p>
<p> </p>
</td>
<td>
<p>“Effect of CdCl2 and antimycin A (AA) on H2O2 production in isolated mitochondria from potato. H2O2 production was measured as scopoletin oxidation. Mitochondria were incubated for 30 min in the measuring buffer </p>
<p>(see the Materials and Methods) containing 0.5 mM succinate as an electron donor and 0.2 µM mesoxalonitrile 3‐chlorophenylhydrazone (CCCP) as an uncoupler, 10 U horseradish peroxidase and 5 µM scopoletin.” </p>
</td>
<td>
<p>0, 10, 30 </p>
<p>µM Cd2+ </p>
<p> </p>
<p>2 µM antimycin A </p>
</td>
<td>
<p> </p>
</td>
</tr>
<tr>
<td>
<p>Flow Cytometry ROS & Cell Viability (Kruiderig et al., 1997) </p>
<p> </p>
</td>
<td>
<p>“For determination of ROS, samples taken at the indicated time points were directly transferred to FACScan tubes. Dih123 (10 mM, final concentration) was added and cells were incubated at 37°C in a humidified atmosphere (95% air/5% CO2) for 10 min. At t 5 9, propidium iodide (10 mM, final concentration) was added, and cells were analyzed by flow cytometry at 60 ml/min. Nonfluorescent Dih123 is cleaved by ROS to fluorescent R123 and detected by the FL1 detector as described above for Dc (Van de Water 1995)”“For determination of ROS, samples taken at the indicated time points were directly transferred to FACScan tubes. Dih123 (10 mM, final concentration) was added and cells were incubated at 37°C in a humidified atmosphere (95% air/5% CO2) for 10 min. At t 5 9, propidium iodide (10 mM, final concentration) was added, and cells were analyzed by flow cytometry at 60 ml/min. Nonfluorescent Dih123 is cleaved by ROS to fluorescent R123 and detected by the FL1 detector as described above for Dc (Van de Water 1995)” </p>
</td>
<td>
<p> </p>
</td>
<td>
<p> </p>
<p> </p>
<p> </p>
<p> </p>
<p> </p>
<p>Strong/easy medium </p>
</td>
</tr>
<tr>
<td>
<p>DCFH-DA </p>
<p>Assay Detection of hydrogen peroxide production (Yuan et al., </p>
<p>2016) </p>
</td>
<td>
<p>Intracellular ROS production was measured using DCFH-DA as a probe. Hydrogen peroxide oxidizes DCFH to DCF. The probe is hydrolyzed intracellularly to DCFH carboxylate anion. No direct reaction with H2O2 to form fluorescent production. </p>
<p> </p>
</td>
<td>
<p>0-400 </p>
<p>µM </p>
</td>
<td>
<p>Long/ Easy High accuracy </p>
</td>
</tr>
<tr>
<td>
<p>H2-DCF-DAAssay Detection of superoxide production (Thiebault etal., 2007) </p>
<p> </p>
</td>
<td>
<p>This dye is a stable nonpolar compound which diffuses readily into the cells and yields H2-DCF. Intracellular OH or ONOO- react with H2-DCF when cells contain peroxides, to form the highly fluorescent compound DCF, which effluxes the cell. Fluorescence intensity of DCF is measured using a fluorescence spectrophotometer. </p>
<p>The dye (CM-H2DCFDA) diffuses into the cell and is cleaved by esterases, the thiol reactive chlormethyl group reacts with intracellular glutathione which can be detected using flow cytometry. </p>
</td>
<td>
<p> </p>
</td>
<td>
<p>Long/Easy/ High Accuracy </p>
</td>
</tr>
</tbody>
</table>
<p>- Western blot for increased Nrf2 protein levels; </p>
<p> </p>
<p>- Western blot of cytoplasmic and nuclear fractions to observe translocation of Nrf2 protein from the cytoplasm to the nucleus; </p>
<table border="1">
<tbody>
<tr>
<td>
<p><strong>Method of Measurement </strong></p>
</td>
<td>
<p><strong>References </strong></p>
</td>
<td>
<p><strong>Description </strong></p>
</td>
<td colspan="2">
<p><strong>OECD-Approved Assay </strong></p>
</td>
</tr>
<tr>
<td>
<p>Chemiluminescence </p>
</td>
<td>
<p>(Lu, C. et al., 2006; </p>
<p>Griendling, K. K., et al., 2016) </p>
</td>
<td>
<p>ROS can induce electron transitions in molecules, leading to electronically excited products. When the electrons transition back to ground state, chemiluminescence is emitted and can be measured. Reagents such as luminol and lucigenin are commonly used to amplify the signal. </p>
</td>
<td colspan="2">
<p>No </p>
<p> </p>
</td>
</tr>
<tr>
<td>
<p>Spectrophotometry </p>
</td>
<td>
<p>(Griendling, K. K., et al., 2016) </p>
</td>
<td>
<p>NO has a short half-life. However, if it has been reduced to nitrite (NO2-), stable azocompounds can be formed via the Griess Reaction, and further measured by spectrophotometry. </p>
</td>
<td colspan="2">
<p>No </p>
</td>
</tr>
<tr>
<td>
<p>Direct or Spin Trapping-Based electron paramagnetic resonance (EPR) Spectroscopy </p>
</td>
<td>
<p>(Griendling, K. K., et al., 2016) </p>
</td>
<td>
<p>The unpaired electrons (free radicals) found in ROS can be detected with EPR and is known as electron paramagnetic resonance. A variety of spin traps can be used. </p>
</td>
<td colspan="2">
<p>No </p>
</td>
</tr>
<tr>
<td>
<p>Nitroblue Tetrazolium Assay </p>
</td>
<td>
<p>(Griendling, K. K., et al., 2016) </p>
</td>
<td>
<p>The Nitroblue Tetrazolium assay is used to measure O2.− levels. O2.− reduces nitroblue tetrazolium (a yellow dye) to formazan (a blue dye), and can be measured at 620 nm. </p>
</td>
<td colspan="2">
<p>No </p>
</td>
</tr>
<tr>
<td>
<p>Fluorescence analysis of dihydroethidium (DHE) or Hydrocyans </p>
</td>
<td>
<p>(Griendling, K. K., et al., 2016) </p>
</td>
<td>
<p>Fluorescence analysis of DHE is used to measure O2.− levels. O2.− is reduced to O2 as DHE is oxidized to 2-hydroxyethidium, and this reaction can be measured by fluorescence. Similarly, hydrocyans can be oxidized by any ROS, and measured via fluorescence. </p>
</td>
<td colspan="2">
<p>No </p>
</td>
</tr>
<tr>
<td>
<p>Amplex Red Assay </p>
</td>
<td>
<p>(Griendling, K. K., et al., 2016) </p>
</td>
<td>
<p>Fluorescence analysis to measure extramitochondrial or extracellular H2O2 levels. In the presence of horseradish peroxidase and H2O2, Amplex Red is oxidized to resorufin, a fluorescent molecule measurable by plate reader. </p>
<p>An indirect fluorescence analysis to measure intracellular H2O2 levels. H2O2 interacts with peroxidase or heme proteins, which further react with DCFH, oxidizing it to dichlorofluorescein (DCF), a fluorescent product. </p>
</td>
<td colspan="2">
<p>No </p>
</td>
</tr>
<tr>
<td>
<p>HyPer Probe </p>
</td>
<td>
<p>(Griendling, K. K., et al., 2016) </p>
</td>
<td>
<p>Fluorescent measurement of intracellular H2O2 levels. HyPer is a genetically encoded fluorescent sensor that can be used for in vivo and in situ imaging. </p>
</td>
<td colspan="2">
<p>No </p>
</td>
</tr>
<tr>
<td>
<p>Cytochrome c Reduction Assay </p>
</td>
<td>
<p>(Griendling, K. K., et al., 2016) </p>
</td>
<td>
<p>The cytochrome c reduction assay is used to measure O2.− levels. O O2.− is reduced to O2 as ferricytochrome c is oxidized to ferrocytochrome c, and this reaction can be measured by an absorbance increase at 550 nm. </p>
<p>The redox state of tissue is detected through nuclear magnetic resonance/magnetic resonance imaging, with the use of a nitroxide spin probe or biradical molecule. </p>
</td>
<td colspan="2">
<p>No </p>
<p> </p>
<p> </p>
<p> </p>
<p> </p>
<p> </p>
<p> </p>
</td>
</tr>
<tr>
<td>
<p>Glutathione (GSH) depletion </p>
</td>
<td>
<p>(Biesemann, N. et al., 2018) </p>
</td>
<td>
<p>A downstream target of the Nrf2 pathway is involved in GSH synthesis. As an indication of oxidation status, GSH can be measured by assaying the ratio of reduced to oxidized glutathione (GSH:GSSG) using a commercially available kit (e.g., <a href="http://www.abcam.com/gshgssg-ratio-detection-assay-kit-fluorometric-green-ab138881.html" rel="noreferrer noopener" target="_blank">http://www.abcam.com/gshgssg-ratio-detection-assay-kit-fluorometric-green-ab138881.html</a>). </p>
<p>Oxidative damage to lipids can be measured by assaying for lipid peroxidation with TBARS using a commercially available kit. </p>
</td>
<td colspan="2">
<p>No </p>
</td>
</tr>
<tr>
<td>
<p>Protein oxidation (carbonylation) </p>
</td>
<td>
<p>(Azimzadeh et al., 2017; Azimzadeh et al., 2015; Ping et al., 2020) </p>
</td>
<td>
<p>Can be determined with ELISA or a commercial assay kit. Protein oxidation can indicate the level of oxidative stress. </p>
</td>
<td colspan="2">
<p>No </p>
</td>
</tr>
<tr>
<td>
<p>Seahorse XFp Analyzer </p>
</td>
<td>
<p>Leung et al. 2018 </p>
</td>
<td>
<p>The Seahorse XFp Analyzer provides information on mitochondrial function, oxidative stress, and metabolic dysfunction of viable cells by measuring respiration (oxygen consumption rate; OCR) and extracellular pH (extracellular acidification rate; ECAR). </p>
</td>
<td>
<p>No </p>
</td>
</tr>
</tbody>
</table>
<p>- qPCR of Nrf2 target genes (e.g., Nqo1, Hmox-1, Gcl, Gst, Prx, TrxR, Srxn), or by commercially available pathway-based qPCR array (e.g., oxidative stress array from SABiosciences);</p>
<p> </p>
<p>- Whole transcriptome profiling by microarray or RNA-seq followed by pathway analysis (in IPA, DAVID, metacore, etc.) for enrichment of the Nrf2 oxidative stress response pathway (e.g., Jackson et al. 2014);</p>
<p>Molecular Biology: Nrf2. Nrf2’s transcriptional activity is controlled post-translationally by oxidation of Keap1. Assays for Nrf2 activity include: </p>
<p>- OECD TG422D describes an ARE-Nrf2 Luciferase test method;</p>
<table border="1">
<tbody>
<tr>
<td>
<p>Method of Measurement </p>
</td>
<td>
<p>References </p>
</td>
<td>
<p>Description </p>
</td>
<td>
<p>OECD-Approved Assay </p>
</td>
</tr>
<tr>
<td>
<p>Immunohistochemistry </p>
</td>
<td>
<p>(Amsen, D., de Visser, K. E., and Town, T., 2009) </p>
</td>
<td>
<p>Immunohistochemistry for increases in Nrf2 protein levels and translocation into the nucleus </p>
</td>
<td>
<p>No </p>
</td>
</tr>
<tr>
<td>
<p>qPCR </p>
</td>
<td>
<p>(Forlenza et al., 2012) </p>
</td>
<td>
<p>qPCR of Nrf2 target genes (e.g., Nqo1, Hmox-1, Gcl, Gst, Prx, TrxR, Srxn), or by commercially available pathway-based qPCR array (e.g., oxidative stress array from SABiosciences) </p>
</td>
<td>
<p>No </p>
</td>
</tr>
<tr>
<td>
<p>Whole transcriptome profiling via microarray or via RNA-seq followed by a pathway analysis </p>
</td>
<td>
<p>(Jackson, A. F. et al., 2014) </p>
</td>
<td>
<p>Whole transcriptome profiling by microarray or RNA-seq followed by pathway analysis (in IPA, DAVID, metacore, etc.) for enrichment of the Nrf2 oxidative stress response pathway </p>
</td>
<td>
<p>No </p>
</td>
</tr>
</tbody>
</table>
<p>- In general, there are a variety of commercially available colorimetric or fluorescent kits for detecting Nrf2 activation.</p>
<p>Ahmad, S. et al. (2021), “60Co-γ Radiation Alters Developmental Stages of Zeugodacus cucurbitae (Diptera: Tephritidae) Through Apoptosis Pathways Gene Expression”, Journal Insect Science, Vol. 21/5, Oxford University Press, Oxford, <a href="https://doi.org/10.1093/jisesa/ieab080" rel="noreferrer noopener" target="_blank">https://doi.org/10.1093/jisesa/ieab080</a> </p>
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<p>Amsen, D., de Visser, K. E., and Town, T. (2009), “Approaches to determine expression of inflammatory cytokines”, in Inflammation and Cancer, Humana Press, Totowa, <a href="https://doi.org/10.1007/978-1-59745-447-6_5" rel="noreferrer noopener" target="_blank">https://doi.org/10.1007/978-1-59745-447-6_5</a> </p>
<p>Azimzadeh, O. et al. (2015), “Integrative Proteomics and Targeted Transcriptomics Analyses in Cardiac Endothelial Cells Unravel Mechanisms of Long-Term Radiation-Induced Vascular Dysfunction”, Journal of Proteome Research, Vol. 14/2, American Chemical Society, Washington, <a href="https://doi.org/10.1021/pr501141b" rel="noreferrer noopener" target="_blank">https://doi.org/10.1021/pr501141b</a> </p>
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<p>Azzam, E. I. et al. (2012), “Ionizing radiation-induced metabolic oxidative stress and prolonged cell injury”, Cancer Letters, Vol. 327/1-2, Elsevier, Ireland, https://doi.org/10.1016/j.canlet.2011.12.012 </p>
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<p>Balasubramanian, D (2000), “Ultraviolet radiation and cataract”, Journal of ocular pharmacology and therapeutics, Vol. 16/3, Mary Ann Liebert Inc., Larchmont, <a href="https://doi.org/10.1089/jop.2000.16.285.%22%20/t%20%22_blank" rel="noreferrer noopener" target="_blank">https://doi.org/10.1089/jop.2000.16.285.</a> </p>
<p>Biesemann, N. et al., (2018), “High Throughput Screening of Mitochondrial Bioenergetics in Human Differentiated Myotubes Identifies Novel Enhancers of Muscle Performance in Aged Mice”, Scientific Reports, Vol. 8/1, Nature Portfolio, London, <a href="https://doi.org/10.1038/s41598-018-27614-8" rel="noreferrer noopener" target="_blank">https://doi.org/10.1038/s41598-018-27614-8</a>. </p>
<p>Elgazzar, A. and N. Kazem. (2015), “Chapter 23: Biological effects of ionizing radiation” in The Pathophysiologic Basis of Nuclear Medicine, Springer, New York, pp. 540-548 </p>
<p>Eruslanov, E., & Kusmartsev, S. (2010). Identification of ROS using oxidized DCFDA and flow-cytometry. Methods in molecular biology ,N.J., Vol. 594, https://doi.org/10.1007/978-1-60761-411-1_4 </p>
<p>Fletcher, A. E (2010), “Free radicals, antioxidants and eye diseases: evidence from epidemiological studies on cataract and age-related macular degeneration”, Ophthalmic Research, Vol. 44, Karger International, Basel, <a href="https://doi.org/10.1159/000316476.%22%20/t%20%22_blank" rel="noreferrer noopener" target="_blank">https://doi.org/10.1159/000316476.</a> </p>
<p>Forlenza, M. et al. (2012), “The use of real-time quantitative PCR for the analysis of cytokine mRNA levels” in Cytokine Protocols, Springer, New York, https://doi.org/10.1007/978-1-61779-439-1_2 </p>
<p>Forrester, S.J. et al. (2018), “Angiotensin II Signal Transduction: An Update on Mechanisms of Physiology and Pathophysiology”, Physiological Reviews, Vol. 98/3, American Physiological Society, Rockville, <a href="https://doi.org/10.1152/physrev.00038.201" rel="noreferrer noopener" target="_blank">https://doi.org/10.1152/physrev.00038.201</a> </p>
<p>Foyer, C. H., A. V. Ruban, and G. Noctor (2017), “Viewing oxidative stress through the lens of oxidative signalling rather than damage”, Biochemical Journal, Vol. 474/6, Portland Press, England, https://doi.org/10.1042/BCJ20160814 </p>
<p>Ganea, E. and J. J. Harding (2006), “Glutathione-related enzymes and the eye”, Current eye research, Vol. 31/1, Informa, London, <a href="https://doi.org/10.1080/02713680500477347.%22%20/t%20%22_blank" rel="noreferrer noopener" target="_blank">https://doi.org/10.1080/02713680500477347.</a> </p>
<p>Griendling, K. K. et al. (2016), “Measurement of reactive oxygen species, reactive nitrogen species, and redox-dependent signaling in the cardiovascular system: a scientific statement from the American Heart Association”, Circulation research, Vol. 119/5, Lippincott Williams & Wilkins, Philadelphia, <a href="https://doi.org/10.1161/RES.0000000000000110" rel="noreferrer noopener" target="_blank">https://doi.org/10.1161/RES.0000000000000110</a> </p>
<p>Guo, C. et al. (2013), “Oxidative stress, mitochondrial damage and neurodegenerative diseases”, Neural regeneration research, Vol. 8/21, Publishing House of Neural Regeneration Research, China, <a href="https://doi.org/10.3969/j.issn.1673-5374.2013.21.009" rel="noreferrer noopener" target="_blank">https://doi.org/10.3969/j.issn.1673-5374.2013.21.009</a> </p>
<p>Hargreaves, M., and L. L. Spriet (2020), “Skeletal muscle energy metabolism during exercise.”, Nature Metabolism, Vol. 2, Nature Portfolio, London, <a href="https://doi.org/10.1038/s42255-020-0251-4" rel="noreferrer noopener" target="_blank">https://doi.org/10.1038/s42255-020-0251-4</a> </p>
<p>Hladik, D. and S. Tapio (2016), “Effects of ionizing radiation on the mammalian brain”, Mutation Research/Reviews in Mutation Research, Vol. 770, Elsevier, Amsterdam, <a href="https://doi.org/10.1016/j.mrrev.2016.08.003" rel="noreferrer noopener" target="_blank">https://doi.org/10.1016/j.mrrev.2016.08.003</a> </p>
<p>Itoh, K., J. Mimura and M. Yamamoto (2010), “Discovery of the negative regulator of Nrf2, Keap1: a historical overview”, Antioxidants & Redox Signaling, Vol. 13/11, Mary Ann Leibert Inc., Larchmont, <a href="https://doi.org/10.1089/ars.2010.3222" rel="noreferrer noopener" target="_blank">https://doi.org/10.1089/ars.2010.3222</a> </p>
<p>Jackson, A.F. et al. (2014), “Case study on the utility of hepatic global gene expression profiling in the risk assessment of the carcinogen furan.”, Toxicology and Applied Pharmacology, Vol. 274/11, Elsevier, Amsterdam, <a href="https://doi.org/10.1016/j.taap.2013.10.019" rel="noreferrer noopener" target="_blank">https://doi.org/10.1016/j.taap.2013.10.019</a> </p>
<p>Jacobsen, N.R. et al. (2008), “Genotoxicity, cytotoxicity, and reactive oxygen species induced by single-walled carbon nanotubes and C60 fullerenes in the FE1-MutaTM Mouse lung epithelial cells”, Environmental and Molecular Mutagenesis, Vol. 49/6, John Wiley & Sons, Inc., Hoboken, <a href="https://doi.org/10.1002/em.20406" rel="noreferrer noopener" target="_blank">https://doi.org/10.1002/em.20406</a> </p>
<p>Karimi, N. et al. (2017), “Radioprotective effect of hesperidin on reducing oxidative stress in the lens tissue of rats”, International Journal of Pharmaceutical Investigation, Vol. 7/3, Phcog Net, Bengaluru, <a href="https://doi.org/10.4103/jphi.JPHI_60_17.%E2%80%AF" rel="noreferrer noopener" target="_blank">https://doi.org/10.4103/jphi.JPHI_60_17. </a> </p>
<p>Leung, D.T.H., and Chu, S. (2018), “Measurement of Oxidative Stress: Mitochondrial Function Using the Seahorse System” In: Murthi, P., Vaillancourt, C. (eds) Preeclampsia. Methods in Molecular Biology, vol 1710. Humana Press, New York, NY. https://doi.org/10.1007/978-1-4939-7498-6_22 </p>
<p>Chepelev, N.L., Kennedy, D.A., Gagne, R., White, T., Long, A.S., Yauk, C.L., White, P.A., 2015. HPLC Measurement of the DNA Oxidation Biomarker, 8-oxo-7,8-dihydro-2'-deoxyguanosine, in Cultured Cells and Animal Tissues. J. Vis. Exp. (102):e52697. doi, e52697.</p>
<p>Lu, C., G. Song, and J. Lin (2006), “Reactive oxygen species and their chemiluminescence-detection methods”, TrAC Trends in Analytical Chemistry, Vol. 25/10, Elsevier, Amsterdam, <a href="https://doi.org/10.1016/j.trac.2006.07.007" rel="noreferrer noopener" target="_blank">https://doi.org/10.1016/j.trac.2006.07.007</a> </p>
<p>Emerit, J., Edeas, M., Bricaire, F., 2004. Neurodegenerative diseases and oxidative stress. Biomed. Pharmacotherapy. 58(1): 39-46.</p>
<p>Nguyen Dinh Cat, A. et al. (2013), “Angiotensin II, NADPH oxidase, and redox signaling in the vasculature”, Antioxidants & redox signaling, Vol. 19/10, Mary Ann Liebert, Larchmont, <a href="https://doi.org/10.1089/ars.2012.4641" rel="noreferrer noopener" target="_blank">https://doi.org/10.1089/ars.2012.4641</a> </p>
<p>Frauenberger, E.A., Scola, G., Laliberté, V.L.M., Duong, A., Andreazza, A.C., 2015. Redox modulations, Antioxidants, and Neuropsychitrica Disorders. Ox. Med. Cell. Longevity. Vol. 2016, Article ID 4729192.</p>
<p>Ping, Z. et al. (2020), “Oxidative Stress in Radiation-Induced Cardiotoxicity”, Oxidative Medicine and Cellular Longevity, Vol. 2020, Hindawi, <a href="https://doi.org/10.1155/2020/3579143" rel="noreferrer noopener" target="_blank">https://doi.org/10.1155/2020/3579143</a> </p>
<p>Halliwell, B., 2006. Oxidative stress and neurodegeneration: where are we now? J. Neurochem. 97(6):1634-1658.</p>
<p>Powers, S.K. and M.J. Jackson. (2008), “Exercise-Induced Oxidative Stress: Cellular Mechanisms and Impact on Muscle Force Production”, Physiological Reviews, Vol. 88/4, American Physiological Society, Rockville, <a href="https://doi.org/10.1152/physrev.00031.2007" rel="noreferrer noopener" target="_blank">https://doi.org/10.1152/physrev.00031.2007</a> </p>
<p>Itoh, K., Mimura, J., Yamamoto, M., 2010. Discovery of the negative regulator of Nrf2, Keap1: a historical overview. Antioxid. Redox Signal. 13, 1665-1678.</p>
<p>Raimondi, V., F. Ciccarese and V. Ciminale. (2020), “Oncogenic pathways and the electron transport chain: a dangeROS liason”, British Journal of Cancer, Vol. 122/2, Nature Portfolio, London, <a href="https://doi.org/10.1038/s41416-019-0651-y" rel="noreferrer noopener" target="_blank">https://doi.org/10.1038/s41416-019-0651-y</a> </p>
<p>Jackson, A.F., Williams, A., Recio, L., Waters, M.D., Lambert, I.B., Yauk, C.L., 2014. Case study on the utility of hepatic global gene expression profiling in the risk assessment of the carcinogen furan. Toxicol. Applied Pharmacol.274, 63-77.</p>
<p>Seen, S. and L. Tong. (2018), “Dry eye disease and oxidative stress”, Acta Ophthalmologica, Vol. 96/4, John Wiley & Sons, Inc., Hoboken, <a href="https://doi.org/10.1111/aos.13526" rel="noreferrer noopener" target="_blank">https://doi.org/10.1111/aos.13526</a> </p>
<p>Nguyen, T., Nioi, P., Pickett, C.B., 2009. The Nrf2-antioxidant response element signaling pathway and its activation by oxidative stress. J. Biol. Chem. 284, 13291-13295.</p>
<p>Ungvari, Z. et al. (2013), “Ionizing Radiation Promotes the Acquisition of a Senescence-Associated Secretory Phenotype and Impairs Angiogenic Capacity in Cerebromicrovascular Endothelial Cells: Role of Increased DNA Damage and Decreased DNA Repair Capacity in Microvascular Radiosensitivity”, The Journals of Gerontology Series A: Biological Sciences and Medical Sciences, Vol. 68/12, Oxford University Press, Oxford, <a href="https://doi.org/10.1093/gerona/glt057." rel="noreferrer noopener" target="_blank">https://doi.org/10.1093/gerona/glt057.</a> </p>
<p>OECD (2018), Test No. 442D: In Vitro Skin Sensitisation: ARE-Nrf2 Luciferase Test Method, OECD Guidelines for the Testing of Chemicals, Section 4, OECD Publishing, Paris, https://doi.org/10.1787/9789264229822-en.</p>
<p> </p>
<p>Vargas-Mendoza, N. et al. (2021), “Oxidative Stress, Mitochondrial Function and Adaptation to Exercise: New Perspectives in Nutrition”, Life, Vol. 11/11, Multidisciplinary Digital Publishing Institute, Basel, <a href="https://doi.org/10.3390/life11111269" rel="noreferrer noopener" target="_blank">https://doi.org/10.3390/life11111269</a> </p>
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<p>OECD (2019), Test No. 495: Ros (Reactive Oxygen Species) Assay for Photoreactivity, OECD Guidelines for the Testing of Chemicals, Section 4, OECD Publishing, Paris, https://doi.org/10.1787/915e00ac-en.</p>
<p>Zhang, R. et al. (2009), “Blockade of AT1 receptor partially restores vasoreactivity, NOS expression, and superoxide levels in cerebral and carotid arteries of hindlimb unweighting rats”, Journal of applied physiology, Vol. 106/1, American Physiological Society, Rockville, <a href="https://doi.org/10.1152/japplphysiol.01278.2007" rel="noreferrer noopener" target="_blank">https://doi.org/10.1152/japplphysiol.01278.2007</a>. </p>
<p>Zhao, R. Z. et al. (2019), “Mitochondrial electron transport chain, ROS generation and uncoupling”, International journal of molecular medicine, Vol. 44/1, Spandidos Publishing Ltd., Athens, <a href="https://doi.org/10.3892/ijmm.2019.4188" rel="noreferrer noopener" target="_blank">https://doi.org/10.3892/ijmm.2019.4188</a> </p>
<p><p>Treatment of human sinonasal epithelial cells with cigarette smoke condensate for 3 minutes significantly reduced forskolin-stimulated CBF (Cohen et al., 2009). CBF was also decreased in differentiated normal human bronchial epithelial cells exposed to whole cigarette smoke (Schmid et al., 2015), in cilia-bearing explant adenoid tissues treated with 5 and 10% cigarette smoke extract (Wang et al., 2012), in hamster oviducts treated various mainstream cigarette smoke fractions (Knoll et al., 1995), and in nasal epithelial cells fom smokers with moderate and severe chronic obstructive pulmonary disease (COPD) (Yaghi et al., 2012).</p>
</p>
<h4>Acetaldehyde</h4>
<p><p>A concentration-dependent decrease in CBF has been observed after treatment with aldehydes. For example inhibition of cilia ATPase activity was observed after treatment with acetaldehyde, in ciliated bovine bronchial epithelial cells (Sisson et al., 1991). </p>
</p>
<h4>Acrolein</h4>
<p><p>Acrolein, an aldehyde in the gas phase of cigarette smoke, induced ciliostasis at high concentrations (> 1 mM), after 5 min of treatment, and cellular necrosis after 3 hr. However, at lower concentrations (from 0.5‒1 mM), acrolein transiently reduced the CBF to 4 Hz (Romet et al., 1990).</p>
</p>
<h4>Nicotine</h4>
<p><p>Normal human bronchial epithelial cells exposed to aerosolized nicotine showed decreased CFTR and BK conductance, CBF, ASL volume, and decreased expression of FOXJ1 and KCNMA1 (Garcia-Arcos et al., 2016). </p>
</p>
<h4>Ozone</h4>
<p><p>Continuous, exposure of human nasal epithelial cells to different concentrations of ozone at 37°C for up to 4 weeks slightly (but not significantly) reduced CBF in healthy mucosa (7.1% at 500 µg/m3 and 10.3% at 1000 µg/m3), and significantly in chronically inflamed mucosa (20.5/16.4%) at 2 weeks. During the third and fourth week of exposure at these higher concentrations CBF was significantly reduced in both healthy (after 3 weeks: 18.7/37.5%; after 4 weeks: 11.1/33.3%) and chronically inflamed mucosa (after 3 weeks: 33.8/26.8%; after 4 weeks: 21.4/38.6%). Low ozone concentrations (100 µg/m3) appeared to not have an effect on CBF (Gosepath et al., 2000).</p>
<p> </p>
</p>
<h4>Sex hormone</h4>
<p><p>Female hormones, i.e. progesterone and estrogen, have been shown to have direct effect on CBF, i.e., progesterone reduces CBF, 17β-estradiol and progesterone receptor antagonists counteract progesterone effects, but estradiol alone has also been shown to have no effect on CBF. However, the mechanism by which these hormones modulate CBF is yet to be elucidated (Jain et al., 2012; Jia et al., 2011).</p>
<p>Age-dependent decreases in CBF have been demonstrated in several species (e.g. guinea pigs, mice, and human) (Bailey et al., 2014; Grubb et al., 2016; Ho et al., 2001; Joki and Saano, 1997; Paul et al., 2013). In a study with 46 healthy subjects with a wide age distribution (mean 42, range 19–81 years), age was found to be negatively associated with airway clearance of inhaled 6-μm Teflon particles (Svartengren et al., 2005).</p>
<p>Female hormones, i.e. progesterone and estrogen, have been shown to have direct effect on CBF, i.e., progesterone reduces CBF, 17β-estradiol and progesterone receptor antagonists counteract progesterone effects, but estradiol alone has also been shown to have no effect on CBF. However, the mechanism by which these hormones modulate CBF is yet to be elucidated (Jain et al., 2012; Jia et al., 2011).</p>
<h4>Key Event Description</h4>
<p>Cohesive beating of cilia lining the upper and lower respiratory tract is critical for efficient MCC. CBF is influenced by several factors including changes in the physical and chemical properties of the ASL (especially the periciliary fluid), structural modulation in the cilia, concentration of cyclic nucleotides cAMP and cGMP, and intracellular calcium (Ca<sup>2+</sup>). Formation of cyclic nucleotides such as cGMP is mediated by nitric oxide (NO), which is released by an enzyme family of nitric oxide synthases (NOSs) when the substrate L-arginine (L-Arg) is transformed to L-citrulline. NO activates its receptor protein, soluble guanylate cyclase (sGC), which catalyzes formation of cGMP from guanosine triphosphate (GTP). cGMP then activates protein kinase G (PKG) which has been implicated in the regulation of CBF (Jiao et al., 2011; Li et al., 2000). NO-dependent stimulation of CBF has also been associated with an increase in cAMP-dependent protein kinase A (PKA) (Di Benedetto et al., 1991; Lansley et al., 1992; Salathe et al., 1993; Sanderson and Dirksen, 1989; Schmid et al., 2007; Sisson et al., 1999; Uzlaner and Priel, 1999). An increase in intracellular endogenous cAMP was observed after treatment with isobutyl-1-methylxanthine that also increased CBF (Tamaoki et al., 1989). cAMP accumulation in the airway cilia has been shown to be dependent on Ca2+–calmodulin-dependent PDE1A and indirectly regulates CBF (Kogiso et al., 2018). Increase in CBF after treatment with NO substrate, L-arginine and inhibition of CBF by a NOS inhibitor, N-omega-nitro-L-arginine methyl ester (L-NAME) further provides evidence for the role of NO in increasing CBF (Jiao J. et al., 2011; Sisson J. H., 1995; Uzlaner and Priel, 1999; Yang et al., 1997). <br />
Modulation of CBF is not always accompanied by changes in cAMP levels. PKC activators, phorbol 12-myristate 13-acetate and L-o~-dioctanoylglycerol have been shown to decrease CBF in a concentration- and time-dependent manner in rabbit tracheal epithelial cells (Kobayashi et al., 1989). CBF has been shown to decrease after exposure to inhaled oxidants such as cigarette smoke across different species. A study with 120 subjects showed a significant decrease in nasal CBF following exposure to tobacco smoke (Agius et al., 1998). Exposure to cigarette smoke extract lead to reduction in forskolin-induced CBF in human sinonasal epithelium (Cohen et al., 2009) and isoproterenol- and methacholine-induced CBF in human adenoid tissues (Wang et al., 2012). This decrease in CBF and unresponsiveness to beta-agonist stimulation occurs in parallel to PKC activation and has been shown to be dependent on the duration of exposure to cigarette smoke in mice (Simet et al., 2010). Normal human bronchial epithelial cells exposed to aerosolized nicotine showed decreased CFTR and BK conductance, impaired CBF, ASL volume, and decreased expression of FOXJ1 and KCNMA1 (Garcia-Arcos et al., 2016). <br />
A concentration-dependent decrease in CBF has been observed after treatment with aldehydes. For example inhibition of cilia ATPase activity was observed after treatment with acetaldehyde, in ciliated bovine bronchial epithelial cells (Sisson et al., 1991). Acrolein, an aldehyde in the gas phase of cigarette smoke, induced ciliostasis at high concentrations (> 1 mM), after 5 min of treatment, and cellular necrosis after 3 hr. However, at lower concentrations (from 0.5‒1 mM), acrolein transiently reduced the CBF to 4 Hz (Romet et al., 1990).<br />
</p>
<h4>How it is Measured or Detected</h4>
<p>There is no standardized method for measuring CBF. Digital high-speed video imaging with a manual count of CBF in slow motion video play is the most commonly used method for CBF measurement (Kim et al., 2011; Peabody et al., 2018). Photometry and video-microscopy have been used to measure CBF in vitro and ex vivo, including in ciliated bovine bronchial epithelial cells (Allen-Gipson et al., 2011; Sisson et al., 2003; Uzlaner and Priel, 1999), normal human bronchial epithelial cells (Feriani et al., 2017), human nasal epithelial cells (Dimova et al., 2005; Min et al., 1999b), human nasal ciliated epithelium (nasal brushings) (Agius et al., 1998), and mouse tracheal rings (Simet et al., 2010).<br />
CBF measurement in vitro generally involves mounting the tissue at the air-liquid interface on a stage followed by microscopic analysis and acquisition of images and/or video recordings of beating cilia. For in vivo and ex vivo measurements, Doppler optical coherence tomography (D-OCT) can also be applied, a mesoscopic non-contact imaging modality that provides high-resolution tomographic images and detects micromotion simultaneously (Jing et al., 2017). D-OCT has been used to quantitatively measure CBF in ex vivo rabbit tracheal cultures (Lemieux et al., 2015).</p>
<h4>References</h4>
<p>Agius, A. M., L. A. Smallman, and A. L. Pahor (1998). Age, smoking and nasal ciliary beat frequency. Clin. Otolaryngol. Allied Sci. 23, 227-230.</p>
<p>Allen-Gipson, D.S., Blackburn, M.R., Schneider, D.J., Zhang, H., Bluitt, D.L., Jarrell, J.C., et al. (2011). Adenosine activation of A(2B) receptor(s) is essential for stimulated epithelial ciliary motility and clearance. Am. J. Physiol. Lung Cell. Mol. Physiol. 301, L171-L180.</p>
<p>Bailey, K.L., Bonasera, S.J., Wilderdyke, M., Hanisch, B.W., Pavlik, J.A., Devasure, J., et al. (2014). Aging causes a slowing in ciliary beat frequency, mediated by PKCε. Am. J. Physiol. Lung Cell. Mol. Physiol. 306, L584-L589.</p>
<p>Cohen, N.A., Zhang, S., Sharp, D.B., Tamashiro, E., Chen, B., Sorscher, E.J., et al. (2009). Cigarette smoke condensate inhibits transepithelial chloride transport and ciliary beat frequency. Laryngoscope 119, 2269-2274.</p>
<p>Di Benedetto, G., Manara-Shediac, F.S. and Mehta, A. (1991). Effect of cyclic AMP on ciliary activity of human respiratory epithelium. Eur. Respir. J. 4, 789-795.</p>
<p>Dimova, S., Maes, F., Brewster, M.E., Jorissen, M., Noppe, M. and Augustijns, P. (2005). High-speed digital imaging method for ciliary beat frequency measurement. J. Pharmacy Pharmacol 57, 521-526.</p>
<p>Feriani, L., Juenet, M., Fowler, C.J., Bruot, N., Chioccioli, M., Holland, S.M., et al. (2017). Assessing the Collective Dynamics of Motile Cilia in Cultures of Human Airway Cells by Multiscale DDM. Biophys. J. 113, 109-119.</p>
<p>Garcia-Arcos, I., Geraghty, P., Baumlin, N., Campos, M., Dabo, A.J., Jundi, B., et al. (2016). Chronic electronic cigarette exposure in mice induces features of COPD in a nicotine-dependent manner. Thorax 71, 1119-1129.</p>
<p>Gosepath, J., Schaefer, D., Brommer, C., Klimek, L., Amedee, R.G., and Mann, W.J. (2000). Subacute Effects of Ozone Exposure on Cultivated Human Respiratory Mucosa. Am. J. Rhinol. 14, 411-418. </p>
<p>Grubb, B.R., Livraghi-Butrico, A., Rogers, T.D., Yin, W., Button, B. and Ostrowski, L.E. (2016). Reduced mucociliary clearance in old mice is associated with a decrease in Muc5b mucin. Am. J. Physiol. Lung Cell. Mol. Physiol. 310, L860-L867.</p>
<p>Ho, J.C., Chan, K.N., Hu, W.H., Lam, W.K., Zheng, L., Tipoe, G.L., et al. (2001). The Effect of Aging on Nasal Mucociliary Clearance, Beat Frequency, and Ultrastructure of Respiratory Cilia. Am. J. Respir. Crit. Care Med. 163, 983-988.</p>
<p>Jain, R., Ray, J.M., Pan, J.-H. and Brody, S.L. (2012). Sex hormone-dependent regulation of cilia beat frequency in airway epithelium. Am. J. Respir. Crit. Care Med. 46, 446-453.</p>
<p>Jia, S., Zhang, X., He, D.Z., Segal, M., Berro, A., Gerson, T., et al., 2011. Expression and Function of a Novel Variant of Estrogen Receptor–α36 in Murine Airways. Am. J. Respir. Cell Mol. Biol. 45, 1084-1089.</p>
<p>Jiao, J., Wang, H., Lou, W., Jin, S., Fan, E., Li, Y., et al. (2011). Regulation of ciliary beat frequency by the nitric oxide signaling pathway in mouse nasal and tracheal epithelial cells. Exp. Cell Res. 317, 2548-2553.</p>
<p>Jing, J.C., Chen, J.J., Chou, L., Wong, B.J.F. and Chen, Z. (2017). Visualization and Detection of Ciliary Beating Pattern and Frequency in the Upper Airway using Phase Resolved Doppler Optical Coherence Tomography. Sci. Rep. 7, 8522-8522.</p>
<p>Joki, S. and Saano, V. (1997). Influence of ageing on ciliary beat frequency and on ciliary response to leukotriene D4 in guinea‐pig tracheal epithelium. Clin. Exp. Pharmacol. Physiol. 24, 166-169.</p>
<p>Kim, W., Han, T.H., Kim, H.J., Park, M.Y., Kim, K.S. and Park, R.W. (2011). An Automated Measurement of Ciliary Beating Frequency using a Combined Optical Flow and Peak Detection. J. Healthc. Inform. Res. 17, 111-119.</p>
<p>Knoll, M., Shaoulian, R., Magers, T. and Talbot, P. (1995). Ciliary beat frequency of hamster oviducts is decreased in vitro by exposure to solutions of mainstream and sidestream cigarette smoke. Biol. Reprod. 53, 29-37.</p>
<p>Kobayashi, K., Tamaoki, J., Sakai, N., Chiyotani, A. and Takizawa, T. (1989). Inhibition of ciliary activity by phorbol esters in rabbit tracheal epithelial cells. Lung 167, 277-284.</p>
<p>Kogiso, H., Hosogi, S., Ikeuchi, Y., Tanaka, S., Inui, T., Marunaka, Y., et al. (2018). [Ca(2+) ]i modulation of cAMP-stimulated ciliary beat frequency via PDE1 in airway ciliary cells of mice. Exp. Physiol. 103, 381-390.</p>
<p>Lansley, A.B., Sanderson, M.J. and Dirksen, E.R. (1992). Control of the beat cycle of respiratory tract cilia by Ca2+ and cAMP. Am. J. Physiol. 263, L232-242.</p>
<p>Lemieux, B.T., Chen, J.J., Jing, J., Chen, Z. and Wong, B.J.F. (2015). Measurement of ciliary beat frequency using Doppler optical coherence tomography. Int. Forum Allergy Rhinol. 5, 1048-1054.</p>
<p>Li, D., Shirakami, G., Zhan, X. and Johns, R.A. (2000). Regulation of ciliary beat frequency by the nitric oxide-cyclic guanosine monophosphate signaling pathway in rat airway epithelial cells. Am. J. Respir. Cell Mol. Biol. 23, 175-181.</p>
<p>Min, Y.-G., Ohyama, M., Lee, K.S., Rhee, C.-S., Oh, S.H., Sung, M.-W., et al. (1999). Effects of free radicals on ciliary movement in the human nasal epithelial cells. Auris Nasus Larynx 26, 159-163.</p>
<p>Paul, P., Johnson, P., Ramaswamy, P., Ramadoss, S., Geetha, B. and Subhashini, A.S. (2013). The Effect of Ageing on Nasal Mucociliary Clearance in Women: A Pilot Study. ISRN Pulmonology 2013, 5.</p>
<p>Romet, S., Dubreuil, A., Baeza, A., Moreau, A., Schoevaert, D. and Marano, F. (1990). Respiratory tract epithelium in primary culture: Effects of. Toxicol. In Vitro 4, 399-402.</p>
<p>Salathe, M., Pratt, M.M. and Wanner, A. (1993). Cyclic AMP-dependent phosphorylation of a 26 kD axonemal protein in ovine cilia isolated from small tissue pieces. Am. J. Respir. Cell Mol. Biol. 9, 306-314.</p>
<p>Sanderson, M.J. and Dirksen, E.R. (1989). Mechanosensitive and beta-adrenergic control of the ciliary beat frequency of mammalian respiratory tract cells in culture. Am. Rev. Respir. Dis. 139, 432-440.</p>
<p>Schmid, A., Sutto, Z., Nlend, M.-C., Horvath, G., Schmid, N., Buck, J., et al. (2007). Soluble Adenylyl Cyclase Is Localized to Cilia and Contributes to Ciliary Beat Frequency Regulation via Production of cAMP. J. Gen. Physiol. 130, 99-109.</p>
<p>Schmid, A., Baumlin, N., Ivonnet, P., Dennis, J.S., Campos, M., Krick, S., et al. (2015). Roflumilast partially reverses smoke-induced mucociliary dysfunction. Respir. Res. 16, 135.</p>
<p>Simet, S.M., Sisson, J.H., Pavlik, J.A., Devasure, J.M., Boyer, C., Liu, X., et al. (2010). Long-term cigarette smoke exposure in a mouse model of ciliated epithelial cell function. Am. J. Respir. Cell Mol. Biol. 43, 635-640.</p>
<p>Sisson, J.H. (1995). Ethanol stimulates apparent nitric oxide-dependent ciliary beat frequency in bovine airway epithelial cells. Am. J. Physiol. 268, L596-600.</p>
<p>Sisson, J.H., May, K. and Wyatt, T.A. (1999). Nitric oxide-dependent ethanol stimulation of ciliary motility is linked to cAMP-dependent protein kinase (PKA) activation in bovine bronchial epithelium. Alcohol Clin. Exp. Res. 23, 1528-1533.</p>
<p>Sisson, J.H., Stoner, J., Ammons, B. and Wyatt, T. (2003). All‐digital image capture and whole‐field analysis of ciliary beat frequency. J. Microsc. 211, 103-111.</p>
<p>Sisson, J.H., Tuma, D.J. and Rennard, S.I. (1991). Acetaldehyde-mediated cilia dysfunction in bovine bronchial epithelial cells. Am. J. Physiol. 260, L29-36.</p>
<p>Svartengren, M., Falk, R. and Philipson, K. (2005). Long-term clearance from small airways decreases with age. Eur. Respir. J. 26, 609-615.</p>
<p>Tamaoki, J., Kondo, M. and Takizawa, T. (1989). Effect of cAMP on ciliary function in rabbit tracheal epithelial cells. J. Appl. Physiol. 66, 1035-1039.</p>
<p>Uzlaner, N. and Priel, Z. (1999). Interplay between the NO pathway and elevated [Ca(2+)](i) enhances ciliary activity in rabbit trachea. J. Physiol. 516, 179-190.</p>
<p>Wang, L.F., White, D.R., Andreoli, S.M., Mulligan, R.M., Discolo, C.M. and Schlosser, R.J. (2012). Cigarette smoke inhibits dynamic ciliary beat frequency in pediatric adenoid explants. Otolaryngol. Head Neck Surg. 146, 659-663.</p>
<p>Yaghi, A., Zaman, A., Cox, G. and Dolovich, M.B. (2012). Ciliary beating is depressed in nasal cilia from chronic obstructive pulmonary disease subjects. Respir. Med. 106, 1139-1147.</p>
<p>Yang, B., Schlosser, R.J. and Mccaffrey, T.V. (1997). Signal transduction pathways in modulation of ciliary beat frequency by methacholine. Ann. Otol. Rhinol. Laryngol. 106, 230-236.</p>
<tr><th scope="col">Level of Biological Organization</th></tr>
</thead>
<tbody class="tbody-striped">
<tr><td>Individual</td></tr>
</tbody>
</table>
</div>
<h3>Evidence for Perturbation by Stressor</h3>
<h4>Sulfur dioxide</h4>
<p><p>SO<sub>2</sub> exposure of dogs dose-dependently decreased CBF and also caused a marked decrease in mean bronchial mucociliary clearance (from 53.7 ± 5.7% to 32.8 ± 7.7%) after 90 min (Yeates et al., 1997). In guinea pig tracheas, SO<sub>2</sub> exposure affected CBF, albeit non-significantly, and mucociliary activity (Knorst et al., 1994).</p>
</p>
<h4>Formaldehyde</h4>
<p><p>Treatment of frog palate epithelium with different concentrations of formaldehyde induced significant decreases in CBF and MCC (Fló-Neyret et al., 2001; Morgan et al., 1984). Exposure of F344 rats to formaldehyde caused epithelial adaptation of the nasal epithelium, effectively reducing the number of ciliated cells (and hence cilia beating activity) through squamous metaplasia. At the same time, formaldehyde exposure resulted in “ciliastasis” or loss of ciliary activity in a concentration- and exposure duration-dependent manner as well as in a slowing of mucus flow rates (Morgan et al., 1986). </p>
</p>
<h4>PM10</h4>
<p><p>Incubation of frog palates with PM10 from Sao Paolo, Brazil, for up to 120 min decreased mucociliary transport at concentrations ≥1000 pg/m<sup>3</sup> (Macchione et al., 1999).</p>
</p>
<h4>Nitric oxide</h4>
<p><p>In New Zealand white rabbits exposed to 3 ppm NO<sub>2</sub> for 24 h, the average CBF decreased from 764 beats/min to 692 beats/min and the transport velocity decreased from 5.23 mm/min to 3.03 mm/min (Kakinoki, 1998).</p>
</p>
<h4>Ozone</h4>
<p><p>Acute exposure (2 h) of adult ewes to 1.0 ppm ozone significantly reduced tracheal mucus transport velocity (TMV) at 40 min and 2 h post-exposure. Repeated exposure to 1.0 ppm ozone for 5 hper day, for 4 consecutive days showed a progressively significant decrease in TMV on the first and second days, and stabilized over the third and fourth days, around values ranging from -42% to -55% of the initial baseline. TMV remained depressed even after the end of exposure, persisting up to 5 days post-exposure (Allegra et al., 1991).<br />
</p>
</p>
<h4>Cigarette smoke</h4>
<p><p>Nasomuciliary clearance time (determined by saccharin transit test) was significantly higher in smokers than in non-smokers 8 h after smoking (16 ± 6 min vs 10 ± 4 min) and insignicantly higher immediately after smoking (11 ± 6 min vs 10 ± 4 min). Nasomuciliary clearance time correlated positively with cigarettes per day and packs/year index (Proença et al., 2011).</p>
<p>In a small Indian cross-sectional study, the mean nasomuciliary clearance (determined by saccharin transit test) in smokers was significantly higher than that of nonsmokers (481.2 ± 29.83 s vs 300.32 ± 17.4 s). In addition, mean nasomuciliary clearance increased as the duration of smoking increased (NMC in smoking <1 year = 492.25 ± 79.93 s, NMC in smoking for 1-5 years = 516.7 ± 34.01 s, and NMC in smoking >5 years = 637.5 ± 28.49 s) (Baby et al., 2014).</p>
<p>Nasomuciliary clearance (determined by saccharin transit test) in active and passive smokers was significantly higher than in non-smokers (23.08 ± 4.60 min; 20.31 ± 2.51 min vs 8.57 ± 2.12 min) (Yadav et al., 2014).</p>
<p>Nasomuciliary clearance (determined by saccharin transit test) was significantly higher in active smokers than in passive smokers and non-smokers (23.59 ± 12.41 min vs 12.6 ± 4.67 min; 6.4 ± 1.55 min) (Habesoglu et al., 2012).</p>
<p>Nasomuciliary clearance time (determined by saccharin transit test) in smokers was significantly higher than in former smokers and non-smokers (15.6 min vs 11.77 min and 11.71 min, respectively) (Pagliuca et al., 2015).</p>
<p>Moderate and heavy smokers had higher saccharin transit test times than light smokers and non-smokers, and there was a positive correlation between STT and cigarettes/day (Xavier et al., 2013).</p>
<p>The median nasal mucociliary clearance time (determined by saccharin transit test) was significantly higher in smokers (who smoked a mean of 20.6 cigarettes (median: 20) per day) than in nonsmokers (12 (interquartile range: 5–33) min vs 9 (interquartile range: 4–12) min) (Dülger et al., 2018). </p>
<p>Nasal mucociliary clearance time (determined by saccharin transit test) in smokers was significantly higher than in non-smokers (536.19 ± 254.81 s vs 320.43 ± 184.98 s) and correlated with the numbers of cigarettes per day, pack-years and smoking duration (Solak et al., 2018).</p>
<p>Current smokers had a median (IQR) mucociliary clearance transit time (determined by saccharin transit test) of 13.15 (9.89–16.08) min, which was significantly longer compared with that of never smokers at 7.24 (5.73–8.73) min, former smokers at 7.26 (6.18–9.17) min, exclusive e-cigarette users at 7.00 (6.38–9.00) min, and exclusive heated tobacco product users at 8.00 (6.00–8.00) min (Polosa et al., 2021).</p>
<p>In healthy adults, tracheal mucus movement varies from 4 to >20 mm/min (Stannard and O'Callaghan, 2006), whereas mucociliary clearance (MCC) in the small airways is slower due to the lower number of ciliated cells (fewer cilia) and their shorter length (Foster et al., 1980; Iravani, 1969; Wanner et al., 1996).<br />
Since optimal MCC is dependent in multiple factors, including cilia number and structure as well as ASL and mucus properties, any disturbances of these can lead to impaired MCC. While high humidity or infection can enhance MCC, long-term exposure to noxious substances (e.g. cigarette smoke) lead to decreased mucus clearance from the airways. In most instances this is reflected by decreased mucus transport rates or velocities.<br />
</p>
<h4>How it is Measured or Detected</h4>
<p>In humans, MCC has been assessed traditionally following inhalation of radio-labeled particles such as 99Tcm-labeled polystyrene particles, resin particles or serum albumin and following their clearance at regular intervals by radioimaging using gamma cameras (Agnew et al., 1986; Kärjä et al., 1982). Taking into account inhalation volumes and flow rates, lung airflow, particle deposition and retention, clearance rates can be calculated and effects of e.g. drugs on MCC can be examined. Alternatively, since MCC occurs at a similar rate in the nose to that in trachea and bronchi (Andersen and Proctor, 1983; Rutland and Cole, 1981) and for ease of use, measurements of MCC can be restricted to that of nasal MCC only. Probably one of the simplest methods is the saccharin transit test (STT). For this test, a small particle of saccharin is placed behind the anterior end of the inferior turbinate. The saccharin will be transported by mucociliary action toward the nasopharynx, where its sweet taste is perceived. When MCC is impaired, saccharin transit times will increase, with a 10- to 20-minute delay being considered a clinical sign of decreased MCC. Using the same principle, the test can also be performed or complemented with dyes such as indigo carmine or methylene blue (Deborah and Prathibha, 2014).</p>
<p>In experimental animals, MCC has been evaluated by gamma-scintigraphy (Greiff et al., 1990; Hua et al., 2010; Read et al., 1992), fluorescence videography/fluoroscopy (in explanted tracheas etc.) (Grubb et al., 2016; Rogers et al., 2018), or by 3D-SPECT (Ortiz Belda et al., 2016). Direct observation of particle movement across airway epithelia to determine mucus velocity or transport rates by using a fiberoptic bronchoscope may be helpful when working in larger animals such as dogs (King, 1998).<br />
In vitro, freshly excised frog palate preparations have been used to assess cilia function and mucociliary transport by videomicroscopy (Macchione et al., 1995; Macchione et al., 1999; Trindade et al., 2007). Murine and human nasal, bronchial and small airway epithelial models grown at the air-liquid interface are also suitable in vitro test systems for determining mucus transport by tracing inert particle movement with a set-up similar to that used for assessing CBF (Benam et al., 2018; Fliegauf et al., 2013; Knowles and Boucher, 2002; Sears et al., 2015).<br />
</p>
<h4>References</h4>
<p>Agnew, J., Sutton, P., Pavia, D. and Clarke, S. (1986). Radioaerosol assessment of mucociliary clearance: towards definition of a normal range. Brit. J. Radiol. 59, 147-151.</p>
<p>Allegra, L., Moavero, N., and Rampoldi, C. (1991). Ozone-induced impairment of mucociliary transport and its prevention with N-acetylcysteine. Am. J. Med. 91, S67-S71.</p>
<p>Andersen, I. and Proctor, D. (1983). Measurement of nasal mucociliary clearance. Eur. J. Respir. Dis. Suppl. 127, 37-40.</p>
<p>Baby, M.K., Muthu, P.K., Johnson, P., and Kannan, S. (2014). Effect of cigarette smoking on nasal mucociliary clearance: A comparative analysis using saccharin test. Lung India 31, 39-42. </p>
<p>Benam, K.H., Vladar, E.K., Janssen, W.J. and Evans, C.M. (2018). Mucociliary defense: emerging cellular, molecular, and animal models. Ann. Am. Thorac. Soc. 15, S210-S215.</p>
<p>Deborah, S. and Prathibha, K., 2014. Measurement of nasal mucociliary clearance. Clin. Res. Pulmonol. 2, 1019.</p>
<p>Dülger, S., Akdeniz, Ö., Solmaz, F., Şengören Dikiş, Ö., and Yildiz, T. (2018). Evaluation of nasal mucociliary clearance using saccharin test in smokers: A prospective study. Clin. Respir. J. 12, 1706-1710. </p>
<p>Fliegauf, M., Sonnen, A.F.P., Kremer, B. and Henneke, P. (2013). Mucociliary Clearance Defects in a Murine In Vitro Model of Pneumococcal Airway Infection. PloS ONE 8, e59925.</p>
<p>Fló-Neyret, C., Lorenzi-Filho, G., Macchione, M., Garcia, M.L.B. and Saldiva, P.H.N. (2001). Effects of formaldehyde on the frog's mucociliary epithelium as a surrogate to evaluate air pollution effects on the respiratory epithelium. Braz. J. Med. Biol. Res. 34, 639-643.</p>
<p>Foster, W., Langenback, E. and Bergofsky, E. (1980). Measurement of tracheal and bronchial mucus velocities in man: relation to lung clearance. J. Appl. Physiol. 48, 965-971.</p>
<p>Greiff, L., Wollmer, P., Erjefält, I., Pipkorn, U. and Persson, C. (1990). Clearance of 99mTc DTPA from guinea pig nasal, tracheobronchial, and bronchoalveolar airways. Thorax 45, 841-845.</p>
<p>Grubb, B.R., Livraghi-Butrico, A., Rogers, T.D., Yin, W., Button, B. and Ostrowski, L.E. (2016). Reduced mucociliary clearance in old mice is associated with a decrease in Muc5b mucin. Am. J. Physiol. Lung Cell. Mol. Physiol. 310, L860-L867.</p>
<p>Habesoglu, M., Demir, K., Yumusakhuylu, A.C., Sahin Yilmaz, A., and Oysu, C. (2012). Does passive smoking have an effect on nasal mucociliary clearance? Otolaryngol Head Neck Surg. 147, 152-156.</p>
<p>Hua, X., Zeman, K.L., Zhou, B., Hua, Q., Senior, B.A., Tilley, S.L., et al. (2010). Noninvasive real-time measurement of nasal mucociliary clearance in mice by pinhole gamma scintigraphy. J. Appl. Physiol. 108, 189-196.</p>
<p>Iravani, J. (1969). Zum Mechanismus der Ortsabhängigkeit der Flimmeraktivität im Bronchialbaum/Location-Dependent Activity of the Ciliary Movement in the Bronchial Tree and its Possible Mechanism. In: Habermann E. et al. (eds) Naunyn Schmiedebergs Archiv für Pharmakologie. Springer, Berlin, Heidelberg.</p>
<p>Kakinoki Y, Ohashi Y, Tanaka A, Washio Y, Yamada K, Nakai Y, Morimoto K. (1998). Nitrogen dioxide compromises defence functions of the airway epithelium. Acta Oto-Laryngol. 118, 221-226.</p>
<p>Kärjä, J., Nuutinen, J. and Karjalainen, P. (1982). Radioisotopic Method for Measurement of Nasal Mucociliary Activity. Arch. Otolaryngol. 108, 99-101.</p>
<p>King, M. (1998). Experimental models for studying mucociliary clearance. Eur. Respir. J. 11, 222-228.</p>
<p>Knorst, M.M., Kienast, K., Riechelmann, H., Müller-Quernheim, J. and Ferlinz, R. (1994). Effect of sulfur dioxide on mucociliary activity and ciliary beat frequency in guinea pig trachea. Int. Arch. Occup. Environm. Health 65, 325-328.</p>
<p>Knowles, M.R. and Boucher, R.C. (2002). Mucus clearance as a primary innate defense mechanism for mammalian airways. J. Clin. Invest. 109, 571-577.</p>
<p>Macchione, M., Guimarães, E., Saldiva, P. and Lorenzi-Filho, G. (1995). Methods for studying respiratory mucus and mucus clearance. Braz. J. Med. Biol Res. 28, 1347.</p>
<p>Macchione, M., Oliveira, A.P., Gallafrio, C.T., Muchão, F.P., Obara, M.T., Guimarães, E.T., et al. (1999). Acute effects of inhalable particles on the frog palate mucociliary epithelium. Environm. Health Persp. 107, 829-833.</p>
<p>Morgan, K., Patterson, D. and Gross, E. (1986). Responses of the nasal mucociliary apparatus of F-344 rats to formaldehyde gas. Toxicol. Appl. Pharmacol. 82, 1-13.</p>
<p>Morgan, K.T., Patterson, D.L. and Gross, E.A. (1984). Frog palate mucociliary apparatus: structure, function, and response to formaldehyde gas. Fund. Appl. Toxicol. 4, 58-68.</p>
<p>Ortiz Belda, J.L., Ortiz, A., Milara Payá, J., Armengot Carceller, M., Sanz García, C., Compañ Quilis, D., et al. (2016). Evaluation of Mucociliary Clearance by Three Dimension Micro-CT-SPECT in Guinea Pig: Role of Bitter Taste Agonists. Plos ONE 11, e0164399.</p>
<p>Pagliuca, G., Rosato, C., Martellucci, S., De Vincentiis, M., Greco, A., Fusconi, M., et al. (2015). Cytologic and functional alterations of nasal mucosa in smokers: temporary or permanent damage? Otolaryngol Head Neck Surg 152, 740-745.</p>
<p>Proença, M., Xavier, R.F., Ramos, D., Cavalheri, V., Pitta, F., and Ramos, E.C. (2011). Immediate and short term effects of smoking on nasal mucociliary clearance in smokers. Revista Portuguesa de Pneumologia (English Edition) 17), 172-176.</p>
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<p>Rutland, J. and Cole, P.J. (1981). Nasal mucociliary clearance and ciliary beat frequency in cystic fibrosis compared with sinusitis and bronchiectasis. Thorax 36, 654-658.</p>
<p>Sears, P.R., Yin, W.-N. and Ostrowski, L.E. (2015). Continuous mucociliary transport by primary human airway epithelial cells in vitro. Am. J. Physiol. Lung Cell. Mol. Physiol. 309, L99-L108.</p>
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<p>Stannard, W. and O'callaghan, C. (2006). Ciliary function and the role of cilia in clearance. J. Aerosol Med. 19, 110-115.</p>
<p>Trindade, S.H.K., De Mello Júnior, J.F., De Godoy Mion, O., Lorenzi-Filho, G., Macchione, M., Guimarães, E.T., et al. (2007). Methods for Studying Mucociliary Transport. Braz. J. Otorhinolaryngol. 73, 704-712.</p>
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<p>Yeates, D.B., Katwala, S.P., Daugird, J., Daza, A.V. and Wong, L.B. (1997). Excitatory and inhibitory neural regulation of tracheal ciliary beat frequency (CBF) activated by ammonia vapour and SO2. Ann. Occup. Hyg. 41, 736-744.</p>
<tr><th scope="col">Level of Biological Organization</th></tr>
</thead>
<tbody class="tbody-striped">
<tr><td>Individual</td></tr>
</tbody>
</table>
</div>
<h3>Evidence for Perturbation by Stressor</h3>
<h4>Ozone</h4>
<p><p>Acute exposure of healthy young adult subjects (aged 19 to 35 years, non-smokers) to 0.06 ppm ozone for 6.6 h resulted in a 1.71 <u>+</u> 0.50% (mean <u>+</u> SEM) decrease in FEV1 and a 2.32 <u>+</u> 0.41% decrease in FVC compared with air exposure (Kim et al., 2011).</p>
<p>A US-based study found inverse associations between increasing lifetime exposure to ozone (estimated median: 36; interquartile range 29–45; range 19–64) and FEF75 and FEF25–75 in adolescents (aged 18–20 years) (Tager et al., 2005).</p>
</p>
<h4>Nitric oxide</h4>
<p><p>In a Dutch cross-sectional study in school children (aged 7–13 years), NOx exposure from industrial emissions per interquartile range of 7.43 μg/m<sup>3</sup> had a significantly lower percent predicted peak expiratory flow (PEF) (-3.67%, 95%CI -6.93% to -0.42%). Children exposed to NOx (per interquartile range of 7.43 μg/m<sup>3</sup>) also had a significantly lower percent forced vital capacity (FVC) and percent predicted 1-s forced expiratory volume (FEV1) (− 2.73 95%CI -5.21 to -0.25) (Bergstra et al., 2018). </p>
<p>The European Study of Cohorts for Air Pollution Effects (ESCAPE), a meta-analysis of 5 cohort studies on the association of air pollution with lung function, found that a 10 μg/m<sup>3</sup> increase in NO<sub>2</sub> exposure was associated with lower levels of FEV1 (−14.0 mL, 95% CI −25.8 to −2.1) and FVC (−14.9 mL, 95% CI −28.7 to −1.1), and an increase of 20 μg/m<sup>3</sup> in NOx exposure was associated with a lower level of FEV1, by −12.9 mL (95% CI −23.87 to −2.0) and of FVC, by −13.3 mL (95% CI −25.9 to −0.7) (Adam et al., 2015).</p>
</p>
<h4>Cigarette smoke</h4>
<p><p>A smoking history of > 20 pack-years decreased pulmonary function including forced vital capacity (FVC), forced expiratory volume in one second (FEV1), FEV1/FVC, and forced expiratory flow at 25–75% (FEF25–75%) (Kuperman and Riker, 1973).</p>
<p>In the Framingham Heart Study, cigarette smoking showed an inverse association with FVC and FEV1% (Ashley et al., 1975).</p>
<p>In the international Seven Countries Study, there was a dose-effect relationship between pack-years and forced expiratory volume in 0.75 s (FEV0.75) in continuous smokers without chronic bronchitis (Pelkonen et al., 2006).</p>
<p>In 34 male subjects aged between 15–18 years who smoked FVC was lower than in an age-matched male group that did not smoke. The most common duration of cigarette smoking was 1–3 years (47%) and the maximal number of cigarettes smoked per day was less than or equal to 10 cigarette(s) per day (88%) (Tantisuwat and Thaveeratitham, 2014). </p>
<p>A dose–response relation was found between smoking and lower levels of FEV1/FVC and FEF25–75 in children between 10-18 years of age (Gold et al., 1996).</p>
<p>In a study of 147 asthmatics, FEV1%predicted was significantly lower in ex-smokers and current smokers compared with never-smokers (Broekema et al., 2009).</p>
<p>In a 6-year longitudinal study in Japanese-American men, FEV1 was lowest in current smokers (2702 mL) and in former smokers (2817 mL) at baseline. These 2 groups experienced a steeper annual decline in FEV1 (-34.4 and -22.8 mL/year, respectively, adjusted by height and age at baseline) compared with never-smokers (-20.3 mL/year) (Burchfiel et al., 1995).</p>
</p>
<h4>Diesel engine exhaust</h4>
<p><p>In a study of 733 adult females who had lived in the Tokyo metropolitan area for more than 3 years, the higher the level of air pollution, the more significantly the FEV1 was reduced (Sekine et al., 2004).</p>
<p>In a study in 29 healthy subjects, exposure to DE inside diesel-powered trains for 3 days was associated with reduced lung function (Andersen et al., 2019).</p>
<p>In workers who tested diesel engines in an assembly unit of a manufacturing plant, FEV1, FEV1/FVC, FEV25-75 and MEF were significantly reduced compared to non-exposed workers (Zhang et al., 2017).</p>
</p>
<h4>PM10</h4>
<p><p>A Taiwanese study in 1016 children between 6 and 15 years of age reported that lifetime exposure to to 25–85 μg/m<sup>3</sup> PM10 were associated with lower FEV1, FVC, and FEF25-75 (Tsui et al., 2018).</p>
<p>The Swiss Study on Air Pollution and Lung Diseases in Adults (SAPALDIA) found that an increase of 10 μg/m<sup>3</sup> in annual mean concentration of PM10 was associated with 3.4% lower FVC and 1.6% lower FEV1 (Ackermann-Liebrich et al., 1997).</p>
<p>In the Health Survey for England, a 10 mg/m<sup>3</sup> difference in PM10 across postcode sectors was associated with a lower FEV1 by 111 mL, independent of active and passive smoking, social class, region and month of testing (Forbes et al., 2009). </p>
<p>A 7 μg/m<sup>3</sup> increase in five year means of PM10 (interquartile range) was associated with a 5.1% (95% CI: 2.5%–7.7%) decrease in FEV1, a 3.7% (95% CI: 1.8%–5.5%) decrease in FVC in the German SALIA study (Schikowski et al., 2005).</p>
<p>The ESCAPE project, a meta-analysis of 5 European cohorts/studies from 8 countries, reported that an increase of 10 μg/m<sup>3</sup> in PM10 was associated with a lower level of FEV1 (−44.6 mL, 95% CI:−85.4– −3.8) and FVC (−59.0 mL, 95% CI: −112.3– −5.7) (Adam et al., 2015).</p>
<p><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Pulmonary function tests reflect the physiological working of the lungs. Therefore, the AO is applicable to a variety of species, including (but not limited to) rodents, rabbits, pigs, cats, dogs, horses and humans, independent of life stage and gender.</span></span></span></span></span></p>
<h4>Key Event Description</h4>
<p style="text-align:justify"><span style="font-size:11pt"><span style="font-family:Calibri,sans-serif"><span style="font-size:12.0pt"><span style="font-family:"Segoe UI",sans-serif">Lung function is a clinical term referring to the physiological functioning of the lungs, most often in association with the tests used to assess it. Lung function loss can be caused by acute or chronic exposure to airborne toxicants or by an intrinsic disease of the respiratory system. </span></span></span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Although signs of cellular injury are typically exhibited first in the nose and larynx, alveolar-capillary barrier breakdown may ultimately arise and result in local edema (Miller and Chang, 2003). Clinically, bronchoconstriction and hypoxia are seen in the acute phase, with affected subjects exhibiting shortness of breath (dyspnea) and low blood oxygen saturation, and with reduced lung function indices of airflow, lung volume and gas exchange (Hert and Albert, 1994; and How it is Measured or Detected;). When alveolar damage is extensive, the reduced lung function can develop into acute respiratory distress syndrome (ARDS). This severe compromise of lung function is reflected by decreased gas exchange indices (PaO<sub>2</sub>/FIO<sub>2</sub> ≤200 mmHg, due to hypoxemia and impaired excretion of carbon dioxide), increased pulmonary dead space and decreased respiratory compliance (Matthay et al., 2019). Acute inhalation exposures to chemical irritants such as ammonia, hydrogen chloride, nitrogen oxides and ozone typically cause local edema that manifests as dyspnea and hypoxia. In cases where a breakdown of the alveolar capillary function ensues, ARDS develops. ARDS has a particularly high risk of mortality, estimated to be 30-40% (Gorguner and Akgun, 2010; Matthay et al., 2018; Reilly et al., 2019).</span></span></span></span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Lung function decrease due to reduction in lung volume is seen in pulmonary fibrosis, which can be linked to chronic exposures to e.g. silica, asbestos, metals, agricultural and animal dusts (Meltzer and Noble, 2008; Cheresh et al., 2013; Cosgrove, 2015; Trethewey and Walters, 2018). Additionally. decreased lung function occurs in pleural disease, chest wall and neuromuscular disorders, because of obesity and following pneumectomy (Moore, 2012). Decreased lung function can also be a result of narrowing of the airways by inflammation and mucus plugging resulting in airflow limitation. Decreased lung function is a feature of obstructive pulmonary diseases (e.g. asthma, COPD) and linked to a multitude of causes, including chronic exposure to cigarette smoke, dust, metals, organic solvents, asbestos, pathogens or genetic factors. </span></span></span></span></span></p>
<p style="text-align:justify"> </p>
<h4>How it is Measured or Detected</h4>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Pulmonary function tests are a group of tests that evaluate several parameters indicative of lung size, air flow and gas exchange. Decreased lung function can manifest in different ways, and individual circumstances, including potential exposure scenarios, determine which test is used. The section outlines the tests used to evaluate lung function in humans (https://www.nhlbi.nih.gov/health-topics/pulmonary-function-tests, accessed 22 March 2021) and in experimental animals.</span></span></span></span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><strong><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Lung function tests used to evaluate human lung function</span></span></strong></span></span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">The most common (“gold standard”) lung function test in human subjects is spirometry. Spirometry results are primarily used for diagnostic purposes, e.g. to discriminate between obstructive and restrictive lung diseases, and for determining the degree of lung function impairment. Specific criteria for spirometry tests have been outlined in the American Thoracic Society (ATS) and the European Respiratory Society (ERS) Task Force guidelines (Graham et al., 2019). These guidelines consist of detailed recommendations for the preparation and conduct of the test, instruction of the person tested, as well as indications and contraindications, and are complemented by additional guidance documents on how to interpret and report the test results (Pellegrino et al., 2005; Culver et al., 2017).</span></span></span></span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Spirometry measures several different parameters during forceful exhalation, including:</span></span></span></span></span></p>
<ul>
<li style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Forced expiratory volume in 1 s (FEV1), the maximum volume of air that can forcibly be exhaled during the first second following maximal inhalation</span></span></span></span></span></li>
<li style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Forced vital capacity (FVC), the maximum volume of air that can forcibly be exhaled following maximal inhalation </span></span></span></span></span></li>
<li style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Vital capacity (VC), the maximum volume of air that can be exhaled when exhaling as fast as possible</span></span></span></span></span></li>
<li style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Peak expiratory flow (PEF), the maximal flow that can be exhaled when exhaling at a steady rate</span></span></span></span></span></li>
<li style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Forced expiratory flow, also known as mid-expiratory flow; the rates at 25%, 50% and 75% FVC are given</span></span></span></span></span></li>
<li style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Inspiratory vital capacity (IVC), the maximum volume of air that can be inhaled after a full expiration</span></span></span></span></span></li>
</ul>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">A reduced FEV1, with normal or reduced VC, normal or reduced FVC, and a reduced FEV1/FVC ratio are indices of airflow limitation, i.e., airway obstruction as seen in COPD (Moore, 2012). In contrast, airway restriction is demonstrated by a reduction in FVC, normal or increased FEV1/FVC ratio, a normal spirometry trace and potentially a high PEF (Moore, 2012).</span></span></span></span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Lung capacity or lung volumes can be measured using one of three basic techniques: 1) plethysmography, 2) nitrogen washout, or 3) helium dilution. Plethysmography consists of a series of sequential measurements in a body plethysmograph, starting with the measurement of functional residual capacity (FRC),</span></span> <span style="font-family:"Segoe UI",sans-serif"><span style="color:black">the volume of gas present in the lung at end-expiration during tidal breathing. Once the FRC is known, expiratory reserve volume (ERV; the volume of gas that can be maximally exhaled from the end-expiratory level during tidal breathing, i.e., the FRC), vital capacity (VC; the volume change at the mouth between the positions of full inspiration and complete expiration), and inspiratory capacity (IC; the maximum volume of air that can be inhaled from FRC) are determined, and total lung capacity (TLC;</span></span> <span style="font-family:"Segoe UI",sans-serif"><span style="color:black">the volume of gas in the lungs after maximal inspiration, or the sum of all volume compartments) and residual volume (RV; the volume of gas remaining in the lung after maximal exhalation) are calculated (Weinstock and McCannon, 2017).</span></span></span></span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">The other two techniques used to measure lung volumes—helium dilution and nitrogen washout—are based on the principle of conservation of mass: [initial gas concentration] x [initial volume of the system] = [final gas concentration] x [final volume of the system]. The nitrogen washout method is based on the fact that nitrogen is present in the air, at a relatively constant amount. The subject is given 100% oxygen to breathe, and the expired gas, which contains nitrogen in the lung at the beginning of the test, is collected. When no more nitrogen is noted in the expirate, the volume of air expired and the entire amount of nitrogen in that volume are measured, and the initial volume of the system (FRC) can be calculated. In the helium dilution method, a known volume and concentration of helium is inhaled by the subject. Helium, an inert gas that is not absorbed significantly from the lungs, is diluted in proportion to the lung volume to which it is added. The final concentration of helium is then measured and FRC calculated (Weinstock and McCannon, 2017).</span></span></span></span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Measurements of lung volumes in humans are technically more challenging than spirometry. However, they complement spirometry (which cannot determine lung volumes) and may be a preferred means of lung function assessment when subject compliance cannot be reasonably expected (e.g. in pediatric subjects) or where forced expiratory maneuvers are not possible (e.g. in patients with advanced pulmonary fibrosis). There are recommended standards for lung volume measurements and their interpretation in clinical practice, issued by the ATS/ERS Task Force (Wanger et al., 2005; Criée et al., 2011). </span></span></span></span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Finally, indices of gas exchange across the alveolar-capillary barrier are tested by diffusion capacity of carbon monoxide (DLCO) studies (also referred to as transfer capacity of carbon monoxide, TLCO). The principle of the test is the increased affinity of hemoglobin to preferentially bind carbon monoxide over oxygen (Weinstock and McCannon, 2017). Complementary to spirometry and lung volume measurements, DLCO provides information about the lung surface area available for gas diffusion. Therefore, it is sensitive to any structural changes affecting the alveoli, such as those accompanying emphysema, pulmonary fibrosis, pulmonary edema, and ARDS. Recommendations for the standardization of the test and its evaluation have been outlined by the ATS/ERS Task Force (Graham et al., 2017). An isolated reduction in DLCO with normal spirometry and in absence of anemia suggests an injury to the alveolar-capillary barrier, as for example seen in the presence of pulmonary emboli or in patients with pulmonary hypertension (Weinstock and McCannon, 2017; Lettieri et al., 2006; Seeger et al., 2013). Reduced DLCO together with airflow obstruction (i.e., reduced FEV1) indicates lung parenchymal damage and is commonly observed in smokers and in COPD patients (Matheson et al., 2007; Harvey et al., 2016), whereas reduced DLCO with airflow restriction is seen in patients with interstitial lung diseases (Dias et al., 2014; Kandhare et al., 2016).</span></span></span></span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><strong><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Lung function tests used to evaluate experimental animal lung function</span></span></strong></span></span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Because spirometry requires active participation and compliance of the subject, it is not commonly used in animal studies. However, specialized equipment such as the flexiVent system (SCIREQ<sup>®</sup>) are available for measuring FEV, FVC and PEF in anesthetized and tracheotomized small laboratory animals. Other techniques such as plethysmography or forced oscillation are increasingly preferred for lung function assessment in small laboratory animals (McGovern et al., 2013; Bates, 2017). </span></span></span></span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">In small laboratory animals, plethysmography can be used to determine respiratory physiology parameters (minute volume, respiratory rate, time of pause and time of break), lung volume and airway resistance of conscious animals. Both whole body and head-out plethysmography can be applied, although there is a preference for the latter in the context of inhalation toxicity studies, because of its higher accuracy and reliability (OECD, 2018a; Hoymann, 2012).</span></span></span></span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Gas diffusion tests are not frequently performed in animals, because reproducible samplings of alveolar gas are difficult and technically challenging (Reinhard et al., 2002; Fallica et al., 2011). Modifications to the procedure employed in humans have, however, open possibilities to obtain a human-equivalent DLCO measure or the diffusion factor for carbon monoxide (DFCO)—a variable closely related to DLCO, which can inform on potential structural changes in the lungs that have an effect on gas exchange indices (Takezawa et al., 1980; Dalbey et al., 1987; Fallica et al., 2011; Limjunyawong et al., 2015).</span></span></span></span></span></p>
<p> </p>
<h4>Regulatory Significance of the AO</h4>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Established regulatory guideline studies for inhalation toxicity focus on evident clinical signs of systemic toxicity, including death, or organ-specific toxicity following acute and (sub)chronic exposure respectively. In toxicological and safety pharmacological studies with airborne test items targeting the airways or the lungs as a whole, lung function is a relevant endpoint for the characterization of potential adverse events (OECD, 2018a; Hoymann, 2012). Hence, the AO “decreased lung function” is relevant for regulatory decision-making in the context of (sub)chronic exposure (OECD, 2018b; OECD, 2018c).</span></span></span></span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Regulatory relevance of the AO “decreased lung function” is evident when looking at the increased risk of diseases in humans following inhalation exposure, and because of its links to other comorbidities and mortality.</span></span></span></span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">To aid diagnosis and monitoring of fibrosis, current recommendations include both the recording of potential environmental and occupational exposures as well as an assessment of lung function (Baumgartner et al., 2000). The latter typically confirms decreased lung function as demonstrated by a loss of lung volume. As the disease progresses, dyspnea and lung function worsen, and the prognosis is directly linked to the decline in FVC (Meltzer and Noble, 2008). </span></span></span></span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="background-color:white"><span style="font-family:"Times New Roman",serif"><span style="font-family:"Segoe UI",sans-serif"><span style="color:black">Chronic exposure to cigarette smoke and other combustion-derived particles results in the development of COPD. COPD is diagnosed on the basis of spirometry results as laid out in the ATS/ERS Task Force documents on the standardization of lung function tests and their interpretation (Pellegrino et al., 2005; Culver et al., 2017, Graham et al., 2019). Rapid rates of decline in the lung function parameter FEV1 are linked to higher risk of exacerbations, increased hospitalization and early death (Wise et al., 2006; Celli, 2010). Reduced FEV1 also poses a risk for serious cardiovascular events and mortality associated with cardiovascular disease (Sin et al., 2005; Lee et al., 2015).</span></span></span></span></span></p>
<p> </p>
<h4>References</h4>
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</div>
<h2>Appendix 2</h2>
<h2>List of Key Event Relationships in the AOP</h2>
<div id="evidence_supporting_links">
<h3>List of Adjacent Key Event Relationships</h3>
<div>
<h4><a href="/relationships/2451">Relationship: 2451: Oxidative Stress leads to CBF, Decreased</a></h4>
<h4><a href="/relationships/2451">Relationship: 2451: Increase, Oxidative Stress leads to CBF, Decreased</a></h4>
<p>Age-dependent decreases in CBF have been demonstrated in several species (e.g. guinea pigs, mice, and human) (Bailey et al., 2014; Grubb et al., 2016; Ho et al., 2001; Joki and Saano, 1997; Paul et al., 2013). </p>
<p>Female hormones, i.e. progesterone and estrogen, have been shown to have direct effect on CBF, i.e., progesterone reduces CBF, 17β-estradiol and progesterone receptor antagonists counteract progesterone effects, but estradiol alone has also been shown to have no effect on CBF. However, the mechanism by which these hormones modulate CBF is yet to be elucidated (Jain et al., 2012; Jia et al., 2011).</p>
<h4>Key Event Relationship Description</h4>
<p>Because the lung interfaces with the external environment, it is frequently exposed to airborne oxidant gases and particulates, and thus prone to oxidant-mediated cellular damage (Ciencewicki et al., 2008). Oxidant stress—through the action of exogenous and endogenous free radicals, such as super oxides, hydroxyl radicals, and hydrogen peroxides—is a common factor in lung inflammation and various respiratory diseases. The presence of redox-sensitive proteins in motile cilia suggests that oxidant stresses may impact ciliary function negatively (Price and Sisson, 2019). Indeed, exposure of human or rodent ciliated airway epithelial cells to hydrogen peroxide, acetaldehyde, ozone or cigarette smoke—all of which are known to cause oxidative stress—decreases CBF in a dose- and time-dependent manner (Bayram et al., 1998; Burman and Martin, 1986; Gosepath et al., 2000; Hastie et al., 1990; Helleday et al., 1995; Kienast et al., 1994; Knorst et al., 1994a; Min et al., 1999; Simet et al., 2010).</p>
<h4>Evidence Supporting this KER</h4>
<p>Experimental studies in vitro have shown that exposure of ciliated respiratory cells directly or indirectly to sources of oxidative stress leads to decreased CBF (Burman and Martin, 1986; Wilson et al., 1987; Feldman et al., 1994; Yoshitsugu et al., 1995; Min et al., 1999), which can be reversed by treatment with antioxidants (Schmid et al., 2015). Cigarette smoke condensate, a known inducer of oxidative stress, also causes a decrease in CBF in vitro (Cohen et al., 2009), while, in human subjects exposed to different oxygen levels, oxygen stress causes a decrease in nasal CBF (Stanek et al., 1998).<br />
</p>
<strong>Biological Plausibility</strong>
<p>One mode of antimicrobial defense in the airway epithelium is generation of free radicals by neutrophils and monocytes/macrophages. Some microbes have also been shown to produce oxidants in significant amounts, e.g. H<sub>2</sub>O<sub>2</sub> production by pneumococcus. Several studies have shown that oxidants, irrespective of the source (microbial or host-derived) inhibit ciliary function. Additionally, there is a large body of experimental evidence demonstrating that exposures to environmental oxidants, including volatile aldehydes, peroxides, sulfur dioxide, nitric dioxide and Diesel exhaust particles have a detrimental impact on ciliary function. Therefore, this KER is highly plausible.</p>
<strong>Empirical Evidence</strong>
<p>Treatment with H<sub>2</sub>O<sub>2</sub> causes a dose-dependent decline in ciliary beat frequency (CBF) in tracheal rings from male Sprague-Dawley rats (Burman and Martin, 1986).</p>
<p>Exposure of nasal ciliated epithelial cells to enzymatically generated oxidants (xanthine/xanthine oxidase) was accompanied by ciliary slowing, which was maximal at the end of the experiment (4 h). After 4 h, the reduction in CBF was 37.4%. Catalase alone, or in combination with superoxide dismutase (SOD), completely protected the epithelial strips from oxidant-mediated ciliary dyskinesia. Exposure of respiratory epithelium to glucose/glucose oxidase resulted in similar effects to those obtained with the xanthine/xanthine oxidase system, with 38% and 57% CBF reduction after 4 h in systems containing 25 and 100 mU of glucose oxidase, respectively. Treatments with H<sub>2</sub>O<sub>2</sub> and HOCl at concentrations of ≥100 µM caused dose-dependent ciliary dyskinesia after 4 h (Feldman et al., 1994).</p>
<p>Marked ciliary slowing was observed after exposure of human nasal epithelial cells to free radicals within the first 5 min. There was a significant difference in CBF between experimental and control groups. Pretreatment with 300 U/mL SOD or with 5 mM 3-ABA prevented CBF changes. (Min et al., 1999).</p>
<p>Pyocyanin dissolved in PBS produced gradual slowing of CBF in strips of normal human nasal ciliated epithelium without recovery. Ciliary dyskinesia was observed only late in the experiments when ciliostasis and epithelial disruption were also noted. In contrast, 1-hydroxyphenazine produced rapid onset of ciliary slowing, dyskinesia, and immediate ciliostasis but also some recovery of ciliary beating during the experiment (Wilson et al., 1987).</p>
<p>Baseline CBF was analyzed in human sinonasal epithelial cells for 4 min (time 0) followed by administration of either cigarette smoke condensate (CSC; not further specified) or DMSO for 3 min. Forskolin was then administered to the apical surface and stimulated CBF measured. Following addition of forskolin, stimulated CBF was significantly decreased in the CSC-exposed group compared to DMSO controls (Cohen et al., 2009).</p>
<p>Treatment of fully differentiated normal human bronchial epithelial cells with 100 nM roflumilast raised the CBF at 1 h after exposure. Subsequent air or cigarette smoke exposure increased CBF significantly in roflumilast-treated cultures (Schmid et al., 2015).</p>
<p>In superficial mucosae of the inferior nasal turbinates from non-smoking healthy volunteers exposed to three different oxygen concentrations—21%, 60% and 95%—for 2 h, CBF dose-dependently increased. Extending exposure to extreme oxygen concentrations to 4 h, however, decreased CBF, which is indicative of “oxygen stress” (Stanek et al., 1998).</p>
<p>Within 2 min of exposure of human respiratory epithelial cell monolayers to 10 µM H<sub>2</sub>O<sub>2</sub>, a decrease in CBF was observed. All cilia stopped moving within 10 min without obvious surface structural change in the ciliated cells. Catalase significantly reduced the ciliotoxic effect of H<sub>2</sub>O<sub>2 </sub>(Yoshitsugu et al., 1995).</p>
<strong>Uncertainties and Inconsistencies</strong>
<p>Several studies show that oxidants decrease CBF which can be reversed by addition of antioxidants, suggesting a direct effect. However, there is evidence suggesting that oxidant-mediated decreases in CBF cannot be prevented by addition of antioxidants. For example, a polycyanin-induced decrease in CBF in human nasal epithelium could be reversed by treatment with isobutylmethylxanthine and forskolin, both of which increase intracellular cAMP, and also by the cAMP analog dibutyryl cAMP, while antioxidants did not seem to have any effect on CBF (Kanthakumar et al., 1993). Like polycyanin, two other <em>P. aeruginosa</em> toxins, 1-hydroxyphenazine (1-HP) and rhamnolipid reduced CBF which was associated with a decrease in intracellular adenosine nucleotides (Kanthakumar et al., 1996). </p>
<p>Inconsistent with several studies, there are studies that suggest that exposure to cigarette smoke does not inhibit CBF. A study involving 56 human subjects (27 non-smokers and 29 smokers) showed no differences in CBF between the 2 groups. However, a decrease in nasal mucociliary clearance was observed in smokers who exhaled smoke through their noses (Stanley et al., 1986). </p>
<p>While several studies have shown age dependence of CBF, there is evidence that suggests otherwise (Agius et al., 1998). </p>
<h4>Quantitative Understanding of the Linkage</h4>
<p>Several studies in various species, including humans and rodents, provide evidence in support of this KER. The empirical evidence confirms both a dose- and time-dependence between the upstream KE/MIE and the downstream KE. Our quantitative understanding of this KER is therefore strong.</p>
<strong>Response-response relationship</strong>
<p>Treatment of human nasal ciliated epithelial cells with 0.4 mM xanthine + 100 mU/mL xanthine oxidase—producing 159 ± 4.0 µM/h H<sub>2</sub>O<sub>2</sub>—decreased CBF by ca. 1 Hz at 1 h and ca. 2.5 Hz (37.4%) at 4 h. Catalase alone (500 U/mL), or in combination with SOD (300 U/mL ) completely protected the cells from oxidant-mediated ciliary dyskinesia (Feldman et al., 1994).</p>
<p>Treatment of human nasal ciliated epithelial cells with 5 mM glucose + 25 mU/mL glucose oxidase— producing 114 ± 7.7 µM/h H<sub>2</sub>O<sub>2</sub>—decreased CBF by ca. 2 Hz at 1 h and ca. 4 Hz (38%) at 4 h. The decline in CBF was even larger with 57% (approx. 6 Hz) at 4 h when 100 mU/mL glucose oxidase was used (producing 322 ± 11.5 µM/h H<sub>2</sub>O<sub>2</sub>). Catalase alone (500 U/mL) completely protected the cells from oxidant-mediated ciliary dyskinesia (Feldman et al., 1994).</p>
<p>Treatment of human nasal ciliated epithelial cells with H<sub>2</sub>O<sub>2</sub> at concentrations ≥100 µM dose-dependently decreased CBF in human nasal ciliated epithelial cells, with 100 µM causing a 22.4% reduction and the maximal decrease (51.6%) seen with 500 µM H<sub>2</sub>O<sub>2</sub> at 4 h. Adding 100 mU/mL MPO to 150 µM H<sub>2</sub>O<sub>2</sub> enhanced the H<sub>2</sub>O<sub>2</sub>-mediated decrease in CBF (control:11.7 ±0.6 Hz; H<sub>2</sub>O<sub>2</sub>: 8.2 ± 1.1 Hz, 30% decrease; H<sub>2</sub>O<sub>2</sub> + MPO: 5.4±0.2 Hz, 53.8% decrease). (Feldman et al., 1994).</p>
<p>Treatment of human nasal ciliated epithelial cells with HOCl at concentrations ≥100 µM dose-dependently decreased CBF in human nasal ciliated epithelial cells, with 100 µM causing a 26.1% reduction and 500 µM causing the maximal decrease (100%) at 4 h (Feldman et al., 1994).</p>
<p>Treatment of human nasal epithelial cells with 0.4 mM xanthine + 100 mU/mL xanthine oxidase decreased CBF by ca. 50% within 2 min. Addition of 300 U/mL SOD abolished this effect (Min et al., 1999).</p>
<p>Treatment of human nasal epithelial cells with 10 mM H<sub>2</sub>O<sub>2</sub> decreased CBF to 36.5 ±4.4% of baseline within 5 min, with a maximal decrease in CBF of 100% seen after 10 min, whereas 1 mM H<sub>2</sub>O<sub>2</sub> had no effect on CBF. Treatment of human nasal ciliated epithelial cells with 0.8 mM xanthine + 100 mU/mL xanthine oxidase transiently increased CBF by 12.1±1.0% from baseline. When xanthine concentration was increased to 4 and 8 mM, CBF decreased by 26.8±1.7 and 25.6±1.5%, respectively (Yoshitsugu et al., 1995).</p>
<p>Treatment of bovine ciliated bronchial epithelial cells with acetaldehyde, an oxidative stressor, decreased CBF in a dose-dependent manner. Significant slowing of ciliary beating by ca. 50% was observed with concentrations as low as 15-30 µM, and ciliary beating was completely abrogated at concentrations > 250 µM. Ciliary beating also decreased following treatment with 15-30 µM propionaldehyde (40-50% of control), butyraldehyde (35-65% of control), isobutyraldehyde (20-40% of control), and benzaldehyde (80-90% of control) (Sisson et al., 1991).</p>
<p>Exposure of rabbit tracheal explants to formaldehyde dose-dependently decreased CBF. At 66 µg formaldehyde/cm<sup>3</sup>, CBF decreased from 12.6 to 11.8 Hz; at 33 µg formaldehyde/cm<sup>3</sup>, CBF decreased from 13.0 to 10.9 Hz (Hastie et al., 1990).</p>
<p>Exposure of guinea pig trachea to SO<sub>2</sub> at concentrations of 2.5-12.5 ppm for 30 min dose-dependently decreased CBF. Exposure to 2.5 ppm SO<sub>2</sub> caused a small, non-significant decrease in mean CBF, and exposure to 5 ppm SO<sub>2</sub> caused a 45% decrease. The greatest decrease (72 %) in mean CBF was recorded after exposure to 12.5 ppm SO<sub>2</sub> (Knorst et al., 1994a).</p>
<p>Exposure of human nasal epithelial cells (cultured in Ringer’s solution) to SO<sub>2</sub> at concentrations of 2.5-12.5 ppm for 30 min dose-dependently decreased CBF. Exposure to 2.5 ppm yielded a 42.8% decrease, whereas exposure to 12.5 ppm yielded a 96.5% decrease in CBF (Kienast et al., 1994).</p>
<p>A 20-min exposure to NO<sub>2</sub>, a known air pollutant, at concentrations of 1.5 or 3.5 ppm did not affect CBF in healthy human subjects at 45 min post-exposure (Helleday et al., 1995).</p>
<p>Exposure of human bronchial epithelial cells from healthy volunteers to 10, 50, and 100 µg/mL Diesel exhaust particles (DEP) significantly decreased CBF by 15.9%, 31.0%, and 55.5%, respectively, from baseline after 24 h (Bayram et al., 1998).</p>
<p>A 4-week exposure of human nasal epithelial cells to 100 µg/m<sup>3</sup> ozone had no effect on CBF, whereas 5- and 10-times that concentration significantly decreased CBF (-11.1% at 500 µg/m<sup>3</sup>; -33.3% at 1000 µg/m<sup>3</sup>) (Gosepath et al., 2000).</p>
<p>Baseline CBF in tracheal rings from C57Bl/6 mice exposed to cigarette smoke (whole body exposure to mainstream and sidestream cigarette smoke via inhalation from 1R1 reference cigarettes, at 150 mg/m<sup>3</sup> total particulate matter for 2 h/day, 5 days/week, for up to 1 year) for 1.5 to 3 months was slightly, but not significantly, increased (∼1 Hz). After 6 months of smoke exposure, however, baseline CBF significantly decreased (∼2–3 Hz) (Simet et al., 2010).<br />
</p>
<strong>Time-scale</strong>
<p>Treatment of human nasal ciliated epithelial cells with 0.4 mM xanthine + 100 mU/mL xanthine oxidase decreased CBF over time, with a noticeable decrease by ca. 1 Hz at 1 h and a maximal decrease of 37.4% reached at 4 h (Feldman et al., 1994).</p>
<p>Treatment of human nasal ciliated epithelial cells with 5 mM glucose + 25 mU/mL glucose oxidase decreased CBF by ca. 2 Hz at 1 h and a maximal decrease of ca. 4 Hz (38%) at 2 h, that did not change until the end of the experiment at 4 h. When 100 mU/mL glucose oxidase was used, CBF decreased by ca. 2 Hz at 1 h, 4 Hz at 2 h, 5.5 Hz at 3 h, reaching a maximum of 57% (approx. 6 Hz) at 4 h (Feldman et al., 1994).</p>
<p>Treatment of human nasal epithelial cells with 0.4 mM xanthine + 100 mU/mL xanthine oxidase decreased CBF maximally by ca. 50% within 2 min, after which it began to increase again, reaching approx. 80% of the baseline value after 30 min (Min et al., 1999).</p>
<p>Treatment of human nasal ciliated epithelial cells with 0.8 mM xanthine + 100 mU/mL xanthine oxidase transiently increased CBF by 12.1±1.0% from baseline within 15 s, after which it rapidly returned to baseline levels (within 30 min). When xanthine concentrations were increased to 4 and 8 mM, CBF decreased by 26.8±1.7 and 25.6±1.5%, respectively (Yoshitsugu et al., 1995).</p>
<p>Treatment of bovine ciliated bronchial epithelial cells with acetaldehyde reduced CBF rapidly, with a significant drop in CBF occurring within 30 s and a maximal decrease by 3 min (Sisson et al., 1991).</p>
<p>Exposure of rabbit tracheal explants to formaldehyde time-dependently decreased CBF: At 66 µg/cm<sup>3</sup>, CBF decreased from 12.6 to 11.8 Hz immediately upon addition of HCHO to complete cessation of beating by 10 min. At 33 µg/cm<sup>3</sup>, CBF decreased from 13.0 to 10.9 Hz by 30 min (Hastie et al., 1990).</p>
<p>At 24 h following a 4-h exposure of healthy human subjects to 3.5 ppm NO<sub>2</sub>, there was a significant elevation in CBF from 12.4±0.9 Hz (at baseline, pre-exposure) to 13.8±0.8 Hz (Helleday et al., 1995).</p>
<p>Exposure of human bronchial epithelial cells to DEP significantly decreased CBF from 2 h onward after incubation with 50 to 100 µg/mL DEP and from 6 hours onward after incubation with 10 µg/mL DEP (Bayram et al., 1998).</p>
<p>A 4-week exposure of human nasal epithelial cells to ozone significantly reduced CBF, with effects becoming noticeable at the higher concentrations (-7.1% at 500 µg/m<sup>3</sup>; <br />
-10.3% at 1000 µg/m<sup>3</sup>) after 2 weeks of exposure and a maximal decrease after 4 weeks (-11.1% at 500 µg/m<sup>3</sup>; -33.3% at 1000 µg/m<sup>3</sup>) (Gosepath et al., 2000).</p>
<td>Mutations in CFTR gene and decreased CFTR levels lead in reduced Cl<sup>–</sup>channel activity and to compromised fluid and electrolyte transport, consequently to reduced airway surface liquid levels, thus hindering coordinated ciliary beating.</td>
<td>Decrease in CFTR protein activity decreases CBF (see also KERs 2440 and 2441).</td>
<td><span style="font-size:11.0pt"><span style="background-color:white"><span style="font-family:"Segoe UI",sans-serif"><span style="color:#212529">Van Goor et al., 2009</span></span></span></span></td>
</tr>
</tbody>
</table>
<strong>Known Feedforward/Feedback loops influencing this KER</strong>
<p>Bailey, K.L., Bonasera, S.J., Wilderdyke, M., Hanisch, B.W., Pavlik, J.A., Devasure, J., et al. (2014). Aging causes a slowing in ciliary beat frequency, mediated by PKCε. Am. J. Physiol. Lung Cell. Mol. Physiol. 306, L584-L589.</p>
<p>Bayram, H., Devalia, J.L., Khair, O.A., Abdelaziz, M.M., Sapsford, R.J., Sagai, M., et al. (1998). Comparison of ciliary activity and inflammatory mediator release from bronchial epithelial cells of nonatopic nonasthmatic subjects and atopic asthmatic patients and the effect of diesel exhaust particles in vitro. J. Allergy Clin. Immunol. 102, 771-782.</p>
<p>Burman, W.J., and Martin, W.J. (1986). Oxidant-Mediated Ciliary Dysfunction: Possible Role in Airway Disease. Chest 89, 410-413. </p>
<p>Ciencewicki, J., Trivedi, S., and Kleeberger, S.R. (2008). Oxidants and the pathogenesis of lung diseases. J. Allergy Clin. Immunol. 122, 456-470. </p>
<p>Cohen, N.A., Zhang, S., Sharp, D.B., Tamashiro, E., Chen, B., Sorscher, E.J., et al. (2009). Cigarette smoke condensate inhibits transepithelial chloride transport and ciliary beat frequency. Laryngoscope 119, 2269-2274.</p>
<p>Feldman, C., Anderson, R., Kanthakumar, K., Vargas, A., Cole, P.J., and Wilson, R. (1994). Oxidant-mediated ciliary dysfunction in human respiratory epithelium. Free Radic. Biol. Med. 17, 1-10. </p>
<p>Gosepath, J., Schaefer, D., Brommer, C., Klimek, L., Amedee, R.G., and Mann, W.J. (2000). Subacute effects of ozone exposure on cultivated human respiratory mucosa. Am. J. Rhinol. 14, 411-418.</p>
<p>Grubb, B.R., Livraghi-Butrico, A., Rogers, T.D., Yin, W., Button, B. and Ostrowski, L.E. (2016). Reduced mucociliary clearance in old mice is associated with a decrease in Muc5b mucin. Am. J. Physiol. Lung Cell. Mol. Physiol. 310, L860-L867.</p>
<p>Hastie, A.T., Patrick, H., and Fish, J.E. (1990). Inhibition and recovery of mammalian respiratory ciliary function after formaldehyde exposure. Toxicol. Appl. Pharmacol. 102, 282-291.</p>
<p>Helleday, R., Huberman, D., Blomberg, A., Stjernberg, N., and Sandstrom, T. (1995). Nitrogen dioxide exposure impairs the frequency of the mucociliary activity in healthy subjects. Eur. Respir. J. 8, 1664-1668.</p>
<p>Ho, J.C., Chan, K.N., Hu, W.H., Lam, W.K., Zheng, L., Tipoe, G.L., et al. (2001). The Effect of Aging on Nasal Mucociliary Clearance, Beat Frequency, and Ultrastructure of Respiratory Cilia. Am. J. Respir. Crit. Care Med. 163, 983-988.</p>
<p>Jain, R., Ray, J.M., Pan, J.-H. and Brody, S.L. (2012). Sex hormone-dependent regulation of cilia beat frequency in airway epithelium. Am. J. Respir. Crit. Care Med. 46, 446-453.</p>
<p>Jia, S., Zhang, X., He, D.Z., Segal, M., Berro, A., Gerson, T., et al., 2011. Expression and Function of a Novel Variant of Estrogen Receptor–α36 in Murine Airways. Am. J. Respir. Cell Mol. Biol. 45, 1084-1089.</p>
<p>Joki, S. and Saano, V. (1997). Influence of ageing on ciliary beat frequency and on ciliary response to leukotriene D4 in guinea‐pig tracheal epithelium. Clin. Exp. Pharmacol. Physiol. 24, 166-169.</p>
<p>Kanthakumar, K., Taylor, G., Cundell, D., Dowling, R., Johnson, M., Cole, P., et al. (1996). The effect of bacterial toxins on levels of intracellular adenosine nucleotides and human ciliary beat frequency. Pulm. Pharmacol. 9, 223-230.</p>
<p>Kanthakumar, K., Taylor, G., Tsang, K., Cundell, D., Rutman, A., Smith, S., et al. (1993). Mechanisms of action of Pseudomonas aeruginosa pyocyanin on human ciliary beat in vitro. Infect. Immun. 61, 2848-2853.</p>
<p>Kienast, K., Riechelmann, H., Knorst, M., Schlegel, J., Müller-Quernheim, J., Schellenbergt, J., et al. (1994). An experimental model for the exposure of human ciliated cells to sulfur dioxide at different concentrations. The clinical investigator 72, 215-219. </p>
<p>Knorst, M.M., Kienast, K., Riechelmann, H., Müller-Quernheim, J., and Ferlinz, R. (1994). Effect of sulfur dioxide on mucociliary activity and ciliary beat frequency in guinea pig trachea. Int. Arch. Occup. Environ. Health 65, 325-328. </p>
<p>Min, Y.-G., Ohyama, M., Lee, K.S., Rhee, C.-S., Oh, S.H., Sung, M.-W., et al. (1999). Effects of free radicals on ciliary movement in the human nasal epithelial cells. Auris Nasus Larynx 26, 159-163.</p>
<p>Paul, P., Johnson, P., Ramaswamy, P., Ramadoss, S., Geetha, B. and Subhashini, A.S. (2013). The Effect of Ageing on Nasal Mucociliary Clearance in Women: A Pilot Study. ISRN Pulmonology 2013, 5.</p>
<p>Price, M.E., and Sisson, J.H. (2019). Redox regulation of motile cilia in airway disease. Redox. Biol. 27, 101146-101146. </p>
<p>Schmid, A., Baumlin, N., Ivonnet, P., Dennis, J.S., Campos, M., Krick, S., et al. (2015). Roflumilast partially reverses smoke-induced mucociliary dysfunction. Respir. Res. 16, 135. </p>
<p>Simet, S.M., Sisson, J.H., Pavlik, J.A., DeVasure, J.M., Boyer, C., Liu, X., et al. (2010). Long-term cigarette smoke exposure in a mouse model of ciliated epithelial cell function. Am. J. Respir. Cell Mol. Biol. 43, 635-640.</p>
<p>Sisson, J.H., Tuma, D.J., and Rennard, S.I. (1991). Acetaldehyde-mediated cilia dysfunction in bovine bronchial epithelial cells. Am. J. Physiol. 260, L29-36. </p>
<p>Stanek, A., Brambrink, A., Latorre, F., Bender, B., and Kleemann, P. (1998). Effects of normobaric oxygen on ciliary beat frequency of human respiratory epithelium. Br. J. Anaesth. 80, 660-664.</p>
<p>Stanley, P., Wilson, R., Greenstone, M., MacWilliam, L., and Cole, P. (1986). Effect of cigarette smoking on nasal mucociliary clearance and ciliary beat frequency. Thorax 41, 519-523.</p>
<p><span style="font-size:11pt"><span style="font-family:Calibri,sans-serif">Van Goor, F., S. Hadida, P. D. Grootenhuis, B. Burton, D. Cao, T. Neuberger, A. Turnbull, A. Singh, J. Joubran, A. Hazlewood, J. Zhou, J. McCartney, V. Arumugam, C. Decker, J. Yang, C. Young, E. R. Olson, J. J. Wine, R. A. Frizzell, M. Ashlock, and P. Negulescu. 2009. "Rescue of CF airway epithelial cell function in vitro by a CFTR potentiator, VX-770." <em>Proc Natl Acad Sci U S A</em> 106 (44): 18825-30. <a href="https://doi.org/10.1073/pnas.0904709106" style="color:blue; text-decoration:underline">https://doi.org/10.1073/pnas.0904709106</a>.</span></span></p>
<p>Wilson, R., Pitt, T., Taylor, G., Watson, D., MacDermot, J., Sykes, D., et al. (1987). Pyocyanin and 1-hydroxyphenazine produced by Pseudomonas aeruginosa inhibit the beating of human respiratory cilia in vitro. J. Clin. Investig. 79, 221-229.</p>
<p>Yoshitsugu, M., Matsunaga, S., Hanamure, Y., Rautiainen, M., Ueno, K., Miyanohara, T., et al. (1995). Effects of oxygen radicals on ciliary motility in cultured human respiratory epithelial cells. Auris Nasus Larynx 22, 178-185.</p>
</div>
<div>
<h4><a href="/relationships/2442">Relationship: 2442: CBF, Decreased leads to MCC, Decreased</a></h4>
<p>Evidences for this KER are derived from studies carried out in dog, gunea pig, rat, frog, sheep, rabbit model systems as well as in human epithelial cell cultures. MCC and CBF were observed to decrease with age in several species (e.g. guinea pigs, mice, and human) (Bailey et al., 2014; Grubb et al., 2016; Ho et al., 2001; Joki and Saano, 1997; Paul et al., 2013; Yager et al., 1978), but evidence by (Agius et al., 1998) suggests that age does not have a major effect on CBF.</p>
<h4>Key Event Relationship Description</h4>
<p>Synchronized ciliary action transports mucus from the distal lung to the mouth, where it is swallowed or expectorated (Munkholm and Mortensen, 2014). In addition to ASL and mucus properties, the speed of ciliary movement, and hence the effectiveness of mucociliary clearance (MCC), is dependent on ciliary amplitude and beat frequency (Rubin, 2002). CBF itself is influenced by several factors, including changes in the physical and chemical properties of the ASL (especially the periciliary fluid), structural modulation in the cilia, concentration of cyclic nucleotides cAMP and cGMP, and intracellular calcium (Ca<sup>2+</sup>). Aside from genetic defects leading to ciliopathies, there is ample evidence for prolonged exposure to noxious agents, such as cigarette smoke, nitrogen oxide and sulfur dioxide, playing a major role in decreasing CBF and hampering efficient MCC.</p>
<h4>Evidence Supporting this KER</h4>
<p>A decrease in CBF resulting from sulfur dioxide exposure reduced mucociliary clearance in dogs (Yeates et al., 1997) and mucociliary activity in guinea pig tracheas (Knorst et al., 1994). In rats, formaldehyde inhalation exposure resulted in lower numbers of ciliated cells, while ciliary activity and mucus flow rates were decreased in a dose and time-dependent manner (Morgan et al., 1986). In humans, CBF positively correlates with nasal mucociliary clearance time (Ho et al., 2001), and bronchiectasis patients have lower nasal CBF and slower mucociliary transport (MCT) (Rutland and Cole, 1981). Administration of nebulized CBF inhibitors and enhancers quantifiably decreased or increased mucociliary clearance, respectively (Boek et al., 1999; Boek et al., 2002). Increased CBF and MCT was also noted in human sinonasal epithelial cell cultures treated with Myrtol®, an essential oil distillate (Lai et al., 2014) and in sheep tracheas and human airway epithelial cultures subjected to temperature changes (Kilgour et al., 2004; Sears et al., 2015). Exposures of frog palate epithelia to formaldehyde and PM10 reduced MCC and mucociliary transport, but only formaldehyde-treated epithelia showed decreases in CBF (Morgan et al., 1984; Macchione et al., 1999; Fló-Neyret et al., 2001).<br />
Ex vivo treatment of sheep trachea with acetylcholine and epinephrine increased CBF, but only acetylcholine increased surface liquid velocity, while both parameters were decreased upon incubation with platelet-activating factor (Seybold et al., 1990). </p>
<strong>Biological Plausibility</strong>
<p>Ciliary function and mucus transport are invariably linked to effective mucus transport along the mucociliary escalator (Bustamante-Marin and Ostrowski, 2017; Mall, 2008). Therefore, this KER is biologically plausible. </p>
<strong>Empirical Evidence</strong>
<p>Studies in animal models of ciliopathies and in individuals with genetic disorders causing cilia defects demonstrate that absent or asynchronous cilia beating results in defective mucus clearance from the lungs, consequently leading to respiratory infections that may be chronic recurrent in nature and ultimately lead to declining lung function (Knowles et al., 2013; Munkholm and Mortensen, 2014; Tilley et al., 2015). Similarly, indirect effects of airway inflammation, caused for example by respiratory infections or allergies, are known to be responsible for changes in cilia beating and hence mucus clearance (Almeida-Reis et al., 2010; Hisamatsu and Nakajima, 2000; Maurer et al., 1982). Finally, airway epithelial injury following exposure to inhalation toxicants can also damage cilia and inhibit cilia function and thereby impair MCC (Iravani and Van As, 1972; Wanner et al., 1996).</p>
<p>A decrease in CBF resulting from sulfur dioxide exposure reduced mucociliary clearance in dogs (Yeates et al., 1997) and mucociliary activity in guinea pig tracheas (Knorst et al., 1994). In rats, formaldehyde inhalation exposure resulted in lower numbers of ciliated cells, while ciliary activity and mucus flow rates were decreased in a dose and time-dependent manner (Morgan et al., 1986). In humans, CBF positively correlates with nasal mucociliary clearance time (Ho et al., 2001), and bronchiectasis patients have lower nasal CBF and slower mucociliary transport (MCT) (Rutland and Cole, 1981). Administration of nebulized CBF inhibitors and enhancers quantifiably decreased or increased mucociliary clearance, respectively (Boek et al., 1999; Boek et al., 2002). Increased CBF and MCT was also noted in human sinonasal epithelial cell cultures treated with Myrtol®, an essential oil distillate (Lai et al., 2014) and in sheep tracheas and human airway epithelial cultures subjected to temperature changes (Kilgour et al., 2004; Sears et al., 2015). Exposures of frog palate epithelia to formaldehyde and PM10 reduced MCC and mucociliary transport, but only formaldehyde-treated epithelia showed decreases in CBF (Morgan et al., 1984; Macchione et al., 1999; Fló-Neyret et al., 2001).</p>
<p>The available evidence does not interrogate the direct relationship between CBF and MCC, but rather evaluates both outcomes in parallel. However, because of the intrinsic linkage of cilia function and MCC, we find the empirical evidence in support of this KER to be moderate.<br />
Ex vivo treatment of sheep trachea with acetylcholine and epinephrine increased CBF, but only acetylcholine increased surface liquid velocity, while both parameters were decreased upon incubation with platelet-activating factor (Seybold et al., 1990). </p>
<strong>Uncertainties and Inconsistencies</strong>
<p>Although ciliary function is considered a primary determinant for effective MCC (Duchateau et al., 1985; Gizurarson, 2015), there is evidence that suggests that MCC can be impeded by other factors that do not affect CBF. For example, nasal CBF in cigarette smokers regularly exhaling through the nose was not significantly different from that of nonsmokers, although they exhibited significantly longer nasomuciliary clearance times compared to nonsmokers. Possible explanations offered for this discrepancy were a potential loss of cilia in the nasal epithelium or increased mucus viscoelasticity (Stanley et al., 1986). Similarly, formaldehyde exposure of rats resulted in decreased cilia numbers and slower mucus flow rates (Morgan KT et al., 1986). On the other hand, there are a number of pharmacological compounds that improve mucociliary clearance through reductions in mucus viscosity, but have no effect on CBF (Jiao and Zhang, 2019), or through increases in CBF, but have no effect on mucociliary clearance (Phillips et al., 1990).</p>
<h4>Quantitative Understanding of the Linkage</h4>
<p>There are several studies providing insights into the negative effect of inhalation exposures on CBF and MCC, that are in line with the current thinking on how these two KEs connect. Additionally, pharmacological studies demonstrated that stimulation of CBF typically results in stimulation of MCC. However, since most studies usually evaluated the KEs in parallel, and even though some results support both dose response and temporal sequence of the KEs, none of the available data affirms causal linkage between CBF and MCC. Our understanding of the evidence is therefore moderate. </p>
<strong>Response-response relationship</strong>
<p>CBF decreased sequentially with increasing SO<sub>2</sub> doses in dogs. CBF decreased from 6.3 ± 0.2 (SE) Hz at baseline to 5.7 ± 0.2 Hz at 5.5 ppm SO<sub>2</sub>. Five ppm SO<sub>2</sub> delivered to both the trachea and tracheobronchial airways for 20 min also caused a marked decrease in mean bronchial mucociliary clearance from 53.7 ± 5.7% to 32.8 ± 7.7% after 90 min (Yeates et al., 1997).</p>
<p>The effects of 30-min exposure to SO<sub>2</sub> on mucociliary activity (MCA) and ciliary beat frequency (CBF) were studied in 31 guinea pig tracheas. A 63% reduction in mean MCA and statistically insignificant changes in CBF were recorded at concentrations of 2.5 ppm SO<sub>2</sub>. Higher SO<sub>2</sub> concentrations caused further impairment of MCA as well as a dose-dependent decrease in CBF: At 5 ppm SO2, CBF decreased by 45%, at 12.5 ppm by 72%. The maximum decrease in MCA (81%) was observed with 7.5 ppm SO<sub>2</sub>; the highest SO<sub>2</sub> concentration did not decrease MCA further. The decrease in MCA was associated with an impairment of CBF only at SO<sub>2</sub> concentrations ≥5.0 ppm (Knorst et al., 1994b).</p>
<p>Administration of a nebulized CBF inhibitor (0.9% NaCl) to 15 healthy volunteers significantly decreased mucociliary transport (MCT) from 7.9±1.5 mm/min (SEM) to 4.5±1.6 mm/min. Salbutamol, a CBF enhancer, significantly increased MCT from 8.0±1.4 to 12.5±1.1 mm/min (Boek et al., 2002; Boek et al., 1999).<br />
<br />
Cooling human airway epithelial cultures grown at the air-liquid interface from 37°C to 25°C over the course of approx. 20 min decreased CBF from 12 to 6 Hz and mucociliary transport (MCT) from 140 to 90 µm/s. Extending the range of temperature tested, CBF was found to increase by 0.49±0.06 Hz for every temperature increase by 1°C, and this was mirrored by an increase in MCT. MCT increased on average between 5 and 11 µm/s for every Hz increase in CBF. This study also showed that CBF decreased with increasing mucin concentration, dropping from 12.4 Hz at 2% bovine submaxillary mucin (BSM) to 10.1 Hz at 8% BSM, concurrent with a ca. 70% reduction in MCT. In addition, treatment with 10 µM basolateral forskolin reproducibly increased CBF by 19.3±2.1% and MCT by 24.4±3.1% over baseline (Sears et al., 2015). In sheep trachea CBF and mucus transport velocity (MTV) were 9.8±2.7 beats/s and 5.7±2.6 mm/min, respectively, at baseline. Temperature reductions from 37°C to 34°C caused a progressive decline in CBF (ca. –20% at 2 h and –90% at 4 h) and MTV (ca. –50% at 2 h and –90% at 4 h), which was further exacerbated by additional temperature decreases (30°C; CBF: ca. –75% at 2 h; MTV: –80% at 2 h) (Kilgour et al., 2004).</p>
<p>Frog palate preparations were incubated with 1.25, 2.5 and 5.0 ppm formaldehyde. At formaldehyde doses of 2.5 and 5 ppm, CBF decreased by ca. 25% compared to baseline within 30 min and by 35-50% within 60 min (Fló-Neyret et al., 2001).<br />
Incubation of frog palates with PM10 from Sao Paolo, Brazil, for up to 120 min did not affect CBF but decreased MCT at concentrations ≥1000 pg/m3 (Macchione et al., 1999)<br />
In freshly excised sheep tracheas, a 60-min incubation with 10 µM platelet-activating factor caused a 6% decrease in CBF and a dose-dependent decrease in surface liquid velocity, reaching a maximum of 63% (Seybold et al., 1990).</p>
<p>In patients with bronchiectasis, nasal CBF was 12.8±1.3 Hz and nasal clearance time was 31.8± 18.4 min. In comparison, in healthy controls, nasal CBF was 14.0±1.3 Hz and nasal clearance time was 17.6± 8.3 min (Rutland and Cole, 1981).</p>
<p>Following basolateral treatment of human sinonasal epithelial cell cultures grown at the air-liquid interface with Myrtol®, a phytopharmaceutical mixture of distillates of rectified essential oils of eucalyptus, sweet orange, myrtle, and lemon as the active ingredients, increased CBF in a dose-dependent manner, with a maximum stimulation with 0.1% of 48±7% after 30 min. The same concentration caused a 46±16% increase in MCT at 40 min (Lai et al., 2014).</p>
<p>In New Zealand white rabbits exposed to 3 ppm NO<sub>2</sub> for 24 h, the average CBF decreased from 764 beats/min to 692 beats/min, and the transport velocity decreased from 5.23 mm/min to 3.03 mm/min (Kakinoki, 1998).</p>
<strong>Time-scale</strong>
<p>A 20-minute exposure of dogs to SO<sub>2</sub> caused a decrease in mean bronchial MCC after 90 min (Yeates et al., 1997).</p>
<p>Frog palate epithelia were incubated with 1.25, 2.5 and 5.0 ppm formaldehyde. At formaldehyde doses of 2.5 and 5 ppm, CBF decreased by ca. 25% compared to baseline within 30 min and by 35-50% within 60 min (Fló-Neyret et al., 2001).</p>
<p>Incubation of freshly excised sheep tracheas with 10 µM platelet-activating factor caused a maximal decrease in CBF of 6% after 60 min and decrease in surface liquid velocity of ca. 30% at 20 min, ca. 50% at 40 min and 63% after 60 min (Seybold et al., 1990).</p>
<p>Following basolateral treatment of human sinonasal epithelial cell cultures grown at the air-liquid interface with different concentrations of Myrtol®, CBF increased rapidly within the first 30 min and then declined thereafter. The maximum response for MCT was seen after 40 min (Lai et al., 2014).</p>
<strong>Known modulating factors</strong>
<p>Physiological factors such as age, sex, posture, sleep, and exercise were shown to affect MCC, although study findings are not always concordant (Houtmeyers et al., 1999). MCC and CBF, for example, were observed to decrease with age in several species in numerous studies (e.g. guinea pigs, mice, and human) (Bailey et al., 2014; Grubb et al., 2016; Ho et al., 2001; Joki and Saano, 1997; Paul et al., 2013; Yager et al., 1978), but evidence by (Agius et al., 1998) suggests that age does not have a major effect on CBF.</p>
<strong>Known Feedforward/Feedback loops influencing this KER</strong>
<p>Almeida-Reis, R., Toledo, A.C., Reis, F.G., Marques, R.H., Prado, C.M., Dolhnikoff, M., et al. (2010). Repeated stress reduces mucociliary clearance in animals with chronic allergic airway inflammation. Respir. Physiol. Neurobiol. 173, 79-85.</p>
<p>Bailey, K.L., Bonasera, S.J., Wilderdyke, M., Hanisch, B.W., Pavlik, J.A., DeVasure, J., et al. (2014). Aging causes a slowing in ciliary beat frequency, mediated by PKCε. Am. J. Physiol. Lung Cell. Mol. Physiol. 306, L584-L589. </p>
<p>Boek, W.M., Graamans, K., Natzijl, H., van Rijk, P.P., and Huizing, E.H. (2002). Nasal Mucociliary Transport: New Evidence for a Key Role of Ciliary Beat Frequency. Laryngoscope 112, 570-573. </p>
<p>Boek, W.M., Keleş, N., Graamans, K., and Huizing, E.H. (1999). Physiologic and hypertonic saline solutions impair ciliary activity in vitro. Laryngoscope 109, 396-399.</p>
<p>Bustamante-Marin, X.M. and Ostrowski, L.E. (2017). Cilia and Mucociliary Clearance. Cold Spring Harb. Persp. Biol. 9, a028241.</p>
<p>Duchateau, G.S., Merkus, F.W., Zuidema, J., and Graamans, K. (1985). Correlation between nasal ciliary beat frequency and mucus transport rate in volunteers. The Laryngoscope 95, 854-859.</p>
<p>Fló-Neyret, C., Lorenzi-Filho, G., Macchione, M., Garcia, M.L.B., and Saldiva, P.H.N. (2001). Effects of formaldehyde on the frog's mucociliary epithelium as a surrogate to evaluate air pollution effects on the respiratory epithelium. Braz. J. Med. Biol. Res. 34, 639-643.</p>
<p>Gizurarson, S. (2015). The effect of cilia and the mucociliary clearance on successful drug delivery. Biol. Pharmaceut. Bull. b14-00398.</p>
<p>Grubb, B.R., Livraghi-Butrico, A., Rogers, T.D., Yin, W., Button, B., and Ostrowski, L.E. (2016). Reduced mucociliary clearance in old mice is associated with a decrease in Muc5b mucin. Am. J. Physiol. Lung Cell. Mol. Physiol. 310, L860-L867.</p>
<p>Hisamatsu, K.-i., and Nakajima, M. (2000). Pranlukast protects leukotriene C4- and D4-induced epithelial cell impairment of the nasal mucosa in vitro. Life Sci. 67, 2767-2773. </p>
<p>Ho, J.C., Chan, K.N., Hu, W.H., Lam, W.K., Zheng, L., Tipoe, G.L., et al. (2001). The Effect of Aging on Nasal Mucociliary Clearance, Beat Frequency, and Ultrastructure of Respiratory Cilia. Am. J. Respir. Crit. Care Med. 163, 983-988. </p>
<p>Houtmeyers, E., Gosselink, R., Gayan-Ramirez, G., and Decramer, M. (1999). Regulation of mucociliary clearance in health and disease. Eur. Respir. J. 13, 1177-1188.</p>
<p>Iravani, J., and Van As, A. (1972). Mucus transport in the tracheobronchial tree of normal and bronchitic rats. J. Pathol. 106, 81-93.</p>
<p>Jiao, J., and Zhang, L. (2019). Influence of Intranasal Drugs on Human Nasal Mucociliary Clearance and Ciliary Beat Frequency. Allergy Asthma Immunol. Res. 11, 306-319. </p>
<p>Joki, S., and Saano, V. (1997). Influence of ageing on ciliary beat frequency and on ciliary response to leukotriene D4 in guinea‐pig tracheal epithelium. Clin. Exp. Pharmacol. Physiol. 24, 166-169.</p>
<p>Kakinoki, Y.O., Ayaki Tanaka, Yushi Washio, Koji Yamada, Yoshiaki Nakai, Kazuhiro Morimoto, Yasushi (1998). Nitrogen dioxide compromises defence functions of the airway epithelium. Acta Otolaryngol. 118, 221-226.</p>
<p>Kilgour, E., Rankin, N., Ryan, S., and Pack, R. (2004). Mucociliary function deteriorates in the clinical range of inspired air temperature and humidity. Intensive Care Med. 30, 1491-1494.</p>
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<p>Knowles, M.R., Daniels, L.A., Davis, S.D., Zariwala, M.A., and Leigh, M.W. (2013). Primary ciliary dyskinesia. Recent advances in diagnostics, genetics, and characterization of clinical disease. Am. J. Respir. Crit. Care Med. 188, 913-922.</p>
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<p>Rutland, J., and Cole, P.J. (1981). Nasal mucociliary clearance and ciliary beat frequency in cystic fibrosis compared with sinusitis and bronchiectasis. Thorax 36, 654-658.</p>
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<p>Wanner, A., Salathe, M., and O'Riordan, T.G. (1996). Mucociliary clearance in the airways. Am. J. Respir. Crit. Care Med. 154, 1868-1902. </p>
<p>Yager, J., Chen, T.-M., and Dulfano, M.J. (1978). Measurement of frequency of ciliary beats of human respiratory epithelium. Chest 73, 627-633.</p>
<p>Yeates, D.B., Katwala, S.P., Daugird, J., Daza, A.V., and Wong, L.B. (1997). Excitatory and inhibitory neural regulation of tracheal ciliary beat frequency (CBF) activated by ammonia vapour and SO2. Ann. Occup. Hyg. 41, 736-744.</p>
</div>
<div>
<h4><a href="/relationships/2443">Relationship: 2443: MCC, Decreased leads to Decreased lung function</a></h4>
<p>The evidences for this KER come from and therefore apply to humans.</p>
<h4>Key Event Relationship Description</h4>
<p>It is very well known that patients suffering from motile ciliopathies, such as primary ciliary dyskinesia, have impaired or absent MCC and lower lung function (reduced FEV1 and FVC) compared to their healthy counterparts (Halbeisen et al., 2018; Marthin et al., 2010; Wallmeier et al., 2020). In cystic fibrosis patients, decreased MCC (due to reduced airway hydration and changes in mucus chemical and viscoelastic properties) causes mucus build-up leading to mucus plugging in the airways and consequently to decreased lung function over time (Kerem et al., 2014; Mossberg et al., 1978; Regnis et al., 1994; Robinson and Bye, 2002; Szczesniak et al., 2017; Wanner et al., 1996). Mucus plugging due to decreased MCC is also considered a major cause of airway obstruction and airflow limitation in COPD patients (Dunican et al., 2021; Okajima et al., 2020) and asthmatics (Kuyper et al., 2003; Maxwell, 1985).</p>
<h4>Evidence Supporting this KER</h4>
<p>Changes in MCC rate are typically paralleled by effects on lung function in several studies where both endpoints have been assessed. In patients with primary ciliary dyskinesia, absence of cilia motion prevents normal MCC and consequently, lung function is reduced (Denizoglu Kulli et al., 2020). In cystic fibrosis patients, the ASL is depleted resulting in impaired MCC (Boucher, 2004). Although the known CFTR genotypes can result in a variety of phenotypes (Derichs, 2013), clinical data indicate that some specific gene defects, such as the p.Phe508del variant, are more frequently associated with decreased lung function indices (e.g. FEV1 % predicted, FVC % predicted, FEF25-75) (Kerem et al., 1990; Johansen et al., 1991; Schaedel et al., 2002). Both cigarette smoking and occupational exposure to biomass fumes led to slower MCC and reduced FEV1 % predicted and FEV1/FVC (Ferreira et al., 2018). Nasomucociliary clearance was slower in COPD smokers compared to former smokers with COPD or to nonsmokers (Ito et al., 2015). Allergen challenge in asthma patients resulted in both reduced MCC and FEV1, which could be reversed by inhalation of hypertonic saline solution (Alexis et al., 2017). In cystic fibrosis patients, treatment with mucolytic agents (Laube et al., 1996; McCoy et al., 1996; Quan et al., 2001; Elkins et al., 2006; Amin et al., 2011; Donaldson et al., 2018) or a CFTR potentiator (Rowe et al., 2014) improved both MCC and lung function (FEV1, FVC and FEF25-75).</p>
<strong>Biological Plausibility</strong>
<p>Lung function is known to decrease with age, and several studies showed that mucus transport rates also decrease in older compared to younger individuals (Goodman et al., 1978; Uzeloto et al., 2021). Impaired MCC is also seen in chronic smokers, even prior to a clinically significant drop in lung function and the detection of small airway disease (Clunes et al., 2012a; Goodman et al., 1978; Lourenço et al., 1971; Uzeloto et al., 2021; Vastag et al., 1986), and in patients with obstructive lung disease and hence, poor lung function (Cruz et al., 1974; Vastag et al., 1986). Adult asthmatics also displayed decreased mucus transport rates/velocities in addition to decreased lung function (Ahmed et al., 1981; Bateman et al., 1983; Foster et al., 1982; Mezey et al., 1978).<br />
In patients with primary ciliary dyskinesia, absence of cilia motion prevents normal MCC and consequently, lung function is reduced (Denizoglu Kulli et al., 2020). In cystic fibrosis patients, the ASL is depleted resulting in impaired MCC (Boucher, 2004a). Although the known CFTR genotypes can result in a variety of phenotypes (Derichs, 2013), clinical data indicate that some specific gene defects, such as the p.Phe508del variant, are more frequently associated with decreased lung function indices (e.g. FEV1 % predicted, FVC % predicted, FEF25-75) (Johansen et al., 1991; Kerem et al., 1990; Schaedel et al., 2002). Unsurprisingly, results from studies with pharmacological agents aimed at restoring CFTR function do not only indicate enhanced MCC but also support improvements in lung function (Bennett et al., 2018; Donaldson et al., 2018; Rowe S. M. et al., 2014a). While the available data link these two KEs, causal evidence is not always available, and some inference is present. Therefore, we judge the biological plausibility of this KER as moderate.</p>
<strong>Empirical Evidence</strong>
<p>Occupational exposure to biomass combustion products resulted in slower MCC and reduced FEV1 % predicted and FEV1/FVC (Ferreira et al., 2018).</p>
<p>Compared to healthy controls, current smokers without airway obstruction and current smokers with COPD exhibited longer saccharin transit times, indicative of impaired MCC, and lower FEV1 % predicted and FEV1/FVC (Uzeloto et al., 2021). Similarly, nasomucociliary clearance was slower in COPD smokers compared to former smokers with COPD or to nonsmokers (Ito et al., 2015). Additionally, mucus plug density—assessed by CT imaging—and mucoid (rather than watery) consistency were inversely related to FEF25–75% and associated with increased RV/TLV (Kesimer et al., 2018).</p>
<p>Asthma patients responded to allergen challenge with a reduction in both MCC and FEV1 (Bennett et al., 2011; Mezey et al., 1978), which could be rescued by inhalation with hypertonic saline solution (Alexis et al., 2017).</p>
<p>Multiple studies interrogating the effect of mucolytic agents such as hypertonic saline solution or recombinant DNase on mucus transport rates or mucus clearance in patients with cystic fibrosis report improvements in both, mucus transport velocities or rates and lung function indices, including FEV1, FVC and FEF25-75 (Amin et al., 2011; Donaldson et al., 2018; Elkins et al., 2006; Laube et al., 1996; McCoy et al., 1996; Quan et al., 2001).</p>
<p>Both MCC and lung function (FEV1, FVC and FEF25-75) improved in cystic fibrosis patients treated with ivacaftor, a CFTR potentiator that increases the channel open probability (Rowe et al., 2014b).</p>
<p>Some studies with mucolytics such as N-acetylcysteine, bromhexine, theophylline/ambroxol or sorebrol demonstrated improved MCC was connected with small improvements in lung function (FEV1, FVC and FEV1/FVC) in patients with chronic bronchitis (Aylward et al., 1980; Castigiioni and Gramolini, 1986; Thomson et al., 1974; Würtemberger et al., 1988).<br />
</p>
<strong>Uncertainties and Inconsistencies</strong>
<p>Genetic defects leading to motile ciliopathies or defects in CFTR function are linked to impaired MCC. However, because of the genetic variety, not every defect, for example in the CFTR gene, also expresses an overt pulmonary phenotype. Other factors, such as low-level chronic inflammation may drive lung pathology by pathways independent of MCC. This might also explain the absence of differences in MCC between healthy smokers and smokers with COPD (Fleming et al., 2019).<br />
Not all studies looking to elucidate the effect of mucolytics on MCC report an improvement of lung function, even though mucus transport rates or tracheobronchial clearance significantly improve. These studies include, for example, some on the effects of hypertonic saline solution, NAC, ambroxol and 2-mercapto-ethane sulphonate (Clarke et al., 1979; Ericsson et al., 1987; Millar et al., 1985; Robinson et al., 1997; Würtemberger et al., 1988). This could be, at least in part, related to the fact that a sudden drop in lung function served as an indicator of patient distress in these studies, and interventions were halted when they occurred to ensure patient safety (Robinson et al., 1996). Another reason could be related to the mechanisms underlying mucus solubilization that may be completely independent of lung function.<br />
MCC is only one means by which mucus can be cleared from the lungs. Another one is cough clearance, and it is highly dependent on the properties of the ASL, in particular the ASL height (Knowles and Boucher, 2002).</p>
<h4>Quantitative Understanding of the Linkage</h4>
<p>The available data, though not causally linking decreases in MCC with decreased lung function, provide a good insight into the importance of the physiological role of MCC in maintaining normal lung function. In at least some studies, impairment of MCC correlated with the drop in FEV1 or FEF25-75. Although clinically valuable benefits can be seen in studies with pharmacological agents such as mucolytics and CFTR modifying drugs, they do not cover a wide range of dose responses nor are they supportive of the KER causality. Therefore, we judge our quantitative understanding as moderate.</p>
<strong>Response-response relationship</strong>
<p>Sixteen Brazilian sugarcane workers aged 25±4 years, with a BMI of 24±3 kg/m2, with exhaled CO of 2.1±1.5 ppm, were examined during the non-harvest season and during the sugarcane burning harvest season. There was a non-significant decrease in saccharin transit time (from 8±1 min to 3±1 min) and a significant decrease FEV1/FVC ratio (from 88.62±5.68 to 84.90±6.47) and %FEV1 (from 92.19±13.24 to 90.44±12.76) during harvest compared with the non-harvest season (Ferreira et al., 2018).</p>
<p>12 (6M/6F) mild allergic, non-smoking asthmatics ages 20–39 with skin sensitivity to house dust mites (HDM) and normal baseline lung function (FEV1 %pred > 80, FEV1/FVC ratio >0.70) inhaled sequential doses of inhaled HDM extract (D. farinae, Greer®, Lenoir, NC) delivered as 5 inhalations from a Devilbiss 646 nebulizer (mass median aerodynamic diameter of 5 um, GSD = 2.0). Five of the 12 patients responded to the allergen challenge with >10% reductions in FEV1 % predicted and reduction in whole lung MCC as evidenced by increased retention rates (mean Central TB Ave120Ret increased from 0.69 to 0.79 for baseline vs. allergen challenge respectively). This reduction in MCC significantly correlated with the post challenge 24 hour FEV1 (Bennett et al., 2011).</p>
<p>Treatment of patients with chronic bronchitis with bromhexine (3 x 16 mg/day) for 14 days resulted in mean changes in FEV1, FVC and FEV1/FVC of + 0.047 L + 0.033 L and +0.6%, respectively, with MCC at 6 h being 6.8% greater after treatment compared to baseline (Thomson et al., 1974).<br />
<br />
Treatment of patients with chronic bronchitis with ambroxol alone (2 x 30 mg/day) or with theophyllin (2 x 400 mg/day) and ambroxol (2 x 30 mg/day) for 7 days MCC/h improved from 18.3 ± 11.1% to 23.3 ± 13% and 29.6 ± 15.7%, respectively whereas lung function remained nearly unchanged with FEV1 predicted of 86.0 ± 9.78 at baseline vs 83.7 ± 9.27 (ambroxol only) and 83.1 ± 11.07 (combination) (Würtemberger et al., 1988).</p>
<p>Treatment of chronic bronchitics with N-acetylcysteine (4 mg/day by metered dose inhaler) for 16 weeks significantly improved sputum viscosity (-0.53 vs -0.67; differences between medians to placebo: 0-14 (-0.77 0.64)) and minimally improved FVC (3.0±0.21 vs 2.9± 0.18 L/s) and PEF (356.7 ±29.64 L/min vs 354.6±25.07) but not FEV1 (1.9±0.18 vs 2.0± 0.13 L/s) (Dueholm et al., 1992).</p>
<p>Treatment of asthmatics with salmeterol improved tracheobronchial clearance rates (AUC: 333±24%h vs 347±30%h in placebo) as well as FEV1 (76 ± 8), FVC (100 ± 5) and PEF % predicted (100 ± 7) compared to placebo (73 ± 8; 95 ± 5; 94 ± 7) (Hasani et al., 2003).</p>
<p>Treatment of mild-to-moderate bronchitics with 42 µg salmeterol slightly enhanced whole lung clearance in 2 hr (not significant; C10–2= 25±11% vs 22±10% in placebo), significantly increased mean peripheral lung clearance (C10–2= 22±9% vs 17±10% in placebo) and significantly increased FEV1 %pred and FEF25–75 at 2 h compared to baseline (93±18%predicted, 2.45 ± 1.08 L/s vs 88±19%predicted, 2.27 ± 0.98 L/s in placebo) (Bennett et al., 2006).</p>
<p>Sputum induction by inhalation of hypertonic saline solution (5%) in asthmatics at 6 hr following challenge with LPS significantly improved FEV % predicted by approx. 20% and was accompanied by a ca. 6-fold increase in whole lung clearance (from 0.1 %/min to 0.6%/min) (Alexis et al., 2017).</p>
<p>133 cystic fibrosis patients (age (mean [SD]) was 21.1 (11.4) years and 46.4% were female. All participants had one copy of the G551D mutation, and 72.2% were compound heterozygous with F508del on the other allele.) completed a 6-month course of ivacaftor. Lung function improved from baseline FEV1% predicted of 82.6 (25.6) to 90.1 (25.0) (mean change, 6.7; 95% CI, 4.9–8.5). In a subgroup of 22 patients, particle clearance from the whole right lung was markedly increased. Average clearance through 60 minutes at 1 month post-treatment was more than twice the baseline value, reflecting substantially improved MCC (Rowe et al., 2014b).<br />
Inhalation of hypertonic saline solution (7%, 4 mL twice daily for 48 weeks) by cystic fibrosis patients improved FVC (by 82 mL; 95 percent confidence interval, 12 to 153) and FEV1 (by 68 mL; 95 percent confidence interval, 3 to 132) values, but not FEF25–75 (Elkins et al., 2006).</p>
<p>In cystic fibrosis patients that inhaled hypertonic saline solution without amiloride twice a day over a period of 14 days one-hour mucus clearance rates improved from baseline (9.3±1.6%) to 14.0±2.0% and increased FEV1 by 6.2%. FVC and FEF25-75 also improved by 1.8% and 13.1%, respectively (Donaldson et al., 2006).</p>
<p>Dornase alfa (recombinant human DNase) is currently used as a mucolytic to treat pulmonary disease in cystic fibrosis. It reduces mucus viscosity in the lungs, promoting improved clearance of secretions (Yang and Montgomery, 2021). In children with cystic fibrosis (mean: 8.4 yrs of age with FEV1 ≥95% predicted) treated with dornase alfa for 96 weeks, FEV1 % predicted improved by 3.2 ± 1.2, FVC % predicted improved by 0.7 ± 1.0, and FEF25-75 % predicted improved by 7.9 ± 2.3 compared to placebo (Quan et al., 2001). In young patients with cystic fibrosis (6-18 yrs of age with FEV1 ≥80% predicted) treated with dornase alfa for 96 weeks, FEF25-75 % predicted improved by 6.1±10.34 compared to placebo (Amin et al., 2011). In 10 adult cystic fibrosis patients receiving 2.5 mg rhDNase twice a day for 6 days, FEV1 and FVC increased by an average of 9.4 ± 3.5% and 12.7 ± 2.6%, respectively, as compared with a decrease of 1.8 ± 1.7% and an increase of 0.4±1.1% in the placebo group, respectively, although there were no significant changes in MCC (Laube et al., 1996). In 320 cystic fibrosis patients (7 to 57 yrs of age), dornase alfa treatment at 2.5 mg/day for 12 weeks (McCoy et al., 1996).</p>
<p>Saccharin transit times (a marker of nasal MCC) were higher in healthy current smokers and COPD smokers than in healthy controls (10.87 [7.29–17] min and 16.47 [8.25–20.15] min, respectively, vs 8.52 [5.54–13.91] min). These groups also differed in their lung function indices: FEV1 % predicted was 101.4 ± 12.37 in healthy controls, 96.41 ± 12.3 in healthy current smokers, and 67.96 ± 24.02 in COPD smokers. FVC % predicted was 103.1 ± 13.45 in healthy controls, 97.51 ± 12.88 in healthy current smokers, and 90.33 ± 29.27 in COPD smokers. FEV1/FVC % predicted was 82.15 [78.5–85] in healthy controls, 82.20 [79.2–84.1] in healthy current smokers, and 61.1 [55.3–67.2] in COPD smokers (Uzeloto et al., 2021).<br />
Saccharin transit time of smokers with COPD (16.5 [11–28] min, median [interquartile range 25–75%]) was slightly longer than that of current smokers (15.9 [10 –27] min), and both were longer compared with exsmokers with COPD (10.2 [6 –12] min) and nonsmokers (8 [6 –16] min). Lung function parameters for the groups were as follows: nonsmokers, FEV1/FVC 0.84 ± 0.09, FEV1 % predicted 103.2 ± 11.5, FVC % predicted 102.2 ± 13.3; current smokers, FEV1/FVC 0.76 ± 0.05, FEV1 % predicted 90.7 ± 7.4, FVC % predicted 96.3 ± 13.9; former smokers with COPD, FEV1/FVC 0.49 ± 0.08, FEV1 % predicted 46.8 ± 12.6, FVC % predicted 76.8 ± 18.5; current smokers with COPD, FEV1/FVC 0.66 ± 0.16, FEV1 % predicted 48.7 ± 16.8, FVC % predicted 71.7 ± 13.0 (Ito et al., 2015).<br />
</p>
<strong>Time-scale</strong>
<p>Six asymptomatic patients with bronchial asthma and a history of allergic pollenosis and episodic bronchospasm consistent with ragweed hypersensitivity wer challenged by inhalation of an aqueous, short ragweed antigen extract (Greer Laboratories, Lenoir, N.C.), diluted with a phosphate-buffered saline solution. Mean tracheal mucus velocity (TMV) decreased to 72% of baseline immediately after challenge when specific airway conductance (SGaw), and FEV1 showed a maximal decrease, with a further decrease to 47% of baseline after 1 h, when SGaw and FEV1 had returned to baseline values (Mezey et al., 1978).</p>
<p>Treatment of chronic bronchitics with N-acetylcysteine (3 x 200 mg/day) for 4 weeks significantly decreased sputum thickness, increased sputum pourability from 650% glycerol time (at baseline) to 320% glycerol time on day 21 and PEFR on days 28 (+5%), 35 (+6%) and 42 (+7%) and FEV1 on days 21 (+2%), 28(+3%), 35 (+4%) and 42 (+5%) compared to baseline (ca. 33% predicted and 28% predicted, respectively) (Aylward et al., 1980).</p>
<p>Treatment of mild-to-moderate bronchitics with 42 µg salmeterol slightly enhanced whole lung clearance in 2 hr (not significant; C10–2= 25±11% vs 22±10% in placebo), significantly increased mean peripheral lung clearance (C10–2= 22±9% vs 17±10% in placebo) and significantly increased FEV1 %pred and FEF25–75 at 2 h compared to baseline (93±18%predicted, 2.45 ± 1.08 L/s vs 88±19%predicted, 2.27 ± 0.98 L/s in placebo) , and significantly increased FEV1 %pred and FEF25–75 at both 1 (92±19%predicted, 2.44 ± 1.14 L/s) and 2 h (93±18%predicted, 2.45 ± 1.08 L/s) compared to baseline (pre-dose; 90 ± 20%predicted, 2.16 ± 0.92 L/s) (Bennett et al., 2006).</p>
<p>In cystic fibrosis patients on a 6-month ivacaftor regimen, FEV1% improvement was detectable as soon as the 1-month follow-up visit (mean change, 6.7; 95% CI, 5.2–8.3) (Rowe et al., 2014b). MCC remained at elevated level at the month 3 visit (Donaldson et al., 2018).</p>
<p>One-hour mucus-clearance rates in cystic fibrosis patients receiving hypertonic saline with placebo were significantly faster than in the group receiving hypertonic saline with amiloride (14.0±2.0 vs. 7.0±1.5 %), and the durability of response following the inhalation of hypertonic saline with placebo was ≥8 hours (Donaldson et al., 2006).<br />
</p>
<strong>Known modulating factors</strong>
<p>Invariably, if mucus viscosity increases (independent of whether that results from increased mucus production (hypersecretion), depletion of the ASL or another cause) and MCC decreases, another mechanism comes into action to clear excess mucus: cough clearance. Cough constitutes a “backup” host defense by which acutely or chronically accumulated mucus is expelled through forceful, high-velocity airflow (Button et al., 2018; King, 2006). Our current understanding of the mechanical principles and biology of cough suggest that failure of cough clearance may also be a contributor to decreased lung function.</p>
<strong>Known Feedforward/Feedback loops influencing this KER</strong>
<p>Unknown</p>
<h4>References</h4>
<p>Ahmed, T., Greenblatt, D.W., Birch, S., Marchette, B., and Wanner, A. (1981). Abnormal mucociliary transport in allergic patients with antigen-induced bronchospasm: role of slow reacting substance of anaphylaxis. Am. Rev. Respir. Dis. 124, 110-114.</p>
<p>Alexis, N.E., Bennett, W., and Peden, D.B. (2017). Safety and benefits of inhaled hypertonic saline following airway challenges with endotoxin and allergen in asthmatics. J. Asthma 54, 957-960.</p>
<p>Amin, R., Subbarao, P., Lou, W., Jabar, A., Balkovec, S., Jensen, R., et al. (2011). The effect of dornase alfa on ventilation inhomogeneity in patients with cystic fibrosis. Eur. Respir. J. 37, 806-812.</p>
<p>Aylward, M., Maddock, J., and Dewland, P. (1980). Clinical evaluation of acetylcysteine in the treatment of patients with chronic obstructive bronchitis: a balanced double-blind trial with placebo control. Eur. J. Respir. Dis. Suppl. 111, 81-89.</p>
<p>Bateman, J., Pavia, D., Sheahan, N., Agnew, J., and Clarke, S. (1983). Impaired tracheobronchial clearance in patients with mild stable asthma. Thorax 38, 463-467.</p>
<p>Bennett, W.D., Zeman, K.L., Laube, B.L., Wu, J., Sharpless, G., Mogayzel, P.J., Jr., et al. (2018). Homogeneity of Aerosol Deposition and Mucociliary Clearance are Improved Following Ivacaftor Treatment in Cystic Fibrosis. J. Aerosol Med. Pulm. Drug Delivery 31, 204-211. </p>
<p>Bennett, W.D., Almond, M.A., Zeman, K.L., Johnson, J.G., and Donohue, J.F. (2006). Effect of salmeterol on mucociliary and cough clearance in chronic bronchitis. Pulmon. Pharmacol. Therap. 19, 96-100.</p>
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