Most urban residents around the world are breathing unhealthy levels of air pollution. Air pollutants impair mucociliary clearance (MCC), a vital defense mechanism against respiratory pathogens, primarily due to generating reactive oxygen species (ROS). However, the mechanisms governing MCC restoration remain largely unexplored. Here, we revealed that oxidative airway injury induces the formation of reactive aldehyde species. We identified aldehyde dehydrogenase ALDH1A1, which is selectively expressed in the airway epithelium, as a critical enzyme for detoxifying reactive aldehydes. The deficiency of ALDH1A1 led to aberrant regeneration of cilia, resulting in impaired MCC function. Consequently, ALDH1A1-deficient mice showed an increased susceptibility to pneumonia when pre-exposed to oxidative stress. We further demonstrated the critical role of ALDH1A1 upon exposure to diesel exhaust particles (DEP), a significant source of particulate matter with a diameter ≦ 2.5 µm (PM2.5) in ambient air. Moreover, pharmacological enhancement of ALDH1A1 activity facilitated MCC restoration. These findings highlight that aldehyde metabolism confers ciliary resilience against oxidative airway injury, offering a potential target to mitigate the negative health consequences of air pollution.
Research Article
Aldehyde metabolism ensures mucociliary clearance in oxidative airway injury
https://doi.org/10.21203/rs.3.rs-5067166/v1
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Mucociliary clearance
Aldehyde dehydrogenase
cilia
airway
The respiratory system is constantly exposed to pathogens and other harmful substances that enter the body through breathing. The airway serves as the first line of defense in the respiratory system, functioning not only as a physical barrier but also actively eliminating inhaled pathogens and particles through MCC1. MCC consists of two essential components: mucus secreted by goblet cells, which traps pathogens and particles, and ciliated cells, which provide the force necessary for mucus movement by the coordinated and rhythmic beating of their cilia2–4. This collaborative functioning of mucus and cilia allows to transport pathogens and particles toward pharynx, where it is subsequently eliminated by swallowing. Thus, MCC is crucial for maintaining the sterility of the respiratory system.
However, due to their direct exposure to the environment, cilia are highly susceptible to damage from environmental pollutants such as ultrafine particles, fine particles, ozone, nitrogen oxides, and transition metals5,6. Many of these pollutants are potent oxidants or able to generate reactive oxygen species (ROS), which significantly contribute to the initial pathophysiological mechanism7,8. For instance, PM2.5, a major hazardous component of air pollutants that exceed World Health Organization (WHO) annual guideline levels in most urban cities around the world (7,061 out of 7,239 cities), induces oxidative stress leading to ciliary loss and subsequent impairment of MCC function9–11. The impaired NCC is strongly associated with an increased risk of respiratory infection, as evidenced by the positive correlation between PM2.5 exposure and respiratory infections12,13. Notably, pneumonia accounts for approximately 7% of deaths attributed to PM2.5 exposure14. Therefore, the ability to restore MCC function is crucial for mitigating the negative consequence of PM2.5 exposure. While the respiratory system exhibits a remarkable regenerative capacity, ciliary regeneration in human typically requires several weeks15. Moreover, prolonged or high concentration of air pollution can cause irreversible damage to the airway epithelium 6. In any case, the mechanisms governing cilia regeneration following airway injury remain poorly understood.
Membrane phospholipids and triglycerides are the primarily targets of oxidative stress, and oxidation of polyunsaturated fatty acids (PUFAs) in membrane lipid bilayers leads to the formation of reactive aldehydes, such as 4-hydroxy-2-nenenal (4-HNE), acrolein, and malondialdehyde16,17. Meanwhile, each ciliated cell is equipped with 200–300 cilia with diameters ranging from 0.2 to 0.3 µm and lengths in 4–7 µm, which are contiguous with the plasma membrane3. Since all cilia face the luminal surface of the respiratory tract, the plasma membrane facing the lumen is remarkably enlarged, estimated to be hundreds of times greater than that of conventional cell types. This led us to hypothesize that ciliated cells are particularly susceptible to forming reactive aldehydes when subjected to oxidative airway injury caused by air pollution. Here, we show that oxidative stress induces the formation of reactive aldehyde species in the airway. We found that aldehyde dehydrogenase ALDH1A1, which is selectively expressed in airway epithelium, is essential for degrading these aldehydes. Moreover, we unveiled that absence of ALDH1A1 leads to aberrant cilia formation upon exposure to oxidative stress, resulting in the impaired MCC and increased susceptibility to Streptococcus pneumoniae infection. We also found ALDH1A1-mediated aldehyde detoxification is crucial for cilia regeneration upon exposure to DEP. Lastly, we revealed that enhancement of endogenous ALDH activity by the administration of a small-molecule ALDH1A1 activator, Alda-1, facilitated MCC restoration upon oxidative airway injury. Altogether, these results demonstrate aldehyde metabolism ensures ciliary resilience and illuminate its therapeutic potential in mitigating the negative health effects associated with air pollution.
Oxidative stress generates reactive aldehydes in the airway
Although PM2.5 consists of a diverse array of chemical components depending on their sources, polycyclic aromatic hydrocarbons (PAHs) are key driver to induce oxidative stress and significantly contribute to the adverse effects18. To investigate the airway injury caused by oxidative stress, we utilized a murine model for the administration of naphthalene, the most abundant PAH in ambient urban air19. Toxicity of naphthalene requires metabolic catalyzation mediated by cytochrome P450 enzyme CYP2F2, which is abundantly expressed in the airway epithelium, thereby causing airway-selective injury through the generation of naphthalene oxide20. Consistent with the previous study21, intraperitoneal injection of naphthalene caused the majority of the airway epithelium to slough off within a day and many areas were regenerated by a week post injection (Supplementary Fig. 1a). We observed that naphthalene acutely increased the levels of ROS and lipid peroxidation in the airway epithelium (Supplementary Fig. 1b and 1c). Additionally, quantitative PCR of lungs showed that naphthalene induced the expression of oxidative stress-responsive genes such as Hmox1 and Txnrd1 (Supplementary Fig. 1d). These results indicate, in line with the previous observation, naphthalene induces oxidative stress and lipid peroxidation in airway epithelium.
Given that reactive aldehyde species are peroxidation end-products of membrane phospholipid PUFAs22, we next examined the formation of aldehydes resulting from naphthalene exposure. Notably, live imaging analysis of tracheas revealed the significant generation of acrolein, the most reactive lipid peroxide-derived aldehyde species, upon naphthalene exposure (Fig. 1a). To specifically interrogate the aldehyde formation in airway ciliated cells, we cultured mouse primary tracheal cells using the air-liquid interface (ALI) method. By employing this technique, we successfully induced ciliated cell differentiation, as demonstrated by the appearance of cells displaying acetylated a-tubulin (TUBA), a structural component of cilia (Fig. 1b). Although exposure to naphthalene slightly induced acrolein in ciliated cells, incorporating a pan-inhibitor for aldehyde dehydrogenase (ALDH), disulfiram, significantly elevated the level of acrolein (Fig. 1b and 1c). This suggests that ALDHs exert a vital function in degrading reactive aldehydes that are formed during oxidative stress. The ALDH superfamily comprises 21 and 19 putatively functional genes in mouse and human genomes, respectively23. The quantitative PCR analysis of mouse tissues demonstrated that, among ALDH family members, lung and trachea displayed the highest expression of Aldh1a1 mRNA (Fig. 1d). Immunofluorescence staining of lung revealed that ALDH1A1 protein was selectively expressed in airway epithelium, but it was absent or low in alveolar epithelium and interstitial region (Fig. 1e). Furthermore, available human RNA-seq and scRNA-seq data demonstrated the highest expression of Aldh1a1 mRNA in lung where it was primarily expressed in airway epithelial cells24 (Fig. 1f and 1g). These results suggest that ALDH1A1 plays a pivotal role in metabolizing reactive aldehydes in the airways of both mice and humans.
ALDH1A1 deficiency causes aberrant cilia formation in response to oxidative injury
In line with the previous observation25, ALDH1A1-deficient mice showed apparently normal development, including their gross lung histology and the appearance of TUBA+ ciliated cells in the airway (Supplementary Fig. 2a, 2b, and 2c). Apart from its role in the detoxification of harmful aldehydes, ALDH1A1 also facilitates the conversion of retinaldehyde into retinoic acid (RA), which functions as a ligand for retinoic acid nuclear receptors to regulate gene expression program26,27. Nonetheless, lung RNA-seq analysis of Aldh1a1+/+ and Aldh1a1−/− mice demonstrated that ALDH1A1 deficiency resulted in marginal changes in the expression of putative RA-responsive genes (Fig. 2a), suggesting that ALDH1A1 has only a minimal impact on RA generation in the airway. Therefore, our focus shifted towards exploring the significance of ALDH1A1 in oxidative airway injury. Aldh1a1+/+ and Aldh1a1−/− mice were intraperitoneally injected with naphthalene at two-week intervals for a total of two times, and deposition of aldehyde in airways was examined two weeks after the second naphthalene injection. Whereas aldehyde signal was not observed in either Aldh1a1+/+ or Aldh1a1−/− mice under steady-state conditions, naphthalene exposure induced the accumulation of 4-HNE, major aldehyde end product of lipid peroxidation22, in CYP2F2+ airway epithelium of Aldh1a1−/− mice but not in that of Aldh1a1+/+ mice (Fig. 2b). This indicates that ALDH1A1 plays a critical role in metabolizing reactive aldehydes in airway epithelium. Simultaneously, we also witnessed naphthalene-exposed Aldh1a1−/− mice displayed irregular arrangement and orientation of TUBA+ cilia (Fig. 2c and Supplementary Fig. 2d). Statistical determination of longitudinal sections demonstrated decreased ciliary area on the luminal surface of large and small airway epithelium in naphthalene-exposed Aldh1a1−/− mice (Fig. 2d). A scanning electron microscope (SEM) analysis of luminal surface of airway also showed a reduction in ciliary area of naphthalene-exposed Aldh1a1−/− mice (Fig. 2e and 2f). Additionally, flow cytometry analysis revealed that ciliated cells of Aldh1a1−/− mice displayed reduced TUBA expression (Supplementary Fig. 2e, 2f, and 2g). In contrast, there was no significant difference between Aldh1a1+/+ and Aldh1a1−/− mice in the cell numbers of airway epithelial cells, ciliated cells, and club cells (Supplementary Fig. 2h). These results suggest that the reduction in the ciliary area of ALDH1A1-deficient mice is likely due to aberrant cilia formation, rather than a reduction in the number of ciliated cells.
To further investigate the aberrant cilia formation in ALDH1A1-deficient ciliated cells, we cultured Aldh1a1+/+ and Aldh1a1−/− tracheal cells using the ALI method and subsequently exposed them to naphthalene. Whereas Aldh1a1+/+ ciliated cells showed no noticeable alteration of cilia morphology regardless of naphthalene exposure, we noticed that TUBA+ cilia of Aldh1a1−/− ciliated cells were not elongated as those of Aldh1a1+/+ ciliated cells after naphthalene exposure (Fig. 2g). Naphthalene-exposed Aldh1a1−/− ciliated cells displayed a curled and twisted appearance of cilia and exhibited lower cilia height (Fig. 2h and 2i). Consequently, cilia of Aldh1a1−/− ciliated cells were not as upright as those of Aldh1a1+/+ cells after naphthalene exposure, whereas the structural arrangements of ODF2+ basal bodies, that anchor the individual cilium, appears to be similar (Fig. 2j). Lastly, the majority of cilia in naphthalene-exposed Aldh1a1−/− ciliated cells displayed irregular wave patterns marked by low or absent beating frequency (Supplementary Video 1). Together, these results indicate that ALDH1A1 deficiency results in the aberrant cilia formation and function in response to oxidative damage.
ALDH1A1 deficiency leads to impaired MCC
To determine if aberrant cilia formation is associated with the dysregulation of MCC, we conducted an ex vivo live imaging that enables visualization of fluorescent beads drifting on the isolated tracheas as a proxy for the functioning of MCC28. Without naphthalene exposure, both control (Aldh1a1+/+ or +/−) and Aldh1a1−/− tracheas generated the uniform movement and consistent directionality of fluorescent beads flow (Fig. 3a, 3b, and Supplementary Video 2). However, the beads movement on naphthalene-exposed Aldh1a1−/− tracheas became erratic and moved in various directions, while that on naphthalene-exposed wildtype (WT) tracheas maintained uniformity and consistency. The MCC of naphthalene-exposed Aldh1a1−/− tracheas displayed aberrant behaviour for at least six weeks after naphthalene exposure (Supplementary Fig. 3a and 3b). Furthermore, intranasally injected fluorescent beads significantly remained in lungs of naphthalene-exposed Aldh1a1−/− mice (Fig. 3c). These results indicate that oxidative airway injury leads to impaired MCC in ALDH1A1-deficient mice.
Given the crucial function of MCC in maintaining sterility of lower respiratory tract, we next determined whether ALDH1A1-deficiency influences susceptibility to respiratory infection. We carried out infection of S. pneumoniae, the most common causative bacteria for community-acquired pneumonia. Without exposure to naphthalene, we found that both control and Aldh1a1−/− mice exhibited similar susceptibility to S. pneumoniae. (Fig. 3d and 3e). In contrast, pre-exposure to naphthalene led to a higher mortality rate for S. pneumoniae infection in Aldh1a1−/− mice compared to control mice (Fig. 3f and 3g). Although numbers of S. pneumoniae in bronchoalveolar lavage fluid (BALF) did not show significant difference between control and Aldh1a1−/− mice (Fig. 3h), histological assessment showed increased tissue damages and local S. pneumoniae burden in lungs of Aldh1a1−/− mice (Fig. 3i and 3j). Taken together, these results indicate that ALDH1A1 functions as a protective mechanism against pneumonia by regulating MCC resilience in response to oxidative airway injury.
Role of ALDH1A1 in PM2.5-induced airway injury
Accumulating evidence indicates oxidative stress significantly contributes to the adverse consequence of PM2.5 exposure7,8. To delineate the role of ALDH1A1 in exposure to PM2.5, we employed intranasal administration of DEP, which account for roughly 20% of PM2.5 present in ambient air29. DEP comprise a center core of elemental carbon, along with PAHs and other adsorbed organic and non-organic compontents29–31. Repetitive intranasal administration of DEP resulted in their deposition in respiratory tracts and caused epithelial injury in the airway (Supplementary Fig. 4a). Consistent with the previous observations32, DEP induced the expression of oxidative stress-responsive genes (Hmox1 and Txnrd1) in lung (Supplementary Fig. 4b). We also observed that DEP elevated the level of ROS, lipid peroxidation, and formation of acrolein in the airway epithelium (Fig. 4a and Supplementary Fig. 4c). These results indicate DEP, similar to naphthalene, induce oxidative stress and formation of reactive aldehydes in airway epithelium.
We next investigated the function of ALDH1A1 in eliminating reactive aldehydes generated by DEP exposure. Immunofluorescence staining showed exposure to DEP induced the accumulation of 4-HNE in bronchial epithelium of Aldh1a1−/− mice, but not in that of Aldh1a1+/+ mice (Fig. 4b). Furthermore, exposure to DEP resulted in the disruption of uniform movement and consistent directionality of fluorescent beads on the tracheas isolated from Aldh1a1−/− mice (Fig. 4c, 4d, and Supplementary Video 3), indicating that ALDH1A1 is crucial for maintaining the proper function of MCC upon DEP exposure. We observed that exposure to DEP, but not naphthalene, led to the significant increase in the numbers of immune cells including neutrophils, CD4 T and CD8 T cells in lungs (Supplementary Fig. 4d), which is consistent with the previous study33. Although ALDH1A1 deficiency did not affect the recruitment of these immune cells (Supplementary Fig. 4e), prior exposure to DEP led to the greater mortality in the following S. pneumoniae infection in Aldh1a1−/− mice compared to control mice (Fig. 4e and 4f). These results suggest that ALDH1A1 deficiency increases pneumonia susceptibility due to the impaired tolerance mechanism instead of altered resistant immunity34.
Enhancing ALDH1A1 activity promotes regeneration of cilia and MCC
People who survived acute respiratory distress syndrome (ARDS), which is the major cause of mortality in patients with pneumonia, tend to have elevated levels of ALDH proteins in their bronchoalveolar lavage fluid (BALF) when compared to non-survivors35. This implicates the potential benefits of enhancing ALDH activity in the airway. Alda-1, a small compound originally identified as an activator of ALDH2, was recently demonstrated the ability to activate ALDH1A136,37. Alda-1 augments ALDH2 activity through binding near its Glu285 and Cys319 residues, which are conserved in both human and mouse ALDH1A138. To investigate the therapeutic potential of enhancement of ALDH1A1 activity, we utilized osmotic pump drug delivery system for the administration of Alda-1 in a sustained fashion (Fig. 5a). The administration of Alda-1 to C57BL/6 mice resulted in the significantly better mucociliary transport activity compared to vehicle control group after naphthalene exposure, while Alda-1 administration per se had no impact (Fig. 5b, 5c, and Supplementary Video 4). Additionally, the levels of TUBA in the lung tissues were higher in mice administrated with Alda-1 compared to vehicle control (Fig. 5d and 5e). These results suggest enhancing ALDH1A1 activity can promote the regeneration of cilia and MCC upon oxidative airway injury.
PM2.5 is one of the largest environmental threats to public health39. It is a leading driver of multiple adverse health effects, including respiratory diseases, lung cancer, and heart diseases, with an estimated annual global mortality of up to 8.9 million deaths 40,41. The airway serves as the interface where inhaled PM2.5 comes into first contact. Therefore, defense strategies against noxious insults at this interface is crucial to prevent adverse effects of PM2.5 exposure. In this study, we demonstrate that ALDH1A1-dependent aldehyde metabolism in the airway plays a crucial role in ciliary resilience upon PM2.5 exposure. Importantly, enhancement of ALDH1A1 activity improved the reversibility of MCC function in response to oxidative airway injury. Targeting this mechanism may offer novel preventive and therapeutic strategies to mitigate the negative health consequences associated with PM2.5 exposure.
In our in vitro observation, ciliated cells displayed greater levels of acrolein than non-ciliated cells, regardless of the exposure to oxidative stress (Fig. 1b). This suggests that ciliated cells are prone to forming reactive aldehyde species as a result of their increased vulnerability to lipid peroxidation, which could be due to their structural nature of extended plasma membrane42. Meanwhile, membrane lipid peroxidation is often associated with ferroptosis, a form of non-apoptotic cell death43. However, aldehyde accumulation in airway epithelium of Aldh1a1−/− mice resulted in abnormal cilia formation despite not affecting the number of ciliated cells (Supplementary Fig. 2e and 2f), which implies that aldehydes primarily injure cilia rather than causing cell death. Although the mechanism by which aldehydes selectively impair cilia remains unclear, it is possible that aldehydes, which are generated in proximity to the plasma membrane of cilia, may disrupt the essential components necessary for the cilium’s architecture.
Although this study illustrated the significance of aldehyde metabolism concerning PM2.5 in the context of pneumonia mortality, it remains to investigate the impact of aldehyde metabolism in other respiratory diseases, including asthma, chronic obstructive pulmonary disease (COPD), and cystic fibrosis, which are also linked to PM2.5 exposure44–46. Additionally, it will be important to determine the role of ALDH1A1 in airway chronic inflammation, as inflammation can trigger oxidative stress through the activation of immune cells. Moreover, people living in modern society directly inhale harmful aldehydes from various sources, such as cigarette smoke/vapor and building materials that cause sick building syndrome47–49. Further studies may also look to explore the significance of aldehyde metabolism against these endogenously and exogenously derived aldehydes in maintaining MCC homeostasis and in disease contexts.
Reagents
Naphthalene, ovalbumin (OVA, low endotoxin), and disulfiram were purchased from FUJIFILM Wako Pure Chemical Corporation (Japan). Cholera toxin B subunit (CTB) was purchased from Bio Academia (Japan). Diesel exhaust particles (DEP, SRM2975, NIST, Gaithersburg, MD, USA) were purchased from Sigma-Aldrich.
Animals
All procedures were approved by the Institutional Animal Care and Use Committee of Osaka University. Mice were bred and housed under specific pathogen-free conditions. All mice were on the C57BL/6J background and littermate controls were used for experiments when feasible. Mice deficient for Aldh1a1 were by CRISPR/Cas-mediated gene targeting in C57BL/6 zygotes by using CRISPR RNA: 5’- agttcttaaccctgcaactg-3’ as targeting guide and STOP sequences were inserted by electroporation. Mice were group-housed under a standard chow at ambient temperature 25 ˚C, 50 % humidity on average and a 12 h light-dark cycle in individually ventilated cage.
Naphthalene exposure
Eight- to nine-week-old mice (C57BL/6, Aldh1a1-/-, Aldh1a1+/+, and Aldh1a1+/- littermates or age-matched pairs) received intraperitoneal injection of naphthalene (200 mg/kg). For trachea tissue live imaging, naphthalene was administered 4 h prior to imaging. For histology, immunofluorescence, electron microscopy, flow cytometry, mucociliary transport assay, RNA-seq, and pneumonia experiments, mice received the second naphthalene injection two weeks after the first injection. Two weeks after the second injection, mice were sacrificed, or used for pneumonia experiments as described below.
DEP exposure
DEP powder was suspended in phosphate-buffered saline (PBS) at 2 mg/mL (w/v), vortexed for 2 min, sonicated for 10 min in a cooled water bath, aliquoted, and stored at -80 ˚C until it is used. Mice (C57BL/6, Aldh1a1-/- vs WT or Aldh1a1+/- littermates or age-matched pairs, aged 10 - 12 weeks) were lightly anaesthetized with isoflurane and exposed to either 100 µg DEP in 50 µL of PBS or 50 µL PBS without DEP. For trachea tissue live imaging, DEP was administered twice at 2 h and 16 h prior to imaging. For the other experiments, DEP was administered six times every second day. Mice were sacrificed 24 h after the last exposure for quantitative PCR, or three days after the last exposure for histology, immunofluorescence, flow cytometry, and mucociliary transport assay. Pneumonia experiments were performed one day after the sixth exposure as described below.
Histology and immunofluorescence staining of lung tissue sections
Mice were sacrificed using a CO2 chamber. Isolated lung tissues were fixed with 4 % paraformaldehyde (PFA) overnight at 4 ˚C, embedded in paraffin. Tissue sections (6 µm thickness) were prepared using a microtome (SLEE medical, Germany) and mounted onto adhesive glass slides (MATSUNAMI, Japan). Rehydration was performed with xylene, followed by a standard ethanol dilution series. For histology, sections were stained with hematoxylin and eosin (H&E) solutions (Fujifilm Wako). For immunostaining, antigen retrieval was performed at 98 ˚C for 45 min using ImmunoSaver (Fujifilm Wako). Sections were permeabilized with 0.1 % Triton X-100 for 10 min and then blocked with Blocking One Histo (nacalai tesque, Japan) at room temperature for 1 h. After blocking, sections were incubated with primary antibodies diluted in Can Get Signal Immunoreaction Enhancer Solution (Toyobo, Japan) overnight at 4 ˚C: ALDH1A1 (D4R9V), 4-HNE (HNEJ-2), CC10 (EPR19846), and S. pneumoniae (Abcam, ab20429). Nuclear staining with 4’,6-diamidino-2-phenylindole (DAPI, 100 nM in PBS, 5 min) was performed after the final antibody application. After DAPI staining, slides were washed four times in PBS, and sealed in mounting medium (Prolong Diamond Antifade Mountant; Thermo Fisher Scientific) for microscopic observation.
ROS, Lipid peroxide, acrolein labelling, and plasma membrane staining of isolated trachea tissues for live imaging
Mice were euthanized in CO2 chamber and tracheas were dissected. For ROS and lipid peroxide labelling, isolated trachea tissues were incubated with LipiRADICAL Green (2.5 µM; Funakoshi, Japan), CellROX Deep Red (10 µM; Thermo Fisher Scientific), and CellTracker Red CMTPX (1:1000 dilution; Thermo Fisher Scientific) in DMEM/F12 without phenol red (Thermo Fisher Scientific) for 30 min at 37 ˚C. For acrolein labelling, trachea tissues were incubated with AcroleinRED (10 µM; Funakoshi), and CellMask Plasma Membrane Deep Red (1:1000 dilution; Thermo Fisher Scientific) for 20 min at 37 ˚C. For time-lapse recording of cilia movement, trachea tissues were incubated with CellMask Plasma Membrane Orange (Thermo Fisher Scientific) at 1:1000 dilution for 30 min at room temperature. After labelling, tissues were rinsed with DMEM/F12 twice, and mounted using medical adhesive (Daiichi Sankyo, Japan) onto glass slides with 0.4 mm – height ridges, covered with DMEM/F12, and sealed with cover glasses for microscopic observation.
Airway epithelial injury model in vitro
C57BL/6 male mice (6-8 weeks) were euthanized in a CO2 chamber and air-liquid interface (ALI) cultures of mouse airway epithelial cells are prepared from trachea and main bronchi without expansion, as described previously50,51. Briefly, after enzymatic digestion and fibroblast-deprivation50,51, collected non-adherent cells were re-suspended in MTEC proliferation medium: DMEM/F12 supplemented with 5 % (v/v) FBS, Insulin-Transferrin-Selenium (Thermo Fisher Scientific), 1.5 mM L-Glutamine, 0.1 µg/mL Cholera Toxin, 0.025 µg/mL murine EGF (PeproTech, Inc), 0.03 mg/mL Bovine Pituitary Extract (Thermo Fisher Scientific), Y-27632 (Cayman Chemical), 0.05 µM retinoic acid (Sigma), and antibiotics (Penicillin-Streptomycin-Amphotericin B mixture, Lonza), seeded onto 6.5 mm Transwell 0.4 µm pore polyester membrane insert (Corning) at 8 x 104 cells / cm2, and incubated at 37˚C with 5 % CO2. At 100 % confluent, differentiation was induced by removing apical media and replacing the basal media with differentiation medium: DMEM/F12 supplemented with 0.1 % (w/v) Bovine Albumin Fraction V (Thermo Fisher Scientific), Insulin-Transferrin-Selenium (Thermo Fisher Scientific), 1.5 mM L-Glutamine, 0.025 µg/mL Cholera Toxin, 0.005 µg/mL murine EGF (PeproTech, Inc), 0.03 mg/mL Bovine Pituitary Extract (Thermo Fisher Scientific), 0.1 µM retinoic acid (Sigma), and antibiotics (Penicillin-Streptomycin-Amphotericin B mixture, Lonza). For naphthalene-induced injury, naphthalene was applied in the basal differentiation medium at 10 µM for 10 days from differentiation day 7. After naphthalene exposure, cells were incubated in normal differentiation medium for four days, in the presence or absence of pan-ALDH inhibitor disulfiram (2 µM), depending on the experimental design. Ethanol and DMSO were used as carrier controls for naphthalene and disulfiram, respectively.
After exposure, cells on the membrane insert were fixed with 4 % (w/v) PFA in PBS for 15 min at room temperature. After fixation, membranes were removed from the inserts and placed on glass slide for immuno-staining. After wash with PBS containing 0.05 % (v/v) Triton X-100 (PBS-T), cells on the membrane were blocked with 5 % (w/v) BSA in PBS-T for 1 h, incubated with a combination of primary antibodies against Acrolein (10A10), ALDH1A1 (D4R9V), ZO-1 (ZO1-1A12), ODF2 (Abcam, ab43840), or TUBA (6-11B-1) in PBS-T containing 2 % (w/v) BSA overnight at 4˚C. After nuclear staining with DAPI in PBS-T for 5 min, membranes were washed with PBS-T, and sealed with a drop of mounting medium (Prolong Diamond Antifade Mountant; Thermo Fisher Scientific) for microscopic observation.
Microscopic imaging and analyses
H&E images and some of immunofluorescence images were captured using BZ-X800 (Keyence corp., Japan). Confocal imaging was performed using Leica confocal microscope (STELLARIS 5 WLL, Leica microsystems) equipped with LAS X software.
For trachea tissue live imaging, z-stack images were captured using Leica confocal microscope and a confocal image with maximal fluorescent intensity was selected for each analysis. CellROX positive and LipiRADICAL Green positive cells per 200 x 200 (40,000) µm2 were counted using ImageJ. For quantitative analyses of acrolein-adducts in ALI culture (in vitro), z-stack images were captured using Leica confocal microscope every 1 µm for a total of 34 sections, and merged grayscale images were analyzed using ImageJ software (National Institutes of Health, Bethesda, MD, USA). For cilia height measurement, ALI culture samples from Aldh1a1+/+ and Aldh1a1-/- with or without naphthalene exposure were immunolabelled for TUBA and ZO-1 and fluorescence images were captured using Leica confocal microscope (STELLARIS 5 WLL) as described above. 3D reconstitutions of Z-stack images were used for cilia height measurement. For each sample, two images (96.88 x 96.88 µm2, each containing 20-30 ciliated cells) were captured, and cilia height was determined as the distance from ZO-1 to the top of TUBA signal in each ciliated cell. Approximately 50 ciliated cells per sample were analysed. Image analysis was performed using ImageJ.
For quantifying the percentage of ciliated epithelial surfaces in lung tissue sections, immunofluorescence images were captured using BZ-X800 from the top to the bottom of 6 µm sections every 0.6 µm, merged, and total epithelial length and ciliated surface length of tissue samples were manually measured using the line tool of ImageJ.
For detailed morphological observation of cilia, Super-Resolution microscopy was performed using Nikon AX R Confocal Microscope System with a Nikon Spatial Array Confocal equipped with 100 x objective lens (PLAN APO λD 100 x / 1.45 Oil) and image acquisition software NIS-Elements (Nikon, Japan). Imaging of fixed samples were performed using Galvano mode.For recording cilia movement, time-lapse imaging was performed using resonant mode at 29.3 fps.
Scanning Electron Microscopy (SEM)
Tissue samples were fixed by reflux with 2 % formaldehyde and 2.5 % glutaraldehyde in 0.1 M phosphate buffer (pH 7.4), sliced into 2 mm pieces and immersed in the same fix buffer. After washing, the specimens ware post-fixed with 1 % osmium tetroxide in 0.1 M phosphate buffer (pH 7.4) containing 1 % potassium ferrocyanide, and conductive-stained with 1 % tannic acid solution and 1 % osmium tetroxide solution. The specimens were dehydrated in graded series of ethanol, substituted with 100 % ethanol, dried by critical point drying method and coated with osmium tetroxide by vacuum deposition method. Electron micrographs were captured with S-4800 field emission scanning electron microscope (Hitachi High-Technologies Corp., Japan).
Mucociliary transport assay
Mucociliary transport was analyzed using fluorescent beads (Fluoresbrite 0.5 µm; Polysciences, Warrington, PA, USA) as described previously28. Briefly, after removing unnecessary surrounding tissues, the isolated trachea was opened from the dorsal side. With luminal surface facing upward, the tissue was placed in a rectangular space surrounded by 0.4 mm2 – height vinyl ridges on a glass slide. The tissue was secured using medical adhesive (Daiichi Sankyo), as described above. Immediately after applying a drop of pre-warmed fluorescent beads (1:500 dilution in DMEM/F12; 37 ˚C) onto the luminal surface, a cover glass was placed on top for microscopic observation. Time-lapse images were recorded at 136 ms interval using Leica confocal microscope STELLARIS 5 WLL with 20 x objective lens (Leica microsystems). Beads were tracked and analyzed using the TrackMate plugin for Fiji52-54. Traveling linearity and directional uniformity was calculated by analysing 10 individual beads per record. Two recordings were performed for each trachea. In each record, 300-600 beads were detected within the area (258.33 µm x 64.20 µm) and 10 beads with recording of more than continuous 6 frames (0.816 s) were randomly selected for calculations. For individual beads, displacement a was defined as the direct distance from initial position (frame t=0, coordinate: xt=0, yt=0) to the endpoint (frame t=6, coordinate: xt=6, yt=6), and total track distance b was defined as the total traveling distance within the same timeframe (from t=0 to t=6). Traveling linearity was calculated by dividing a by b.
Traveling linearity = a / b
For calculating directional uniformity, individual trajectory vector f was determined as displacement from frame 0 to frame 6, and average trajectory vector F was defined as the average of 10 individual trajectory vectors within the same record. Directional uniformity was calculated by dividing the length of F (displacement as a group) by the average size of 10 individual f vectors (individual displacement).
Directional uniformity = F / faverage
In vivo Imaging
Mice were anesthetized using an anaesthetic combination (medetomidine, midazolam, and butorphanol), and fluorescent carboxylate-modified microspheres (1:100 dilution; Invitrogen) were administered intranasally. At 1, 6, and 24h after administration, in vivo imaging was performed using IVIS Lumina III ver4.7. Settings for imaging were as follows: lamp level: low; excitation: 680 nm; emission: 790 nm; epi fluorescence; binning: medium; FOV: A; F-stop = 2; and acquisition time = 1 s. Total flux (photons/s) was measured within equally sized rectangular regions of interest using Living Image software (PerkinElmer).
Flow cytometry
After removal of blood cells by cardiac perfusion with 5 mL of ice-cold phosphate buffered saline (PBS) via the right ventricle and instillation of dissociation buffer (Hanks' Balanced Salt Solution [HBSS] containing 0.2 U/mL Liberase TM [Roche] and 20 µg/mL DNase I [Roche]) via a trachea catheter, lung lobes were isolated. Minced lung tissues were digested in dissociation buffer at 37 ˚C for 30 min. The reaction was stopped by adding excess volume of buffer containing 10 % (v/v) fetal bovine serum (FBS), and the mixtures were filtered through 70 µm to remove undigested parts and debris. After blocking with anti-mouse CD16/CD32 (Fc Block, BD Pharmingen), leukocytes were labelled using combinations of fluorophore-conjugated antibodies: anti CD45 – Brilliant Violet 421 (Bv421), anti CD11c – Alexa Fluor 488, anti SiglecF – Alexa Fluor 647, anti CD4 – PE/Cyanine7 (Cy7), and anti CD8a – PE.
For labelling epithelial populations, digested tissues were cleaned using Debris Removal kit (Myltenyi) and then blocked with anti-mouse CD16/CD32 (Fc Block; BD Pharmingen). Cell surface markers were labelled using anti CD45 – PE/Cy7 (leukocytes), anti CD31 – PE (endothelial cells), and anti CD326 – Bv421 (epithelial cells). After labelling dead cells using Zombie Aqua (BioLegend, CA, USA), cells were fixed and permeabilized using Cytofix/Cytoperm Fixation/Permeabilization kit (BD Biosciences). Intracellular airway epithelial marker proteins were labelled using anti CYP2F2 – Alexa Fluor 488 and anti TUBA – Alexa Fluor 647. After staining, cell suspensions were filtered through 70-µm strainer. Flow cytometry was performed on an LSRFortessa (BD Biosciences). Aldehyde dehydrogenase activity was determined by ALDEFLUOR (StemCell Technologies).
Real-time RT-PCR
Total RNA extraction was performed using RNeasy Mini (Qiagen) for trachea and Trizol Reagent (Thermo Fisher Scientific) for the other tissues. cDNA was prepared using PrimeScript RT Master Mix (Clonetech) containing random hexamer and oligo (dT) primer. Alternatively, SMART MMLV Reverse Transcriptase (Clonetech) with oligo (dT) primer was used. Real-time PCR was performed using KAPA SYBR FAST (KAPA BioSystems, MA, USA) and CFX Connect Real-Time System (Bio-Rad). The primers used in this study are listed in table S2. The primers used for the mouse genes were as follows: Gapdh, 5’-gttgtctcctgcgacttcaac-3’ and 5’-ccagggtttcttactccttgg-3’; Rpl13a, 5’-ggctgaagcctaccagaaagt-3’ and 5’-tcttttctgcctgtttccgta-3’; b-actin, 5’-tgttaccaactgggacgaca-3’ and 5’-ggggtgttgaaggtctcaaa-3’; Aldh1a1, 5’-ggcttaatccaacagattcattcacct-3’ and 5’-acacctggggaacagagca-3’; Aldh1a2, 5’-cacaggagagcaagtgtgtga-3’ and 5’-tagttgcaagagttgccctgt-3’; Aldh1a3, 5’-aaacccacggtcttctcagat-3’ and 5’-ctttgtccaggtttttggtga-3’; Aldh1a7, 5’-agcttaatctggcagaatcagagtct-3’ and 5’-tcagaggaataaccccgaggaat-3’; Aldh2, 5’-tttatgaacagtggccagacc -3’ and 5’-tcgttgatgatcctcccatag-3’; Aldh3a1, 5’-gatcctaactccaaggtgatgc-3’ and 5’-acccgtttgatgagcttattgt-3’; Aldh3a2, 5’-gatcctaactccaaggtgatgc-3’ and 5’-acccgtttgatgagcttattgt-3’; Aldh3b1, 5’-gaagcatttcaagcgactcc-3’ and 5’-caggcttctcacagtcacca-3’; Aldh3b2, 5’-gcaacgatggcttcctctac-3’ and 5’-agcctatggcccagcttatc-3’; Aldh3b3, 5’-agcgctttatgcctattcca-3’ and 5’-acggaggccattaagcttct-3’; Aldh4a1, 5’-tggaagcacacctcctctct-3’ and 5’-aagggcgacaactggtactg-3’; Aldh5a1, 5’-ttactggctcaacagcaacg-3’ and 5’-tgtttgagcaaacgcaagtc-3’; Aldh6a1, 5’-atcctcgtaggggaggctaa-3’ and 5’-ttaattcttcgcccatccag-3’; Aldh7a1, 5’-ggaaggaataggcgaggttc-3’ and 5’-agtgatgattcccaccaagc-3’; Aldh8a1, 5’-gcaaagcacatttggagaaag-3’ and 5’-agcgggactcatccttaatgt-3’; Aldh9a1, 5’-ggccagtttctgtgtcatcat-3’ and 5’-cccttcacagcattctccata-3’; Aldh16a1, 5’-cttctcctttccgcacagtc-3’ and 5’-ccatgagcattgatccacac-3’; Aldh18a1, 5’-atggttaccgctttggactg-3’ and 5’-cttccatgctcggagaagtc-3’; Txnrd1, 5’-cagttcgtcccaacgaaaat-3’ and 5’-gcacattggtctgctcttca-3’; Hmox1, 5’-tgctcgaatgaacactctgg-3’ and 5’-tctctgcaggggcagtatct-3’.
RNA-seq
Lung tissues were isolated from two mice per group (Aldh1a1+/+ and Aldh1a1-/-) and digested as described above for RNA extraction and purification. RNA-seq sample preparation was performed using RNeasy Mini Kit (QIAGEN). Sequencing libraries were constructed through library preparation following the recommended protocol for the TruSeq stranded mRNA Library Prep kit (Illumina). Fragment size of the libraries was confirmed with an LabChip DNA High Sensitivity Reagent Kit (PerkinElmer). Libraries were sequenced on NovaSeq 6000 (Illumina) in the 101-base single-read mode. The accession numbers for the RNA-seq dataset reported in this paper are GSE267105. Among known RA responsive 532 genes55,56, a total of 153 genes with more than 100 reads were selected as shown in table S3 and plotted against all genes (log 2 [fold change] [x-axis] against average FPKM [y-axis]).
Open source data exploration
RNA-seq data for Human ALDH family genes57 were downloaded via ENCODE Expression Atlas58 on July 10, 2023. Human lung ALDH1A1 single cell data are sourced from LungMAP Consortium [U01HL122642] (LungMAP ID: LMEX0000004396) and downloaded from www.lungmap.net (LungMAP Data Coordinating Center [1U01HL122638]), on September 02, 2023.
Bacterial pneumonia
For bacterial pneumonia model, S. pneumoniae strain TIGR4 was used59. Male C57BL/6 mice aged 10 - 13 weeks, pre-exposed to either naphthalene or DEP, were used for bacterial pneumonia experiments. Mice were anaesthetized by intraperitoneal injection of anaesthetic mixtures (medetomidine, midazolam, and butorphanol) before infection, and intranasally instilled with 1-2 x 108 colony-forming units (CFU) of TIGR4 in 20 µL PBS. Bacterial culture, instillation, and CFU determination were performed as described previously59.
Western Blot
For ALDH1A1 detection, lung lysates were prepared by using SDS sample buffer (62.5 mM Tris-HCl (pH6.8), 2 % SDS, 10 % glycerol, 5 % 2-mercaptoethanol, and 0.001 % bromophenol blue), separated by electrophoresis on 4-15 % polyacrylamide gel, and transferred onto a PVDF membrane. The membrane was incubated with antibodies against ALDH1A1 or b-actin in EveryBlot (Bio-rad), followed by incubation with antibodies against rabbit IgG conjugated with horseradish peroxidase (HRP). For TUBA detection, lung lysates were prepared using RIPA buffer, and protein concentrations were determined by bicinchoninic acid method for loading normalization. Lysates were heat-denatured in SDS sample buffer, separated on 10 % polyacrylamide and transferred onto PVDF membrane. The membranes were incubated with anti-TUBA (D20G3, 1:1000) followed by HRP-conjugated anti rabbit IgG, or with HRP-conjugated anti b-actin (13E5, 1:1000) in Can Get Signal Reagent (Toyobo). The peroxidase activity was detected by ImmunoStar Zeta (Fujifilm Wako).
Statistical analysis
All experiments were conducted at least twice, and statistical analyses (ANOVA followed by post-hoc tests or Student’s t test) were performed using GraphPad Prism 9. Each data points and mean values are presented unless otherwise noted. Mouse survival curves were compared with the log-rank test. P values < 0.05 were considered statistically significant. Samples sizes are detailed in figure legends.
Alda-1 in vivo administration
Mice were divided into four groups: vehicle control without injury, Alda-1 without injury, vehicle control with naphthalene exposure, and Alda-1 with naphthalene exposure. Alda-1 (in 50% DMSO: 50% PEG, at 0.8 mg/kg/h) or vehicle control (50% DMSO: 50% PEG) were administered using ALZET osmotic pump (DURECT Corp., Cupertino, CA) as described previously60. Osmotic pumps were implanted three days before naphthalene exposure to minimize the impact of surgical procedure on the outcome of experiments. Four days after intraperitoneal injection of naphthalene (200 mg/kg), mice were sacrificed and trachea and lung tissues were isolated for mucociliary transport assay and western blotting, respectively.
Acknowledgements
We thank all members of the Y.O. laboratory. We thank K. Ohara (Osaka University) for discussions. We thank H. Miyachi (Kyoto University) and S. Kitano (Kyoto University) for the generation of Aldh1a1-deficient mice, Genome Information Research Center (Osaka University) for RNA-seq analysis, Core Instrumentation Facility of Research Institute of Microbial Diseases (Osaka University) for electron microscope analysis, and Nikon Imaging Center (Osaka University) for imaging analysis. We also thank M. Morimoto (RIKEN) and S. Nakayama (RIKEN) for constructive suggestions. This work was supported by Grant Program for Next Generation Principal Investigators at Immunology Frontier Research Center (IFReC) at Osaka University, "The Nippon Foundation - Osaka University Project for Infectious Disease Prevention", and Japan Society for the Promotion of Science and Technology Agency grant number 22K07134.
Author contributions
N.S. and Y.O. conceived the project idea and designed experiments. N.S. carried out most of the experiments, with assistance from H.K. and Y.O. T.Y. contributed to the initial implementation of the study. J.S. contributed to the expertise and funding acquisition. M.Y. and S.K. provided S. pneumoniae. N.S. and Y.O. wrote the manuscript. All authors participated in editing the manuscript.
Competing interests
The authors declare no competing interests.
Data and materials availability
Sequencing data are available from the National Center for Biotechnology Information Gene Expression Omnibus (GSE GSE267105). All data are available in the main text or the supplementary materials.
- Whitsett, J. A. & Alenghat, T. Respiratory epithelial cells orchestrate pulmonary innate immunity. Nat Immunol16, 27-35 (2015). https://doi.org:10.1038/ni.3045
- Bustamante-Marin, X. M. & Ostrowski, L. E. Cilia and Mucociliary Clearance. Cold Spring Harb Perspect Biol9 (2017). https://doi.org:10.1101/cshperspect.a028241
- Tilley, A. E., Walters, M. S., Shaykhiev, R. & Crystal, R. G. Cilia dysfunction in lung disease. Annu Rev Physiol77, 379-406 (2015). https://doi.org:10.1146/annurev-physiol-021014-071931
- Spassky, N. & Meunier, A. The development and functions of multiciliated epithelia. Nat Rev Mol Cell Biol18, 423-436 (2017). https://doi.org:10.1038/nrm.2017.21
- Cao, Y., Chen, M., Dong, D., Xie, S. & Liu, M. Environmental pollutants damage airway epithelial cell cilia: Implications for the prevention of obstructive lung diseases. Thorac Cancer11, 505-510 (2020). https://doi.org:10.1111/1759-7714.13323
- Aghapour, M. et al. Role of air pollutants in airway epithelial barrier dysfunction in asthma and COPD. Eur Respir Rev31 (2022). https://doi.org:10.1183/16000617.0112-2021
- Lodovici, M. & Bigagli, E. Oxidative stress and air pollution exposure. J Toxicol2011, 487074 (2011). https://doi.org:10.1155/2011/487074
- Gangwar, R. S., Bevan, G. H., Palanivel, R., Das, L. & Rajagopalan, S. Oxidative stress pathways of air pollution mediated toxicity: Recent insights. Redox Biol34, 101545 (2020). https://doi.org:10.1016/j.redox.2020.101545
- Shairsingh, K. et al. WHO air quality database: relevance, history and future developments. Bull World Health Organ101, 800-807 (2023). https://doi.org:10.2471/BLT.23.290188
- Air Quality, E. a. H. A., Environment, Climate Change and Health (ECH). Improving the capacity of countries to report on air quality in cities. World Health Organization, 1-46 (2023).
- Institute, H. E. Air quality and health in cities. State of Global Air, 1-18 (2022).
- Pun, V. C., Kazemiparkouhi, F., Manjourides, J. & Suh, H. H. Long-Term PM2.5 Exposure and Respiratory, Cancer, and Cardiovascular Mortality in Older US Adults. Am J Epidemiol186, 961-969 (2017). https://doi.org:10.1093/aje/kwx166
- Yee, J., Cho, Y. A., Yoo, H. J., Yun, H. & Gwak, H. S. Short-term exposure to air pollution and hospital admission for pneumonia: a systematic review and meta-analysis. Environ Health20, 6 (2021). https://doi.org:10.1186/s12940-020-00687-7
- Yang, L., Li, C. & Tang, X. The Impact of PM(2.5) on the Host Defense of Respiratory System. Front Cell Dev Biol8, 91 (2020). https://doi.org:10.3389/fcell.2020.00091
- Heguy, A. et al. Responses of the human airway epithelium transcriptome to in vivo injury. Physiol Genomics29, 139-148 (2007). https://doi.org:10.1152/physiolgenomics.00167.2006
- Ayala, A., Munoz, M. F. & Arguelles, S. Lipid peroxidation: production, metabolism, and signaling mechanisms of malondialdehyde and 4-hydroxy-2-nonenal. Oxid Med Cell Longev2014, 360438 (2014). https://doi.org:10.1155/2014/360438
- Altomare, A. et al. Lipid peroxidation derived reactive carbonyl species in free and conjugated forms as an index of lipid peroxidation: limits and perspectives. Redox Biol42, 101899 (2021). https://doi.org:10.1016/j.redox.2021.101899
- Idowu, O. et al. Beyond the obvious: Environmental health implications of polar polycyclic aromatic hydrocarbons. Environ Int123, 543-557 (2019). https://doi.org:10.1016/j.envint.2018.12.051
- Rappaport, S. M., Waidyanatha, S. & Serdar, B. Naphthalene and its biomarkers as measures of occupational exposure to polycyclic aromatic hydrocarbons. J Environ Monit6, 413-416 (2004). https://doi.org:10.1039/b314088c
- Shultz, M. A., Choudary, P. V. & Buckpitt, A. R. Role of murine cytochrome P-450 2F2 in metabolic activation of naphthalene and metabolism of other xenobiotics. J Pharmacol Exp Ther290, 281-288 (1999).
- Hsu, H. S. et al. Repair of naphthalene-induced acute tracheal injury by basal cells depends on beta-catenin. J Thorac Cardiovasc Surg148, 322-332 (2014). https://doi.org:10.1016/j.jtcvs.2013.10.039
- Pizzimenti, S. et al. Interaction of aldehydes derived from lipid peroxidation and membrane proteins. Front Physiol4, 242 (2013). https://doi.org:10.3389/fphys.2013.00242
- Jackson, B. et al. Update on the aldehyde dehydrogenase gene (ALDH) superfamily. Hum Genomics5, 283-303 (2011). https://doi.org:10.1186/1479-7364-5-4-283
- Guo, M. et al. Guided construction of single cell reference for human and mouse lung. Nat Commun14, 4566 (2023). https://doi.org:10.1038/s41467-023-40173-5
- Fan, X. et al. Targeted disruption of Aldh1a1 (Raldh1) provides evidence for a complex mechanism of retinoic acid synthesis in the developing retina. Mol Cell Biol23, 4637-4648 (2003). https://doi.org:10.1128/MCB.23.13.4637-4648.2003
- Duester, G. Retinoic acid synthesis and signaling during early organogenesis. Cell134, 921-931 (2008). https://doi.org:10.1016/j.cell.2008.09.002
- Yoshihara, T. & Okabe, Y. Aldh1a2 + fibroblastic reticular cells regulate lymphocyte recruitment in omental milky spots. J Exp Med220 (2023). https://doi.org:10.1084/jem.20221813
- Nakayama, S. et al. Planar cell polarity induces local microtubule bundling for coordinated ciliary beating. J Cell Biol220 (2021). https://doi.org:10.1083/jcb.202010034
- Wichmann, H. E. Diesel exhaust particles. Inhal Toxicol19 Suppl 1, 241-244 (2007). https://doi.org:10.1080/08958370701498075
- McClellan, R. O. Health effects of exposure to diesel exhaust particles. Annu Rev Pharmacol Toxicol27, 279-300 (1987). https://doi.org:10.1146/annurev.pa.27.040187.001431
- van den Brule, S. et al. Diesel exhaust particles alter the profile and function of the gut microbiota upon subchronic oral administration in mice. Part Fibre Toxicol18, 7 (2021). https://doi.org:10.1186/s12989-021-00400-7
- Li, N. et al. Nrf2 is a key transcription factor that regulates antioxidant defense in macrophages and epithelial cells: protecting against the proinflammatory and oxidizing effects of diesel exhaust chemicals. J Immunol173, 3467-3481 (2004). https://doi.org:10.4049/jimmunol.173.5.3467
- Shears, R. K. et al. Exposure to diesel exhaust particles increases susceptibility to invasive pneumococcal disease. J Allergy Clin Immunol145, 1272-1284 e1276 (2020). https://doi.org:10.1016/j.jaci.2019.11.039
- Medzhitov, R., Schneider, D. S. & Soares, M. P. Disease tolerance as a defense strategy. Science335, 936-941 (2012). https://doi.org:10.1126/science.1214935
- Bhargava, M. et al. Bronchoalveolar Lavage Fluid Protein Expression in Acute Respiratory Distress Syndrome Provides Insights into Pathways Activated in Subjects with Different Outcomes. Sci Rep7, 7464 (2017). https://doi.org:10.1038/s41598-017-07791-8
- Kotraiah, V., Pallares, D., Toema, D., Kong, D. & Beausoleil, E. Identification of aldehyde dehydrogenase 1A1 modulators using virtual screening. J Enzyme Inhib Med Chem28, 489-494 (2013). https://doi.org:10.3109/14756366.2011.653353
- Belmont-Diaz, J. A., Yoval-Sanchez, B., Calleja-Castaneda, L. F., Pardo Vazquez, J. P. & Rodriguez-Zavala, J. S. Alda-1 modulates the kinetic properties of mitochondrial aldehyde dehydrogenase (ALDH2). FEBS J283, 3637-3650 (2016). https://doi.org:10.1111/febs.13833
- Beretta, M. et al. Characterization of the East Asian variant of aldehyde dehydrogenase-2: bioactivation of nitroglycerin and effects of Alda-1. J Biol Chem285, 943-952 (2010). https://doi.org:10.1074/jbc.M109.014548
- Collaborators, G. B. D. R. F. Global burden of 87 risk factors in 204 countries and territories, 1990-2019: a systematic analysis for the Global Burden of Disease Study 2019. Lancet396, 1223-1249 (2020). https://doi.org:10.1016/S0140-6736(20)30752-2
- Burnett, R. et al. Global estimates of mortality associated with long-term exposure to outdoor fine particulate matter. Proc Natl Acad Sci U S A115, 9592-9597 (2018). https://doi.org:10.1073/pnas.1803222115
- Cohen, A. J. et al. Estimates and 25-year trends of the global burden of disease attributable to ambient air pollution: an analysis of data from the Global Burden of Diseases Study 2015. Lancet389, 1907-1918 (2017). https://doi.org:10.1016/S0140-6736(17)30505-6
- Nechipurenko, I. V. The Enigmatic Role of Lipids in Cilia Signaling. Front Cell Dev Biol8, 777 (2020). https://doi.org:10.3389/fcell.2020.00777
- Stockwell, B. R. Ferroptosis turns 10: Emerging mechanisms, physiological functions, and therapeutic applications. Cell185, 2401-2421 (2022). https://doi.org:10.1016/j.cell.2022.06.003
- Khreis, H. et al. Exposure to traffic-related air pollution and risk of development of childhood asthma: A systematic review and meta-analysis. Environ Int100, 1-31 (2017). https://doi.org:10.1016/j.envint.2016.11.012
- Burnett, R. T. et al. An integrated risk function for estimating the global burden of disease attributable to ambient fine particulate matter exposure. Environ Health Perspect122, 397-403 (2014). https://doi.org:10.1289/ehp.1307049
- Goss, C. H., Newsom, S. A., Schildcrout, J. S., Sheppard, L. & Kaufman, J. D. Effect of ambient air pollution on pulmonary exacerbations and lung function in cystic fibrosis. Am J Respir Crit Care Med169, 816-821 (2004). https://doi.org:10.1164/rccm.200306-779OC
- Margham, J. et al. Chemical Composition of Aerosol from an E-Cigarette: A Quantitative Comparison with Cigarette Smoke. Chem Res Toxicol29, 1662-1678 (2016). https://doi.org:10.1021/acs.chemrestox.6b00188
- Jensen, R. P., Luo, W., Pankow, J. F., Strongin, R. M. & Peyton, D. H. Hidden formaldehyde in e-cigarette aerosols. N Engl J Med372, 392-394 (2015). https://doi.org:10.1056/NEJMc1413069
- Salthammer, T., Mentese, S. & Marutzky, R. Formaldehyde in the indoor environment. Chem Rev110, 2536-2572 (2010). https://doi.org:10.1021/cr800399g
- Eenjes, E. et al. A novel method for expansion and differentiation of mouse tracheal epithelial cells in culture. Sci Rep8, 7349 (2018). https://doi.org:10.1038/s41598-018-25799-6
- Lam, H. C., Choi, A. M. & Ryter, S. W. Isolation of mouse respiratory epithelial cells and exposure to experimental cigarette smoke at air liquid interface. J Vis Exp (2011). https://doi.org:10.3791/2513
- Ershov, D. et al. TrackMate 7: integrating state-of-the-art segmentation algorithms into tracking pipelines. Nat Methods19, 829-832 (2022). https://doi.org:10.1038/s41592-022-01507-1
- Schindelin, J. et al. Fiji: an open-source platform for biological-image analysis. Nat Methods9, 676-682 (2012). https://doi.org:10.1038/nmeth.2019
- Tinevez, J. Y. et al. TrackMate: An open and extensible platform for single-particle tracking. Methods115, 80-90 (2017). https://doi.org:10.1016/j.ymeth.2016.09.016
- Balmer, J. E. & Blomhoff, R. Gene expression regulation by retinoic acid. J Lipid Res43, 1773-1808 (2002). https://doi.org:10.1194/jlr.r100015-jlr200
- Blake, J. A. et al. Vol. 49 D981-D987 (Nucleic Acids Res., 2021).
- Lin, S. et al. Comparison of the transcriptional landscapes between human and mouse tissues. Proc Natl Acad Sci U S A111, 17224-17229 (2014). https://doi.org:10.1073/pnas.1413624111
- Consortium, E. P. An integrated encyclopedia of DNA elements in the human genome. Nature489, 57-74 (2012). https://doi.org:10.1038/nature11247
- Yamaguchi, M. et al. Zinc metalloproteinase ZmpC suppresses experimental pneumococcal meningitis by inhibiting bacterial invasion of central nervous systems. Virulence8, 1516-1524 (2017). https://doi.org:10.1080/21505594.2017.1328333
- Sun, L., Ferreira, J. C. & Mochly-Rosen, D. ALDH2 activator inhibits increased myocardial infarction injury by nitroglycerin tolerance. Sci Transl Med3, 107ra111 (2011). https://doi.org:10.1126/scitranslmed.3002067
The authors declare no competing interests.
- FigureS120240828.tif
Supplementary Fig. 1: Aldehyde generation in the airway WT mice were intraperitoneally injected with oil or 200 mg/kg naphthalene. a lungs were isolated at one, three, seven days post naphthalene injection. Experimental design (left) and haematoxylin and eosin (H&E) staining of large (upper right) and lower (lower right) airways are shown. b Four hours post naphthalene injection, the luminal surface of isolated tracheas was determined for cell shape (CellTracker), oxidative stress (CellROX), and lipid peroxidation (LipiRADICAL) and representative live images are shown. c The proportions of the area that were positive for CellROX (left) and LipiRADICAL (right) in (b) were determined (n=3). d Quantitative PCR for oxidative stress-responsive genes (Hmox1 and Txnrd1) in lung at twenty-four hours post oil or naphthalene injection (n=4-5). Each point represents one mouse and the mean values are shown by red horizontal lines (c and d). *****P<0.0001, ****P<0.001, and *P<0.05. Data are represented by at least three independent experiments with similar results (a and b).
- FigureS220240828.tif
Supplementary Fig. 2: ALDH1A1 deficiency causes aberrant cilia formation a Lung lysates of indicated genotypes were analyzed by Western Blot for ALDH1A1 and GAPDH. b Hematoxylin and eosin (H&E) staining of lungs of indicated genotypes. Gated area in left are shown at a higher magnification in right. c Bronchial epithelium of indicated genotypes were stained for Cilia (TUBA), ALDH1A1, and nuclei (DAPI), and representative immunofluorescence images are shown. d-h Mice with indicated genotypes were intraperitoneally injected with 200 mg/kg naphthalene at two-week intervals for a total of two times. Two weeks after the second injection, lungs were isolated. d Immunofluorescence staining of longitudinal sections of the small airway epithelium was performed for cilia (TUBA), airway epithelial cells (CC10), and nuclei (DAPI) and representative images are shown. e low cytometry analysis of lung cells was conducted. f TUBA expression of ciliated cells (CD45-CD31-CD326+TUBA+CYP2F2+) in (e) was determined. g Mean Flourescence Intensity (MFI) of TUBA in (f) is shown (n=4). h The numbers of total epithelial cells (CD45-CD31-CD326+), ciliated cells, and Club cells (CD45-CD31-CD326+TUBA-CYP2F2+) in (e) are shown (n=7). Each point represents one mouse and the mean values are shown by red horizontal lines (g and h). *P<0.05, and n.s., not significant. Data are represented by at least three independent experiments with similar results (a-f).
- FigureS320240828.tif
Supplementary Fig. 3: Impaired MCC in ALDH1A1-deficient mice a Mice with indicated genotypes (Cont; Aldh1a1+/+ or Aldh1a1+/-, KO; Aldh1a1-/-) mice were intraperitoneally injected with oil or 200 mg/kg naphthalene at two-week intervals for a total of two times. b 14, 28, or 42 days after the second naphthalene injection, mucociliary transport in isolated tracheas were determined using live imaging of fluorescent beads and trajectory uniformity calculated from 10 beads per recorded area (left) and path linearity of individual beads (right) are presented (n=3-8). ****P<0.001, ***P<0.01, **P<0.02, *P<0.05, and n.s., not significant.
- FigureS420240828.tif
Supplementary Fig. 4: Protective role of ALDH1A1 in PM2.5-induced injury a WT mice were intranasally administered with PBS or 100 µg of DEP every other day for a total of six times and lungs were harvested three days after the last injection. Left; Experimental design. Right; Gross lung photos and representative images of Hematoxylin and eosin (H&E) staining for longitudinal sections of the large airway are shown. Arrow, cilia; Arrowhead, deposition of DEP. b Quantitative PCR for oxidative stress-responsive genes (Hmox1 and Txnrd1) in lung at twenty-four hours post DEP injection (n=5-6). c WT mice were intranasally administered with 100 µg of DEP at 2 h and 16 h prior to dissection. Luminal surface of isolated tracheas was determined for cell shape (CellTracker), oxidative stress (CellROX), and lipid peroxidation (LipiRADICAL) and representative live images are shown. d WT mice were intraperitoneally injected with oil or 200 mg/kg naphthalene at two-week intervals for a total of two times. Alternatively, WT mice were intranasally administered with PBS or 100 µg of DEP every other day for a total of six times. Numbers of macrophages (Mac), neutrophils (Neu), B cells (B), CD4 T cells (CD4 T), and CD8 T cells (CD8 T) in lungs at fourteen days after the last injection of oil or naphthalene or at three days after the last injection of PBS or DEP were determined (n=4-7). e Leukocyte cell numbers in lungs were determined at three days after the last DEP injection are shown (n=3-4). Each point represents one mouse and the mean values are shown by red horizontal lines (b, d and e). *****P<0.0001, ****P<0.001, *P<0.05, and n.s., not significant. Data are represented by at least two independent experiments with similar results (a and c).
- SupplementaryVideo1.mp4
Supplementary video 1 Aldh1a1+/+ and Aldh1a1-/- mice were intraperitoneally injected with 200 mg/kg naphthalene at two-week intervals for a total of two times. Tracheas were harvested two weeks after the second injection, and ciliary beating was recorded.
- SupplementaryVideo2.mp4
Supplementary video 2 Aldh1a1+/+, Aldh1a1+/-, and Aldh1a1-/- mice were intraperitoneally injected with or without 200 mg/kg naphthalene at two-week intervals for a total of two times. Tracheas were harvested two weeks after the second injection, and the movement of fluorescent beads on the isolated tracheas was recorded.
- SupplementaryVideo3.mp4
Supplementary video 3 Aldh1a1+/- and Aldh1a1-/- mice were intranasally injected with PBS or 100 mg DEP every other day for a total of six times. Tracheas were harvested three days after the last injection, and the movement of fluorescent beads on the isolated tracheas was recorded.
- SupplementaryVideo4.mp4
Supplementary video 4 WT mice were infused with vehicle or Alda-1 (0.8mg/kg/h) via implanted osmotic pump. Three days post infusion, mice were intraperitoneally injected with or without 200 mg/kg naphthalene. Tracheas were harvested four days post naphthalene injection, and the movement of fluorescent beads on the isolated tracheas was recorded.