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. 2022 Sep 15;190(2):215–226. doi: 10.1093/toxsci/kfac096

Perfluorooctane Sulfonic Acid Disrupts Protective Tight Junction Proteins via Protein Kinase D in Airway Epithelial Cells

Joseph H Lucas 1, Qixin Wang 2, Irfan Rahman 3,✉
PMCID: PMC9960012  PMID: 36106993

Abstract

Perfluorooctane sulfonic acid (PFOS) is a long chain per- and polyfluoroalklyl substance (PFAS) that has been used in aqueous film-forming foams. Emerging epidemiological evidence indicates that PFOS may be associated with chronic lung diseases such as asthma and analysis of human tissues demonstrates that the lungs carry a significant body burden of PFOS. Deficits in barrier function are a major risk factor for asthma. Thus, we hypothesized that PFOS exposure will lead to impaired epithelial barrier function through dysregulated tight junctions. Hence, we assessed the impact of PFOS on epithelial barrier integrity. Bronchial epithelial cells (16HBE) were grown on collagen-coated transwells and treated to 5–25 μM PFOS, and assessed for changes in barrier function and tight junction proteins. Rescue experiments were performed using the protein kinase D (PKD) inhibitor, CID755673. PFOS treatment reduced transepithelial electrical resistance (TEER) and increased 4 kDa FITC-dextran flux. Additionally, PFOS significantly decreased protein levels and the tight junction organization rate of occludin and zonula occludens 1. Increased phosphorylation (Ser744/Ser748) of PKD was observed 3 h following PFOS treatment. Pretreatment with the PKD inhibitor attenuated PFOS-mediated changes in TEER and FITC-dextran flux and restored occludin protein levels. In conclusion, PFOS causes loss of airway barrier integrity and the disruption of tight junctions in bronchial epithelial cells, which was partly attenuated through the inhibition of PKD. These findings demonstrate that PFOS is capable of disrupting airway barrier function, a potentially driving factor underlying associations between PFOS and respiratory diseases such as asthma.

Keywords: PFOS, pulmonary epithelial barrier, tight junctions

Perfluorooctane sulfonic acid (PFOS) is a long-chain per and polyfluoroalklyl substance (PFAS) that has been widely used in stain repellants, aqueous film-forming foams, and food packaging materials for decades (Panieri et al., 2022). PFOS, similar to other PFAS, is mobile, bioaccumulative in humans, and extremely resistant to both environmental degradation and metabolism (Shahsavari et al., 2021). Although PFOS is no longer produced in the United States, exposure continues to be a concern due to its long biological half-life (human serum t1/2 = 4.8 years) (Olsen et al., 2007), continued use in several industry sectors in Asia (Liu et al., 2021), and the emergence of alternative or short-chain PFAS, which may elicit similar toxicity to PFOS (Chu et al., 2020; Corsini et al., 2012; Gaballah et al., 2020). Toxicological studies provide evidence of adverse human health effects such as changes in liver enzymes (Jin et al., 2020), increased serum cholesterol (Li et al., 2020), decreased birth weights (Wikström et al., 2020), and possible carcinogenicity (Consonni et al., 2013). Despite recent advances in research, the impacts of PFOS on respiratory health are poorly understood. Emerging epidemiological evidence suggests that there may be an association between PFAS exposure and allergic asthma. Higher serum PFOS levels were positively correlated with reported asthmatic symptoms. (Averina et al., 2019; Dong et al., 2013; Jackson-Browne et al., 2020). Biodistribution studies indicate that PFOS accumulates in the lungs, particularly during fetal development in both humans and rodents (Borg et al., 2010; Cui et al., 2009; Mamsen et al., 2019). Moreover, PFOS exposure has been associated with increased apoptosis and decreased urinary clara cell protein (CC16), a biomarker of epithelial cell damage and asthma development (Kropski et al., 2009; Zhou et al., 2017a). PFOS exposure impairs lung development (Grasty et al., 2003; Zhang et al., 2021) in rodents and induces the release of IL-1β, an effector cytokine of the innate immune response in macrophages (Wang et al., 2021). Collectively, these studies indicate that PFOS exposure may disrupt homeostatic respiratory function and influence allergic asthma pathogenesis, yet little is known about how PFOS affects the pulmonary epithelial barrier.

Growing evidence supports the key role of epithelial barriers in the pathology of allergic airway diseases because the main role of the airway epithelial barrier is to provide protection against inhaled toxicants and allergens (Ganesan et al., 2013). Decreased abundance of junction proteins such as E-cadherin and zonula occludens 1 (ZO-1) and reduced transepithelial electrical resistance (TEER) in bronchial biopsies were found in asthmatic patients when compared with healthy controls (de Boer et al., 2008; Hackett et al., 2013; Xiao et al., 2011). Moreover, allergens such as house dust mite (HDM) contain proteolytic activity that directly cleaves tight junction proteins (Herbert et al., 1995). Defects in barrier function may (1) increase the penetration of inhaled allergens and xenobiotics, (2) facilitate immunosurveillance by intra-epithelial dendritic cells, or (3) may activate downstream signaling cascades that promote the secretion of Th2 cytokines and epithelial differentiation (Georas and Rezaee, 2014). Pulmonary barrier function is primarily mediated by apical junction complexes (AJCs), which are composed of a mesh of membrane-associated proteins that seal neighboring cells together. These cell contacts are formed by apical tight junctions, adherens junctions, and desmosomes. Tight junctions are the primary regulators of paracellular permeability and are mainly composed of members of the claudin and tight junction-associated MARVEL protein (TAMP) families (occludin, tricelluin, and MARVEL proteins) as well as scaffolding proteins, such as ZO-1. Claudins are key structural transmembrane proteins that form dimers with claudins on neighboring cells. Occludin forms similar interactions, but knockout studies in rodents demonstrated that it is not required for barrier formation and may play a larger role in tight junction stability as opposed to tight junction formation (Schulzke et al., 2005). Recent evidence also suggests that occludin plays a regulatory role within the tight junction, mediating changes in barrier function in response to environmental factors (Richter et al., 2019). Studies also show that ZO-1 aids in tight junction stability by anchoring components of the apical junction complex including occludin to the actin cytoskeleton (Van Itallie et al., 2009).

Previous studies have shown that PFOS has been implicated in the loss of barrier function in both cellular models of the blood-brain (Yu et al., 2020) and the blood-testis barrier (Gao et al., 2017; Qiu et al., 2016). The disruption of tight junctions was accompanied by alterations in the cytoskeletal organization in these studies. Furthermore, sub-acute exposure to PFOS decreased ZO-1 and occludin mRNA abundance in the colon of mice (Wang et al., 2020). Similar reductions in tight junction protein levels were also observed in the brain following sub-acute PFOS exposure (Yu et al., 2020). Taken together, these studies indicate that exposure to PFOS has systemic effects on tissue barrier function. However, no studies have examined the pulmonary epithelial barrier as a target of PFOS toxicity. Thus, we sought to determine the impact of PFOS on the airway barrier integrity. Barrier permeability, tight junction expression, and localization were assessed. Furthermore, the involvement of protein kinase D (PKD), a major regulator of epithelial barrier function was also interrogated.

MATERIALS AND METHODS

 

 

Cell culture and treatment

Human bronchial epithelial 16HBE14o- cells (16HBE; no. SCC150) were obtained from Millipore Sigma (Burlington, Massachusetts). Cells were cultured in high glucose Dulbecco modified medium (DMEM; no. 10566) supplemented with 15 mM HEPES (no. 15630), 1 mM sodium pyruvate (no. 113600), 10% fetal bovine serum (FBS; no. 10082), and 1% penicillin/streptomycin (no. 15140) obtained from Gibco, Inc. (Billings, Montana). 16HBE cells were maintained at 37°C and 5% CO2. For cell treatments, 16HBE cells were seeded either onto polyester transwell inserts with 0.4 μm pore and 1.12 cm2 growth area (no. 3460) or standard cell culture plates (6 or 12 well; no. 3513 and no. 3516) from Corning (Corning, New York). Perfluorooctane sulfonic acid (PFOS, >98% purity, no. 77282) and the PKD inhibitor, CID755673 (CID, no. 476495) were obtained from Sigma-Aldrich (St Louis, Missouri). Cytotoxicity was determined by measuring lactate dehydrogenase release in the conditioned media using a kit (no. 04744934001, Roche). IL-8 release was measured using a kit from Invitrogen (no. CHC1303). The highest PFOS dose (25 µM) used in this study was selected based on the maximum reported serum concentrations in the literature and scaled down (Olsen et al., 1999), because lung PFOS levels are not well characterized. Following initial dose response experiments, 15 µM PFOS was selected for subsequent experiments for 2 reasons: (1) This dose corresponded to the elevated mean serum PFOS concentrations observed in a non-occupational cohort (Zhou et al., 2014) and (2) produced a robust adverse response, which allowed for further interrogation in mechanistic studies.

Barrier function assays

16HBE cells were cultured on polyester transwell inserts with a 0.4-μm pore and 1.12 cm2 growth area (no. 3460, Corning) for 3–5 days until TEER stabilized (at least 500 Ω × cm2). Cells were serum deprived (1% FBS) overnight and then treated with PFOS or CID for the indicated time points. TEER was measured using an EVOM2 volt-ohm-meter (World Precision Instruments, Sarasota, Florida). Cell-free transwells were used for blank subtraction. Paracellular permeability was assessed using a fluorescent tracer, 4 kDa FITC-dextran. FITC-dextran was diluted to a concentration of 1.5 mg/ml in phenol-free media and added to the apical chamber. About 50 μl of media was collected from the basolateral chamber 24 and 72 h following treatment, and fluorescence (485-nm excitation/528-nm emission) was measured using a Cytation 5 multimode reader. Standard curves were generated to calculate FITC-dextran concentrations.

Immunoblotting

For protein analysis, 16HBE cells were plated in 12 or 6 well culture dishes and treated with either PFOS (5–15 μM) and/or CID (10 μM). Cells were lysed with RIPA buffer containing Halt Protease and Phosphatase inhibitor cocktail (Thermo Fischer Scientific, no. 78440, Waltham, Massachusetts). Protein concentrations were measured in whole cell lysates by Pierce BCA Protein Assay Kit (no. 23225, Thermo Scientific). The protein was separated on a 10% or 12.5% SDS-PAGE gel and transferred to a nitrocellulose membrane. Anti-occludin (ab242202 1:1000, Abcam), anti-ZO-1 (ab216880, 1:1000, Abcam), anti-pPKD (2054s 1:1000, Cell Signaling Technologies [CST]), and anti-PKD (90039s 1:1000, CST) primary antibodies were diluted in 5% BSA or non-fat dry milk and incubated with membranes overnight at 4°C. Membranes were incubated with HRP-conjugated secondary anti-rabbit (no. 1706515, 1:5000, BioRad), or anti-mouse (no. 1706516 1:4000, BioRad) for chemiluminescence detection using the Bio-Rad ChemiDoc MP imaging system. Image Lab software (v4.1, BioRad, Hercules, California) was used for densitometric quantification of band intensities. Ponceau S Staining Solution (59803S, CST) or β-actin (ab20272, 1:2000, Abcam) was used as an endogenous control. Unedited blots are located in the Supplementary Files (Figures S1, S3–S4).

mRNA isolation and quantitative real-time PCR

RNA was isolated from cells using a Qiagen RNeasy Mini Kit (no. 74106) and quantified by Nano-drop spectrophotometer (ND-1000, NanoDrop Technologies, Wilmington, Delaware). Reverse transcription was done by RT2 First Strand Kit (no. 330401, Qiagen), and qRT-PCR was conducted with SYBR green master mix (no. 330559, Qiagen) on Bio-Rad CFX96 qPCR instrument. RNA expression was determined by 2-ΔΔCT method for TJP1 (forward: 5′-CGCGTCTCTCCACATACATTC-3′, reverse: 5′-GCTGGCTTATTCTGAGATGGA-3′) and OCLN (forward: 5′-ATGGCAAAGTGAATGACAAGC-3′, reverse: 5’AGGCGAAGTTAATGGAAGCTC-3′). The mRNA expression was normalized using GAPDH (forward: 5' - AAGGTGAAGGTCGGAGTCAAC -3', reverse: 5' - GGGGTCATTGATGGCAACAATA - 3') and ACTB (forward: 5'- ACAGAGCCTCGCCTTTG-3', reverse: 5'-CCTTGCACATGCCGGAG-3') as housekeeping controls.

Immunofluorescence

Cells were grown on glass coverslips sterilized with 70% ethanol and dried overnight under UV radiation. Once the cells reached confluence, they were treated as described above. At specified time points, cells were washed twice with 1× PBS and then fixed with ice-cold methanol for 10 min at 4°C. Cells were then washed twice with 1× TBS for 5 min each and then blocked with 10% normal goat serum for 1 h on a rocker. Cells were then incubated with primary antibodies against occludin (ab242202, Abcam) and ZO-1 (ab216880, Abcam) sequentially overnight. Secondary antibodies (anti-mouse Alex Fluor 488 and anti-rabbit Alexa Fluor 568) were used to visualize proteins by immunofluorescence on a Nikon fluorescence microscope. The DNA stain, 4′,6-diamidino-2-phenylindole (DAPI) was used as a counterstain. The tight junction organization rate (TiJOR) was determined for each image in ImageJ via macro plugin kindly provided by Dr Christine Terryn (Terryn et al., 2013). Briefly, in ImageJ, background subtraction was used (to remove cytoplasmic staining of tight junctions), and then polygons were created on the image with increasing step size and the number of fluorescent peaks along the perimeter was divided by the perimeter of each polygon. Ten polygons were used and an average was calculated for each image. A minimum of 3 images per slide were analyzed to determine the TiJOR for each sample. Representative images are shown with yellow overlay (ZO-1) or green overlay (occludin). Immunofluorescence controls are provided in Supplementary Files (Figure S2).

Statistical analysis

Statistical significance of parameters was determined by repeated measures analysis of variance (ANOVA) for endpoints with repeated collections or 2-way ANOVAs to analyze time by treatment effects when appropriate. For parameters collected at a single time point, 1-way ANOVA or t tests were conducted followed by Tukey’s or Sidak’s post hoc tests. All statistical analysis was conducted in GraphPad Prism (La Jolla, California). Data are represented as means ± standard error of the mean (SEM). A p value <.05 was considered statistically significant.

RESULTS

 

PFOS Treatment Augments Airway Epithelial Barrier Function

To determine the effects of PFOS treatment, 16HBE cells were seeded on collagen-coated transwells and allowed to reach confluency. PFOS treatment at 5, 15, and 25 μM did not significantly increase LDH release at 24 h post-treatment, whereas 25 μM treatment led to significant LDH release (41%) at 72 post-treatment (Figure 1A). A significant decrease in interleukin 8 (IL-8) release was observed between controls (329.203 pg/ml) and 15 µM PFOS (250.947 pg/ml) at 24 h post-treatment (Figure 1B). There was a dose and time-dependent loss of barrier function in PFOS-treated cells. PFOS treatment reduced TEER at doses as low as 5 µM PFOS (84% of t = 0 h) compared with controls (103% of t = 0 h) and as early as 4 h post-treatment with more significant reductions 72 h post-treatment (Figure 1C). A statistically significant increased permeability, measured by FITC-dextran permeability assay, was observed between controls (55.74 μg/ml), 5 μM PFOS (75.94 μg/ml), and 15 μM PFOS (100.42 μg/ml) at 72 h post-treatment, whereas there was no change at 24 h post-treatment (Figure 1D).

Figure 1.

Figure 1.

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Perfluorooctane sulfonic acid (PFOS) treatment augments airway epithelial barrier function. 16HBE cells were treated to 5–25 μM PFOS in transwells. (A) LDH release and (B) IL-8 were measured in apical-derived conditioned media. (C) TEER and (D) FITC-dextran permeability were assessed at the indicated time points. Bars represent mean ± SEM. n = 3–6, *p < .05, **p < .01, ***p < .001 indicate significance compared with control (CTRL).

PFOS Downregulates Tight Junction Proteins

As shown in Figure 2A, 15 μM PFOS significantly increased mRNA expression of TJP1 (1.69-fold expression) and OCLN (2.12-fold expression) compared with control. However, protein abundances of ZO-1 and occludin were reduced with 15 μM PFOS treatment (0.48 and 0.64-fold change, respectively) at 24 h (Figs. 2B and 2C). Similar reductions in protein levels were observed at 15 μM PFOS treatment after 72 h (Figs. 2B and 2D). There was no significant change in protein or mRNA expression at 5 μM PFOS at either time point.

Figure 2.

Figure 2.

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PFOS treatment downregulates tight junction proteins. 16HBE cells were treated to 5-15 μM PFOS for 24–72 h. The cells were collected for mRNA expression or protein analysis. (A) TJP1 and OCLN mRNA expression after 24 h PFOS exposure. (B) Representative blots of zonula occludens 1 (ZO-1) and occludin are shown, and protein levels of ZO-1 and occludin were calculated as fold change at (C) 24 h and (D) 72 h post treatment. Bars represent mean ± SEM, n = 4–10. **p < .01, ***p < .001 indicate significance compared with control (CTRL).

Immunofluorescence of ZO-1 and occludin was also conducted to assess protein organization. There was a significant reduction in ZO-1 TiJOR calculations between the control (0.033 TiJOR) and PFOS (15 μM) treated cells (0.018 TiJOR) (Figs. 3A and 3B). Occludin TiJOR was also significantly reduced (0.035 vs 0.02 TiJOR) compared with control (Figs. 3A and 3C), indicating a loss of tight junction network integrity. The distribution of ZO-1 and occludin along the cell periphery was observed in control cells. However, PFOS (15 μM) treatment disrupted the integrity of the cellular boundaries as shown by ZO-1 immunofluorescence, and completely diminished the expression of occludin in 16HBE cells.

Figure 3.

Figure 3.

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PFOS perturbs the organization of tight junction proteins. 16HBE cells were treated to 15 μM PFOS for 72 h and fixed in ice-cold methanol. (A) Representative images of ZO-1(yellow) and occludin (green) are shown with DAPI merged images displayed below. (B) ZO-1 and (C) occludin TiJORs were calculated. Bars represent mean ± SEM, n = 3. *p < .05, **p < .01, indicate significance compared with control (CTRL). DAPI was used as a nuclear stain. Scale bar = 100 μm.

PKD Inhibition Prevents PFOS-Mediated Disorganization of Tight Junction Proteins

PKD phosphorylation (Ser744/Ser748) was used as a marker of PKD activation. PKD phosphorylation was increased by PFOS at 3 h when normalized to total PKD or total protein levels (Figs. 4A and 4B). There was no significant change in PKD activity at 6 or 24 h using either normalization method.

Figure 4.

Figure 4.

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PFOS increases PKD activity. 16HBE cells were treated to 15 μM PFOS for 3–24 h. Protein was isolated and analyzed by Western blot. (A) Representative blots are shown here and (B) pPKD (Ser744/Ser748) protein levels normalized to total protein or total PKD levels are shown. Bars represent mean ± SEM, n = 6–7, 2 independent experiments. **p < .01, ***p < .001 indicate significance compared with control (CTRL).

To determine the role of PKD, we pretreated cells with the PKD inhibitor CID755673 (CID). Pretreatment with CID did not significantly affect barrier function parameters, except for a small increase in TEER at 72 h. We found CID pretreatment helped to restore the TEER which was decreased after PFOS treatment at either 24 or 72 h post-PFOS treatment (Figure 5A). We also performed FITC-dextran permeability assays to confirm the TEER results. Similarly, CID pretreatment alone showed no significant difference in permeability either at 24 or 72 h post-treatment. However, CID pretreatment attenuated PFOS-induced permeability (54.69 μg/ml in the PFOS + CID group) compared with PFOS only group (79.81 μg/ml), but was still significantly increased compared with vehicle (39.07 μg/ml). At 24 h post-treatment, there was a trend of restoring PFOS-induced permeability by CID pretreatment, however, there is no statistical difference (Figure 5B).

Figure 5.

Figure 5.

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Protein kinase D (PKD) inhibition attenuates PFOS-mediated barrier dysfunction. 16HBE cells were pretreated with 10 μM CID 755673 (CID) for 1 h, then treated to 15 μM PFOS for 24–72 h. (A) TEER and (B) FITC-dextran flux were assessed. Bars represent mean ± SEM, n = 3. *p < .05, **p < .01, ***p < .001 indicate significance compared with vehicle (VEH). CID pretreatment + PFOS (P+C). Non-significant (n.s.) comparisons.

Compared with vehicle, PFOS caused a significant decrease in occludin (0.73-fold change), which was attenuated by CID pretreatment (0.98-fold change vs VEH group) (Figure 6). ZO-1 protein level was significantly reduced by PFOS treatment (0.59-fold change) and CID pretreatment did not prevent ZO-1 downregulation (0.66-fold change) (Figure 6). The distribution of occludin was significantly disrupted by PFOS, whereas CID pretreatment restored occludin protein expression and localization (Figure 7). PFOS treatment also reduced ZO-1 TiJOR, but CID pretreatment did not attenuate protein disruption (Figure 7). In line with our Western blot results, we have found that the integrity of occludin was dysregulated by PFOS treatment but rescued by CID pretreatment, whereas ZO-1 expression was not attenuated.

Figure 6.

Figure 6.

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PKD inhibition attenuates PFOS-disrupted occludin, but not ZO-1 protein. 16HBE cells were pretreated with DMSO or 10 μM CID 755673 (CID) for 1 h, then treated to 15 μM PFOS for 24 h. The cells were collected for protein and analyzed by Western blot. (A) Blot images of ZO-1 and occludin protein are shown and (B) fold change was determined by densitometric analysis. Bars represent mean ± SEM, n = 3. *p < .05, **p < .01, ***p < .001 indicate significance compared with vehicle (VEH). CID pretreatment + PFOS (P+C). Non-significant (n.s.) comparisons.

Figure 7.

Figure 7.

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PKD inhibition attenuates PFOS-disrupted occludin localization. 16HBE cells were pretreated with DMSO or 10 μM CID 755673 (CID) for 1 h, then treated to 15 μM PFOS for 24 h. Cells were fixed with methanol and processed for immunofluorescence. (A) Representative images of cells treated with PFOS and/or CID are shown. ZO-1 (yellow) and occludin (green) are shown alongside DAPI merged images. (B) ZO-1 and occludin TiJOR were determined. Bars represent mean ± SEM, n = 3. *p < .05, **p < .01, indicate significance compared with vehicle (VEH). CID pretreatment + PFOS (P+C). Scale bar = 100 μm.

DISCUSSION

There is growing evidence that PFOS is associated with adverse respiratory outcomes such as asthma (Dong et al., 2013; Zhou et al., 2017b; Zhu et al., 2016), impaired lung function (Kung et al., 2021), and an increased incidence of respiratory viral infections (Dalsager et al., 2021). Biodistribution studies indicate that PFOS accumulates in the lungs (Mamsen et al., 2019; Pérez et al., 2013) further implicating PFOS in lung disease development. Pulmonary barrier dysfunction is a noted feature in atopic asthma (Xiao et al., 2011). The pulmonary barrier restricts access to environmental toxicants and supports innate immune function and repair mechanisms (Crosby and Waters, 2010; Georas and Rezaee, 2014; Parker and Prince, 2011). As such, even small changes in barrier function can have large implications for asthma disease risk and severity. However, few studies have examined the impact of PFOS on respiratory barrier function. In this study, we used the maximum reported PFOS concentration in serum as the basis for our dose response experiments, because the PFOS lung bioaccumulation factor is not well characterized (Olsen et al., 1999). PFOS treatment disrupted barrier function in airway epithelial cells by dysregulating tight junction networks. Moreover, deficits in barrier function were accompanied by the activation of PKD, and pretreatment with the PKD inhibitor (CID) attenuated PFOS-induced barrier disruption. This is the first study to investigate the effects of PFOS on pulmonary barrier function, and link PFOS exposure with PKD signaling.

Initially, we observed that high concentrations of PFOS treatment induced significant cytotoxicity. However, non-toxic concentrations of PFOS decreased the secretion of IL-8, a potent chemokine that recruits neutrophils to injured areas (Reynolds et al., 2018). Previously, studies have shown that PFOS exposure suppresses humoral immune responses (Peden-Adams et al., 2008), in agreement with our data. Other studies have shown that inflammatory cytokines contribute to barrier dysfunction; however, it is cell type specific (Capaldo and Nusrat, 2009). IFNγ treatment decreased permeability in lung epithelial cells while increased permeability was observed in endothelial cells along with decreased E-cadherin and occludin (Ahdieh et al., 2001; Ng et al., 2015). Detailed studies are needed to understand the crosstalk between inflammatory cytokines and barrier dysfunction.

This study demonstrates that PFOS disrupts airway epithelial barrier function as measured by decreased TEER and increased permeability to FITC-dextran. We found that PFOS significantly reduced epithelial barrier function as early as 4 h following treatment. Although there are no studies on the effects of PFOS on pulmonary barrier function, our results are consistent with an increasing number of studies that have found disruption of tissue barrier integrity at the blood-testis barrier and blood-brain barrier (BBB) with PFOS. In a 28-day oral PFOS exposure, mice displayed morphological changes to the BBB histologically (Yu et al., 2020). Additionally, PFOS treatment resulted in decreased TEER in sertoli cells (Chen et al., 2017a) and other reports show that intestinal barrier function is negatively affected by PFOS (Diaz et al., 2021; Glynn et al., 2017; Wang et al., 2020). In our study, TEER reduction preceded changes in FITC-dextran permeability by at least 20 h, which may indicate dynamic changes in barrier function over time. Initial TEER loss suggests “pore” pathways are affected due to disturbances in ion transport, whereas permeability to FITC-dextran at later time points indicates increased macromolecule transport/diffusion by “leak” pathways (Shen et al., 2011). Further work is required to understand the potential interplay between physical interactions and downstream signaling pathways, and the kinetics involved in PFOS-mediated barrier dysfunction.

In our study, we observed that PFOS treatment disrupted the protein levels and localization of ZO-1 and occludin. Both proteins are implicated in the maintenance of the paracellular “leak” pathway and mediate responses to environmental cues (Marchiando et al., 2010; Shen et al., 2011; Van Itallie et al., 2010; Yu et al., 2005). Multiple studies have reported the loss and/or reorganization of tight junction proteins with PFOS exposure. Primary mouse sertoli cells had reduced expression of occludin protein following treatment to 30 μM PFOS for 12 h (Qiu et al., 2016). Additionally, another report shows that 20 μM PFOS treatment resulted in disruption of occludin localization. However, only ZO-1 protein levels decreased—though occludin protein remained unchanged in partial agreement with our study (Gao et al., 2017). These data support that the loss of barrier function by PFOS is driven both by changes in tight junction recycling (production and degradation) and protein trafficking/organization. However, our data do not preclude the possibility that other tight junctions besides occludin and ZO-1 such as claudins, tricelluin, and junctional adhesion proteins (JAMs) are involved. This is likely the case, given that evidence suggests there is functional redundancy between various members of the TAMP and claudin families (Anderson and Van Itallie, 2009; Schulzke et al., 2005).

Recently, studies have demonstrated the importance of the small molecule, PKD, in the regulation of pulmonary epithelial barrier function (Gan et al., 2013; Rezaee et al., 2013). PKD is a serine/threonine kinase that is implicated in many biological processes including angiogenesis (Ren, 2016), immune modulation (Veazey et al., 2020), protein trafficking (Hausser et al., 2005), cellular proliferation (Poli et al., 2014), and epithelial barrier function (Tinsley et al., 2004; Veazey et al., 2020). We have observed increased phosphorylation of PKD at 3 h PFOS post-treatment. Limited studies are available regarding the interactions of PFOS and PKD activity, but some reports indicate that PFOS can activate protein kinase C (PKC), a kinase upstream of PKD. Incidentally, PKC is also implicated in the regulation of barrier function (Koizumi et al., 2008; Waschke et al., 2006). In one report, gene profiling of fetal lungs from developmentally exposed mice showed elevated PKCα transcript (Rosen et al., 2009). Others have shown that PFOS activates PKC in cerebella granule cells (Lee et al., 2012). Pretreatment with CID partially attenuated PFOS-mediated barrier dysfunction with improved TEER and occludin protein levels. However, PKD inhibition did not ameliorate ZO-1 protein levels. Thus, our data show that PFOS mediates barrier dysfunction in part by activating PKD, but that downregulation of ZO-1 is not a crucial element of the barrier. ZO-1 knockout studies in mice also show normal barrier function in the GI tract. However, these mice were more susceptible to secondary insults (Kuo et al., 2021). Given the recent epidemiological evidence associating PFOS exposure with allergic asthma, and our findings, it is possible that PFOS may augment asthma disease risk or severity by modulating barrier function. PFOS-mediated barrier dysfunction could drive Th2 cytokine signaling or facilitate access to inhaled allergens (Figure 8). Future studies are needed to examine the effects of PFOS on barrier function in combination with additional insults or within asthma disease models to better understand the relationship between PFOS and allergic airway responses.

Figure 8.

Figure 8.

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Schematic for potential implications of PFOS-mediated barrier dysfunction on allergic airway disease risk. Non-exposed individuals have functional tight junctions that sequester inhaled environmental contaminants and allergens in the airway lumen, preventing further exposure. In exposed lungs, we hypothesize that PFOS causes a breakdown and internalization of tight junctions, impairing the ability of the respiratory system to filter out allergenic xenobiotics, resulting in augmented asthma risk. Figure was created with BioRender.com.

It is important to consider the limitations of our study. Our doses ranged from 5 to 25 μM (2500–12 500 ng/ml). The average serum concentration of PFOS in the United States from the 2017–2018 NHANES cohort was 4.25 ng/ml (CDC, 2018). Non-occupational exposure has been reported to be as high as 759.2 ng/ml in the United States (Steenland et al., 2009). However, extremely elevated levels have been observed in China, with employees from a fishery and their family members reaching average serum levels of 11 400 ng/ml and 3150 ng/ml respectively, with 1 individual recording a serum concentration of 31 400 ng/ml (Zhou et al., 2014). Taken together, our study more closely models occupational exposure or people who live in highly contaminated sites. Furthermore, serum concentrations may not reflect levels found in the lung. Two human studies have found that PFOS accumulates to similar levels in the lungs when compared with the liver (a target organ highly suspected of bioaccumulation). In one study, average adult lung PFOS levels were 29.1 ng/g of lung tissue (Pérez et al., 2013) and in another report, average fetal lung tissue levels were 2.59 ng/g of lung tissue (Mamsen et al., 2019). Neither study measured PFOS levels in plasma or serum, making it difficult to assess bioaccumulation. However, the lung PFOS concentrations were similar to serum PFOS levels measured in NHANES cohorts during similar periods (CDC, 2018). In a single dose gestational rodent exposure, PFOS levels in the lung were approximately 2 times higher than serum levels (Borg et al., 2010). However, others have demonstrated lung concentrations that were lower than serum measurements in rodents (Cui et al., 2009). More studies are needed to look at PFOS biodistribution to more accurately assess lung toxicity at relevant levels.

Another limitation of our study is that we did not fully elucidate the mechanisms underlying PFOS-mediated barrier dysfunction. PFOS is known to be a promiscuous pollutant that activates numerous nuclear receptors and signaling pathways within multiple organ systems, which makes it challenging to identify single points of regulation. Some notable effects of PFOS include activation of extracellular receptor kinase (ERK), peroxisome proliferator-activated receptors (PPARs), and inhibition of protein kinase B (AKT) activity. In lung epithelial cells, ERK has been shown to mediate PKC-dependent tight junction organization and cortactin degradation (Aggarwal et al., 2011; Zhao et al., 2012). Cortactin, an actin-binding protein, facilitates cell motility and cytoskeletal dynamics (Kirkbride et al., 2011). PKD-induced barrier dysfunction also involves cortactin-dependent actin remodeling (Rezaee et al., 2013), suggesting that PFOS mediated signaling may converge on the regulation of the actin cytoskeleton. However, more studies are required to understand these interactions. In contrast to our findings, activation of PPARs and inhibition of AKT are often associated with the enhancement of barrier function in the pulmonary epithelium. In one study, PPARγ agonists enhanced barrier function in the nasal epithelium (Ogasawara et al., 2010). However, other studies have shown that PPARγ agonists promote Th2 inflammation which is often associated with barrier dysfunction (Chen et al., 2017b). Lastly, activation of AKT is largely associated with inflammation and internalization of cell-cell junctions (He et al., 2020; Ye et al., 2019). Complex crosstalk between these pathways and others probably mediates PFOS-induced barrier function. The use of high-throughput screening assays or “omics” analysis may be necessary to tease apart the contributions of each pathway, especially as future research incorporates in vivo and human exposure data. Nonetheless, our data show that PKD is an important mediator of PFOS toxicity in bronchial epithelial cells and should be investigated further.

CONCLUSION

In conclusion, PFOS treatment to airway epithelial cells caused a loss of barrier integrity along with dysregulation of tight junction proteins. Moreover, inhibiting PKD preserved barrier integrity and maintained occludin, but not ZO-1 protein levels and organization. Our findings are the first to demonstrate that the airway epithelial barrier is a target of PFOS exposure, which has important implications on asthma disease pathogenesis More research is needed to assess the impact of PFOS in in vivo models and evaluate the contribution of PFOS exposure to allergic airway disease risk and its exacerbations.

SUPPLEMENTARY DATA

Supplementary data are available at Toxicological Sciences online.

AUTHOR CONTRIBUTIONS

J.H.L. and I.R. conceived and designed the experiments. J.H.L. conducted the experiments. J.H.L. analyzed the data. J.H.L. wrote and J.H.L., Q.W., and I.R. revised the manuscript.

Supplementary Material

kfac096_Supplementary_Data
Click here for additional data file. (1.2MB, zip)

ACKNOWLEDGMENTS

Figures 1–7 were created with PowerPoint and GraphPad Prism. Figure 8 was created using BioRender software. We thank Cortney Pang for assisting in the proofing of the final manuscript.

FUNDING

National Institutes of Health Toxicology Training (T32ES007026). The funding body has no role in the design of the study, data collection, analysis, interpretation of data, or writing of the manuscript.

DECLARATION OF CONFLICTING INTERESTS

The authors declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Contributor Information

Joseph H Lucas, Department of Environmental Medicine, University of Rochester Medical Center, Rochester, New York 14642, USA.

Qixin Wang, Department of Environmental Medicine, University of Rochester Medical Center, Rochester, New York 14642, USA.

Irfan Rahman, Department of Environmental Medicine, University of Rochester Medical Center, Rochester, New York 14642, USA.

Data Availability

All data and materials are described in the manuscript.

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Associated Data

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Supplementary Materials

kfac096_Supplementary_Data
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Data Availability Statement

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