
==== Front
iScience
iScience
iScience
2589-0042
Elsevier

S2589-0042(24)01928-X
10.1016/j.isci.2024.110703
110703
Article
Defective CFTR modulates mechanosensitive channels TRPV4 and PIEZO1 and drives endothelial barrier failure
Amoakon Jean-Pierre amoakoja@mail.uc.edu
12∗
Lee Jesun 2
Liyanage Pramodha 2
Arora Kavisha 2
Karlstaedt Anja 3
Mylavarapu Goutham 4
Amin Raouf 4
Naren. Anjaparavanda P. apnaren@cshs.org
1245∗∗
1 Department of Systems Biology and Physiology, University of Cincinnati College of Medicine, Cincinnati, OH, USA
2 Division of Pulmonary Medicine and Critical Care, Cedars-Sinai Medical Center, Los Angeles, CA, USA
3 Department of Cardiology, Smidt Heart Institute, Cedars-Sinai Medical Center, Los Angeles, CA, USA
4 Division of Pulmonary Medicine, Cincinnati Children’s Hospital Medical Center, Cincinnati, OH, USA
∗ Corresponding author amoakoja@mail.uc.edu
∗∗ Corresponding author apnaren@cshs.org
5 Lead contact

09 8 2024
20 9 2024
09 8 2024
27 9 1107032 6 2024
25 6 2024
6 8 2024
© 2024 The Author(s)
2024
https://creativecommons.org/licenses/by-nc-nd/4.0/ This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).
Summary

Cystic fibrosis (CF) is a genetic disease caused by a mutation in the cystic fibrosis transmembrane conductance regulator (CFTR) gene. Despite reports of CFTR expression on endothelial cells, pulmonary vascular perturbations, and perfusion deficits in CF patients, the mechanism of pulmonary vascular disease in CF remains unclear. Here, our pilot study of 40 CF patients reveals a loss of small pulmonary blood vessels in patients with severe lung disease. Using a vessel-on-a-chip model, we establish a shear-stress-dependent mechanism of endothelial barrier failure in CF involving TRPV4, a mechanosensitive channel. Furthermore, we demonstrate that CFTR deficiency downregulates the function of PIEZO1, another mechanosensitive channel involved in angiogenesis and wound repair, and exacerbates loss of small pulmonary blood vessel. We also show that CFTR directly interacts with PIEZO1 and enhances its function. Our study identifies key cellular targets to mitigate loss of small pulmonary blood vessels in CF.

Graphical abstract

Highlights

• CFTR deficiency promotes barrier failure in the pulmonary microvascular endothelium

• CFTR deficiency alters endothelial cells lipid metabolism and membrane fluidity

• CFTR deficiency sensitizes TRPV4 to fluid shear stress

• CFTR directly interacts with PIEZO1

Cell biology; Functional aspects of cell biology; Pathophysiology

Subject areas

Cell biology
Functional aspects of cell biology
Pathophysiology
Published: August 9, 2024
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pmcIntroduction

Cystic fibrosis (CF) is a life-threatening genetic disorder caused by a deleterious mutation in the cystic fibrosis transmembrane conductance regulator (CFTR) gene. The CFTR protein is a cAMP-dependent chloride channel that is highly expressed in epithelial cells lining the lungs, the digestive tract, and reproductive organs.1 Respiratory failure is the main cause of mortality in CF and often results from the collective effects of alveolar mucus plugging, lung fibrosis, chronic inflammation, progressive destruction of the structural airway, and poor ventilation.

The pathophysiology underlying CF lung disease may be related to multiple underlying processes. Historically, pathology in the airway epithelium has been of particular interest. However, The CFTR protein is also expressed on endothelial cells (ECs), among other cell types.2 Available studies indicate that endothelial cell dysfunction plays a role in vascular pathology and perfusion deficits.3,4,5,6 Sub-optimal gas exchange in the alveoli may also occur if endothelial cells in the pulmonary capillaries do not function normally, precluding adequate oxygenation.7,8 Consequently, endothelial cell dysfunction may contribute to CF lung disease; yet the role of CFTR in endothelial cells within the pulmonary vasculature is not fully understood.

Endothelial cells are constantly subject to shear stress (SS), which is one of the most important external stimuli for endothelial homeostasis.9,10,11,12,13 Small pulmonary blood vessels (SPBVs) must withstand physiologically high SS.14,15 In non-CF lung diseases, such as emphysema and other obstructive pulmonary diseases, pulmonary vascular remodeling and pruning of SPBVs occurs; these changes coincide with reduced CFTR expression and blood flow changes.16,17,18,19,20,21,22,23 This association suggests CFTR may play a role in microvascular homeostasis. Therefore, we set out to investigate how CFTR dysfunction affects the response of ECs to SS in the lung microvasculature using a vessel-on-a-chip model.24

We demonstrate that dysfunctional CFTR leads to loss of SPBVs by promoting endothelial barrier failure and impairing angiogenesis and wound repair. We identify a clear mechanism of endothelial barrier failure that is SS dependent and that involves TRPV4, a calcium-permeable mechanosensitive channel (MsC), and its associated downstream calcium signaling. Furthermore, we show that CFTR deficiency negatively affects the function of PIEZO1, another calcium-permeable MsC that is known to play a role in angiogenesis and wound repair,25,26,27 two processes that can mitigate loss of SPBVs. We present evidence that CFTR directly interacts with and enhances the function of PIEZO1 and that CFTR deficiency reduces PIEZO1 channel activity. Overall, we establish previously unknown mechanisms underlying the progression of CF vascular disease and identify key cellular targets relevant to the loss of SPBVs in CF.

Results

CFTR deficiency promotes barrier failure in lung microvascular endothelial cells in the presence of shear stress

Since CFTR is expressed and functional in ECs (Figure S1), and vascular perturbations and perfusion deficits are reported in CF patients,2,3,4,5,6 we hypothesized that pulmonary vascular remodeling may occur in CF. In a pilot study, we performed pulmonary vascular morphometry and reconstructed the pulmonary vasculature of 40 individuals with CF. We found that early in the clinical course of CF when pulmonary function is normal, most of the total blood volume (TBV) is in small blood vessels (cross-sectional surface area [CSA] < 10 mm2). However, in CF patients with severe lung disease, most of the TBV was in larger blood vessels (Figures 1A and S2). We suspected that this extensive remodeling was due to loss of SPBVs and progressive redistribution of total blood volume to larger vessels.Figure 1 Shear stress stimulation on CFTR-defective lung microvascular ECs leads to loss of endothelial monolayer integrity

(A) Representative images of reconstructed pulmonary vasculatures of CF patients using pulmonary vascular morphometry extracted from volumetric computed tomography scans of the chest at maximum inspiration (n > 10 patients).

(B and C) Multi-channel vessel-on-a-chip microfluidic device (B) connected to a peristaltic pump for shear flow studies (C). Shear stress (SS) ranges from 10 to 25 dyne/cm2 in the microvasculature.15,28 Some of the channels remain in static conditions as controls for each treatment (C).

(D) Representative images used to quantify data in (E) (scale bar, 1,000 μm). Human lung microvascular ECs (HMVEC-Ls) were treated with DMSO or 20 μM CFTRinh-172. For each treatment, cells remained in static conditions or were subjected to SS (15 dyne/cm2) for 24 h.

(E) Cell counts per image of DMSO and CFTRinh-172-treated HMVEC-Ls in static or flow conditions (15 dyne/cm2) after 24 h (n > 10; data are mean ± SD). Statistical analyses: two-way ANOVA: interaction (p < 0.001).

(F) Change in cell counts per image of negative-control-siRNA- and CFTR-siRNA-treated HMVEC-Ls before and after SS for 24 h (n = 14; data are mean ± SD). Statistical analyses: two-tailed Student’s t test.

(G) Representative images of live recording of HMVEC-Ls subjected to SS (15 dyne/cm2) showing progressive cell retraction and formation of gaps in the endothelial monolayer (scale bar: 1,000 μm).

(H) Representative immunofluorescence images showing vascular endothelial cadherin (VE-cad) expression (green) of DMSO and CFTRinh-172-treated HMVEC-Ls in static or flow conditions (15 dyne/cm2) after 24 h (scale bar: 100 μm). See also Figures S1–S4 and Videos S1 and S2. ∗p ≤ 0.05; ∗∗p ≤ 0.01; ∗∗∗p ≤ 0.001; ∗∗∗∗p ≤ 0.0001.

Hemodynamics varies between small and large blood vessels, especially SS, which plays a critical role in EC migration, proliferation, flow alignment, and NO production.9,10,11,12,13 Depending on the severity of CF lung disease, patients may develop pulmonary hypertension,29,30,31 which can result in higher pressure and increased SS.32,33 Therefore, we decided to investigate the response of CFTR-defective (by knockdown or inhibition) HMVEC-Ls under high SS using a vessel-on-a-chip model (Figures 1B, 1C, and S3). We used SS of 15 dyne/cm2, which is within the physiological range for healthy adults (10–25 dyne/cm2).15,28

In static conditions (e.g., without SS), inhibiting CFTR with CFTR-specific inhibitor CFTRinh-172,34 or knocking down CFTR, had no adverse effects on cell viability compared to control. However, applying SS to CFTR-defective ECs for 24 h caused significant apoptosis compared to control (Figures 1D–1F and S4).

Live recordings revealed that high SS causes a progressive retraction of ECs from one another and subsequent formation of gaps in the monolayer (Figure 1G and Videos S1 and S2). A recent CFTR study reported similar observations under low SS (5 dyne/cm2).6

Video S1. Live recording of HMVEC-Ls treated with DMSO and subjected to shear stress, related to Figure 1

Recording shows progressive cell retraction and formation of small gaps in the endothelial monolayer. Timestamp: (H:MM:SS).

Video S2. Live recording of HMVEC-Ls treated with 20 μM CFTRinh-172 and subjected to shear stress, related to Figure 1

Recording shows faster cell retraction and formation of larger gaps in the endothelial monolayer compared to control DMSO. Timestamp: (H:MM:SS).

We speculated that the cell retraction was due to degradation of adherens junctions (AJs) and loss of cell-cell contact. Staining for vascular endothelial cadherin (VE-cad), the main component of AJs, revealed strong expression of VE-cad in static conditions in both control and CFTRinh-172-treated cells. However, under SS, inhibiting CFTR caused a significant reduction in VE-cad expression; this reduction was more pronounced in cells displaying a greater degree of retraction (Figure 1H). These findings suggest that loss of AJs leads to reduced cell viability when CFTR is defective under SS.

Overactivation of mechanosensitive channel TRPV4 causes shear-stress-dependent barrier failure when CFTR is defective

AJs have been shown to be directly cleaved by calcium-dependent ubiquitous protease calpains.27,35,36,37 Only two calpain isoforms are expressed in ECs: μ-calpain and m-calpain.38 μ-calpain requires only micromolar concentration of calcium for activation, whereas m-calpain requires almost three orders of magnitude more calcium.39 Only m-calpain specifically can cleave AJs.36,40 Therefore, we hypothesized that there must be a more robust SS-dependent intracellular calcium increase in ECs when CFTR is defective to chronically activate m-calpain.

Live calcium imaging revealed a low static baseline level of intracellular calcium that rose sharply in response to a 1-min SS stimulus (Videos S3 and S4).41 The rise was more pronounced and sustained when CFTR was knocked down compared to control (Figures 2B–2D).Figure 2 Overactivation of TRPV4 causes shear-stress-dependent loss of endothelial monolayer integrity when CFTR is defective

(A) Representative images of live recording of intracellular calcium levels (green fluorescence) of HMVEC-Ls under shear stress (SS) (15 dyne/cm2) used to quantify data in (B), (C) and (D) (scale bar: 1,000 μm).

(B–D) (B) Time courses used to quantify data in (C) and (D). Change in intracellular calcium levels of negative-control-siRNA- and CFTR-siRNA-treated HMVEC-Ls subjected to SS (15 dyne/cm2) for only 1 min out of the 10-min total recording time (n = 131 cells; data are mean ± SE). (C and D) Change in intracellular calcium levels from (B) at 1 min (C) and at 3 min (D) (n = 131 cells; data are mean ± SD). Statistical analyses: two-tailed Student’s t test.

(E) Representative images of live recording of intracellular calpain activity (blue fluorescence) of HMVEC-Ls under SS (15 dyne/cm2) used to quantify data in (F) and (G) (scale bar: 1,000 μm).

(F) Representative time courses used to quantify data in (G). Change in intracellular calpain activity of negative-control-siRNA- and CFTR-siRNA-treated HMVEC-Ls subjected to SS (15 dyne/cm2) for 15 min (n = 431 cells; data are mean ± SE).

(G) Change in intracellular calpain activity from (F) at 15 min (n = 431 cells; data are mean ± SD). Statistical analyses: two-tailed Student’s t test.

(H) Representative images used to quantify data in (I) of HMVEC-Ls subjected to SS (15 dyne/cm2) for at least 24 h (scale bar: 1,000 μm). HMVEC-Ls were treated with 20 μM CFTRinh-172 only, or with a combination of 20 μM CFTRinh-172 and 500 nM calpastatin.

(I) Cell counts per image of CFTRinh-172- and CFTRinh-172 + calpastatin-treated HMVEC-Ls in static or flow conditions (15 dyne/cm2) after 24 h (n = 15; data are mean ± SD). Statistical analyses: two-way ANOVA: interaction (p < 0.0001).

(J) Representative images used to quantify data in (K) and (L) (scale bar: 1,000 μm). HMVEC-Ls were treated with DMSO, 3 μM calcium ionophore A23187, 2.5 μM Yoda1, and 50 μM GSK1016790A, in regular medium, calcium-depleted medium, or calcium-depleted medium supplemented with 50 mM CaCl2. For all treatments, cells remained in static conditions.

(K and L) Percent cell confluency per image from (J) of DMSO-, calcium ionophore A23187-, Yoda1- (L), and GSK1016790A (K)-treated HMVEC-Ls in regular medium, calcium-depleted medium, or calcium-depleted medium supplemented with CaCl2, in static conditions (n = 3; data are mean ± SD). Statistical analyses: one-way ANOVA with Tukey’s multiple comparison.

(M) Representative images before immunofluorescence in (M) and used to quantify data in (N) (scale bar: 1,000 μm). HMVEC-Ls were treated with DMSO, 20 μM CFTRinh-172 only, or with a combination of 20 μM CFTRinh-172 and 1 μM TRPV4-specific inhibitor HC067047. For each treatment, cells either remained in static conditions or were subjected to SS (15 dyne/cm2) for at least 24 h.

(N) Representative immunofluorescence images from (L) showing VE-cad expression (green) (scale bar: 100 μm).

(O) Cell counts per image of DMSO-, CFTRinh-172-, and CFTRinh-172 + HC067047-treated HMVEC-Ls in static or flow conditions (15 dyne/cm2) after 24 h (n > 10; data are mean ± SD). Statistical analyses: two-way ANOVA with Tukey’s multiple comparison: interaction (p < 0.01). See also Videos S3, S4, S5, S6, S7, and S8. ∗p ≤ 0.05; ∗∗p ≤ 0.01; ∗∗∗p ≤ 0.001; ∗∗∗∗p ≤ 0.0001.

Video S3. Calcium imaging of HMVEC-Ls in static conditions, related to Figure 2

Live recording of intracellular calcium increase (green fluorescence) in static conditions shows that baseline level of intracellular calcium is low and stable. Timestamp: (H:MM:SS).

Video S4. Calcium imaging of HMVEC-Ls in response to shear stress, related to Figure 2

Live recording of intracellular calcium increase (green fluorescence) in response to a 1-min shear stress (15 dyne/cm2). Baseline level of intracellular calcium is low in static conditions but rises sharply in response to shear stress. Timestamp: (H:MM:SS).

We inferred that this higher rise in calcium would lead to higher calpain activity. To measure the real-time activity of calpain in response to SS, we used CMAC-t-BOC-Leu-Met, a cell-permeable compound that is non-fluorescent in the extracellular environment. Once inside the cell, this compound is cleaved by activated calpain, becomes impermeable, and fluoresces (Figure 2E and Video S5).42 We found higher calpain activity when CFTR was knocked down compared to control (Figures 2F and 2G), which can disrupt expression of AJs.27 Next, we inhibited CFTR while also inhibiting calpain using the endogenous inhibitor calpastatin; this inhibitor binds calpain only after it is activated by calcium and has a higher affinity for m-calpain.43 These steps mitigated the detrimental effects of CFTR deficiency under SS (Figures 2H and 2I) and suggest that calpain plays a role in the mechanism of barrier failure in CF.

Video S5. Calpain activity of HMVEC-Ls under shear stress, related to Figure 2

Live recording of intracellular calpain activity (blue fluorescence) of HMVEC-Ls under shear stress (15 dyne/cm2) using CMAC-t-BOC-Leu-Met, a cell-permeable compound that is non-fluorescent in the extracellular environment. However, once inside the cell, this compound gets cleaved by activated calpain, becomes impermeable, and fluoresces. Timestamp: (H:MM:SS).

To study the effects of chronic calcium entry on EC viability, we used calcium ionophore A23187 and found that high intracellular calcium levels led to cell death (Figure 2J and Video S6). Since the effects of CFTR deficiency were only noticeable in the presence of SS, we suspected an upstream involvement of MsCs. Two channels, PIEZO1 and TRPV4, stood out because they are not only directly activated by SS but also permeable to calcium.27,44,45,46,47,48,49,50 Furthermore, PIEZO1 has been shown to act upstream of TRPV4 by activating phospholipase A2, which is required for the activation of TRPV4.41

Video S6. Live recording of HMVEC-Ls treated with 3 μM calcium ionophore A23187 in regular EBM-2 medium under static conditions, related to Figure 2

High intracellular calcium levels led to cell death. Timestamp: (H:MM:SS).

We started by investigating if chronic activation of PIEZO1 by selective agonist Yoda151 would also lead to cell death. We found that chronic activation of PIEZO1 led to rapid cell death (about 1 h); this occurred only in the presence of calcium (Figures 2J and 2K and Video S7). Therefore, we rationalized that chronic activation of PIEZO1 in response to SS might be leading to cell death when CFTR is defective. We inhibited PIEZO1 using a cationic-permeable MsCs inhibitor GsMTx-4 while also inhibiting CFTR under SS52; these steps were not sufficient to rescue the cells (Figures S5A and S5B). It should be noted that GsMTx-4, though non-specific for PIEZO1, does not inhibit TRPV4.41

Video S7. Live recording of HMVEC-Ls treated with 2.5 μM Yoda1 in regular EBM-2 medium under static conditions, related to Figure 2

Chronic activation of PIEZO1 led to rapid cell death (about 1 h). Timestamp: (H:MM:SS).

Interestingly, although PIEZO1 activation reportedly leads to a transient intracellular calcium elevation, TRPV4 activation induces a stronger and more sustained rise.41 Therefore, TRPV4 is better equipped to provide the required calcium levels to activate m-calpain. Besides, calcium entry via TRPV4 specifically increases lung endothelial permeability; when calcium enters ECs via another calcium channel, α1G, endothelial permeability is not altered.53 We reasoned that this difference could be explained by the propensity of TRPV4 to induce stronger intracellular calcium elevation. Additionally, calcium entry via TRPV4 has previously led to the remodeling of AJs in ECs.54 Evidence from animal models also highlights the relevance of TRPV4; when TRPV4 is activated by a specific agonist, GSK1016790A (GSK101), pulmonary edema and alveolar hemorrhage occur in the lungs of rats.55,56,57 For all these reasons, further investigation of TRPV4 was warranted.

Chronic activation of TRPV4 by GSK10156,57 led to progressive cell retraction and formation of intercellular gaps in the endothelial monolayer (about 10 h); these changes occurred only in the presence of calcium (Figures 2J and 2L and Video S8). Next, we inhibited TRPV4 using a TRPV4-specific inhibitor, HC067047,58 while also inhibiting CFTR under SS; similar to the inhibition of calpain, the inhibition of TRPV4 was sufficient to rescue the cells and maintain the integrity of AJs (Figures 2M–2O). These results are comparable to those from a study demonstrating that endothelial activity of PAR1, a TRPV4 sensitizer, leads to sustained calcium elevation and increased vascular permeability through disruption of AJs. Moreover, TRPV4 inhibition mitigated the detrimental effects of PAR1 activation.59 A recent in vivo/ex vivo study in the lungs of rats and mice also showed that CFTR deficiency leads to increased endothelial calcium response, loss of endothelial barrier integrity, and edema formation. TRPV4 inhibition or knockdown (KD) prevented these effects.60 Taken together, these studies suggest that proteins and molecules that lower the activation threshold of TRPV4 could lead to calcium overload and increased vascular permeability. Thus, we rationalized that CFTR could modulate TRPV4 function.

Video S8. Live recording of HMVEC-Ls treated with 50 μM GSK1016790A in regular EBM-2 medium under static conditions, related to Figure 2

Chronic activation of TRPV4 led to progressive cell retraction and formation of intercellular gaps in the endothelial monolayer (about 10 h). Timestamp: (H:MM:SS).

CFTR deficiency alters lipid metabolism and cytoskeleton organization

Proteins and molecules that change the fluidity or curvature of the membrane have been found to modulate MsCs.61,62 In fact, a stiffer membrane inhibits MsCs, whereas a more fluid membrane enhances their activation.63,64,65 Because the membrane is made up of lipids, changes in lipid composition can directly affect membrane mechanical properties, including fluidity.66,67

Defective CFTR expression has been shown to increase sphingolipid synthesis and change membrane composition.68 Moreover, sphingosine-1-phosphate (S1P), a signaling sphingolipid known to increase AJs’ expression and barrier impermeability, may be transported by CFTR.69,70,71,72 In fact, S1P supplementation mitigated loss of endothelial barrier integrity when CFTR was inhibited or knocked down.73 Taken together, these findings suggest CFTR plays a role in lipid regulation.

We performed bulk RNA sequencing on control and CFTR-KD HMVEC-Ls to investigate lipid metabolism. For technical reasons, RNA sequencing was performed on cells cultured in static conditions. Nonetheless, this approach allows us to appreciate gene expression changes caused by CFTR KD without the confounding factor of flow. We found 349 genes to be differentially expressed in CFTR-KD HMVEC-Ls. The top 30 differentially expressed genes are plotted (Figure 3A), and the heatmap shows a consistent signature across samples (Figure 3B).Figure 3 CFTR knockdown leads to lipid metabolic changes and cytoskeleton reorganization

(A) Volcano plot of differentially expressed genes (DEGs) in CFTR-siRNA-treated HMVEC-Ls compared to control. Names of top 30 genes are displayed.

(B) Heatmap of DEGs in negative-control-siRNA- and CFTR-siRNA-treated HMVEC-Ls.

(C) Differentially regulated pathways of CFTR-siRNA-treated HMVEC-Ls predicted by ingenuity pathway analysis with Z score of at least ±0.5. Numbers of DEGs are indicated on bars.

(D) Change in gene expression of select genes involved in sphingolipids and S1P metabolism in CFTR-siRNA-treated HMVEC-Ls compared to control. ∗DEG.

(E) Representative images used to quantify data in (F) of HMVEC-Ls subjected to SS (15 dyne/cm2) for at least 24 h (scale bar: 1,000 μm). HMVEC-Ls were treated with 20 μM CFTRinh-172 only or with a combination of 20 μM CFTRinh-172 and 1 μM S1PL-IN-31.

(F) Cell counts per image of CFTRinh-172- and CFTRinh-172 + S1PL-IN-31-treated HMVEC-Ls in static or flow conditions (15 dyne/cm2) after 24 h (n = 15; data are mean ± SD). Statistical analyses: two-way ANOVA: interaction (p < 0.0001).

(G–I) Change in gene expression of select genes involved in adherens junctions and calpain activity regulations (G), phospholipid metabolism (H), and mechanotransduction (I) in CFTR-siRNA-treated HMVEC-Ls compared to control. ∗DEG.

(J) Representative immunofluorescence images showing TRPV4 surface expression (green) and F-actin expression (red) of HMVEC-Ls in static or flow conditions (15 dyne/cm2) after 24 h (scale bar: 100 μm).

(K) Change in gene expression of select genes involved in cytoskeleton binding and actin regulation in CFTR-siRNA-treated HMVEC-Ls compared to control. ∗DEG.

(L) Representative immunofluorescence images used to quantify data in (M) (scale bar, 100 μm). F-actin staining (red) of negative-control-siRNA- and CFTR-siRNA-treated HMVEC-Ls in static or flow conditions (15 dyne/cm2) after 24 h. White arrow depicts direction of flow.

(M) Modal amount and goodness of fit of actin filaments with direction of flow for negative-control-siRNA- and CFTR-siRNA-treated HMVEC-Ls. Directionality analysis of F-actin filaments was performed using directionality tool in Fiji (ImageJ) with local gradient orientation method. Statistical analyses: two-tailed Student’s t test.

(N) Change in gene expression of select genes involved in focal adhesion in CFTR-siRNA-treated HMVEC-Ls compared to control. ∗DEG. See also Figure S6 and Videos S9 and S10. ∗p ≤ 0.05; ∗∗p ≤ 0.01; ∗∗∗p ≤ 0.001; ∗∗∗∗p ≤ 0.0001.

Ingenuity pathway analysis revealed changes in vascular system and lipid synthesis. For example, there was a significant upregulation of vascular permeability, an overall upregulation of angiogenesis/vasculogenesis (possibly as a compensatory mechanism), and an overall downregulation of lipid synthesis (Figure 3C). Using the DAVID database, we identified several genes enriched in pathways of interest. We decided to focus not only on genes that were significantly differentially expressed but also on the trends of certain important genes (e.g., upregulation or downregulation, though not significant).

Selected genes in the sphingolipids and S1P pathway show that ACER2, a ceramidase responsible for the hydrolysis of ceramides into sphingosine, the precursor of S1P, is significantly downregulated. Expression of SGPL1, the gene that encodes S1P lyase (the enzyme responsible for breakdown of S1P), tends to be upregulated. Additionally, the expression of S1PR1 (the gene that encodes S1P receptor 1) tends to be downregulated (Figure 3D). All of this points to downregulation of S1P signaling. We found that raising intracellular S1P levels by inhibiting S1P lyase activity with S1PL-IN-31 was sufficient to mitigate the detrimental effects of CFTR deficiency under SS (Figures 3E and 3F). As mentioned earlier, S1P increases AJs surface expression.69,71 Therefore, these results indicate that AJs disassembly is pivotal to the loss of endothelial barrier integrity when CFTR is defective under SS. Thus, we also looked at the AJs pathway and found that expression of CDH5 (VE-cad), and of its membrane regulator CTNND1 (p120-catenin),74 tend to be downregulated. Furthermore, the expression of CAPN2 (m-calpain) tends to be upregulated, whereas the expression of CAST (calpastatin), the endogenous inhibitor of calpain, tends to be downregulated (Figure 3G).

We also found an overall downregulation of genes associated with phospholipid synthesis (Figure 3H), which could change membrane composition and dynamics and affect mechanotransduction. We investigated the two main endothelial MsCs and found that neither PIEZO1 nor TRPV4 expression levels were changed (Figure 3I). However, KDR (vascular endothelial growth factor receptor 2 [VEGFR2]) expression level was drastically reduced (Figure 3I). This reduction is significant because VEGFR2 forms a junctional sensory complex with VEGFR3 (FLT4), CD31 (PECAM1), and VE-cad (CDH5) that mediates SS signaling.75,76 We speculate that reduced VEGFR2 expression and loss of VE-cad are likely to alter mechanosensation when CFTR is defective. Besides, the signaling cascade that occurs when VEGF, which is significantly increased in the serum of CF patients, binds to VEGFR2 is critical for EC proliferation, migration, survival, and angiogenesis.77,78 A notable disruption in VEGF/VEGFR2 signaling could prevent new blood vessel formation (Figure S6 and Videos S9 and S10), wound healing, and further exacerbate loss of SPBVs. However, it is important to note that even though angiogenesis appears to be reduced in the CF pulmonary microvasculature, there are several reports of the opposite in larger vessels such as the bronchial vessels.79,80 In fact, the bronchial circulation, thanks to its higher proliferative capacity, undergoes increased angiogenesis in CF.79,80,81,82 This phenomenon is also observed in other inflammatory diseases such as asthma and interstitial pulmonary fibrosis.81,82 Though beneficial in the short term by increasing gas exchange, this increased vascularization poses the risk of massive hemoptysis (blood vessel rupture, coughing up blood).79,81,82 The reason for the difference in angiogenic activity between capillaries and larger vessels is unclear. Nonetheless, we believe it could be due to a difference in SS and VEGF/VEGFR2 signaling.

Video S9. Live recording of tube formation of DMSO-treated HMVEC-Ls in Figure S6B, related to Figure 3

Tubes are intact after 8 h and do not start breaking down until after 10 h. Timestamp: (H:MM:SS).

Video S10. Live recording of tube formation of calcium ionophore A23187-treated HMVEC-Ls in Figure S6B, related to Figure 3

Tubes start breaking down after only 6 h and are completely broken down by 8 h. Timestamp: (H:MM:SS).

Additionally, since (1) the effect of CFTR deficiency is only apparent under SS and (2) transcriptomics data show that TRPV4 expression is unaltered when CFTR is defective, we hypothesized that there must be an SS-dependent regulation of TRPV4 surface expression. Consequently, we performed immunofluorescence staining using an anti-TRPV4 antibody that binds to an extracellular loop of TRPV4. This staining was performed without permeabilization to ensure binding only to TRPV4 on the cell surface. Also, F-actin staining helped highlight the overall structure of the cell. We found that in static conditions, TRPV4 was mainly expressed in clusters at intercellular junctions. However, clustering was abrogated under SS (Figure 3J). A recent study showed that under static conditions, TRPV4 channels cluster and form a complex with β-catenin at the AJs. SS leads to reduced clustering of TRPV4, its dissociation from β-catenin, and its translocation to the basal membrane.83 Thus, we proposed that TRPV4, when in a complex with β-catenin, might stabilize AJs. This protective effect is lost when TRPV4 is translocated to the basal membrane. Consequently, when CFTR is defective and SS is applied, the instability of β-catenin, combined with the cleavage of VE-cad, could result in faster degradation of AJs.

Also, the loss of mechanotransduction has further implications. Impaired mechanotransduction has been associated with reduced EC alignment and actin cytoskeleton disorganization.76,84,85,86 Several genes in the cytoskeleton binding and actin regulation pathway, such as MYO5B and LASP1, are significantly downregulated when CFTR is knocked down (Figure 3K). Interestingly, MYO5B, an actin-based mechanoenzyme, appears to be required for trafficking of CFTR to the plasma membrane and to recycling endosomes.87 Moreover, gene expression of WDR1, an actin-regulating protein that induces actin filament disassembly, is increased.88,89

Since SS induces EC cytoskeleton remodeling and alignment in the direction of flow,84,85,86 we hypothesized that F-actin organization under SS would be disrupted when CFTR was knocked down. We found similar F-actin protein expression for control and CFTR-KD ECs in static conditions. However, F-actin expression was reduced and completely disorganized under SS in CFTR-KD ECs (Figures 3L and 3M).

Given the direct role of actin in the regulation of focal adhesion molecules,90,91,92 we sought to determine if the focal adhesion pathway was affected. We discovered that several key genes were downregulated (Figure 3N). These findings suggest that CFTR-defective ECs are less likely to stay anchored to the extracellular matrix under SS, which further aggravates loss of barrier integrity.

CFTR deficiency sensitizes TRPV4 to fluid SS through changes in AA and EPA metabolism and increased membrane fluidity

Besides phospholipid and sphingolipid imbalances, polyunsaturated fatty acid (PUFA) imbalances have also been identified in CF patients and CFTR-knockout mice. Specifically, an increase in arachidonic acid (AA) and a decrease in docosahexaenoic acid (DHA) levels have been noted.93,94 These imbalances were also reported in nasal epithelial cells of CF patients, regardless of pancreatic sufficiency.95,96 Also, expression of defective CFTR led to increased eicosapentaenoic acid (EPA) levels in human bronchial epithelial cells.97,98,99

In the membrane, AA and EPA are typically part of the acyl chains of phospholipids. However, they can also occur as free fatty acids derived from the hydrolysis of phospholipids by phospholipases.66 When incorporated into the lipid bilayer, AA and EPA can alter the mechanical properties of the cell membrane and modulate the activity of MsCs.63,64,66

Our transcriptomic analysis revealed that expression of FADS2, the enzyme that catalyzes the first and rate-limiting step in the synthesis of AA and EPA from essential fatty acid precursors linoleic acid (LA) and alpha linoleic acid (αLA), tends to be downregulated (Figure 4A). LA and αLA levels are reported to be reduced in the serum of CF patients,99 which correlates with the lower enzymatic activity of FADS2. Nonetheless, PLA2G4A and PLA2G4B, the genes that encode cytosolic phospholipase A2 (the enzyme that catalyzes the release of AA from phospholipids), tend to be upregulated (Figure 4A). ELOVL1, a gene that enables elongase activity, which is crucial in AA, EPA, and sphingolipid syntheses, also tends to be upregulated (Figure 4A). Therefore, based on these results and the aforementioned studies suggesting an association between CF deficiency and increased AA and EPA levels, we rationalized that CFTR-defective HMVEC-Ls would also display increased AA and EPA levels. We quantified free AA and EPA levels using targeted liquid chromatography-tandem mass spectrometry; we discovered that CFTR-KD HMVEC-Ls had significantly higher AA and EPA levels than control cells (Figures 4B and 4C).Figure 4 CFTR deficiency lowers the flow-induced activation threshold of TRPV4 through changes in AA and EPA metabolism and increased membrane fluidity

(A) Change in gene expression of select genes involved in AA and EPA metabolism in CFTR-siRNA-treated HMVEC-Ls compared to control. ∗DEG.

(B and C) AA (B) and EPA (C) content analyzed by liquid chromatography-tandem mass spectrometry in negative-control-siRNA- and CFTR-siRNA-treated HMVEC-Ls. Statistical analyses: two-tailed Student’s t test.

(D and E) Representative images used to quantify data in (F) and (G) (scale bar: 1,000 μm). HMVEC-Ls were pre-treated for 24 h with DMSO (D) or methanol (E), 20 μM AA (D) or 40–80 μM EPA100 (E), or with a combination of 20 μM AA (D) or 40–80 μM EPA (E), and 1 μM TRPV4-specific inhibitor HC067047. For each treatment, cells either remained in static conditions or were subjected to SS (15 dyne/cm2) for at least an additional 24 h.

(F and G) Cell counts per image of DMSO (F) or methanol (G), AA (F) or EPA (G), and AA (F) or EPA (G) + HC067047 24 h pre-treated HMVEC-Ls in static or flow conditions after an additional 24 h (n > 10; data are mean ± SD). Statistical analyses: two-way ANOVA with Tukey’s multiple comparison: interaction (F) and (G) (p < 0.0001).

(H) Excimers to monomers ratio of pyrenedecanoic acid (PDA) in the membrane of negative-control-siRNA- and CFTR-siRNA-treated HMVEC-Ls. The greater the ratio of excimers to monomers, the more fluid the membrane.

(I) Representative immunofluorescence images showing membrane expression of VE-cad (red) and AA-alkyne or EPA-alkyne clicked with azide-488 (green) (scale bar: 100 μm). HMVEC-Ls were treated with DMSO, 20 μM AA-alkyne, or 20 μM EPA-alkyne for 48 h. ∗p ≤ 0.05; ∗∗p ≤ 0.01; ∗∗∗p ≤ 0.001; ∗∗∗∗p ≤ 0.0001.

This increase in AA and EPA when CFTR is defective is critical because AA, EPA, and their respective metabolites, 5′6′-EET and 17′18′-EEQ, have been shown to dramatically lower the activation threshold of TRPV4 or directly bind to and activate it.64,101,102,103 To confirm that AA and EPA can directly upregulate the activity of TRPV4 in HMVECs-L, we supplemented cell culture medium with AA or EPA while inhibiting TRPV4 with HC067047. We found that under static conditions, supplementing AA or EPA had no adverse effects on cell viability compared to control (Figures 4D–4G). However, under SS, there was a drastic reduction in cell viability compared to control when AA and EPA were supplemented, much like what occurs in CFTR deficiency. These negative effects were also prevented by TRPV4 inhibition (Figures 4D–4G). Overall, results indicate that AA and EPA significantly lower the mechanical stress activation threshold of TRPV4.

Changes in PUFAs levels can impact the architecture of the membrane. In fact, PUFAs have been reported to increase membrane structural disorder and fluidity.104,105 Because MsCs are gated by bilayer tension, increased fluidity results in enhanced TRPV4 activation.63,64,65 We performed fluorescence anisotropy and found a significant increase in EC membrane fluidity when CFTR was knocked down compared to control (Figure 4H).

To confirm that AA and EPA were indeed incorporated into the cell membrane where they could enhance TRPV4 activity, we performed click-chemistry-based immunofluorescence. We supplemented the culture medium of HMVEC-Ls with DMSO, AA-alkyne, or EPA-alkyne. We then clicked the cells with azide-488 while also costaining for VE-cad as a membrane indicator. We found that ECs were in fact incorporating supplemented AA and EPA into the membrane (Figure 4I). These results may be clinically relevant since AA is reportedly increased in the blood of CF patients.94 CF ECs could be not only metabolizing more AA and EPA by themselves (Figures 4B and 4C) but also incorporating AA and EPA from the bloodstream directly into membrane-bound phospholipids; this could lead to further elevation of intracellular AA and EPA levels and potentially drive overactivation of TRPV4.

CFTR expression enhances PIEZO1 function

Ideally, loss of SPBVs should be mitigated by angiogenesis and wound repair. However, in CF pulmonary disease, TBV is redistributed to larger blood vessels, which suggests lack of angiogenesis (Figure 1A). Angiogenic sprouting is regulated by binding of VEGF to VEGFR2.77 Impaired angiogenesis in CF could be due to the significant reduction in VEGFR2 expression (Figures 3I and S6), despite reports of elevated serum VEFG levels in CF patients.78 However, PIEZO1 also regulates EC pathfinding in sprouting vessels during angiogenesis.106 In fact, PIEZO1 has been shown to be critical for ECs SS sensing, migration, flow alignment, vasculogenesis, angiogenesis, wound repair, and apoptosis25,26,27,106—all of these functions are important in maintaining vascular integrity. We therefore hypothesized that CFTR deficiency might negatively regulate the activity of PIEZO1.

Whole-cell patch-clamping revealed that PIEZO1 activity was attenuated upon activation by specific agonist Yoda1 in CFTR-KD HMVEC-Ls when compared to control (Figures 5A–5C). To understand if PIEZO1 function could be dependent on CFTR expression, we transfected PIEZO1 by itself or cotransfected it with CFTR in HEK293 cells. We found that CFTR expression greatly enhanced PIEZO1 function (Figures 5D–5F). Because PIEZO1 is a cationic channel, we questioned whether the enhancing effect may occur because CFTR is a chloride channel. Therefore, we cotransfected PIEZO1 with other chloride channels/transporters such as TMEM16A and SLC26A9 and found that only CFTR had an enhancing effect on PIEZO1 function (Figure 5F). We then performed calcium imaging and found that Yoda1 induced a greater intracellular calcium increase in HEK-293 cells cotransfected with PIEZO1 and CFTR compared to cells transfected with PIEZO1 alone (Figures 5G and 5H).Figure 5 CFTR directly interacts with PIEZO1 and enhances its function

(A) Representative traces of whole-cell currents of negative-control-siRNA- and CFTR-siRNA-treated HMVEC-Ls. Bath solution contained 10 μM Yoda1.

(B) Representative I–V curves from (A).

(C) Summary of Yoda1-induced current in negative-control-siRNA- and CFTR-siRNA-treated HMVEC-Ls (n > 5; data are mean ± SD). Statistical analyses: two-tailed Student’s t test.

(D) Representative traces of whole-cell currents of HEK-293 cells transfected with PIEZO1 only, or cotransfected with PIEZO1 and WT CFTR. Bath solution contained 10 μM Yoda1, 20 μM Forskolin, and 20 μM CFTRinh-172.

(E) Representative I–V curves from (D).

(F) Summary of Yoda1-induced current in HEK-293 cells transfected with PIEZO1 only, or cotransfected with PIEZO1 and TMEM16A, or with PIEZO1 and SLC26A9, or with PIEZO1 and CFTR (n > 5; data are mean ± SD). Statistical analyses: one-way ANOVA with Dunnett’s multiple comparison.

(G) Time courses used to quantify data in (H). DMSO or 25 μM Yoda1-induced change in intracellular calcium levels of HEK-293 cells transfected with empty vector only, PIEZO1 only, CFTR only, or cotransfected with PIEZO1 and CFTR (n = 8 wells; data are mean ± SE).

(H and G) Change in intracellular calcium levels from (G) using maximum fluorescence intensity (n = 8 wells; data are mean ± SD). Statistical analyses: one-way ANOVA with Tukey’s multiple comparison.

(I) Representative traces of whole-cell currents of HEK-293 cells cotransfected with PIEZO1 and CFTR mutant F508del, pre-treated with combination of CFTR correctors VX445/VX661 (2 μM each) for 24 h.107 Bath solution contained 10 μM Yoda1, 20 μM Forskolin, and 20 μM CFTRinh-172.

(J) Summary of Yoda1-induced current in HEK-293 cells transfected with PIEZO1 only or cotransfected with PIEZO1 and CFTR mutant F508del. Cells were pre-treated with or without combination of CFTR correctors VX445/VX661 (2 μM each) for 24 h (n > 5; data are mean ± SD). Statistical analyses: two-way ANOVA: interaction (p < 0.05).

(K) Representative time courses used to quantify data in (L). 100 μM Yoda1-induced change in intracellular calcium levels of HEK-293 cells transfected with empty vector only, or cotransfected with PIEZO1 and WT CFTR, or with PIEZO1 and F508del CFTR (n = 4 wells; data are mean ± SE).

(L) Change in intracellular calcium levels from (K) using maximum fluorescence intensity (n = 4 wells; data are mean ± SD). Statistical analyses: one-way ANOVA with Tukey’s multiple comparison.

(M) Coimmunoprecipitation blot of HEK 293 cells transfected with Flag-WT-CFTR only or co-transfected with Flag-WT-CFTR and PIEZO1-sYFP2.

(N) Representative traces of whole-cell currents of HEK-293 cells transfected with PIEZO1 only, or co-transfected with PIEZO1 and WT CFTR, and treated with or without 20 μM CFTRinh-172 in bath solution that also contained 10 μM Yoda1.

(O) Representative I–V curves from (N).

(P) Summary of Yoda1-induced current in HEK-293 transfected with PIEZO1 only, or co-transfected with PIEZO1 and WT CFTR, and treated with or without 20 μM CFTRinh-172 in bath solution (n > 6; data are mean ± SD). Statistical analyses: one-way ANOVA with Tukey’s multiple comparison. ∗p ≤ 0.05; ∗∗p ≤ 0.01; ∗∗∗p ≤ 0.001; ∗∗∗∗p ≤ 0.0001.

Next, we coexpressed PIEZO1 with CFTR mutant F508del (folding mutation) in HEK-293 cells to investigate if the enhancement effect of CFTR was conformation dependent. On its own, expression of this variant did not enhance PIEZO1 function (Figure 5J). However, when treated with a combination of potent CFTR correctors (VX445 and VX661),107 the corrected F508del mutant was able to enhance PIEZO1 function (Figures 5I and 5J). These results suggest that expression of WT-like CFTR is required to enhance PIEZO1 function. Furthermore, calcium imaging indicated that Yoda1 induces a greater intracellular calcium increase in HEK-293 cells cotransfected with PIEZO1 and WT CFTR compared to cells transfected with PIEZO1 and F508del CFTR mutant (Figures 5K and 5L). Therefore, we inferred that CFTR could directly interact with PIEZO1. We performed coimmunoprecipitation and discovered that WT CFTR and PIEZO1 were in a complex together (Figure 5M).

Lastly, we found that the enhancement effect of WT CFTR on PIEZO1 function was attenuated when CFTR was maintained in a closed state by CFTRinh-172 (Figures 5N–5P). This finding suggests that both CFTR expression and activity are required to enhance PIEZO1 function.

We propose that CFTR, by directly interacting with PIEZO1 in a complex, stabilizes and enhances its function. This could have important ramifications because changes in the kinetics of PIEZO1 have been associated with disorders such as xerocytosis.108,109 Therefore, it is crucial to better understand CFTR’s modulatory effects on PIEZO1. The next step in doing so should be identifying which domains of CFTR and PIEZO interact and whether or not a scaffolding protein is involved.

Model of endothelial barrier failure in CF

We propose a mechanism of loss of endothelial barrier integrity in CF, which is detailed in Figure 6.Figure 6 Proposed model of loss of endothelial barrier integrity

(A–C) In the healthy endothelium, functional CFTR and PIEZO1 directly interact at the cell membrane via unknown domains.

(D–M) In the case of CF disease, mutant CFTR, such as F508del-CFTR, is degraded in the ER, which limits interaction with PIEZO1. However, higher membrane content in AA, EPA, and their respective metabolites 5′6′-EET and 17′18′-EEQ68,70,93,95,96,97,98,99 would lower the activation threshold of or even directly activate TRPV4.63,64,101,102,105 This would lead to calcium overload and to overactivation of calcium-dependent m-calpain, which would result in faster degradation of VE-cad.27,35,36,37,40

In the healthy endothelium, functional CFTR and PIEZO1 directly interact at the cell membrane via unknown domains (Figure 6A). This interaction stabilizes PIEZO1 and enhances its function. SS activates PIEZO1, which leads to a transient rise in calcium41 (Figure 6B). Transient calcium elevation activates calcium-dependent cytosolic phospholipase A2 (cPLA2)41,110 (Figure 6C), which results in downstream processes associated with PIEZO1; examples of downstream processes are EC flow alignment and angiogenesis/wound repair in response to injuries.25,26,27,106

In CF disease, mutant CFTR, such as F508del-CFTR, is retained in the ER and quickly degraded, which limits the enhancing interaction with PIEZO1 (Figure 6D). However, we suspect that cPLA2 might still be activated by some other mechanisms such as calcium release from intracellular stores,111,112,113 independently of PIEZO1 (Figure 6E). In fact, it has been reported that AA-induced lipid peroxidation can lead to increased release of calcium from intracellular stores and increased PLA2 activation; this does not depend on increased influx from extracellular calcium113 (e.g., through PIEZO1). Our data report that AA is increased in CFTR-deficient cells (Figure 4B). Furthermore, CF is associated with ER-stress-induced calcium release due to the unfolded protein response (UPR).112 Altogether, these could explain how cPLA2 gets activated independently of PIEZO1.

Activated cPLA2 would preferably hydrolyze AA-containing membrane-bound phospholipids and free AA110,114,115 (Figure 6F). EPA, on the other hand, would preferably be freed by calcium-independent iPLA2100 (Figure 6G). In CF disease, the membrane is richer in AA and EPA compared to a healthy state,68,70,93,95,96,97,98,99 which likely results in increased free AA and EPA. Free AA and EPA would then be converted into their respective metabolites 5′6′-EET and 17′18′-EEQ (Figure 6H) and incorporated back into the membrane (Figure 6I), where they would lower the activation threshold of or even directly activate TRPV4.63,64,101,102,105

Consequently, for a given SS stimulus, an increased number of TRPV4 channels would get activated (Figure 6J). Sustained increase in intracellular calcium likely ensues,41 eventually resulting in calcium overload (Figure 6K). Higher intracellular calcium levels lead to overactivation of calcium-dependent m-calpain (Figure 6L), which results in faster degradation of VE-cad and reduced expression of AJs27,35,36,37,40 (Figure 6M). Loss of focal adhesion is also likely to play a role, and its involvement in the proposed mechanism requires further study.

Model of small pulmonary blood vessel loss in CF

We also propose an integrated mechanism of loss of SPBVs in CF (Figure 7) based on our findings and available literature: (1) altered gene expression, (2) loss of CFTR-PIEZO1 interaction, and (3) altered lipid metabolism leading to overactivation of TRPV4 are the key contributors to loss of barrier integrity. Eventually, barrier failure, coupled with impaired angiogenesis, is likely to be the main driving force behind the loss of SPBVs. This proposed mechanism may be the basis for the extensive vascular remodeling observed in severe CF lung disease (Figure 1A).Figure 7 Proposed model of small pulmonary blood vessel loss in CF

Loss of functional CFTR in the lung vasculature leads to altered gene expression, loss of CFTR-PIEZO1 interaction, and altered lipid metabolism that leads to the overactivation of TRPV4. Consequently, endothelial barrier integrity is compromised, and angiogenesis is impaired; as a result, loss of small pulmonary blood vessels occurs in CF.

Discussion

We have established a mechanism underlying loss of SPBVs in CF by demonstrating that CFTR directly interacts with MsC PIEZO1. Furthermore, by altering membrane lipid composition, CFTR deficiency can indirectly lead to overactivation of another MsC, namely TRPV4, which results in loss of cell-cell contact. This study paves the way for the identification of other MsCs that could be modulated by CFTR through similar mechanisms. Moreover, assessing lipid imbalances could provide insight into the degree of endothelial barrier perturbation in CF; in CF-related lung diseases such as pneumonia, interstitial lung disease, and obstructive pulmonary disease19,20,21; and in cardiac conditions. This study may inform the development of new therapeutics targeted at the endothelium in CF, CF-related, and cardiovascular diseases.

Limitations of the study

By design, this was primarily a human study that used both human primary cells and a vessel-on-a-chip model. Despite some possibilities of allometric scaling between humans and mice, SS is much higher in mice compared to humans, and it is possible that scaling error may occur.116,117,118 On the other hand, our approach limited confounding variables that are typically problematic with in vivo studies, and most importantly, allowed us to impose static conditions. Without the ability to ensure static conditions, we would not have been able to identify the SS-dependent endothelial barrier failure associated with CFTR deficiency. Besides, the multi-channel microfluidic devices allowed us to run control and test samples concurrently in a high-throughput manner; this would not have been feasible in vivo. Moreover, our findings regarding the important role of TRPV4 in CF endothelial barrier failure are likely to translate in vivo; in fact, a very recent in vivo/ex vivo study using the lungs of rats and mice reported that TRPV4 inhibition, as well as CFTR potentiation, helped prevent loss of endothelial barrier integrity in pneumonia, where CFTR expression is reduced.60

STAR★Methods

Key resources table

REAGENT or RESOURCE	SOURCE	IDENTIFIER	
Antibodies	
	
Rabbit anti VE-Cadherin antibody	Abcam	Cat#ab33168; RRID:AB_870662	
Alexa Fluor 488 Goat anti-Rabbit IgG antibody	Invitrogen	Cat#A-11008; RRID:AB_143165	
Rabbit anti-TRPV4 (extracellular) antibody	Alomone Labs	Cat#ACC-124; RRID:AB_2340915	
Alexa Fluor 568 Goat anti-Rabbit IgG antibody	Abcam	Cat#ab175471; RRID:AB_2576207	
GFP-NanoAntibody	Allele Biotechnology	Cat#ABP-nAb-GFPA	
Monoclonal ANTI-FLAG M2 antibody	Sigma	Cat#F3165; RRID:AB_259529	
Monoclonal Anti-Green Fluorescent Protein (GFP) antibody	Sigma	Cat#G1546; RRID:AB_1079024	
	
Chemicals, peptides, and recombinant proteins	
	
CFTRinh-172	Sigma	Cat#C2992	
Calpastatin	Sigma	Cat#208902	
S1PL-IN-31	Aobious	Cat#AOB31664	
HC067047	Tocris	Cat#4100	
GSK1016790A	Tocris	Cat#6433	
7-Amino-4-Chloromethylcoumarin, t-BOC-L-Leucyl-L-Methionine amide (CMAC, t-BOC-Leu-Met)	ThermoFisher	Cat#A6520	
GsMTx-4	Alomone Labs	Cat#STG-100	
Calcium Ionophore A23187	Sigma	Cat#C7522	
Yoda1	Tocris	Cat#5586	
Chelex-100	Sigma	Cat#C7901-25G	
Tezacaftor (VX661)	MedChemExpress	Cat#HY-15448	
Elexacaftor (VX445)	MedChemExpress	Cat#HY-111772	
Rhodamine phalloidin	Cytoskeleton	Cat#PHDR1	
Arachidonic acid	Sigma	Cat#A3611-10MG	
Eicosapentaenoic acid	Sigma	Cat#E2011-10MG	
Arachidonic acid alkyne	Cayman Chemicals	Cat#10538	
Eicosapentaenoic acid alkyne	Cayman Chemicals	Cat#16704	
Azide-fluor 488	Sigma	Cat#760765-1MG	
WT-CFTR	Yarlagadda et al.119	N/A	
ΔF508 CFTR	Yarlagadda et al.119	N/A	
Flag-WT-CFTR	Yarlagadda et al.119	N/A	
mPiezo1-IRES-eGFP	Coste et al.120	Addgene Cat#80925; RRID:Addgene_80925	
PIEZO1-sYFP2	Chuntharpursat-Bon et al.121	N/A	
TMEM16A	Karl Kunzelmann Lab	N/A	
SLC26A9	RESOLUTE Consortium & Giulio Superti-Furga Lab	Addgene Cat#132140; RRID:Addgene_132140	
pcDNA3.1	Invitrogen	Cat#V79020	
	
Critical commercial assays	
	
Lipofectamine RNAiMax	ThermoFisher	Cat#13778150	
Lipofectamine 3000	ThermoFisher	Cat#L3000015	
μ-Slide VI 0.4 Ibitreat surface modification	Ibidi	Cat#80606	
Fluo-8	Abcam	Cat#ab112129	
Click-&-Go Cell Reaction Buffer Kit	Click Chemistry Tools	Cat#1263	
Ambion miRNA Isolation Kit	Invitrogen	Cat#AM1561	
Apoptosis/Necrosis Assay Kit (blue, green, red)	Abcam	Cat#ab176749	
Membrane Fluidity Kit	Abcam	Cat#ab189819	
	
Deposited data	
	
RNA-sequencing data	Gene Expression Omnibus database	Accession numbers: GSE244214	
	
Experimental models: Cell lines	
	
Primary Human Lung Microvascular Endothelial Cells	Lonza	Cat#CC-2527	
HEK-293	ATCC	Cat# CRL-1573	
	
Recombinant DNA	
	
Silencer Select Human CFTR siRNA	ThermoFisher	Cat#4392420/Assay ID:s534180	
Silencer Select Negative Control siRNA	ThermoFisher	Cat#4390843	
	
Software and algorithms	
	
Fiji (ImageJ)	Schindelin et al.122	N/A	
LC Pro plugin	Michael Francis	https://imagej.nih.gov/ij/plugins/lc-pro/index.html	
Database for Annotation, Visualization and Integrated Discovery (DAVID)	Laboratory of Human Retrovirology and Immunoinformatics (LHRI)	https://david.ncifcrf.gov/tools.jsp	
GraphPad Prism 9	Graphpad software, Inc	N/A	
Adobe Illustrator	Adobe	N/A	
Excel	Microsoft	N/A	
PowerPoint	Microsoft	N/A	
BioRender	BioRender	N/A	
R Studio	Posit	N/A	
BaseSpace	Illumina	N/A	
QIAGEN Ingenuity Pathway Analysis	Qiagen	N/A	

Resource availability

Lead contact

Further information and requests for resources and reagents should be directed to the lead contact, AP Naren (APNaren@cshs.org).

Materials availability

• This study did not generate new unique reagents.

Data and code availability

• Bulk RNA sequencing data have been deposited at NCBI’s GEO Database and are publicly available as of the date of publication. Accession numbers are listed in the key resources table.

• This paper does not report original code.

• Any additional information required to reanalyze the data reported in this paper is available from the lead contact upon request.

Experimental model and study participant details

Primary cells

HMVEC-L

HMVEC-L from male and female donors (Lonza, Cat#CC-2527) were cultured in EGM-2 Bulletkit Culture Medium with supplements (Lonza, Cat#CC-3162) at 37°C and 5% CO2. Culture media bottle was supplemented with 1% penicillin-streptomycin (Gibco, Cat#15-140-163) and 1 mL plasmocin prophylactic (InvivoGen, Cat#ant-mpp). Cells were authenticated by quality control testing and tested for mycoplasma contamination by manufacturer.

Cell line

HEK-293 cells

HEK-293 (ATCC, Cat#CRL-1573) cells were maintained in DMEM/F-12 culture medium (Invitrogen, Cat#11330) supplemented with 5% FBS (Sigma, Cat#12103C), 1% penicillin-streptomycin (Gibco, #15-140-163) and 1 mL plasmocin prophylactic (InvivoGen, Cat#ant-mpp) at 37°C and 5% CO2. Cells were authenticated by short tandem repeat (STR) profiling and tested for mycoplasma contamination by manufacturer.

Human subjects

CF patients

In a pilot study approved by and conducted at Cincinnati Children’s Hospital Medical Center, 40 CF patients (age 14.47 ± 4.16 years, 53% female, 100% white) with a range of FEV1% predicted between 27%-121% had CT scans of the lungs at a baseline clinical condition without symptoms of pulmonary exacerbation. Subjects were categorized based on the following FEV1% predicted criteria. Normal = FEV1% ≥ 100 (n = 10), mild lung disease = FEV1% =80-99 (n = 9), Moderate lung disease = FEV1% 60-79 (n = 10) and severe = FEV1% = < 60 (n = 11). There was a significant sociation between the decline in the blood volume in small blood vessels expressed as percent of total lung blood volume after adjustment for age, body mass index (BMI) and gender. Neither BMI nor gender were significant variables in the model. Informed consent was obtained from all subjects.

Method details

Computerized Tomography

CT exams were performed on a Canon Aquilion ONE (Canon Medical Systems Corporation, Tochigi, Japan) 320-detector CT scanner utilizing our standard high resolution CT protocol. All exams were performed without sedation or the use of contrast. Exams were acquired in volumetric mode with low dose technique utilizing 100 kV (120 kv for > 70 kg pt), 11-170 mAs. Volumetric acquisition was obtained at full inspiration and expiration, approximating total lung capacity and functional residual capacity as close as possible. Images were reconstructed in the axial plane using the “body standard volume” reconstruction kernel with lung window settings, slice thickness of 0.5 mm, and slice increment of 0.3 mm.

Pulmonary vascular morphometry

Vascular morphometry was extracted from volumetric computerized tomography (CT) scans of the chest at maximum inspiration. Feature detection algorithms such as scale-space particles methodology were used to detect vessels, which appear as intensity ridge lines on CTs and mathematically defined by the Hessian (second order derivatives) eigenvectors.123,124 Three-dimensional physical space of the volumetric CT domain was expanded to a fourth dimension of scale for computational purposes. This four-dimensional computational domain was initialized with particles along three spatial axes and scale axis. The particles were iteratively moved along the intensity ridge lines until the system energy was minimized. This approach was applied separately for each lung independently and after resampling CT data with isotropic voxel size of 0.5mm. Vessel centerlines were identified by the strength of the intensity ridge lines in the CTs. Vascular radii were analytically estimated based on the value of the scale in the four-dimensional domain at each point along vessel center lines. Total blood vessel volume (TBV) and small blood vessel volume (BV5),125 defined as the volume of all blood vessels with cross-sectional area less than 5 mm2 were calculated for each of the patients and BV5/TBV ratio were compared across patients and controls.

Cell transfection

HMVEC-L siRNA

100,000 HMVEC-Ls per well between passage 3 and 6 were seeded on a 6-well plate (Corning, 3516) and grown for 4 days (<80% confluency). Silencer Select Human CFTR siRNA (ThermoFisher, Cat#4392420/s534180) or Silencer Select Negative Control siRNA (ThermoFisher, Cat#4390843) were transfected (40 nM siRNA final concentration) into cells using Lipofectamine RNAiMax (ThermoFischer, Cat#13778150) in EBM-2 basal medium (Lonza, Cat#CC-3156) containing no supplements according to manufacturer’s protocol for 5h.63 Transfection medium was then replaced with supplemented EGM-2 culture medium (Lonza, Cat#CC-3162). Cells were used 48h post-transfection.

HEK-293 overexpression system

HEK-293 cells were grown to 60-80% confluency in 6-well plate (Corning, Cat#3516). WT-CFTR,119 F508del CFTR,119 Flag-WT-CFTR,119 mPIEZO1 (Addgene, Cat#80925), PIEZO1-sYFP2,121 TMEM16A (kindly provided by Dr. Karl Kunzelmann, University of Regensburg, Regensburg, Germany), SLC26A9 (Addgene, Cat#132140), GFP (Addgene, Cat#74165) or pcDNA3.1 empty vector (Invitrogen, Cat#V79020) were transfected (0.5 μM for GFP, 1 μM for all other constructs) into cells using Lipofectamine 3000 (ThermoFisher, Cat#L3000015) in 3 mL Opti-MEM (Gibco, Cat#31985088) for 24h. Transfection medium was then replaced with DMEM/F-12 culture medium (Invitrogen, Cat#11330). Cells were used within 48h post-transfection.

Microfluidics studies

30 μL of culture medium containing 18,000-25,000 HMVEC-Ls between passage 3-7 were added per channel of μ-Slide VI (Ibidi, 0.4 Ibitreat surface modification, Cat#80606) and incubated at 37°C for 10 min to allow cells to attach. 60 μL of medium were then added to inlet and outlet wells. Slide was returned to incubator overnight for cells to spread and proliferate.

The following day, 10 mL of medium containing test compound(s) such as CFTRinh-172 (Sigma, Cat#C2992), Calpastatin (Sigma, Cat#208902), S1PL-IN-31 (Aobious, Cat#AOB31664), HC067047 (Tocris, Cat#4100), GsMTx-4 (Alomone Labs, Cat#STG-100), Arachidonic acid (Sigma, Cat#A3611-10MG), Eicosapentaenoic acid (Sigma, Cat#E2011-10MG), DMSO or methanol was prepared in 15 mL tubes. Medium containing appropriate test compounds was added to channels so that for each test compound, there was one flow channel and one static channel for control. Inlet and outlet wells of flow channels were filled to their maximum volume to prevent air gaps once connected to tubing.

Flow channels were connected to digital peristaltic pump (VWR, Cat#MFLX78001-72) inside incubator (37°C and 5% CO2) using 2.06 mm 3-stop pump tubing (VWR, Cat#96461-42) and tube adapter set (Ibidi, Cat#10831). To set up on automated microscope (Lionheart FX, Agilent BioTek) for live recording, PVC-tubing (ThermoFisher, Cat#14-387-345) was used for extension. Tubing was completely primed with medium as to remove any air and to prevent air bubble formation. Incubator and gas-injector on automated microscope were used to maintain standard cell culture conditions (37°C and 5% CO2). Static channels were covered to minimize medium evaporation.

Peristaltic pump was set to 11.83 mL/min for each channel, which corresponds to a shear stress of 15 dynes/cm2 inside the channel of the μ-Slide VI 0.4, according to manufacturer (Ibidi). Flow was run for 24-48h. Phase-contrast images were obtained on automated microscope (Lionheart FX, Agilent BioTek). Data analysis (cell count) was performed using Fiji (ImageJ).

Calcium removal and supplementation

5 g/100 mL of Chelex-100 (Sigma, Cat#C7901-25G) was added to EGM-2 medium and gently stirred for 1h at room temperature (RT). Medium was then filtered with a vacuum filter system (Corning, Cat#CLS430758). Chelex-100-treated medium was supplemented with CaCl2 (Sigma, Cat#C1016).

Calcium imaging

Automated microscope

siRNA transfected HMVEC-Ls were seeded on μ-Slide VI 0.4 as described above. The following day, cells were loaded with Fluo-8 (Abcam, Cat#ab112129) by adapting the manufacturer protocol to the microfluidic device. Briefly, loading dye solution was made by mixing (9000 μL of HHBS∗X/96) + (1000 μL of Pluronic F-127∗X/96) + (20 μL of Fluo-8 dye∗X/96) where X is equal to the number of channels to be loaded with Fluo-8. Then, loading dye was mixed in a 1:1 ratio with HHBS. The loading mixture was added to the cells inside the microfluidic device so as to completely replace the culture medium. The cells were incubated at 37°C for 30 min in dark, then for 1h at room temperature (RT) in dark. The loading mixture was then completely exchanged with medium, and the slide mounted on automated microscope as described above. The automated microscope was set to read GFP for a total recording time of 10 min at a rate of 1 image per second. Cells were subjected to only 1 min of flow. Data analysis was performed using LC Pro plugin in Fiji (ImageJ) with a p value of 0.05.

Microplate reader

Transfected HEK-293 cells were seeded on poly-L-lysine (Sigma, Cat#P4707) coated 96-well optical black plate (ThermoFisher, #165305) at 80,000 cells per well in 100 μL DMEM/F-12 medium containing no FBS and no P/S overnight. The following day, cells were loaded with Fluo-8 (Abcam, Cat#ab112129) according to manufacturer protocol. Briefly, loading dye solution was made by mixing (9000 μL of HHBS∗X/96) + (1000 μL of Pluronic F-127∗X/96) + (20 μL of Fluo-8 dye∗X/96) where X is equal to the number of wells to be loaded with Fluo-8. Then, loading dye was mixed in a 1:1 ratio with DMEM/F-12 containing no FBS and no P/S. 100 μL of loading mixture was added to each well. The cells were incubated at 37°C for 30 min in dark, then for 1h at room temperature in dark (RT). No washes were done after incubation. 50 μL of 5X Yoda1 (Tocris, Cat#5586) at final concentration 25-50 μM or DMSO was added to cells and plate was immediately read at Ex/Em = 490/525 nm on a microplate reader (FlexStation 3, VWR; or Synergy Neo2, Agilent BioTek).

Calpain activity assay

siRNA transfected HMVEC-Ls were seeded on μ-Slide VI 0.4 as described above. The following day, the slide was mounted on the automated microscope as described above. 10 μM of 7-Amino-4-Chloromethylcoumarin, t-BOC-L-Leucyl-L-Methionine amide (CMAC, t-BOC-Leu-Met, ThermoFisher, Cat#A6520) was added to the 15 mL tubes to be used to apply flow, but none to the medium inside the channels of the chip. This was important as CMAC, t-BOC-Leu-Met would immediately get cleaved by calpain and give off blue fluorescence42 if added before recording. It was also crucial that after priming the tubing with the media containing CMAC, t-BOC-Leu-Met, the tubing was connected to the chip so as to not introduce any CMAC, t-BOC-Leu-Met in the channel until after recording has started. The automated microscope was set to read DAPI for a total recording time of 15 min at a rate of 1 image per second. The channels should not be displaying significant fluorescence at the start of recording. Cells were subjected to continuous flow for the duration of recording. Data analysis was performed using LC Pro plugin in Fiji (ImageJ) with a p value of 0.05.

Fluorescence Microscopy

Adherens junctions staining

HMVEC-Ls on μ-Slide VI 0.4 were washed 3 times with PBS (Invitrogen, Cat#10010) and fixed with 4% paraformaldehyde (Sigma, Cat#252549-500ML) diluted in PBS, for 10 min at RT. Samples were then washed again 3 times with PBS and permeabilized for 30 min at RT using 10X Permeabilization Buffer (Invitrogen, Cat#00-8333-56) diluted 1:10 in double distilled water. Samples were blocked in 2.5% normal goat serum (Vector Laboratories, Cat#S-1012-50) for 1h at RT and then incubated with primary Rabbit anti VE-Cadherin antibody (Abcam, Cat#ab33168) diluted 1:1000 in antibody diluent buffer (Epredia, Cat#TA-125-ADQ), at 4°C overnight. The next day, samples were washed 3 times for 10 min each on side-to-side shaker with PBS + 0.05% Tween-20 (Sigma, Cat#P9416-100ML) before being incubated with Alexa Fluor 488 Goat anti-Rabbit IgG antibody (Invitrogen, Cat#A-11008) diluted 1:1000 in PBS, for 1h at RT. Samples were then washed 3 times for 10 min each on side-to-side shaker with PBS + 0.05% Tween-20 and incubated with DAPI solution (ThermoFisher, Cat#62248) diluted 1:1000 in PBS, for 5 min at RT. Samples were mounted in vectashield antifade mounting media (Vector Laboratories, Cat#H-1000) and imaging was performed using a fluorescence microscope (Olympus FV1200).

TRPV4 extracellular staining

Staining of HMVEC-Ls was performed exactly as for adherens junctions but permeabilization step was omitted and samples were incubated with primary rabbit anti-TRPV4 (extracellular) antibody (Alomone labs, Cat#ACC-124) diluted 1:50.

F-actin staining and directionality

HMVEC-Ls on μ-Slide VI 0.4 were stained using rhodamine phalloidin (Cytoskeleton, Cat#PHDR1) according to manufacturer’s instructions. Images were obtained using a fluorescence microscope (Olympus FV1200). Directionality analysis of F-actin filaments was performed using directionality tool in Fiji (ImageJ) with local gradient orientation method.

Alkyne labeling and click chemistry

This experiment was modified from a previously described method.126 60,000 HMVEC-Ls in 200 μL of cell culture medium were seeded directly on the glass of a 35 mm glass bottom dish (MatTek, Cat#P35GC-1.5-14-C) and incubated at 37°C for 15 min. 1.5 mL of culture medium was then added and the cells were returned to the incubator to spread and proliferate overnight. The next day, medium was replaced with fresh medium containing 20 μM arachidonic acid alkyne (Cayman Chemicals, Cat#10538), 20 μM eicosapentaenoic acid alkyne (Cayman Chemicals, #16704) or dimethyl sulfoxide (Sigma, Cat#D2650-100ML). The cells were then cultured at 37°C for a further 2 days. Then, cells were washed 3 times with PBS (Invitrogen, Cat#10010) and fixed with 4% paraformaldehyde (Sigma, Cat#252549-500ML) diluted in PBS, for 10 min at RT. Samples were then washed again 3 times with PBS and permeabilized for 30 min at RT using 10X Permeabilization Buffer (Invitrogen, Cat#00-8333-56) diluted 1:10 in double distilled water. Samples were washed 3 times with PBS and clicked using Click-&-Go Cell Reaction Buffer Kit (Click Chemistry Tools, Cat#1263) according to manufacturer’s instructions with 5 μM azide-fluor 488 (Sigma, Cat#760765-1MG) for 30 min at RT in dark on orbital shaker. Samples were then washed 5 times with PBS and blocked in 2.5% normal goat serum (Vector Laboratories, Cat#S-1012-50) for 1h at RT. Samples were then incubated with primary Rabbit Anti VE-Cadherin antibody (Abcam, Cat#ab33168) diluted 1:1000 in antibody diluent buffer (Epredia, Cat#TA-125-ADQ), at 4°C overnight in dark. The next day, samples were washed 3 times for 10 min each on orbital shaker with PBS + 0.05% Tween-20 (Sigma, Cat#P9416-100ML) before being incubated with Alexa Fluor 568 Goat anti-Rabbit IgG antibody (Abcam, Cat#ab175471) diluted 1:1000 in PBS, for 1h at RT in dark. Samples were then washed 3 times for 10 min each on orbital shaker with PBS + 0.05% Tween-20 and incubated with DAPI solution (ThermoFisher, Cat#62248) diluted 1:1000 in PBS, for 5 min at RT. Samples were mounted in vectashield antifade mounting media (Vector Laboratories, Cat#H-1000) and imaging was performed using a fluorescence microscope (Olympus FV1200).

Apoptosis/Necrosis assay

HMVEC-Ls on μ-Slide VI 0.4 were stained with Apoptosis/Necrosis Assay Kit (Abcam, Cat#ab176749) according to manufacturer’s instructions. Images were obtained on automated microscope (Lionheart FX, Agilent BioTek).

Fluorescence anisotropy

siRNA transfected HMVEC-Ls were seeded on poly-L-lysine (Sigma, Cat#P4707) coated 96-well optical black plate (ThermoFisher, #165305) at 20,000 cells per well in 100 μL EGM-2 medium overnight. The following day, cells were loaded with 10 μM pyrenedecanoic acid (PDA) labeling solution from membrane fluidity kit (Abcam, Cat# ab189819) according to manufacturer protocol for 20 min at RT. Following incubation, labeling solution was removed, and cells were washed twice with medium. 100 μL of fresh medium was then added to each well and fluorescence was immediately read at Ex/Em of both 350/400 nm and 350/470 nm to detect emission of the monomers and excimers respectively, using a microplate reader (Synergy Neo2, Agilent BioTek).

RNA Extraction for Bulk-RNA sequencing

RNA from siRNA transfected HMVEC-Ls was isolated using Ambion miRNA Isolation Kit (Invitrogen, Cat#AM1561) according to manufacturer’s instructions. Samples were frozen and sent to the Genomics, Epigenomics and Sequencing Core at University of Cincinnati College of Medicine for analysis. General bioinformatic analysis was performed via BaseSpace SEQUENCE HUB app RNA-Seq Alignment v2.0.2 followed by RNA-Seq Differential Expression version 1.0.1. The analysis used STAR for alignment and Salmon for quantification (Transcripts Per Million, TPM), followed by DESeq2 to identify differentially expressed genes. List of significantly differentially expressed genes were submitted to the Database for Annotation, Visualization and Integrated Discovery (DAVID) to identify enriched pathways of interest.

Ingenuity pathway analysis

QIAGEN’s Ingenuity® Pathway Analysis (IPA®, QIAGEN) was applied to the DEGs (347 mapped, 142 up-regulated and 205 down-regulated) in the above-described dataset to identify enriched pathways of interest.

Lipid analysis

siRNA transfected HMVEC-Ls were detached, pelleted, and resuspended in cell freezing medium (ThermoFisher, Cat#12648010) before being immediately snapped frozen in liquid nitrogen. Samples were sent to the Eicosanoid Core Laboratory at Vanderbilt University Medical Center for analysis. AA and EPA levels were determined by liquid-chromatography tandem mass spectrometry as previously described.127,128,129,130

Patch clamping

HEK293 cells were co-transfected with plasmids encoding mPIEZO1 (Addgene, Cat#80925) and WT-CFTR119 or F508del CFTR,119 respectively, for 24 hours using Lipofectamine 3000 (ThermoFisher, Cat#L3000015). F508del CFTR119 was corrected with combination of 2 μM each of CFTR correctors VX445 (MedChemExpress, Cat#HY-111772) and VX661 (MedChemExpress, Cat#HY-15448)107 in culture medium 24h prior to experiment. SLC26A3 (Addgene, Cat#132140) or TMEM16A (kindly provided by Dr. Karl Kunzelmann, University of Regensburg, Regensburg, Germany) expression plasmids were also used for comparison with WT-CFTR. Co-transfection of GFP-expression plasmid (Addgene, Cat#74165) was performed for positive selection. Whole cell patch clamp recordings were performed using an Axopatch-200B amplifier connected to Axon DigiData 1550B (Molecular Devices, CA, USA). Patch pipettes with resistances of 3 - 6 MΩ after being filled with pipette solution were prepared using a micropipette puller (Sutter Instrument, CA, USA, Cat# P-1000). To simultaneously obtain current traces at -60mV and I/V curves of CFTR, whole-cell currents were consecutively recorded with a 1 s voltage ramp of ± 100 mV applied every 10 s: hold at Vm = - 60 mV and filtered at 1 kHz and sampled at 50 Hz. The pipette solution was composed of (in mM): 133 CsCl-, 5 ethylene glycol tetraacetic acid (EGTA), 1 CaCl2, 1 MgCl2, 10 HEPES, 0.4 GTP and 4 MgATP, titrated to pH 7.3 with CsOH. Bath solution was composed of (in mM): 127 NaCl-, 3 KCl-, 2.5 CaCl2, 1 MgCl2, 10 Glucose, 10 HEPES titrated to pH 7.3 with NaOH.120 Yoda1 (Tocris, Cat#5586) was also made as a 10 mM stock solution dissolved in DMSO and was diluted to 10 μM final concentration in the bath solution. To obtain brief and robust Yoda1-induced Piezo1 currents, we used VCS-6_Perfusion valve system (Warner Instruments, Cat#643088) where mechanical pressure can be applied using a system for carrying out large bath volumes and fast perfusion rates at the same time, which had a similar effect such like a mechanical pressure driven by a piezoelectric-driven glass probe.51

Co-immunoprecipitation

HEK 293 cells co-expressing Flag-WT-CFTR119 and PIEZO1-sYFP2121 and HEK-293 cells expressing Flag-WT-CFTR119 and pcDNA3.1 empty vector (Invitrogen, Cat#V79020) were lysed in lysis buffer (1x PBS, containing 0.2% Triton-X-100 and protease inhibitors phenylmethylsulfonyl fluoride 1 mM, pepstatin-A 1 μg/ml, leupeptin 1 μg/ml, aprotinin 1 μg/ml), and the clear supernatant was subjected to immunoprecipitation using GFP-NanoAntibody (Allele Biotechnology, Cat#ABP-nAb-GFPA). The immunoprecipitated complex was eluted with 100 mM glycine (pH 2.2) and quickly neutralized with 150 mM Tris (pH 8.8). The eluted proteins were mixed with sample buffer (5X; containing 1% β-mercaptoethanol), denatured, subjected to SDS-PAGE, transferred to PVDF membrane, and immunoblotted using anti-Flag antibody (Sigma, Cat#F3165) to detect CFTR and anti-GFP antibody to detect PIEZO1 (Sigma, Cat#G1546).

Tube formation assay

Wells of a 24-well plate (Corning, Cat#3526) were coated with 300 μL of Matrigel (Corning, Cat# 354234). The coated plate was incubated for 10 min at RT and for 30 min at 37°C. 250 μL of culture media containing 2X concentration of test compounds were added per well. HMVEC-Ls between passage 3 and 5 were then detached, pelleted, and resuspended at a concentration of 400,000 cells/mL. 250 μL of cell suspension were added to each test well. Images were obtained on automated microscope (Lionheart FX, Agilent BioTek).

Quantification and statistical analysis

Quantification for each assay is described in the method details section. Data are shown as mean ± SD and are representative of at least two independent experiments. N values depict number of technical replicates, which are described in the figure legends. All statistical analyses were performed on GraphPad Prism 9. Significance of data in flow studies was tested by two-way ANOVA if only two groups, or by two-way ANOVA with Tukey’s multiple comparison if more than two groups. Significance of data on Figure 5J was tested by two-way ANOVA. Significance of all other data was tested by student t-test or by one-way ANOVA with Tukey’s multiple comparison when comparing all groups with one another; or with Dunnett’s multiple comparison when comparing all groups to a control group. A p value <0.05 was deemed significant. Significant differences between groups were indicated as: ∗p ≤ 0.05; ∗∗p ≤ 0.01; ∗∗∗p ≤ 0.001; ∗∗∗∗p ≤ 0.0001.

Supplemental information

Document S1. Figures S1–S6

Acknowledgments

We thank Dr. Milne and the Eicosanoid Core Laboratory at Vanderbilt University for lipid analysis. We thank Dr. Zhang and the Omics core at University of Cincinnati for RNA-sequencing analysis. We thank Dr. Christine Schuler for language editing. This research was supported by the 10.13039/100000968 American Heart Association Pre-Doctoral Fellowship (J-PA, AHA award 903099 ), the 10.13039/100000002 NIH Grants (APN, NHLBI HL147351 ; NIDDK P30-DK117467 ), and the 10.13039/100000897 Cystic Fibrosis Foundation (RA, AMIN19A0 ).

Author contributions

A.P.N. and J.-P.A. designed experiments and wrote the paper. J.L. performed all patch-clamping experiments. R.A. recruited CF patients and collected computerized tomography scans. G.M. performed pulmonary vascular morphometry analysis. P.L. and J.-P.A. performed RNA-sequencing analysis. J.-P.A. performed all other experiments. K.A. performed ingenuity pathway analysis, edited the manuscript, and provided feedback. A.K. provided knowledge of lipid metabolism and suggestions regarding experimental details. All authors read and approved the final manuscript.

Declaration of interests

The authors declare no competing interests.

Supplemental information can be found online at https://doi.org/10.1016/j.isci.2024.110703.
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