1. Biochemistry and Chemical Biology
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Endothelial pannexin 1–TRPV4 channel signaling lowers pulmonary arterial pressure in mice

  1. Zdravka Daneva
  2. Matteo Ottolini
  3. Yen Lin Chen
  4. Eliska Klimentova
  5. Maniselvan Kuppusamy
  6. Soham A Shah
  7. Richard D Minshall
  8. Cheikh I Seye
  9. Victor E Laubach
  10. Brant E Isakson
  11. Swapnil K Sonkusare  Is a corresponding author
  1. Robert M. Berne Cardiovascular Research Center, University of Virginia, United States
  2. Department of Pharmacology, University of Virginia, United States
  3. Department of Biomedical Engineering, University of Virginia, United States
  4. Department of Anesthesiology, Department of Pharmacology, University of Illinois, United States
  5. Department of Biochemistry, University of Missouri-Columbia, United States
  6. Department of Surgery, University of Virginia, United States
  7. Department of Molecular Physiology and Biological Physics, University of Virginia, United States
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Cite this article as: eLife 2021;10:e67777 doi: 10.7554/eLife.67777

Abstract

Pannexin 1 (Panx1), an ATP-efflux pathway, has been linked with inflammation in pulmonary capillaries. However, the physiological roles of endothelial Panx1 in the pulmonary vasculature are unknown. Endothelial transient receptor potential vanilloid 4 (TRPV4) channels lower pulmonary artery (PA) contractility and exogenous ATP activates endothelial TRPV4 channels. We hypothesized that endothelial Panx1–ATP–TRPV4 channel signaling promotes vasodilation and lowers pulmonary arterial pressure (PAP). Endothelial, but not smooth muscle, knockout of Panx1 increased PA contractility and raised PAP in mice. Flow/shear stress increased ATP efflux through endothelial Panx1 in PAs. Panx1-effluxed extracellular ATP signaled through purinergic P2Y2 receptor (P2Y2R) to activate protein kinase Cα (PKCα), which in turn activated endothelial TRPV4 channels. Finally, caveolin-1 provided a signaling scaffold for endothelial Panx1, P2Y2R, PKCα, and TRPV4 channels in PAs, promoting their spatial proximity and enabling signaling interactions. These results indicate that endothelial Panx1–P2Y2R–TRPV4 channel signaling, facilitated by caveolin-1, reduces PA contractility and lowers PAP in mice.

Introduction

The pulmonary endothelium exerts a dilatory influence on small, resistance-sized pulmonary arteries (PAs) and thereby lowers pulmonary arterial pressure (PAP). However, endothelial signaling mechanisms that control PA contractility remain poorly understood. In this regard, pannexin 1 (Panx1), which is expressed in the pulmonary endothelium and epithelium (Navis et al., 2020), has emerged as a crucial controller of endothelial function (Begandt et al., 2017; Good et al., 2015). Panx1, the most studied member of the pannexin family, forms a hexameric transmembrane channel at the cell membrane that allows efflux of ATP from the cytosol (Bao et al., 2004; Lohman et al., 2012). Previous studies indicated that flow/shear stress increases ATP efflux through Panx1 in EC monolayers (Wang et al., 2016). Endothelial Panx1 (Panx1EC) has also been linked to inflammation in pulmonary capillaries (Sharma et al., 2018). Beyond this, however, the physiological roles of Panx1EC in the pulmonary vasculature are largely unknown.

Recent studies show that endothelial transient receptor potential vanilloid 4 (TRPV4EC) channels reduce PA contractility and lower resting PAP (Daneva et al., 2021). Ca2+ influx through TRPV4EC channels activates endothelial nitric oxide synthase (eNOS; Marziano et al., 2017) to dilate PAs. Moreover, exogenous ATP increases the activity of TRPV4EC channels in PAs (Marziano et al., 2017), although the regulation of TRPV4EC channels by endogenously released ATP is not known. We postulated that Panx1EC-effluxed ATP acts through TRPV4EC channels to reduce PA contractility and lower PAP.

Purinergic receptor signaling is an essential regulator of pulmonary vascular function (Lyubchenko et al., 2011; McMillan et al., 1999; Yamamoto et al., 2003; Konduri and Mital, 2000). Extracellular ATP (eATP) is an endogenous activator of purinergic receptor signaling. However, the purinergic receptor subtype involved in eATP-induced activation of TRPV4EC channels has not been identified (Marziano et al., 2017). The pulmonary endothelium expresses both P2Y and P2X receptor subtypes. Konduri et al. showed that eATP dilates PAs through P2Y2 receptor (P2Y2R) activation and subsequent endothelial NO release (Konduri and Mital, 2000). These findings raise the possibility that the endothelial P2Y2 receptor (P2Y2REC) may be the signaling intermediate for Panx1EC–TRPV4EC channel communication in PAs. The physiological roles of P2Y2REC in the pulmonary vasculature remain unknown, mostly due to the lack of studies in PAs from endothelium-specific P2ry2 conditional knockout mice (P2ry2 cKO in EC).

The linkage between Panx1EC-mediated eATP release and subsequent activation of P2Y2REC–TRPV4EC signaling could depend on the spatial proximity of individual elements—Panx1EC, P2Y2REC, and TRPV4EC—a functionality possibly provided by a signaling scaffold. Caveolin-1 (Cav-1) is a structural protein that interacts with and stabilizes other proteins in the pulmonary circulation (Bernatchez et al., 2005). Notably, endothelium-specific Cav1 conditional knockout (Cav1 cKO-EC) mice showed reduced TRPV4EC channel activity and elevated resting PAP (Daneva et al., 2021), supporting a crucial role for Cav-1 in TRPV4EC regulation of PAP. Although Cav-1 has also been shown to co-localize with Panx1 and P2Y2R in other cell types (Goedicke-Fritz et al., 2015; DeLalio et al., 2018; Martinez et al., 2016), its role in endothelial Panx1–P2Y2R signaling is not known.

Here, we tested the hypothesis that Panx1EC–P2Y2REC–TRPV4EC channel signaling, supported by a signaling scaffold provided by Cav-1EC, reduces PA contractility and PAP. Using inducible, EC-specific Panx1, Trpv4, P2ry2, and Cav1 cKO mice, we show that endothelial Panx1–P2Y2R–TRPV4 signaling reduces PA contractility and lowers PAP. Panx1EC-generated eATP acts via P2Y2REC stimulation to activate protein kinase Cα (PKCα) and thereby increase TRPV4EC channel activity. Flow/shear stress is the physiological activator of ATP efflux through Panx1EC in PAs. Panx1EC, P2Y2REC, PKCα, and TRPV4EC channels co-localize with Cav-1EC, ensuring spatial proximity among the individual elements and supporting signaling interactions. Overall, these findings advance our understanding of endothelial mechanisms that control PAP and suggest the possibility of targeting these mechanisms to lower PAP in pulmonary vascular disorders.

Results

Endothelial Panx1-mediated ATP release activates TRPV4EC signaling

The regulation of TRPV4EC channels by endogenously released ATP remains unknown. We postulated that ATP efflux through endothelial Panx1 promotes TRPV4EC channel activity. First, we determined the effect of eATP-hydrolyzing enzyme, apyrase (10 U/mL), on TRPV4EC channel activity in PAs from tamoxifen-inducible, EC-specific Panx1 conditional knockout (Panx1 cKO-EC) mice (Lohman et al., 2015) and tamoxifen-injected Panx1fl/fl Cre- (Panx1fl/fl) control mice (Figure 1A, Figure 1—figure supplement 1; Sharma et al., 2018). En face PAs from Panx1 cKO-EC mice displayed a lack of endothelial (CD31, green) Panx1 immunostaining (red). Localized, unitary Ca2+ influx signals through TRPV4EC channels, termed TRPV4EC sparklets (Sonkusare et al., 2012), were recorded in en face, fourth-order PAs (~50 μm) loaded with Fluo-4. Addition of apyrase reduced the activity of TRPV4EC sparklets in PAs from control mice, confirming the regulation of TRPV4EC channels by endogenous eATP (Figure 1A). However, apyrase was unable to decrease TRPV4EC sparklet activity in PAs from Panx1 cKO-EC mice, suggesting that endothelial Panx1 may be a critical source of eATP in PAs (Figure 1A).

Figure 1 with 2 supplements see all
ATP efflux through Panx1EC ATP activates TRPV4EC channels in pulmonary arteries (PAs) and lowers pulmonary arterial pressure (PAP).

(A) Left: immunofluorescence images of en face fourth-order PAs from Panx1fl/fl and Panx1 cKO-EC mice. CD31 immunofluorescence indicates ECs. Center: representative traces showing TRPV4EC sparklet activity in en face preparations of PAs from Panx1fl/fl mice in the absence or presence of apyrase (10 U/mL). Dotted lines are quantal levels. Experiments were performed in Fluo-4-loaded PAs in the presence of cyclopiazonic acid (CPA; 20 μmol/L CPA, included to eliminate Ca2+ release from intracellular stores). Right: TRPV4EC sparklet activity (NPo) per site in en face preparations of PAs from Panx1fl/fl and Panx1 cKO-EC mice in the presence or absence of apyrase (10 U/mL; n = 5; ***p<0.001 vs. Panx1fl/fl [-apyrase, 10 U/mL]; ns indicates no statistical significance; t-test). ‘N’ is the number of channels per site and ‘PO’ is the open state probability of the channel. (B), measurements of ATP (nmol/L) levels in PAs from Panx1fl/fl, Panx1 cKO-EC, Panx1 cKO-SMC, Trpv4fl/fl, and Trpv4 cKO-EC mice, and endothelium-denuded PAs from Panx1fl/fl and Panx1 cKO-SMC mice (n = 5–6; *p<0.05 vs. Panx1 cKO-EC; *p<0.05 vs. Panx1fl/fl [denuded]; ***p<0.001 vs. Panx1fl/fl; ***p<0.001 vs. Panx1 cKO-SMC; ns indicates no statistical significance; one-way ANOVA). (C) Average resting right ventricular systolic pressure (RVSP) values in Panx1fl/fl, Panx1 cKO-EC, and Panx1 cKO-SMC mice (n = 6; ***p<0.001 vs. Panx1fl/fl; ns indicates no statistical significance; one-way ANOVA). (D) Left grayscale image of a field of view in an en face preparation of Fluo-4-loaded PAs from Panx1fl/fl and Panx1 cKO-EC mice showing approximately 20 ECs. Dotted outlines indicate an EC (20 μmol/L CPA included to eliminate Ca2+ release from intracellular stores). Right: representative traces showing TRPV4EC sparklet activity in en face preparations of PAs from Panx1fl/fl and Panx1 cKO-EC mice in response to GSK1016790A (GSK101; 1 nmol/L). Experiments were performed in Fluo-4-loaded PAs in the presence of CPA (20 μmol/L). (E) TRPV4EC sparklet activity (NPO) per site and sites per cell in en face preparations of PAs from Panx1fl/fl and Panx1 cKO-EC mice under baseline conditions (i.e., 20 μmol/L CPA) and in response to 1 nmol/L GSK101 (n = 6; *p<0.05, **p<0.01 vs. Panx1fl/fl; *p<0.05 vs. Panx1fl/fl; ns indicates no statistical significance; two-way ANOVA). (F) Left: representative GSK101 (10 nmol/L)-induced outward TRPV4EC currents in freshly isolated ECs from Panx1fl/fl and Panx1 cKO-EC mice and effect of GSK2193874 (GSK219, TRPV4 inhibitor, 100 nmol/L) in the presence of GSK101. Currents were elicited by a 200 ms voltage step from –50 mV to +100 mV. Center: scatterplot showing outward currents at +100 mV under baseline conditions, after the addition of GSK101 (10 nmol/L), and after the addition of GSK219 (100 nmol/L; n = 5–6 cells, *p<0.05 vs. Panx1 cKO-EC [+GSK101]; **p<0.01 vs. Panx1 cKO-EC [baseline]; ***p<0.001 vs. Panx1fl/fl [+baseline]; vs. Panx1fl/fl [+GSK101]; and Panx1 cKO-EC [+GSK101] vs. Panx1fl/fl [+GSK101]; two-way ANOVA). Right: scatterplot showing GSK219-sensitive TRPV4EC currents in response to GSK101 (100 nmol/L; ns indicates no statistical significance; n = 5).

Bioluminescence measurements confirmed lower baseline eATP levels in PAs from Panx1 cKO-EC mice than PAs from Panx1fl/fl control mice (Figure 1B), supporting an essential role for Panx1EC as an eATP-release mechanism in PAs. PAs from Trpv4 cKO-EC mice, however, exhibited unaltered basal eATP levels, suggesting that TRPV4EC channels do not regulate Panx1EC activity under basal conditions. Although eATP levels were also reduced in PAs from inducible, smooth muscle cell-specific Panx1 cKO (Panx1 cKO-SMC) (Good et al., 2018) mice, the eATP levels in these mice were higher than Panx1 cKO-EC mice (Figure 1B, Figure 1—figure supplement 1). Endothelial denudation also reduced eATP levels in PAs from control mice, which were reduced further in endothelium-denuded PAs from Panx1 cKO-SMC mice.

We recently demonstrated that right ventricular systolic pressure (RVSP), a commonly used in vivo indicator of PAP, was elevated in inducible EC-specific Trpv4 KO (Trpv4 cKO-EC) mice (Daneva et al., 2021). Similarly, Panx1 cKO-EC mice also showed elevated RVSP (Figure 1C). The Fulton index, a ratio of right ventricular (RV) weight to left ventricle plus septal (LV + S) weight, was not altered in Panx1 cKO-EC mice compared to control mice, suggesting a lack of right ventricular hypertrophy in these mice (Table 1). Baseline RVSP was not altered in Panx1 cKO-SMC mice (Figure 1C), indicating a lack of regulation of resting PAP by SMC Panx1. Functional cardiac MRI studies indicated no alterations in cardiac function in Panx1 cKO-EC mice compared to the control mice (Table 1), confirming that the changes in RVSP were not due to altered cardiac function.

Table 1
Fulton index and functional MRI analysis of cardiac function in Panx1fl/fl and Panx1 cKO-EC mice.
Panx1fl/flPanx1 cKO-EC
Fulton index0.23 ± 0.010.26 ± 0.03
EDV (µL)46.9 ± 2.750.9 ± 2.9
ESV (µL)14.8 ± 1.713.1 ± 1.4
EF (%)68.9 ± 2.074.3 ± 2.3
SV (µL)32.2 ± 1.337.8 ± 2.4
R-R (ms)127.1 ± 5.5130.8 ± 2.5
CO (mL/min)15.2 ± 0.617.3 ± 1.2

Baseline TRPV4EC sparklet activity and that induced by a low concentration (1 nmol/L) of the specific TRPV4 channel agonist, GSK1016790A (hereafter, GSK101), were significantly reduced in PAs from Panx1 cKO-EC mice compared to PAs from Panx1fl/fl mice (Figure 1D and E). Additionally, the number of TRPV4EC sparklet sites per cell was decreased in PAs from Panx1 cKO-EC mice (Figure 1E). At the agonist concentration that maximally activates TRPV4EC sparklets in PAs (30 nmol/L GSK101; Daneva et al., 2021), sparklet activity per site and sparklet sites per cell were not different between Panx1 cKO-EC Panx1 and control mice (Figure 1—figure supplement 2). Outward currents through TRPV4EC channels, elicited by 10 nmol/L GSK101, were also lower in Panx1 cKO-EC than Panx1fl/fl mice (Figure 1F, left and center). However, when maximally activated, TRPV4EC channel currents were not different between Panx1 cKO-EC and Panx1fl/fl mice (Figure 1F, right), suggesting that the maximum number of functional TRPV4EC channels is not altered in Panx1 cKO-EC mice.

Endothelial Panx1–TRPV4 signaling lowers pressure- and agonist-induced PA constriction

Isolated, pressurized PAs (50–100 μm, Figure 2A) from Trpv4 cKO-EC mice exhibited a greater intraluminal pressure-induced (myogenic) constriction than PAs from control mice (Figure 2B, Figure 2—figure supplement 1), providing the first evidence that TRPV4EC channels oppose myogenic constriction in PAs. This finding was further supported by a greater contractile response to the thromboxane A2 receptor agonist U46619 in PAs from Trpv4 cKO-EC mice (1–300 nmol/L; Figure 2C). PAs from Panx1 cKO-EC mice also showed a higher myogenic constriction than PAs from control mice (Figure 2D), offering the first evidence that endothelial Panx1 regulates myogenic constriction of PAs. U46619-induced constriction was also increased in PAs from Panx1 cKO-EC mice compared to PAs from control mice. Pretreatment of PAs from Panx1 cKO-EC mice with a low concentration of TRPV4 agonist (GSK101, 3 nmol/L) reduced the U46619-induced constriction to control levels, indicating that endothelial Panx1 dilates PAs through TRPV4EC channels. The presence of apyrase also increased U46619-induced constriction of PAs from control mice, confirming the dilatory effect of eATP on PAs (Figure 2—figure supplement 2). Further, exogenous ATP-induced dilation was absent in PAs from Trpv4 cKO-EC mice (Figure 2—figure supplement 3, center) but was not affected in PAs from Panx1 cKO-EC mice (Figure 2—figure supplement 3, right), supporting the concept that ATP-TRPV4EC channel signaling occurs downstream of Panx1EC. Together, these data provide the first evidence that Panx1EC–eATP–TRPV4EC channel signaling lowers PA contractility and resting PAP.

Figure 2 with 3 supplements see all
Endothelial Panx1–TRPV4 signaling lowers myogenic and agonist-induced constriction of pulmonary arteries (PAs).

(A) Top: an image showing the left lung and the order system used to isolate fourth-order PAs in this study; bottom: an image of a fourth-order PA cannulated and pressurized at 15 mm Hg. (B) Percentage myogenic constriction of PAs from Trpv4fl/fl and Trpv4 cKO-EC mice (n = 6; *p<0.05; t-test). (C) Percent constriction of PAs from Trpv4fl/fl and Trpv4 cKO-EC mice in response to thromboxane A2 receptor agonist U46619 (U466, 1–300 nmol/L; n = 5; *p<0.05 vs. Trpv4fl/fl [10 nmol/L], **p<0.01 vs. Trpv4fl/fl [30, 100, and 300 nmol/L]; ##p<0.01 vs. Trpv4fl/fl; two-way ANOVA). (D) Percentage myogenic constriction of PAs from Panx1fl/fl and Panx1 cKO-EC mice (n = 6; *p<0.05; t-test). (E) U46619 (U466, 1–300 nmol/L)-induced constriction of PAs from Panx1fl/fl, Panx1 cKO-EC, and Panx1 cKO-EC mice in the absence or presence of GSK101 (3 nmol/L) (n = 5; **p<0.01 vs. Panx1 cKO-EC, ***p<0.01 vs. Panx1fl/fl; two-way ANOVA, between groups). (F) Schematic of flow-induced ATP release from isolated and cannulated fourth-order PAs. Shear stress was calculated using the following equation: τ=4(μQ˙)/(πr3), where μ is viscosity, Q. is volumetric flow, and r is internal radius of the vessel. Outflow was collected every 10 min and ATP was measured using Luciferin-Luciferase ATP Bioluminescence Assay. (G) Release of ATP (nmol/L) from PAs of Panx1fl/fl and Panx1 cKO-EC mice in response to flow/shear stress in the presence of ARL-67156 (ARL; ecto-ATPase inhibitor; 300 μmol/L; 4, 7, and 14 dynes/cm2; n = 6; *p<0.05 vs. Panx1fl/fl [4 dynes/cm2]; **p<0.01 vs. Panx1fl/fl [7 dynes/cm2]; ###p<0.001 vs. Panx1 cKO-EC; two-way ANOVA). (H) Release of ATP (nmol/L) from PAs of Trpv4fl/fl and Trpv4 cKO-EC mice in response to flow/shear stress in the presence of ARL (300 μmol/L; 4, 7, and 14 dynes/cm2; n = 6; *p<0.05 vs. Trpv4fl/fl [4 dynes/cm2]; #p<0.05 vs. Trpv4 cKO-EC [4 dynes/cm2]; two-way ANOVA).

Figure 2—source data 1

Endothelial TRPV4 knockout increases U46619-induced constriction of PAs.

https://cdn.elifesciences.org/articles/67777/elife-67777-fig2-data1-v2.xlsx
Figure 2—source data 2

Endothelial Panx1 knockout increases U46619-induced constriction of PAs.

https://cdn.elifesciences.org/articles/67777/elife-67777-fig2-data2-v2.xlsx
Figure 2—source data 3

Shear stress increases ATP efflux through endothelial Panx1 in PAs.

https://cdn.elifesciences.org/articles/67777/elife-67777-fig2-data3-v2.xlsx
Figure 2—source data 4

Endothelial TRPV4 channel does not contribute to shear stress-induced increase in luminal ATP.

https://cdn.elifesciences.org/articles/67777/elife-67777-fig2-data4-v2.xlsx

To verify the possibility that flow/shear stress activates ATP efflux through endothelial Panx1, we measured luminal eATP levels in PAs following exposure to different intraluminal shear stress levels (4, 7, and 14 dynes/cm2; Figure 2F; Ahn et al., 2017). Increase in shear stress elevated luminal eATP levels in PAs from control mice, but not in PAs from Panx1 cKO-EC mice (Figure 2G), confirming a critical role for Panx1EC in shear stress-induced increase in luminal eATP. Also, shear stress-induced increase in luminal eATP was not altered in PAs from Trpv4 cKO-EC mice compared to control mice (Figure 2H), suggesting that TRPV4EC channels do not influence the efflux of ATP through Panx1EC in response to increase in shear stress. eATP acts through purinergic P2Y2REC stimulation to activate TRPV4EC channels.

The main P2Y receptor subtypes in the pulmonary endothelium are P2Y1R and P2Y2R (Konduri and Mital, 2000; Konduri et al., 2004; Zemskov et al., 2011). The selective P2Y1R inhibitor MRS2179 (MRS, 10 μmol/L) did not alter eATP activation of TRPV4EC sparklets (Figure 3A). In contrast, the selective P2Y2R inhibitor AR-C 118925XX (AR-C; 10 μmol/L) completely abrogated the effect of eATP on TRPV4EC sparklets (Figure 3A). eATP was also unable to activate TRPV4EC sparklets in inducible, endothelium-specific EC-specific P2ry2 cKO-EC mice (Figure 3A), providing further evidence that eATP activates TRPV4EC channels in PAs specifically via P2Y2REC signaling. The general P2X1-5 receptor inhibitor, PPADS (10 μmol/L), and P2X7 receptor inhibitor, JNJ-47965567 (JNJ, 1 μmol/L), did not alter the effect of eATP on TRPV4EC sparklets, ruling out a role for P2X1-5/7 receptors in eATP activation of TRPV4EC channels in PAs (Figure 3B). In ECs freshly isolated from PAs of C57BL6 mice, ATP (10 μmol/L) increased the outward currents through TRPV4EC channels (Figure 3C). Furthermore, the selective P2Y2R agonist, 2-thiouridine-5′-triphosphate (2-thio UTP; 0.5 μmol/L) activated TRPV4EC sparklets in PAs from P2ry2fl/fl mice but not in PAs from P2ry2 cKO-EC mice (Figure 3D).

Endothelial P2Y2R-TRPV4 channel signaling lowers pulmonary artery (PA) contractility and pulmonary arterial pressure (PAP).

(A) Left: immunofluorescence images of en face fourth-order PAs from P2ry2fl/fl and P2ry2 cKO-EC mice. CD31 immunofluorescence indicates ECs. Right: effects of ATP (1 μmol/L) on TRPV4EC sparklet activity in the absence or presence of the P2Y1R inhibitor MRS2179 (MRS; 10 μmol/L) or P2Y2R inhibitor AR-C 118925XX (AR-C; 10 μmol/L) in PAs from P2ry2fl/fl and P2ry2 cKO-EC mice, expressed as NPO per site (n = 5; ***p<0.001 vs. Control [- ATP]; **p<0.01 vs.+ MRS [- ATP]; ns indicates no statistical significance; two-way ANOVA). ‘N’ is the number of channels per site and ‘PO’ is the open state probability of the channel. (B) Effects of ATP (1 μmol/L) on TRPV4EC sparklet activity in the presence of the general P2X1-5/7R inhibitor PPADS (10 μmol/L) and P2X7R inhibitor JNJ-47965567 (JNJ; 1 μmol/L) in PAs of C57BL6/J mice (n = 5; *p<0.05 vs. [-ATP]; one-way ANOVA). (C) Top: representative ATP (10 μmol/L)-induced outward TRPV4 currents in freshly isolated ECs from C57BL6/J mice and the effect of GSK2193874 (GSK219; TRPV4 inhibitor; 100 nmol/L) in the presence of ATP. Currents were elicited by a 200 ms voltage step from –50 mV to +100 mV. Bottom: scatterplot showing outward currents at +100 mV under baseline conditions, after the addition of ATP, and after the addition of GSK219 (100 nmol/L; n = 6 cells; ***p<0.001 vs. baseline; **p<0.01 vs.+ ATP [10 μmol/L]; one-way ANOVA). (D) Left: representative traces showing TRPV4EC sparklet activity in en face preparations of PAs from P2ry2fl/fl mice. Dotted lines are quantal levels. Right: TRPV4EC sparklet activity per site (NPO) in en face preparations of PAs from P2ry2fl/fl and P2ry2 cKO-EC mice under baseline conditions (i.e., 20 μmol/L cyclopiazonic acid [CPA]) and in response to 2-thio UTP (P2Y2R agonist, 0.5 μmol/L; n = 5; *p<0.05 vs. P2ry2fl/fl [-2-thio UTP]; ns indicates no statistical significance; t-test). (E) Left: average resting right ventricular systolic pressure (RVSP) values in P2ry2fl/fl and P2ry2 cKO-EC mice (n = 6; **p<0.01; t-test). Right: average Fulton index values in P2ry2fl/fl and P2ry2 cKO-EC mice (n = 5–6; ns indicates no statistical significance). (F) Right: representative diameter traces showing ATP (1 μmol/L)-induced dilation of PAs from P2ry2fl/fl and P2ry2 cKO-EC mice, pre-constricted with the thromboxane A2 receptor agonist U46619 (U466, 50 nmol/L). Fourth-order PAs were pressurized to 15 mm Hg. Right: percent dilation of PAs from P2ry2fl/fl and P2ry2 cKO-EC mice in response to ATP (1 μmol/L; n = 5–10; ***p<0.01 vs. P2ry2fl/fl [ATP 1 μmol/L]; t-test). (G) Percentage myogenic constriction of PAs from P2ry2fl/fl and P2ry2 cKO-EC mice (n = 5–7; ***p<0.001; t-test). (H) U46619 (U466, 1–300 nmol/L)-induced constriction of PAs from P2ry2fl/fl, P2ry2 cKO-EC, and P2ry2 cKO-EC mice in the absence or presence of GSK101 (3 nmol/L) (n = 5; ***p<0.001 vs. P2ry2 cKO-EC, ***p<0.001 vs. P2ry2fl/fl; two-way ANOVA).

Figure 3—source data 1

Endothelial P2Y2R knockout increases U46619-induced constriction of PAs.

https://cdn.elifesciences.org/articles/67777/elife-67777-fig3-data1-v2.xlsx

Similar to Panx1 cKO-EC mice, P2ry2 cKO-EC mice showed elevated RVSP and unaltered Fulton index (Figure 3E). Exogenous ATP (1 μmol/L)-induced dilation was abolished in PAs from P2ry2 cKO-EC mice (Figure 3F), confirming an essential role of P2Y2REC in ATP-induced dilation of PAs. Further, PAs from P2ry2 cKO-EC mice showed higher myogenic and U46619-induced constriction compared to PAs from control mice (Figure 3G). As observed with PAs from Panx1 cKO-EC mice, pretreatment with a low concentration of TRPV4 channel agonist (GSK101, 3 nmol/L) reduced U46619-induced constriction to control levels in PAs from P2ry2 cKO-EC mice (Figure 3H). Taken together, these findings demonstrate that P2Y2REC is the signaling intermediate for Panx1EC–TRPV4EC channel interaction in PAs.

Cav-1EC provides a scaffold for Panx1EC–P2Y2REC–TRPV4EC signaling

We hypothesized that Cav-1EC provides a signaling scaffold that supports and maintains the spatial proximity among the individual elements in the Panx1EC–P2Y2REC–TRPV4EC pathway. Previous studies demonstrated that endothelium-specific knockout of Cav1 results in reduced TRPV4EC channel current density and elevated PAP (Daneva et al., 2021). Here, we provide evidence that eATP-induced activation of TRPV4EC sparklets is absent in PAs from Cav1 cKO-EC mice (Figure 4A; knockout validation in Daneva et al., 2021). As observed with PAs from Trpv4 cKO-EC and P2ry2 cKO-EC mice, eATP-induced dilation was also abolished in PAs from Cav1 cKO-EC mice (Figure 4B). These results provided the first functional evidence that Cav-1EC is required for eATP–P2Y2REC–TRPV4EC signaling in PAs. To provide additional evidence to support Cav-1EC–dependent co-localization of Panx1EC–P2Y2REC–TRPV4EC signaling elements in PAs, we performed in situ proximity ligation assays (PLAs), which allow the detection of two proteins in close proximity (<40 nm). PLA data confirmed that Cav-1EC exists within nanometer proximity of Panx1EC, P2Y2REC, and TRPV4EC channels in PAs (Figure 4C). Nanometer proximity was also observed between TRPV4EC channels and P2Y2REC and between Panx1EC and P2Y2REC (Figure 4D, Figure 4—figure supplement 1). TRPV4EC:P2Y2R and P2Y2R:Panx1 co-localization was lost in PAs from Cav1 cKO-EC mice, further supporting the crucial scaffolding role of Cav-1EC in Panx1EC–P2Y2REC–TRPV4EC pathway. PA endothelium has also been shown to express another P2Y family receptor, P2Y1 (P2Y1R) (Konduri et al., 2004). The PLA data confirmed that P2Y1R does not occur in nanometer proximity with Cav-1EC in PAs (Figure 4—figure supplement 2). Together, these data confirmed a crucial role for Cav-1EC in facilitating the spatial proximity amongst the individual elements of the Panx1EC–P2Y2REC–TRPV4EC pathway.

Figure 4 with 2 supplements see all
Cav-1EC provides a signaling scaffold for Panx1EC–P2Y2REC–TRPV4EC signaling in pulmonary arteries (PAs).

(A) Left: representative traces showing TRPV4EC sparklets in en face preparations of PAs from Cav1fl/fl and Cav1 cKO-EC mice in the absence or presence of ATP (1 μmol/L). Dotted lines are quantal levels. Right: TRPV4EC sparklet activity (NPO) per site in en face preparations of PAs from Cav1fl/fl and Cav1 cKO-EC mice in the absence or presence of 1 μmol/L ATP (n = 5; *p<0.05 vs. Cav1fl/fl [- ATP]; **p<0.01 vs. Cav1fl/fl [- ATP]; ns indicates no statistical significance; two-way ANOVA). Experiments were performed in Fluo-4-loaded fourth-order PAs in the presence of cyclopiazonic acid (CPA; 20 μmol/L), included to eliminate Ca2+ release from intracellular stores. ‘N’ is the number of channels per site and ‘PO’ is the open state probability of the channel. (B) Percentage dilation of PAs from Cav1fl/fl and Cav1 cKO-EC mice in response to ATP (1 μmol/L). PAs were pre-constricted with the thromboxane A2 receptor analog U46619 (50 nmol/L; n = 5; ***p<0.01 vs. Cav1fl/fl; t-test). (C) Top: representative merged images of proximity ligation assays (PLAs) signal, showing EC nuclei and Cav-1EC:Panx1EC, Cav-1EC:P2Y2REC, and Cav-1EC:TRPV4EC co-localization (white puncta) in fourth-order PAs from Cav1fl/fl and Cav1 cKO-EC mice. Bottom: quantification of Cav-1EC:Panx1EC, Cav-1EC:P2Y2REC, and Cav-1EC:TRPV4EC co-localization in PAs from Cav1fl/fl and Cav1 cKO-EC mice (n = 5; ***p<0.001 vs. Cav1fl/fl; t-test). (D) Representative PLA images showing EC nuclei, TRPV4EC:P2Y2REC and Panx1EC:P2Y2REC co-localization (white puncta) in fourth-order PAs from Cav1fl/fl and Cav1 cKO-EC mice. Bottom: quantification of TRPV4EC:P2Y2REC and Panx1EC:P2Y2REC co-localization in PAs from Cav1fl/fl and Cav1 cKO-EC mice (n = 5; ***p<0.001 vs. Cav1fl/fl; t-test).

Cav-1EC anchoring of PKCα mediates P2Y2REC-dependent activation of TRPV4EC channels in PAs

P2Y2R is a Gq protein-coupled receptor that activates the phospholipase C (PLC)–diacylglycerol (DAG)–PKC signaling pathway. Notably, PKC is known to phosphorylate TRPV4 channels and potentiate its activity (Fan et al., 2009). eATP, the DAG analog OAG (1 μmol/L), and the PKC activator phorbol myristate acetate (PMA; 10 nmol/L) stimulated TRPV4EC sparklet activity in small PAs (Figure 5A, B and C). Inhibition of PLC with U73122 (3 μmol/L) abolished eATP activation of TRPV4EC sparklets, but not OAG- or PMA-induced activation of TRPV4EC sparklets. Moreover, the PKCα/β inhibitor Gö-6976 (1 μmol/L) prevented activation of TRPV4EC sparklets by ATP, OAG, and PMA (Figure 5A, B and C), supporting the concept that eATP activation of P2Y2REC stimulates TRPV4EC channel activity via PLC–DAG–PKC signaling in PAs. TRPV4EC channel activation by PLC–DAG–PKC signaling was further supported by increased activity of TRPV4EC sparklets in PAs from Cdh5-optoα1 adrenergic receptor (Cdh5-optoα1AR) mouse, which expresses light-sensitive α1AR in endothelial cells (Figure 5D). When activated with light (~473 nm), Optoα1AR generates the secondary messengers IP3 and diacylglycerol (DAG) (Airan et al., 2009). Light activation resulted in increased activity of TRPV4EC sparklets (Figure 5D, Figure 5—figure supplement 1), an effect that was abolished by the PKCα/β inhibitor Gö-6976 (1 μmol/L) and in the presence of specific TRPV4 inhibitor GSK2193874 (hereafter GSK219; 100 nmol/L; Figure 5—figure supplement 2).

Figure 5 with 2 supplements see all
ATP activates TRPV4EC channels via phospholipase C–diacylglycerol–protein kinase C (PLC–DAG–PKC) signaling in pulmonary arteries (PAs).

(A) Left: representative traces showing TRPV4EC sparklet activity in en face preparations of PAs from C57BL6/J mice before and after treatment with ATP (1 μmol/L). Right: effects of U73122 (PLC inhibitor; 3 μmol/L) or Gö-6976 (PKCα/β inhibitor; 1 μmol/L) on TRPV4EC sparklet activity in en face preparations of PAs from C57BL6/J mice before and after treatment with ATP (1 μmol/L), expressed as NPO per site. Experiments were performed in Fluo-4-loaded fourth-order PAs in the presence of cyclopiazonic acid (CPA; 20 μmol/L), included to eliminate Ca2+ release from intracellular stores (n = 5; *p<0.05 vs. Control [-ATP]; ns indicates no statistical significance; one-way ANOVA). ‘N’ is the number of channels per site and ‘PO’ is the open state probability of the channel. Dotted lines indicate quantal levels. (B) Left: representative traces showing TRPV4EC sparklet activity in en face preparations of PAs from C57BL6/J mice in the absence or presence of OAG (DAG analog; 1 μmol/L). Right: effects of U73122 (3 μmol/L) or Gö-6976 (1 μmol/L) on TRPV4EC sparklet activity in en face preparations of PAs from C57BL6/J mice before and after treatment with OAG (1 μmol/L, n = 6; **p<0.01 vs. Control [-OAG]; **p<0.01 vs. U73122 [-OAG]; ns indicates no statistical significance; one-way ANOVA). (C) Left: representative traces showing TRPV4EC sparklets in en face preparations of PAs from C57BL6/J mice in the absence or presence of phorbol myristate acetate (PMA) (PKC activator; 10 nmol/L). Right: effects of U73122 (3 μmol/L) or Gö-6976 (1 μmol/L) on TRPV4EC sparklet activity in en face preparations of PAs from C57BL6/J mice before and after treatment with PMA (n = 6; *p<0.05 vs. Control [-PMA]; *p<0.05 vs. U73122 [-PMA]; ns indicates no statistical significance; one-way ANOVA). (D) Top: representative traces showing TRPV4EC sparklet activity in en face preparations of PAs from Cdh5-optoα1AR (adrenergic receptor) mouse before and after light activation (470 nm). Center: scatterplot showing TRPV4 sparklet activity before and after light activation in the absence or presence of PKCα/β inhibitor Gö-6976 (1 μmol/L, n = 4, ***p<0.01 vs. –Gö-6976 [before]; ns indicates no statistical significance; one-way ANOVA). Bottom: scatterplot showing TRPV4 sparklet activity, expressed as sparklet sites per cell, before and after light activation, in the absence or presence of PKCα/β inhibitor Gö-6976 (1 μmol/L; n = 4; ***p<0.001 vs. –Gö-6976 [before]; ns indicates no statistical significance; one-way ANOVA).

Since Cav-1 possesses a PKC-binding domain (Mineo et al., 1998) and exists in nanometer proximity with TRPV4EC channels and P2Y2REC, we tested the hypothesis that Cav-1EC anchoring of PKC mediates P2Y2REC–TRPV4EC channel interaction in PAs. PLA experiments confirmed that PKC also exists in nanometer proximity with Cav-1EC in PAs (Figure 6A). The PKC dependence of Cav-1EC activation of TRPV4EC channels was confirmed by studies in HEK293 cells transfected with TRPV4 alone or TRPV4 channels plus Cav-1 (Figure 6B), which showed that TRPV4 currents were increased in the presence of Cav-1. Further, the PKCα/β inhibitor Gö-6916 (1 μmol/L) reduced TRPV4 channel currents in Cav-1/TRPV4-co-transfected cells to the level of that in cells transfected with TRPV4 alone (Figure 6B and C). These results imply that Cav-1 enhances TRPV4 channel activity via PKCα/β anchoring. Experiments in which TRPV4 channels were co-expressed with PKCα or PKCβ showed that only PKCα increased currents through TRPV4 channels (Figure 6D). Collectively, these results support the conclusion that Panx1EC–P2Y2REC–PKCα–TRPV4EC signaling on a Cav-1EC scaffold reduces PA contractility and lowers resting PAP (Figure 6E).

Localization of PKCα with Cav-1EC increases the activity of TRPV4EC channels in pulmonary arteries (PAs).

(A) Top: representative merged images of proximity ligation assays (PLAs) showing endothelial cell (EC) nuclei and Cav-1EC:PKC co-localization (white puncta) in fourth-order PAs from Cav1fl/fl and Cav1 cKO-EC mice. Bottom: quantification of Cav-1EC:PKC co-localization in PAs from Cav1fl/fl and Cav1 cKO-EC mice (n = 5; ***p<0.001 vs. Cav1fl/fl; t-test). (B) Representative traces showing TRPV4 currents in the absence or presence of Gö-6976 (PKC inhibitor; 1 μmol/L) in HEK293 cells transfected with TRPV4 alone or co-transfected with TRPV4 plus wild-type Cav-1, recorded in the whole-cell patch-clamp configuration. (C) Current density scatterplot of TRPV4 currents at +100 mV in the absence or presence of Gö-6976 (1 μmol/L) and after the addition of GSK2193874 (GSK219; TRPV4 inhibitor; 100 nmol/L) in HEK293 cells transfected with TRPV4 alone or TRPV4 plus wild-type Cav-1 (n = 5; **p<0.01 vs. Control [TRPV4]; **p<0.01 vs. Control [TRPV4+ Cav-1]; ns indicates no statistical significance; one-way ANOVA). (D) Current density plot of TRPV4 currents at +100 mV in HEK293 cells transfected with TRPV4+ PKCα or TRPV4+ PKCβ and in the presence of GSK219 (100 nmol/L; n = 5; ***p<0.001 vs. TRPV4+ PKCα; t-test). (E) Schematic depiction of the Panx1EC–P2Y2REC–TRPV4EC signaling pathway that promotes vasodilation and lowers pulmonary arterial pressure (PAP) in PAs. ATP released from Panx1EC activates P2Y2REC purinergic receptors on the EC membrane. Stimulation of P2Y2REC recruits PKCα, which anchors to the scaffolding protein Cav-1EC in close proximity to TRPV4EC channels. TRPV4EC channel-dependent vasodilation lowers PAP.

Discussion

Regulation of PA contractility and PAP is a complex process involving multiple cell types and signaling elements. In particular, the endothelial signaling mechanisms that control resting PAP remain poorly understood. Our studies identify a Panx1EC, P2Y2REC, and TRPV4EC channel-containing signaling nanodomain that reduces PA contractility and lowers PAP. Although Panx1EC and P2Y2REC have been implicated in the regulation of endothelial function, their impact on PAP remains unknown. We demonstrate critical roles for several key, linked mechanistic, pathways showing that (1) Panx1EC increases eATP levels in small PAs; (2) Panx1EC-generated eATP, in turn, enhances Ca2+ influx through TRPV4EC channels, thereby dilating PAs and lowering PAP; (3) eATP acts through purinergic P2Y2REC–PKCα signaling to activate TRPV4EC channels; and (4) Cav-1EC provides a signaling scaffold that ensures spatial proximity among the elements of the Panx1EC–P2Y2REC–PKCα–TRPV4EC pathway. Our findings reveal a novel signaling axis that can be engaged by physiological stimuli to lower PAP and could also be therapeutically targeted in pulmonary vascular disorders. Moreover, the conclusions in this study may assist in future investigations of the mechanisms underlying pulmonary endothelial dysfunction.

Both ECs and SMCs control vascular contractility and arterial pressure. The expression of Panx1 and TRPV4 channels in both ECs and SMCs (Sharma et al., 2018; DeLalio et al., 2018; Martin et al., 2012; Ottolini et al., 2020a; Yang et al., 2006) makes it challenging to decipher the cell type-specific roles of Panx1 and TRPV4 channels using global knockouts or pharmacological strategies. Indeed, global Trpv4 knockout mice showed no systemic blood pressure or PAP phenotype (Xia et al., 2013; Zhang et al., 2009; Hong et al., 2018). However, inducible, Trpv4 cKO-EC mice had elevated systemic blood pressure and PAP (Daneva et al., 2021; Ottolini et al., 2020b). Lack of a phenotype in global knockout mice could be due to the deletion of TRPV4 channels from multiple cell types or compensatory mechanisms that have developed over time (reviewed by El-Brolosy and Stainier, 2017). Therefore, studies utilizing cell-specific knockout mice are necessary for a definitive assessment of the control of PAP by EC and SMC Panx1 and TRPV4 channels. Although SMC TRPV4 channels have been shown to contribute to hypoxia-induced pulmonary vasoconstriction, resting PAP is not altered in global Trpv4 knockout mice (Xia et al., 2013; Yang et al., 2012). Further, our studies indicate that SMC Panx1 and TRPV4 channels do not influence resting PAP. Taken together with findings from EC-knockout mice, these results provide strong evidence that endothelial, but not SMC, Panx1 and TRPV4 channels maintain low PA contractility and PAP under resting conditions. Despite the elevated PAP in EC-specific Panx1, P2ry2, and Trpv4 cKO mice (Daneva et al., 2021), right ventricular hypertrophy was not observed. These findings could be attributed to a short duration of inducible genetic deletion in our studies. Although the duration of the knockout is sufficient to result in elevated PAP, a longer duration or larger changes in PAP may be required for observing right ventricular hypertrophy in these mouse models.

Recent studies in pulmonary fibroblasts and other cell types suggest that TRPV4 channel-mediated increases in cytosolic Ca2+ can induce eATP release through Panx1 (Baxter et al., 2014; Rahman et al., 2018). However, the reverse interaction, in which Panx1-mediated eATP release activates TRPV4 channels, has not been explored in any cell type. Since Panx1 is activated by cytosolic Ca2+ (Locovei et al., 2006) and eATP has been previously shown to activate TRPV4EC channels (Marziano et al., 2017), bidirectional signaling between Panx1 and TRPV4 channels is conceivable. Our demonstration that baseline eATP levels are unchanged in PAs from Trpv4 cKO-EC mice rules out a role for TRPV4EC channels in controlling eATP release under baseline conditions. Moreover, TRPV4EC channels did not contribute to flow-induced efflux of ATP through Panx1EC. Nevertheless, these data from pulmonary ECs do not rule out potential TRPV4–Ca2+–Panx1 signaling in other cell types.

Elevated capillary TRPV4EC channel activity has been linked to increased endothelial permeability (Thorneloe et al., 2012; Yin et al., 2008), lung injury (Alvarez et al., 2006), and pulmonary edema (Thorneloe et al., 2012; Yin et al., 2008). Moreover, Panx1EC-mediated eATP release is associated with vascular inflammation at the level of capillaries (Sharma et al., 2018). The physiological roles of Panx1EC and TRPV4EC channels in PAs, however, remain unknown. ECs from pulmonary capillaries and arteries are structurally and functionally different. Whereas PAs control pulmonary vascular resistance and PAP, capillaries control vascular permeability. TRPV4EC channels couple with distinct targets in arterial and capillary ECs (Sonkusare et al., 2012; Longden et al., 2017). Our data identify physiological roles of Panx1EC–TRPV4EC channel signaling in PAs, but whether such signaling operates in the capillary endothelium and is essential for its physiological function is unclear.

Purinergic signaling and the endogenous purinergic receptor agonist eATP are essential controllers of pulmonary vascular function (Konduri and Mital, 2000; Konduri et al., 2004; Hennigs et al., 2019; Kylhammar et al., 2014). Our discovery of the Panx1EC–P2Y2REC–TRPV4EC channel pathway establishes a signaling axis in ECs that regulates pulmonary vascular function. The pulmonary vasculature is a high-flow circulation, yet the flow-induced signaling mechanisms are poorly understood in PAs. Our results confirm that flow/shear stress increases ATP efflux through Panx1EC in PAs, which could be a potential mechanism for flow-induced dilation of PAs. Further investigations are needed to verify flow/shear stress-induced, eATP-dependent activation of P2Y2REC–PKCα–TRPV4EC signaling in PAs. Several purinergic receptor subtypes are expressed in the pulmonary vasculature, including P2YRs and P2XRs (Konduri et al., 2004; Hennigs et al., 2019; Syed et al., 2010). Although only P2Y2REC appears to mediate eATP activation of TRPV4EC channels, our studies do not rule out potentially important roles for other P2Y or P2X receptors in the pulmonary endothelium.

Activation of TRPV4EC channels by eATP released through Panx1EC in PAs would be facilitated by spatial localization of TRPV4EC channels with Panx1EC. In keeping with this, several scaffolding proteins are known to promote localization of TRPV4 channels with their regulatory proteins, including A-kinase anchoring protein 150 (AKAP150) and Cav-1 (Ottolini et al., 2020b; Li et al., 2018). Although AKAP150 is not found in the pulmonary endothelium (Marziano et al., 2017), Cav-1 is a key structural protein in the pulmonary vasculature and has a well-established role in controlling TRPV4EC channel activity, pulmonary vascular function, and PAP (Daneva et al., 2021; Zhao et al., 2002; Zhao et al., 2009). Moreover, Cav-1-dependent signaling is impaired in pulmonary hypertension (Daneva et al., 2021; Bakhshi et al., 2013; Maniatis et al., 2008; Nickel et al., 2015). Studies in other cell types have shown that Cav-1 can co-localize with Panx1 and P2Y2Rs (DeLalio et al., 2018; Martinez et al., 2016). Additionally, Cav-1 can interact with PKC at the Cav-1 scaffolding domain (Mineo et al., 1998). Our results demonstrate that Cav-1EC exists in nanometer proximity with Panx1EC, P2Y2REC, PKC, and TRPV4EC channels in PAs. Furthermore, the activation of TRPV4EC channels by Panx1EC, eATP, P2Y2REC, or PKCα requires Cav-1EC. Based on these findings, we conclude that Cav-1EC enables Panx1EC–P2Y2REC–TRPV4EC signaling at EC membranes in PAs. Cav-1 is also a well-known anchor protein for eNOS (Bernatchez et al., 2005), acting by stabilizing eNOS expression and negatively regulating its activity (Bernatchez et al., 2005). We previously showed that TRPV4EC Ca2+ sparklets activate eNOS in PAs (Marziano et al., 2017; Ottolini et al., 2020a). Specifically, TRPV4 channel activation increased endothelial NO levels, an effect that was absent in PAs from eNOS knockout mice (Marziano et al., 2017). Moreover, TRPV4 channel-induced vasodilation was abolished by NOS inhibitor L-NNA. Thus, Cav-1EC enhancement of Ca2+ influx through TRPV4EC channels may represent novel mechanisms for regulating eNOS activity.

Cav-1EC/PKCα-dependent signaling is a novel endogenous mechanism for activating arterial TRPV4EC channels and lowering PAP. Proximity to PKCα appears to be crucial for the normal function of TRPV4 channels. Evidence from the systemic circulation suggests that co-localization of TRPV4 channels with scaffolding proteins enhances their activity (Mercado et al., 2014; Sonkusare et al., 2014), and we specifically demonstrated that PKC anchoring by AKAP150 enhances the activity of TRPV4EC channels in mesenteric arteries (Ottolini et al., 2020b). Here, we show that PKC anchoring by Cav-1EC enables PKC activation of TRPV4EC channels in PAs. This discovery raises the possibility that disruption of PKC anchoring by Cav-1EC could impair the Panx1EC–P2Y2REC–TRPV4EC signaling axis under disease conditions. A lack of PKC anchoring by scaffolding proteins in systemic arteries has been demonstrated in obesity and hypertension (Ottolini et al., 2020b; Sonkusare et al., 2014). Further studies of pulmonary vascular disorders are required to establish whether the Panx1EC–P2Y2REC–PKCα–TRPV4EC signaling axis is impaired in pulmonary vascular disorders.

In conclusion, Panx1EC–P2Y2REC–TRPV4EC channel signaling reduces PA contractility and maintains a low resting PAP. This mechanism is facilitated by eATP released through Panx1EC and subsequent activation of P2Y2REC–PKCα signaling. Cav-1EC ensures the spatial proximity among Panx1EC, P2Y2REC, and TRPV4EC channels and also anchors PKCαclose to TRPV4EC channels. These findings identify a novel endothelial Ca2+ signaling mechanism that reduces PA contractility. Further investigations are needed to determine whether impairment of this pathway contributes to elevated PAP in pulmonary vascular disorders and whether this pathway can be targeted for therapeutic benefit.

Materials and methods

Key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Genetic reagent (Mus musculus)C57BL/6JThe Jackson LaboratoryStock no: 000664
Genetic reagent (M. musculus)Trpv4conditional knockout in ECDr. Swapnil SonkusarePMID:32008372
Genetic reagent (M. musculus)Trpv4 conditional knockout in SMCDr. Swapnil SonkusarePMID:33879616
Genetic reagent (M. musculus)Panx1 conditional knockout in ECDr. Brant IsaksonPMID:26242575
Genetic reagent (M. musculus)Panx1 conditional knockout in SMCDr. Brant IsaksonPMID:25690012
Genetic reagent (M. musculus)Cav1 conditional knockout in ECDr. Swapnil SonkusarePMID:33879616Dr. Richard MinshallPMID:22323292
Genetic reagent (M. musculus)P2ry2fl/fl miceDr. Cheikh SeyePMID:27856454
Genetic reagent (M. musculus)Cdh5-Optoα1AR-
IRES-lacZ
CHROMus
(Cornell University, USA)
AntibodyTRPV4 antibody (aa100-150), (mouse polyclonal)LifeSpan Bioscience IncCat. #: LS-C94498;RRID:AB_2893149(1:200)
AntibodyAnti-caveolin-1 antibody - caveolae marker (rabbit polyclonal)Abcam plcCat. #: Ab2910;RRID:AB_303405(1:500)
AntibodyCaveolin-1 antibody (7C8) (mouse monoclonal)Novus Biologicals, LLCCat. #: NB100-615;RRID:AB_10003431(1:200)
AntibodyPKC (mouse monoclonal)Santa Cruz
Biotechnology, Inc
Cat. #: SC-17769;RRID:AB_628139(1:250)
AntibodyPanx1 (rabbit polyclonal)Alomone LabsCat. #: ACC-234;RRID:AB_2340917(1:100)
AntibodyP2Y2R (rabbit polyclonal)Alomone LabsCat. #: APR-010;RRID:AB_2040078(1:250)
AntibodyP2Y1R (rabbit polyclonal)Alomone LabsCat. #: APR-009;RRID:AB_2040070(1:100)
Chemical compound, drugGSK2193874Tocris BioscienceCat. #: 5106/5
Chemical compound, drugCyclopiazonic acid (CPA)Tocris BioscienceCat. #: 1235/10
Chemical compound, drugGSK1016790ATocris BioscienceCat. #: 6433/10
Chemical compound, drugPhorbol 12-myristate 13-acetate (PMA)Tocris BioscienceCat. #: 1201/1
Chemical compound, drugAR-C 118925XXTocris BioscienceCat. #: 4890/5
Chemical compound, drug2-Thio UTP tetrasodium saltTocris BioscienceCat. #: 3280/1
Chemical compound, drugMRS2179Tocris BioscienceCat. #: 0900/10
Chemical compound, drugU-73122Tocris BioscienceCat. #: 1268/10
Chemical compound, drugNS309Tocris BioscienceCat. #: 3895/10
Chemical compound, drugARL-67156Tocris BioscienceCat. #: 1283/10
OtherFluo-4-AMInvitrogenCat. #: F14201
Chemical compound, drug1-O-9Z-octadecenoyl-2-O-acetyl-sn-glycerol (OAG)Cayman ChemicalsCat. #: 62600
Chemical compound, drugPPADSCayman ChemicalsCat. #: 14537
Chemical compound, drugGö-6976Cayman Chemicals Cat. #: 13310
Chemical compound, drugJNJ-47965567Cayman ChemicalsCat. #: 21895
Chemical compound, drugU46619Cayman ChemicalsCat. #: 16452
Chemical compound, drugTamoxifenSigma-AldrichCat. #: T5648
Peptide, recombinant proteinApyraseSigma-AldrichCat. #: A6535
Software, algorithmLabChart8ADInstruments
https://www.adinstruments.com/products/labchart
RRID:SCR_017551
Software, algorithmSegment version 2.0 R5292Twilio(http://segment.heiberg.se)
Software, algorithmIonOptixIonOptix, LLC (


https://www.ionoptix.com/products/software/ionwizard-core-and-analysis/)
Software, algorithmSparkAnDr. Adrian Bonev,
University of Vermont,
Burlington, VT, USA
PMID:22556255
Software, algorithmClampFit10.3Molecular Devices
(https://www.moleculardevices.com/)
RRID:SCR_011323
Software, algorithmImageJNational Institutes of Health
(https://imagej.nih.gov/ij/)
RRID:SCR_003070
Software, algorithmPatchMaster v2x90 programHarvard Bioscience
https://www.harvardbioscience.com/
RRID:SCR_000034
Software, algorithmFitMaster v2x73.2Harvard Bioscience
https://www.harvardbioscience.com/
RRID:SCR_016233
Software, algorithmMATLAB R2018aMathWorks
https://www.mathworks.com/products/matlab.html
RRID:SCR_013499
Software, algorithmCorelDraw Graphics Suite X7CorelDraw
(https://www.coreldraw.com/en)
RRID:SCR_014235
Software, algorithmGraphPad Prism 8.3.0GraphPad Software, Inc
(https://www.graphpad.com/)
RRID:SCR_002798
Software, algorithmGLIMMPSE software
(https://glimmpse.samplesizeshop.org/)
RRID:SCR_016297
Software, algorithmBiorenderhttp://biorender.comRRID:SCR_018361

Drugs and chemical compounds

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Cyclopiazonic acid (CPA), GSK2193874, GSK1016790A, phorbol 12-myristate 13-acetate (PMA), AR-C 118925XX, 2-Thio UTP tetrasodium salt, MRS2179, U-73122, NS309, and ARL-67156 were purchased from Tocris Bioscience (Minneapolis, MN). Fluo‐4-AM (Ca2+ indicator) were purchased from Invitrogen (Carlsbad, CA). 1-O-9Z-octadecenoyl-2-O-acetyl-sn-glycerol (OAG), PPADS (sodium salt), Gö-6976, JNJ-47965567, and U46619 were purchased from Cayman Chemicals (Ann Arbor, MI). Tamoxifen and apyrase were obtained from Sigma‐Aldrich (St. Louis, MO).

Animal protocols and models

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All animal protocols were approved by the University of Virginia Animal Care and Use Committee (protocols 4100 and 4120). Both male and female mice were used in this study and age- and sex-matched controls were used. No sex differences were observed in RVSPs and TRPV4-induced dilation of PAs. C57BL6/J were obtained from the Jackson Laboratory (Bar Harbor, ME). Inducible endothelial cell (EC)-specific TRPV4 channel knockout (Trpv4 cKO-EC; Lohman et al., 2015; Moore et al., 2013), smooth muscle cell (SMC)-specific TRPV4 channel knockout (Trpv4 cKO-SMC; Billaud et al., 2015), EC-specific caveolin-1 knockout (Cav1 cKO-EC; Chen et al., 2012), EC-specific P2Y2R receptor knockout (P2ry2 cKO-EC; Chen et al., 2017), EC-specific Panx1 channel knockout (Panx1 cKO-EC; Lohman et al., 2015; Poon et al., 2014) and SMC-specific Panx1 channel knockout (Panx1 cKO-SMC; Billaud et al., 2015) mice (10–14 weeks old) were used. Mice were housed in an enriched environment and maintained under a 12:12 hr light/dark photocycle at ∼23°C with fresh tap water and standard chow diet available ad libitum. Mice were euthanized with pentobarbital (90 mg/kg; intraperitoneally; Diamondback Drugs, Scottsdale, AZ) followed by cervical dislocation for harvesting lung tissue. Fourth‐order PAs (~50 μm diameter) were isolated in cold HEPES‐buffered physiological salt solution (HEPES‐PSS, in mmol/L, 10 HEPES, 134 NaCl, 6 KCl, 1 MgCl2 hexahydrate, 2 CaCl2 dihydrate, and 7 dextrose, pH adjusted to 7.4 using 1 mol/L NaOH).

Trpv4fl/fl (Moore et al., 2013), Cav1fl/fl (Chen et al., 2012), Panx1fl/fl (Lohman et al., 2015; Poon et al., 2014) and P2ry2fl/fl 57mice were crossed with VE-cadherin (Cdh5, endothelial) Cre mice (Moore et al., 2013) or SMMHC (smooth muscle) Cre mice (Wirth et al., 2008). EC- or SMC-specific knockout of Trpv4, Cav1, Panx1, or P2ry2 was induced by injecting 6-week-old Trpv4fl/fl Cre+, Cav1fl/fl Cre+, Panx1fl/fl Cre+, and P2ry2fl/fl Cre+ mice with tamoxifen (40 mg/kg intraperitoneally per day for 10 days). Tamoxifen-injected Trpv4fl/fl Cre-, Cav1fl/fl Cre-, Panx1fl/fl Cre-, and P2ry2fl/fl Cre- mice were used as controls. Mice were used for experiments after a 2-week washout period. Genotypes for Cdh5 Cre and SMMHC Cre were confirmed following previously published protocols (Moore et al., 2013; Wirth et al., 2008). Trpv4fl/fl (Moore et al., 2013), Cav1fl/fl (Chen et al., 2012), Panx1fl/fl (Lohman et al., 2015; Poon et al., 2014), and P2ry2fl/fl (Chen et al., 2017) genotyping was performed as described previously. Cdh5-Optoα1AR mice were developed by CHROMus (Cornell University, USA).

RVSP and Fulton index measurement

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Mice were anesthetized with pentobarbital (50 mg/kg bodyweight; intraperitoneally) and bupivacaine HCl (100 μL of 0.25% solution; subcutaneously) was used to numb the dissection site on the mouse. RVSP was measured as an indirect indicator of PAP. A Mikro-Tip pressure catheter (SPR-671; Millar Instruments, Huston, TX), connected to a bridge amp (FE221), and a PowerLab 4/35 4-channel recorder (ADInstruments, Colorado Springs, CO), was inserted through the external jugular vein into the right ventricle. Right ventricular pressure and heart rate were acquired and analyzed using LabChart8 software (ADInstruments). A stable 3 min recording was acquired for all the animals, and 1 min continuous segment was used for data analysis. When necessary, traces were digitally filtered using a low-pass filter at a cutoff frequency of 50 Hz. At the end of the experiments, mice were euthanized, and the hearts were isolated for right ventricular hypertrophy analysis. Right ventricular hypertrophy was determined by calculating the Fulton index, a ratio of the right ventricular (RV) heart weight over the left ventricular (LV) plus septum (S) weight (RV/ LV + S).

Luciferase assay for total ATP release

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ATP assay protocol was adapted from Yang et al., 2020. Fourth‐order PAs (~50 μm diameter) were isolated in cold HEPES‐buffered physiological salt solution (HEPES‐PSS, in mmol/L, 10 HEPES, 134 NaCl, 6 KCl, 1 MgCl2 hexahydrate, 2 CaCl2 dihydrate, and 7 dextrose, pH adjusted to 7.4 using 1 mol/L NaOH). Isolated PAs were pinned down en face on a Sylgard block and cut open. PAs were placed in black, opaque 96-well plates and incubated in HEPES-PSS for 10 min at 37°C, followed by incubation with the ectonucleotidase inhibitor ARL 67156 (300 μmol/L, Tocris Bioscience, Minneapolis, MN) for 30 min at 37°C. 50 μL volume of each sample was transferred to another black, opaque 96-well plate. ATP was measured using ATP bioluminescence assay reagent ATP Bioluminescence HSII kit (Roche Applied Science, Penzberg, Germany). Using a luminometer (FluoStar Omega), 50 μL of luciferin:luciferase reagent (ATP bioluminescence assay kit HSII; Roche Applied Science) was injected into each well and luminescence was recorded following a 5 s orbital mix and sample measurement at 7 s. ATP concentration in each sample was calculated from an ATP standard curve. For some experimental groups, PAs were first mounted on a pressure myography chamber and were denuded by pushing air through the lumen for 1 min.

Cardiac magnetic resonance imaging (MRI)

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MRI studies were conducted under protocols that comply with the Guide for the Care and Use of Laboratory Animals (NIH publication no. 85-23, revised 1996). Mice were positioned in the scanner under 1.25% isoflurane anesthesia and body temperature was maintained at 37°C using thermostatic circulating water. A cylindrical birdcage RF coil (30 mm diameter, Bruker, Ettlingen, Germany) with an active length of 70 mm was used, and heart rate, respiration, and temperature were monitored during imaging using a fiber optic, MR-compatible system (Small Animal Imaging Inc, Stony Brook, NY). MRI was performed on a 7 Tesla (T) Clinscan system (Bruker) equipped with actively shielded gradients with a full strength of 650 mT/m and a slew rate of 6666 mT/m/ms (Vandsburger et al., 2007). Six short-axis slices were acquired from base to apex, with slice thickness of 1 mm, in-plane spatial resolution of 0.2 × 0.2 mm2, and temporal resolution of 8–12 ms. Baseline ejection fraction (EF), end-diastolic volume (EDV), end-systolic volume (ESV), myocardial mass, wall thickness, stroke volume (SV), and cardiac output (CO) were assessed from the cine images using the freely available software Segment version 2.0 R5292 (http://segment.heiberg.se).

Pressure myography

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Isolated mouse PAs (~50 μm) were cannulated on glass micropipettes in a pressure myography chamber (The Instrumentation and Model Facility, University of Vermont, Burlington, VT) at areas lacking branching points and were pressurized at a physiological pressure of 15 mm Hg (Ottolini et al., 2020a). Arteries were superfused with PSS (in mmol/L, 119 NaCl, 4.7 KCl, 1.2 KH2PO4, 1.2 MgCl2 hexahydrate, 2.5 CaCl2 dihydrate, 7 dextrose, and 24 NaHCO3) at 37°C and bubbled with 20% O2/5% CO2 to maintain the pH at 7.4. All drug treatments were added to the superfusing PSS. PAs were pre-constricted with 50 nmol/L U46619 (a thromboxane A2 receptor agonist). All other pharmacological treatments were performed in the presence of U46619. Before measurement of vascular reactivity, arteries were treated with NS309 (1 μmol/L), a direct opener of endothelial IK/SK channels, to assess endothelial health. Arteries that failed to fully dilate to NS309 were discarded. Changes in arterial diameter were recorded at a 60‐ms frame rate using a charge‐coupled device camera and edge‐detection software (IonOptix LLC, Westwood, MA; Sonkusare et al., 2012; Sonkusare et al., 2014). All drug treatments were incubated for 10 min. At the end of each experiment, Ca2+‐free PSS (in mmol/L, 119 NaCl, 4.7 KCl, 1.2 KH2PO4, 1.2 MgCl2 hexahydrate, 7 dextrose, 24 NaHCO3, and 5 EGTA) was applied to assess the maximum passive diameter. Percent constriction was calculated by

(1) [(DiameterbeforeDiameterafter)/Diameterbefore]×100

where Diameterbefore is the diameter of the artery before a treatment and Diameterafter is the diameter after the treatment. Percent dilation was calculated by

(2) [(DiameterdilatedDiameterbasal)/(DiameterCafreeDiameterbasal)]×100

w here Diameterbasal is the stable diameter before drug treatment, Diameterdilated is the diameter after drug treatment, and DiameterCa‐free is the maximum passive diameter.

Flow/shear stress-induced ATP release

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Flow/shear stress was measured using a protocol modified from Ahn et al., 2017. Briefly, isolated PAs (~50 μm) were cannulated on glass micropipettes in a pressure myography chamber (The Instrumentation and Model Facility, University of Vermont) at areas lacking branching points and were pressurized at a physiological pressure of 15 mm Hg (Ottolini et al., 2020a). Arteries were superfused with PSS (in mmol/L, 119 NaCl, 4.7 KCl, 1.2 KH2PO4, 1.2 MgCl2 hexahydrate, 2.5 CaCl2 dihydrate, 7 dextrose, and 24 NaHCO3) at 37°C and bubbled with 20% O2/5% CO2 to maintain the pH at 7.4. The arteries were treated luminally with 300 μmol/L ARL-67156 (ecto-ATPase inhibitor; Sigma‐Aldrich) to avoid ATP degradation throughout the duration of the experiment. The tips of the cannulating pipettes were always arranged with smaller pipettes upstream and larger pipettes downstream. The average tip size was 20.1 ± 0.4 μm at the upstream end and 23.6 ± 0.4 μm at the downstream end. Both ends of the vessel were secured, and the vessel was maintained at an intraluminal pressure of 15 cmH2O by elevating the inflow reservoir. Flow/shear stress was increased by adjusting the height of the reservoir. Flow-induced luminal solution was collected at the outflow pipette end. After a 30 min equilibration period, a baseline sample was collected for luminal ATP measurement. Shear stress was calculated from the flow rate in the vessel lumen and the diameter of the vessels using the equation (Zemskov et al., 2011) : τ=4(μQ˙/(πr3)), where μ is viscosity, Q. is volumetric flow rate, and r is internal radius of the vessel. The volumetric flow rate was measured as the volume of the flowthrough at different pressures. Vessel diameter was measured at each flow rate. The shear stress range was 4–14 dynes/cm2. Luminal outflow samples per shear stress range were obtained every 30 min. The samples were used for luciferase assays for total ATP release, as described above.

Ca2+ imaging

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Measurements of TRPV4EC Ca2+ sparklets in the native endothelium of mouse PAs were performed as previously described (Sonkusare et al., 2012). Briefly, fourth-order (~50 μm) PAs were pinned down en face on a Sylgard block and loaded with Fluo-4-AM (10 μmol/L) in the presence of pluronic acid (0.04%) at 30°C for 30 min. TRPV4EC Ca2+ sparklets were recorded at 30 frames per second with Andor Revolution WD (with Borealis) spinning‐disk confocal imaging system (Oxford Instruments, Abingdon, UK) comprised an upright Nikon microscope with a 60× water dipping objective (numerical aperture 1.0) and an electron multiplying charge coupled device camera (iXon 888, Oxford Instruments). All experiments were carried out in the presence of cyclopiazonic acid (20 μmol/L, a sarco‐endoplasmic reticulum [ER] Ca2+‐ATPase inhibitor) in order to eliminate the interference from Ca2+ release from intracellular stores. Fluo-4 was excited at 488 nm with a solid‐state laser and emitted fluorescence was captured using a 525/36 nm band‐pass filter. TRPV4EC Ca2+ sparklets were recorded before and 5 min after the addition of specific compounds. To generate fractional fluorescence (F/F0) traces, a region of interest defined by a 1.7‐μm2 (5 × 5 pixels) box was placed at a point corresponding to peak sparklet amplitude. Each field of view was ~110 × 110 μm and covered ~15 ECs. Representative F/F0 traces were filtered using a Gaussian filter and a cutoff corner frequency of 4 Hz. Sparklet activity was assessed as described previously using the custom‐designed SparkAn software (Sonkusare et al., 2012; Sonkusare et al., 2014).

Calculation of TRPV4 sparklet activity per site

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Activity of TRPV4 Ca2+ sparklets was analyzed as described previously (Sonkusare et al., 2012; Ottolini et al., 2020b; Sonkusare et al., 2014). Area under the curve for all the events at a site was determined using trapezoidal numerical integration ([F−F0]/F0 over time, in seconds). The average number of active TRPV4 channels, as defined by NPO (where N is the number of channels at a site and PO is the open state probability of the channel), was calculated by

(3) NPO=(Tlevel1+2Tlevel2+3Tlevel3+4Tlevel4)/Ttotal

where T is the dwell time at each quantal level detected at TRPV4 sparklet sites and Ttotal is the duration of the recording. NPO was determined using Single Channel Search module of Clampfit and quantal amplitudes derived from all‐points histograms (Marziano et al., 2017) (ΔF/F0 of 0.29 for Fluo‐4-loaded PAs).

Total number of sparklet sites in a field was divided by the number of cells in that field to obtain sparklet sites per cell.

All-points histograms

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All-points amplitude histograms were constructed as described previously (Sonkusare et al., 2012; Ottolini et al., 2020b). Briefly, images were filtered with a Kalman filter (adopted from an ImageJ plug-in written by Christopher Philip Mauer, Northwestern University, Chicago, IL; acquisition noise variance estimate = 0.05; filter gain = 0.8). The inclusion criteria were a stable baseline containing at least five steady points and a steady peak containing at least five peak points. Sparklet traces were exported to ClampFit10.3 for constructing an all-points histogram, which was fit with the multiple Gaussian function below:

(4) fF/F0=i=1Nai2πσiexp-FF0-μi22σi2

where F/F0 represents fractional fluorescence, a represents the area, μ represents the mean value, and σ2 represents the variance of the Gaussian distribution. While the detected sparklets can have multiple amplitudes corresponding to quantal level 1, 2, 3, or 4, the baseline (level 0) was the same for all the detected sparklets regardless of the amplitude of the sparklets. Therefore, the baseline corresponds to a higher count compared to all other events.

Immunostaining

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Immunostaining was performed on fourth-order PAs (~50 μm) pinned en face on SYLGARD blocks. PAs were fixed with 4% paraformaldehyde (PFA) at room temperature for 15 min and then washed three times with phosphate-buffered saline (PBS). The tissue was permeabilized with 0.2% Triton-X for 30 min, blocked with 5% normal donkey serum (ab7475, Abcam, Cambridge, MA) or normal goat serum (ab7475, Abcam), depending on the host of the secondary antibody used, for 1 hr at room temperature. PAs were incubated with the primary antibodies (Key resources table) overnight at 4°C. Following the overnight incubation, PAs were incubated with secondary antibody 1:500 Alexa Fluor 568-conjugated donkey anti-rabbit (Life Technologies, Carlsbad, CA) for 1 hr at room temperature in the dark room. For nuclear staining, PAs were washed with PBS and then incubated with 0.3 mmol/L DAPI (Invitrogen, Carlsbad, CA) for 10 min at room temperature. Images were acquired along the z-axis from the surface of the endothelium to the bottom where the EC layer encounters the smooth muscle cell layer with a slice size of 0.1 μm using the Andor microscope described above. The internal elastic lamina (IEL) autofluorescence was evaluated using an excitation of 488 nm with a solid-state laser and collecting the emitted fluorescence with a 525/36 nm band-pass filter. Immunostaining for the protein of interest was evaluated using an excitation of 561 nm and collecting the emitted fluorescence with a 607/36 nm band-pass filter. DAPI immunostaining was evaluated using an excitation of 409 nm and collecting the emitted fluorescence with a 447/69 nm band-pass filter. The specificity of Panx1 and P2Y2R antibodies was confirmed by a lack of signal in PAs from endothelial knockout mice. The specificity of TRPV4, Cav-1, and PKC antibodies was confirmed previously (Daneva et al., 2021; Ottolini et al., 2020b).

In situ PLA

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Fourth-order (~50 μm) PAs were pinned en face on SYLGARD blocks. PAs were fixed with 4% PFA for 15 min followed by three washes with PBS. PAs were then permeabilized with 0.2% Triton X for 30 min at room temperature followed by blocking with 5% normal donkey serum (Abcam plc, Cambridge, MA) and 300 mmol/L glycine for 1 hr at room temperature. After three washes with PBS, PAs were incubated with the primary antibodies (Key resources table) overnight at 4°C. The PLA protocol from Duolink PLA Technology kit (Sigma-Aldrich) was followed for the detection of co-localized proteins. Lastly, PAs were incubated with 0.3 μmol/L DAPI nuclear staining (Invitrogen) for 10 min at room temperature in the dark room. PLA images were acquired using the Andor Revolution spinning-disk confocal imaging system along the z-axis at a slice size of 0.1 μm. Images were analyzed by normalizing the number of positive puncta by the number of nuclei in a field of view. The specificity of the PLA antibodies was determined using PAs from endothelial knockout mice for one of the protein pairs.

Plasmid generation and transfection into HEK293 cells

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HEK293 cells authenticated with STR profiling were obtained from ATCC USA. Mycoplasma contamination was not detected as per ATCC website. The TRPV4 coding sequence without stop codons was amplified from mouse heart cDNA. The amplified fragment was inserted into a plasmid backbone containing a CMV promoter region for expression and, in addition, is suitable for lentiviral production by Gibson assembly. The in-frame FLAG tag was inserted into the 3′-primer used for amplification. Constructs were verified by sequencing the regions that had been inserted into the plasmid backbone. HEK293 cells were seeded (7 × 105 cells per 100 mm dish) in Dulbecco’s Modified Eagle Medium with 10% fetal bovine serum (Thermo Fisher Scientific Inc, Waltham, MA) 1 day prior to transfection. Cells were transfected using the LipofectamineLTX protocol (Thermo Fisher Scientific Inc). TRPV4 was co-expressed with PKCα and PKCβ, obtained from Origene Technologies (Montgomery County, MD).

Patch clamp in freshly isolated pulmonary ECs and in HEK293 cells

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Fresh ECs were obtained via enzymatic digestion of fourth-order PAs. Briefly, PAs were incubated in the dissociation solution (in mmol/L, 55 NaCl, 80 Na glutamate, 6 KCl, 2 MgCl2, 0.1 CaCl2, 10 glucose, 10 HEPES, pH 7.3) containing Worthington neutral protease (0.5 mg/mL) for 30 min at 37°C. The extracellular solution consisted of (in mmol/L) 10 HEPES, 134 NaCl, 6 KCl, 2 CaCl2, 10 glucose, and 1 MgCl2 (adjusted to pH 7.4 with NaOH). The intracellular pipette solution for perforated-patch configuration consisted of (in mmol/L) 10 HEPES, 30 KCl, 10 NaCl, 110 K-aspartate, and 1 MgCl2 (adjusted to pH 7.2 with NaOH). Cells were kept at room temperature in a bathing solution consisting of (in mmol/L) 10 HEPES, 134 NaCl, 6 KCl, 2 CaCl2, 10 glucose, and 1 MgCl2 (adjusted to pH 7.4 with NaOH). Narishige PC-100 puller (Narishige International USA, Inc, Amityville, NY) was utilized to pull patch electrodes, which were polished using MicroForge MF-830 polisher (Narishige International USA, Inc). The pipette resistance was (3–5 ΩM). Amphotericin B was dissolved in the intracellular pipette solution to reach a final concentration of 0.3 μmol/L. The data were acquired using HEKA EPC 10 amplifier and PatchMaster v2x90 program (Harvard Bioscience, Holliston, MA) and analyzed using FitMaster v2x73.2 (Harvard Bioscience) and MATLAB R2018a (MathWorks, Natick, MA). TRPV4 channel current was recorded from freshly isolated ECs as described previously (Sonkusare et al., 2012; Ottolini et al., 2020b). Briefly, GSK101-induced outward currents through TRPV4 channels were assessed in response to a 200 ms voltage step from –45 mV to +100 mV in the presence of ruthenium red in order to prevent Ca2+ and activation of IK/SK channels at negative voltages.

TRPV4 channel current was recorded in HEK293 cells using whole-cell patch configuration 48 hr after transfection. The intracellular solution consisted of (in mmol/L) 20 CsCl, 100 Cs-aspartate, 1 MgCl2, 4 ATP, 0.08 CaCl2, 10 BAPTA, 10 HEPES, pH 7.2 (adjusted with CsOH). The extracellular solution consisted of (in mmol/L) 10 HEPES, 134 NaCl, 6 KCl, 2 CaCl2, 10 glucose, and 1 MgCl2 (adjusted to pH 7.4 with NaOH). Currents were measured using a voltage clamp protocol where voltage-ramp pulses (–100 mV to +100 mV) were applied over 200 ms with a holding potential of –50 mV. TRPV4 currents were measured before or 5 min after treatment.

Quantitative polymerase chain reaction (qPCR)

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Mouse mesenteric arteries were denuded by pushing air through the arteries for 1 min. RNA was isolated using a Direct-zol RNA Miniprep (R2051, Zymo Research, Irvine, CA), with an in-column DNA Removal Kit. cDNA was converted with Bio-Rad iScript cDNA Synthesis Kit (1708841, Hercules, CA). The qPCR reaction mixes were prepared using Bio-Rad 2x SsoAdvanced Universal SYBR Green Supermix (1725272, Hercules, CA), 200 nmol/L primers (Panx1_F: 5′ TGCACAAGTTCTTCCCCTACA, Panx1_R: ATGGCGCGGTTGTAGACTTT; GAPDH_F: GGTTGTCTCCTGCGACTTCA; GAPDH_R TAGG GCCTCTCTTGCTCAGT; Eurofins Genomics Louisville, KY), and 20 nmol/L cDNA, then run in a Bio-Rad CFX96 qPCR Detection System. Results were analyzed using the 2-∆∆Ct method.

Statistical analysis

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Results are presented as mean ± SEM. The n = 1 was defined as one artery in the imaging experiments (Ca2+ imaging, PLA), one cell for patch-clamp experiments, one mouse for RVSP measurements, one artery for pressure myography experiments, one mouse for functional MRI, one mouse for ATP measurements, and one mouse for qPCR experiments. The data were obtained from at least three mice in experiments performed in at least two independent batches. The individual data points are shown for each dataset. For in vivo experiments, an independent team member performed random assignment of animals to groups and did not have knowledge of treatment assignment groups. All the in vivo experiments were blinded; information about the groups or treatments was withheld from the experimenter or from the team member who analyzed the data. All data are shown in graphical form using CorelDraw Graphics Suite X7 (Ottawa, ON, Canada) and statistically analyzed using GraphPad Prism 8.3.0 (Sand Diego, CA). A power analysis to determine group sizes and study power (>0.8) was performed using GLIMMPSE software (α = 0.05; >20% change). Using this method, we estimated at least five cells per group for patch-clamp experiments, five arteries per group for imaging and pressure myography experiments, and four mice per group for RVSP measurements and MRI. A Shapiro–Wilk test was performed to determine normality. The data in this article were normally distributed; therefore, parametric statistics were performed. Data were analyzed using two-tailed, paired or independent t-test (for comparison of data collected from two different treatments), one-way ANOVA or two-way ANOVA (to investigate statistical differences among more than two different treatments). Tukey correction was performed for multiple comparisons with one-way ANOVA, and Bonferroni correction was performed for multiple comparisons with two-way ANOVA. Statistical significance was determined as a p-value <0.05.

Data availability

All data generated or analyzed during this study are included in the manuscript. Individual numeric values are shown in the scatterplots for each dataset. An Excel sheet with source data for Figure 1J has been provided.

References

Decision letter

  1. Mark T Nelson
    Reviewing Editor; University of Vermont, United States
  2. Richard W Aldrich
    Senior Editor; The University of Texas at Austin, United States
  3. William Jackson
    Reviewer; Michigan State University, United States

Our editorial process produces two outputs: i) public reviews designed to be posted alongside the preprint for the benefit of readers; ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Acceptance summary:

This study, which makes a connection between several proteins known to regulate endothelial function in pulmonary arteries, may be of interest to vascular, pulmonary and ion channel physiologists. The study provides compelling evidence that ATP released from pulmonary artery endothelial cell pannexin1 channels activates TRPV4 channelsthat is facilitated by the scaffolding protein Caveolin-1 and that this pathway helps to maintain low pulmonary vascular resistance and pulmonary artery pressure. Identification of this pathway provides new drug targets to improve pulmonary endothelial function in disease states such characterized by impaired endothelial function.

Decision letter after peer review:

Thank you for submitting your article "Endothelial Pannexin 1-TRPV4 channel signaling lowers pulmonary arterial pressure" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Richard Aldrich as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: William Jackson (Reviewer #3).

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential revisions:

1. The major concern is related to conceptual significance. The reviewer appreciates that the work presented here connects Cav-1, Panx1, P2Y2R, PKC and TRPV4 into a signaling axis regulating pulmonary artery reactivity. However, this group has already published similar papers implicating a role for this axis in pulmonary arteries (and a similar axis in systemic arteries), and comparable conclusions have been reached by examining members of the pathway independently. Therefore, it is unclear what new conceptual information is gained, other than the link between all the proteins in the complex. Perhaps the authors could highlight more the major gaps in knowledge and novel aspects of their work. It should be clear to the reviewers what is novel and new.

2. A critical step in the proposed pathway is activation of Pannexin 1. The authors should provide evidence or at least clearly discuss how pannexin 1 could be activated to engage the proposed pathway.

3. The physiological role of the proposed pathway is unclear. PAP is normally low (8 – 20 mm Hg). Are the authors proposing that this pathway is always engaged to maintain low PAP? If so, then how is Pannexin 1 being tonically activated? This would also imply that there exists a tonic constrictor pathway which Pannexin1-TRPV4 opposes. Does this exist? Or the proposed Panx1-V4 pathway only engaged in the face of pulmonary hypertension. It is hard to envision a dilatory pathway when the system is already at low pressure, i.e., relaxed.

4. The use of the term, "small, resistance-sized pulmonary arteries" is curious. Pulmonary arteries have low resistance and pressure. What is the basis of using this term?

5. Previous work from this group has shown that endothelial ATP activates TRPV4 channels to cause nitric oxide dependent vasodilation of pulmonary arteries (PMID: 29275372 and PMID: 32463112). The current study has many of the same elements of previous studies. The authors should clearly what is new and what has been shown before.

6. Authors proposed that shear stress and flow can cause release of ATP in physiology condition and cited previous work from Yamamoto et al. (PMID: 12714321). It is important that authors perform some functional experiments to confirm this mechanism with TRPV4 and Panx1 KOs.

7. Authors' measurements show that approximately 400 nM ATP is released from PA endothelial cells, but they have used 1 µM for dilation experiments (Figure 1B) and 10 µM for all other experiments, which is 6-100-fold higher concentration than the measured concentration. Authors needs to provide justification for that.

8. Page 10: authors mentioned that PPADS and JNJ 47965567 did not inhibit TRPV4 sparkets, but Figure 2D shows significant inhibition which potentially suggest involvement of other purinergic (P2Xs) signaling.

9. Why did authors use different concentrations of activators such as GSK101 (1-30 nM) with different KO animals? Justification needed.

10. Figure 5E, authors suggested that PAP regulation is via NO signaling, but they did not provide any experimental evidence using eNOS inhibitors or eNOS KO animals.

11. Authors mentioned that they have used both Male and Female mice. Did they observe any differences between the sexes?

12. It is unclear why 1 nM GSK101 (Figure 1D) or 10 nM GSK101 (Figure 1G) induced reduced TRPV4 activity/current density in Panx1EC-/- compared to Panx1fl/fl. These observations do not support the statement on Page 6, Line 127-129, indicating that "reduced TRPV4EC channel activity in Panx1EC-/- is due to impaired channel regulation…". Since GSK101 is bypassing activation of the Panx1 pathway to stimulate TRPV4 channels directly, the expectation is that channel activity/current density should be similar provided that all other conditions are similar. A deeper consideration of this issue will help clarify concerns. The authors may consider performing immunofluorescence experiments to show that TRPV4-associated fluorescence or distribution of TRPV4 is similar in EC from Panx1fl/fl and Panx1EC-/-.

13. The statement on Page 6, Line 131-132 is not supported by the data presented. Figure 1 does not show the direct regulation of TRPV4 by Panx1. This statement should be revised. The U466 experiments in Figure 1I are interesting but do not link TRPV4 and Panx1. These experiments show that genetic ablation of any of these proteins increases pulmonary artery reactivity to U466. Can the increase in contraction to U466 in Panx1EC-/- reversed by treating the arteries with the TRPV4 agonist?

14. A major conclusion of the study is the formation of a nanocomplex between Cav-1, Panx1, P2Y2R, PKC and TRPV4. However, the interaction between these proteins is only presented for a subset of protein pairs. The study will be strengthened by providing data showing nanometer proximity between Panx1-P2Y2R, Panx1-TRPV4, P2Y2R-TRPV4 and that genetic ablation of Cav-1 disrupt the proximity between all these protein pairs. Based on the authors' data, P2Y1R should not activate TRPV4 channels. Thus, it will be expected that P2Y1R are not part of the complex. Is this the case? Please show antibody validation and negative controls for PLA. The authors should also explain why they think that Cav-1, Panx1, P2Y2R, PKC and TRPV4 are in nanometer proximity of each other given that no super-resolution data was presented.

15. Is ATP-induced dilation of small diameter pulmonary arteries prevented in Panx1EC-/-, P2Y2REC-/- and Cav-1EC-/-?

16. The description of Figure 2A gives the impression that perhaps all the ATP may come from EC. How much of the ATP will be left if the endothelial layer is removed before performing the assay? What are the ATP levels when using Panx1SMC-/- arteries? The study will be strengthened by linking mechanisms mediating basal ATP release with activation of the proposed pathway.

17. The sparklet traces shown in Figure 4D (recorded with X-Rhod in Cdh5-optoα1AR EC) is ascribed to TRPV4 channels, but there is no evidence that this is the case. This is important as data suggest that these sparklet events have a higher activity before light stimulation than other TRPV4 sparklet data presented throughout the manuscript. Are the amplitude and kinetics of sparklet events in this figure match that of known TRPV4 sparklets?

18. Is TRPV4 current density diminished in Cav-1EC-/-?

19. Authors should show evidence that TRPV4 and Panx1 expression is reduced/knockout in TRPV4SMC-/- and Panx1SMC-/-, respectively.

20. The immunofluorescent images in Figures 1A, 1C, 2C and 3A should be better described in the main text.

21. page 5, line 111 – I'm a bit puzzled why there was no right hypertrophy in your models with elevated PAP? Was it just that the duration of elevated PAP was insufficient to cause right heart hypertrophy? Please discuss.

22. page 6, lines 125-127 – Were currents different between control and EC Panx1-/- with 30 nM GSK101 in your patch clamp experiments? Why were higher concentrations of GSK101 used in the patch clamp experiments?

23. page 6, lines 129-131 – Did knock out of EC TRPV4-/- or EC Panx1-/- cause PAs to develop myogenic tone?

24. page 6 line 134 and onward – in your ex vivo ca2+ imaging experiments, what is the stimulus that is leading to Panx1 activity and release of ATP? It would seem important to identify the stimulus. Also, does luminal apyrase in pressure myograph experiments have the same effect as Panx1 knockout? How about P2Y2R-/- in pressure myography experiments? These experiments also would seem to be important to close the loop.

25. page 10, line 227 – What is the physiological stimulus for Panx1 and ATP release?

26. page 11, lines 236-239 – How do you reconcile the lack of effect of global TRPV4 knockout on PAP with your finding that EC TRPV4-/- increases PAP? Shoudln't global do the same thing? This should be better discussed.

27. page 16, lines 351-353 – You did not cannulate the pressure catheter – you cannulated the external jugular vein for access to the right ventricle – please revise to clarify.

28. Table 2 – How did you determine the specificity of the antibodies used?

29. Figure legends – please provide exact n-values for each panel and also please provide exact p-values for all statistical tests.

https://doi.org/10.7554/eLife.67777.sa1

Author response

Essential revisions:

1. The major concern is related to conceptual significance. The reviewer appreciates that the work presented here connects Cav-1, Panx1, P2Y2R, PKC and TRPV4 into a signaling axis regulating pulmonary artery reactivity. However, this group has already published similar papers implicating a role for this axis in pulmonary arteries (and a similar axis in systemic arteries), and comparable conclusions have been reached by examining members of the pathway independently. Therefore, it is unclear what new conceptual information is gained, other than the link between all the proteins in the complex. Perhaps the authors could highlight more the major gaps in knowledge and novel aspects of their work. It should be clear to the reviewers what is novel and new.

We thank the reviewers and the Editor for identifying the strengths of the manuscript and for their constructive feedback. We recently reported that endothelial TRPV4 channels decrease the contractility of small pulmonary arteries (PAs) and lower resting pulmonary arterial pressure (PAP)1. Moreover, exogenous ATP activated endothelial TRPV4 channels to dilate PAs2. However, the regulation of TRPV4 channel activity by endogenously released ATP, the source of endogenously released ATP, and the precise signaling mechanisms for ATP activation of endothelial TRPV4 channels were not known. In the current manuscript, we present a novel signaling axis whereby ATP efflux through endothelial Pannexin 1 (Panx1) activates nearby TRPV4 channels via purinergic receptor signaling to lower PA contractility and PAP. Following key findings contribute to the high conceptual significance and novelty of the study:

1) First evidence, using endothelial knockout mice, that ATP efflux through endothelial Panx1 lowers PA contractility and PAP. Notably, previous studies have shown that endothelial Panx1 activity does not contribute to vasodilation in systemic arteries and systemic blood pressure regulation3.

2) First direct evidence that ATP efflux through Panx1 promotes endothelial TRPV4 channel activity in PAs, but TRPV4 channel activity does not regulate ATP efflux through Panx1 under resting conditions.

3) First evidence that ATP effluxed through endothelial Panx1 stimulates purinergic P2Y2 receptor (P2Y2R) signaling to activate TRPV4 channels and lower PA contractility and resting PAP.

4) Earlier, we showed that endothelial caveolin-1 (Cav-1) lowers the resting PAP1. In the current manuscript, we provide evidence that endothelial Cav-1 provides a signaling scaffold for Panx1, P2Y2R, and TRPV4 channels, ensuring their spatial proximity in PAs. Activation of the endothelial Panx1–P2Y2 receptor–TRPV4 channel pathway, enabled by the Cav-1 scaffold, lowers PA contractility and PAP.

5) PAs are a high-flow vascular bed, yet flow-induced endothelial signaling is poorly understood in PAs. We provide evidence that physiological flow/shear stress increases luminal ATP release through endothelial Panx1 activation.

We have now modified the Introduction and other sections of the manuscript to highlight the conceptual significance and novelty of the results presented in this manuscript.

2. A critical step in the proposed pathway is activation of Pannexin 1. The authors should provide evidence or at least clearly discuss how pannexin 1 could be activated to engage the proposed pathway.

Using pressure myography in small PAs, we now show that endothelial Panx1-dependent signaling lowers pressure-induced (myogenic) constriction of PAs, thus exerting a dilatory effect under resting conditions. Regarding physiological activators of Panx1, flow/shear stress has been shown to activate ATP efflux through Panx14. PAs are a high-flow vascular bed. However, flow/shear stress-activated signaling in PAs remains entirely unknown. In the revised manuscript, we provide evidence that flow/shear stress increases luminal ATP levels in PAs through endothelial Panx1 activation. Specifically, increased flow/shear stress elevated luminal ATP levels in pressurized PAs, an effect that was absent in PAs from endothelial Panx1-/- mice (Figure 2G). Together, these findings support a dilatory effect of endothelial Panx1 under resting conditions and activation of endothelial Panx1 by flow/shear stress.

3. The physiological role of the proposed pathway is unclear. PAP is normally low (8 – 20 mm Hg). Are the authors proposing that this pathway is always engaged to maintain low PAP? If so, then how is Pannexin 1 being tonically activated? This would also imply that there exists a tonic constrictor pathway which Pannexin1-TRPV4 opposes. Does this exist? Or the proposed Panx1-V4 pathway only engaged in the face of pulmonary hypertension. It is hard to envision a dilatory pathway when the system is already at low pressure, i.e., relaxed.

We thank the reviewer for raising these important points. The reviewer has correctly noted that in this manuscript we propose a tonic endothelial Panx1-TRPV4 signaling axis that maintains a low PAP. It has generally been considered that PAs, due to low intraluminal pressure, are relaxed/low-resistance. However, this assumption results from the lack of studies on pressure-induced (myogenic) constriction in small PAs under resting conditions. In this manuscript we provide evidence that small PAs (50-100 microns) show myogenic constriction (~ 20% at 15 mm Hg), whereas large PAs (> 200 microns) do not (Figure 2B, 2D, 3G; Supplemental Figure 2A). Further, we show that PAs from endothelial Panx1-/-, TRPV4-/-, and P2Y2R-/- mice develop significantly higher myogenic constriction compared to PAs from the respective control mice (Figure 2B, 2D and 3G). These data strongly support tonic activation of endothelial Panx1–P2Y2R–TRPV4 channel pathway and its dilatory effect under basal conditions. In addition to myogenic constriction, agonist-induced constriction was also higher in PAs from endothelial Panx1-/-, TRPV4-/-, and P2Y2R-/- mice compared to the control mice (Figure 2C, 2E, and 3H). The detailed studies of myogenic constriction of PAs and mechanisms involved will be published in a separate manuscript.

As the reviewer pointed out, it is plausible that the Panx1-dependent signaling is altered in pulmonary hypertension, a possibility that has not been tested. In this regard we have shown that endothelial TRPV4 channel activity is impaired in PAs from PH patients and mouse models of PH1.

4. The use of the term, "small, resistance-sized pulmonary arteries" is curious. Pulmonary arteries have low resistance and pressure. What is the basis of using this term?

While the general opinion is that PAs are low-resistance or are in a completely relaxed state, there are no detailed studies showing a lack of myogenic constriction in pressurized small PAs under basal conditions. Our new PA pressure myography data show that small PAs (~ 50-100 microns) develop pressure-induced/myogenic constriction, whereas large PAs (~ 200 microns or more) do not (Figure 2B, 2D, and 3G, Supplemental Figure 2A). We use the term “resistance PAs” to describe PAs that show myogenic constriction (~ 50-100 microns, used in this study). We present evidence that PAs develop myogenic constriction at the physiological intraluminal pressure (15 mm Hg, Figure 2B, 2D, and 3G). We also show that PAs from endothelial Panx1-/-, TRPV4-/-, and P2Y2R-/- mice develop significantly higher myogenic constriction compared to PAs from the respective control mice. Thus, endothelial knockout of Panx1, P2Y2R, or TRPV4 channel increases PA contractility and elevates PAP.

5. Previous work from this group has shown that endothelial ATP activates TRPV4 channels to cause nitric oxide dependent vasodilation of pulmonary arteries (PMID: 29275372 and PMID: 32463112). The current study has many of the same elements of previous studies. The authors should clearly what is new and what has been shown before.

We previously showed that exogenous ATP (1-10 µM) dilates PAs through endothelial TRPV4-eNOS signaling1, 2. However, the regulation of TRPV4 channel activity by endogenously released ATP, the source of endogenously released ATP, and the signaling mechanisms for ATP activation of endothelial TRPV4 channels were not known. The crucial new findings in this manuscript (not published elsewhere in the literature) include: (1) endothelial Panx1 is the predominant source of ATP efflux in PAs; (2) efflux of ATP through endothelial Panx1 enhances the activity of endothelial TRPV4 channels; (3) Panx1-effluxed extracellular ATP activates TRPV4 channels via endothelial P2Y2R–protein kinase C signaling; (4) Cav-1 provides a signaling scaffold for endothelial Panx1–P2Y2R–TRPV4 channel signaling; (5) endothelial Panx1–P2Y2R–TRPV4 channel pathway opposes myogenic and agonist-induced constriction of PAs and lowers PAP; and (6) flow/shear stress activates ATP release through endothelial Panx1 in PAs.

We have now revised the Introduction and other sections of the manuscript to highlight the novel findings.

6. Authors proposed that shear stress and flow can cause release of ATP in physiology condition and cited previous work from Yamamoto et al. (PMID: 12714321). It is important that authors perform some functional experiments to confirm this mechanism with TRPV4 and Panx1 KOs.

We performed a series of experiments to address this comment. We studied luminal ATP levels in pressurized PAs (15 mm Hg) in response to different flow/shear stress levels. Increased flow/shear stress elevated the luminal ATP levels in PAs from control mice, but this response was absent in PAs from endothelial Panx1-/- mice (Figure 2G). Interestingly, flow/shear stress-induced increase in luminal ATP levels was not altered in PAs from endothelial TRPV4-/- mice (Figure 2H), suggesting that TRPV4 channels are not involved in flow-induced activation of endothelial Panx1.

7. Authors' measurements show that approximately 400 nM ATP is released from PA endothelial cells, but they have used 1 µM for dilation experiments (Figure 1B) and 10 µM for all other experiments, which is 6-100-fold higher concentration than the measured concentration. Authors needs to provide justification for that.

Thank you for raising this important issue. In our ATP assays (Figure 1B), we measured the ATP levels in the solution containing PAs. We postulated that the concentration of local ATP generated close to TRPV4 channels may be higher than that recorded in the solution. We used 1 μmol/L ATP for PA pressure myography, 10 μmol/L for patch-clamp experiments, and 1 μmol/L ATP for ca2+ imaging (apologies for the typo in the initial submission). 1 μmol/L ATP was sufficient to get a dilation in PAs and an increase in endothelial TRPV4 sparklet activity. For patch-clamp experiments in isolated endothelial cells, some cells responded (increase in TRPV4 currents) to 1 μmol/L ATP whereas others did not. However, 100% of the cells responded to 10 μmol/L ATP in patch-clamp experiments. This could be due to the fact that patch-clamp experiments were performed at room temperature and in enzymatically isolated ECs, which could make them slightly less sensitive to extracellular ATP.

8. Page 10: authors mentioned that PPADS and JNJ 47965567 did not inhibit TRPV4 sparkets, but Figure 2D shows significant inhibition which potentially suggest involvement of other purinergic (P2Xs) signaling.

We performed further statistical analyses in response to this comment. Our data showed that TRPV4 sparklet activity in the presence of ATP was not different amongst control, +PPADS, and + JNJ groups, although the data appeared to be trending towards a decrease in activity with PPADS/JNJ. To explore this possibility further, we increased the n numbers for PPADS and JNJ (previously n=3, Figure 3B). However, the difference was still not significant amongst the groups (n=5). Therefore, we concluded that in PAs, ATP activates TRPV4 sparklets mainly through P2Y2R activation. Also, please note that in endothelial P2Y2 receptor knockout mice, ATP was not able to activate TRPV4 sparklets (Figure 3A). While these data do not completely rule out a role for P2X signaling in ATP-activation of TRPV4 channels in PAs, they do confirm an essential role for endothelial P2Y2R.

9. Why did authors use different concentrations of activators such as GSK101 (1-30 nM) with different KO animals? Justification needed.

TRPV4 sparklet measurements in the intact endothelium at 37°C are more sensitive to GSK101 than patch-clamp experiments in enzymatically isolated ECs at room temperature. Therefore, the concentration of GSK101 is lower in calcium imaging experiments than patch-clamp experiments. For example, 1 nmol/L GSK101 does not activate TRPV4 currents in patch-clamp experiments but shows considerable TRPV4 sparklet activity in the intact endothelium. Below is a detailed description of the concentrations of GSK101 used and the underlying reasons:

1. For determining the effect of a treatment on TRPV4 sparklet activity, we compared baseline activity (in the absence of GSK101) or that in the presence of a low concentration of GSK101 (1 nmol/L). This level of sparklet activation makes it easier to discern a change of activity with a treatment or genetic deletion.

2. The maximum TRPV4 sparklet activity in PAs is seen at 30 nmol/L GSK1011. Therefore, this concentration of GSK101 was used to determine whether the maximum number of functional channels is altered by a genetic deletion.

3. For patch-clamp experiments performed in isolated ECs at room temperature, we used 10 nmol/L GSK101 to consistently get TRPV4 currents without getting maximal channel activation. Next, we used 100 nmol/L GSK101 to obtain near-maximal activation of the channel1 and to rule out the effect of genetic deletion on maximum functional channels.

10. Figure 5E, authors suggested that PAP regulation is via NO signaling, but they did not provide any experimental evidence using eNOS inhibitors or eNOS KO animals.

We apologize for not citing our previous studies on eNOS-NO signaling downstream of TRPV4 channels. We previously reported that endothelial TRPV4 sparklets dilate PAs via eNOS activation. Specifically, TRPV4 channel activation increased NO levels, an effect that was absent in PAs from eNOS-/- mice2. Moreover, TRPV4 channel-induced vasodilation was abolished by NOS inhibitor L-NNA. Also, in PAs from endothelial TRPV4-/- mice, endothelial NO levels were reduced1, 2. We have now cited these studies in the revised manuscript (lines 336 – 339).

11. Authors mentioned that they have used both Male and Female mice. Did they observe any differences between the sexes?

Previous studies reported lower TRPV4 channel activity in cerebral pial and parenchymal myocytes from female mice than male mice5. We used male and female mice for RVSP measurements and vasodilation, and did not observe any sex differences. Therefore, we now specify in the Methods section that no sex differences were observed (lines 377-378).

12. It is unclear why 1 nM GSK101 (Figure 1D) or 10 nM GSK101 (Figure 1G) induced reduced TRPV4 activity/current density in Panx1EC-/- compared to Panx1fl/fl. These observations do not support the statement on Page 6, Line 127-129, indicating that "reduced TRPV4EC channel activity in Panx1EC-/- is due to impaired channel regulation…". Since GSK101 is bypassing activation of the Panx1 pathway to stimulate TRPV4 channels directly, the expectation is that channel activity/current density should be similar provided that all other conditions are similar. A deeper consideration of this issue will help clarify concerns. The authors may consider performing immunofluorescence experiments to show that TRPV4-associated fluorescence or distribution of TRPV4 is similar in EC from Panx1fl/fl and Panx1EC-/-.

Higher concentrations of GSK101 were used to get maximal channel activity in ECs from PAs (30 nmol/L for calcium imaging and 100 nmol/L for patch-clamp1, 6), indicating maximum number of functional channels. Our data show that the maximum number of functional channels are not different between endothelial Panx1-/- and control mice (Supplemental Figure 1B). However, significant differences in activity were observed at lower levels of TRPV4 activation (1 nmol/L GSK101 for calcium imaging and 10 nmol/L for patch-clamp). Importantly, baseline (absence of GSK101) TRPV4 channel activity was also reduced in ECs from endothelial Panx1-/- mice (Figure 1D). Thus, Panx1 deletion reduced TRPV4 channel activity but not maximal functional channels. We agree that these data could also be explained by channel mis-localization and are not a direct evidence for “impaired channel regulation”. Therefore, we have removed the statement on impaired channel regulation in the revised manuscript.

13. The statement on Page 6, Line 131-132 is not supported by the data presented. Figure 1 does not show the direct regulation of TRPV4 by Panx1. This statement should be revised. The U466 experiments in Figure 1I are interesting but do not link TRPV4 and Panx1. These experiments show that genetic ablation of any of these proteins increases pulmonary artery reactivity to U466. Can the increase in contraction to U466 in Panx1EC-/- reversed by treating the arteries with the TRPV4 agonist?

We performed additional myography experiments, where U46619-induced constriction in PAs from endothelial Panx1-/- and P2Y2R-/- mice was studied in the presence of a low concentration of GSK101 (3 nmol/L) (Figure 2E). GSK101 reduced U46619-induced constriction to the control levels seen in PAs from endothelial Panx1fl/fl and P2Y2Rfl/fl mice. These data have been added to Figures 2E and 3H.

14. A major conclusion of the study is the formation of a nanocomplex between Cav-1, Panx1, P2Y2R, PKC and TRPV4. However, the interaction between these proteins is only presented for a subset of protein pairs. The study will be strengthened by providing data showing nanometer proximity between Panx1-P2Y2R, Panx1-TRPV4, P2Y2R-TRPV4 and that genetic ablation of Cav-1 disrupt the proximity between all these protein pairs. Based on the authors' data, P2Y1R should not activate TRPV4 channels. Thus, it will be expected that P2Y1R are not part of the complex. Is this the case? Please show antibody validation and negative controls for PLA. The authors should also explain why they think that Cav-1, Panx1, P2Y2R, PKC and TRPV4 are in nanometer proximity of each other given that no super-resolution data was presented.

PLA puncta indicate that two proteins are present within 40 nm of each other. In the initial version of this manuscript, we showed PLA data confirming the co-localization of endothelial Cav-1 with Panx1, P2Y2R, and TRPV4.

In response to the reviewer’s comment, we have now added PLA data demonstrating co-localization between TRPV4:P2Y2R and Panx1:P2Y2R (Figure 4E). Importantly, the PLA puncta were almost abolished when endothelial Cav-1 was genetically ablated, implying that endothelial Cav-1 is required for TRPV4:P2Y2R and Panx1:P2Y2R proximity. We also attempted PLA experiments for the one remaining protein pair- Panx1:TRPV4 using many different commercial antibodies. However, we could not draw meaningful conclusions due to non-specific signal with these new antibodies.

Our PLA experiments to detect potential co-localization between P2Y1R and Cav-1 (Supplemental Figure 4) demonstrated that P2Y1R does not co-localize with Cav-1. These data stand in agreement with the data that P2Y1R inhibition does not alter ATP activation of TRPV4 sparklets (Figure 1B).

The specificity of TRPV4, Panx1, P2Y2R, and Cav-1 antibodies was determined using PAs from the knockout mice as shown in Figure 1A, 3A, and Daneva et al., PNAS, 2021 1. The specificity of the PKC antibody was tested using a competing peptide, as described earlier7. The negative controls for PLA experiments are now provided in the form of loss of PLA signal in endothelial Cav-1-/- mice for the following pairs: (1) Cav1:TRPV4; (2) Cav-1:Panx1; and (3) Cav-1:P2Y2R. We have also provided negative controls for TRPV4:P2Y2R and Panx1:P2Y2R pairs as loss of signal in PAs from endothelial P2Y2R-/- mice the PLA experiments (Supplemental Figure 4).

15. Is ATP-induced dilation of small diameter pulmonary arteries prevented in Panx1EC-/-, P2Y2REC-/- and Cav-1EC-/-?

We conducted additional pressure myography experiments to address this comment. ATP-induced dilation was reduced in PAs from endothelial TRPV4-/-, P2Y2R-/-, and Cav-1-/- mice (Supplemental Figure 2C; Figure 3F and 4A). These data supported the idea that exogenous ATP dilates PAs through endothelial P2Y2R–TRPV4–signaling facilitated by Cav-1. ATP dilation, however, was not altered in PAs from endothelial Panx1-/- mice (Supplemental Figure 2D), suggesting that Panx1 is upstream of ATP-P2Y2R-TRPV4 signaling.

16. The description of Figure 2A gives the impression that perhaps all the ATP may come from EC. How much of the ATP will be left if the endothelial layer is removed before performing the assay? What are the ATP levels when using Panx1SMC-/- arteries? The study will be strengthened by linking mechanisms mediating basal ATP release with activation of the proposed pathway.

In the new Figure 1B, we demonstrated reduced ATP release in PAs from endothelial Panx1-/- mice compared to the control mice. We have now added data that endothelial denudation shows a similar decrease in ATP levels, suggesting that Panx1 is the predominant source of ATP release from ECs. Notably, the ATP levels in PAs from smooth muscle Panx1-/- mice were significantly higher than those in PAs from endothelial Panx1-/- mice, suggesting that the contribution of ECs to extracellular ATP is higher than that of SMCs. Finally, we show that ATP levels in endothelium-denuded PAs from smooth muscle Panx1-/- mice are lower than endothelium-denuded PAs from control mice. These data confirmed that both EC and SMC Panx1 releases ATP, although ECs appear to be the predominant source of ATP under basal conditions.

17. The sparklet traces shown in Figure 4D (recorded with X-Rhod in Cdh5-optoα1AR EC) is ascribed to TRPV4 channels, but there is no evidence that this is the case. This is important as data suggest that these sparklet events have a higher activity before light stimulation than other TRPV4 sparklet data presented throughout the manuscript. Are the amplitude and kinetics of sparklet events in this figure match that of known TRPV4 sparklets?

In the initial submission we only presented the number of TRPV4 sparklet sites per cell with X-Rhod-1. NPO could not be determined as the quantal level with X-Rhod-1 was not known. To address this concern from the reviewer, we first determined the quantal level of TRPV4 sparklets with X-Rhod-1 (0.21 ΔF/F0, presented in Supplemental Figure 5B). We are now able to present the X-Rhod-1 data as TRPV4 sparklet sites per field and sparklet activity (NPO) per site. Overall, initial TRPV4 sparklet activity was not different between fluo-4 and X-Rhod-1.

Further, in the presence of TRPV4 inhibitor GSK219 (100 nmol/L), light activation did not increase calcium signals (Supplemental Figure 5A), suggesting a specific effect on TRPV4 channels in PA endothelium.

18. Is TRPV4 current density diminished in Cav-1EC-/-?

We recently reported that pulmonary artery ECs from Cav-1EC-/- mice showe reduced TRPV4 current density compared to the ECs from control mice1. We now cite this study and have added the statement reflecting a decrease in TRPV4 current density in Cav-1EC-/- mice (lines 203-204).

19. Authors should show evidence that TRPV4 and Panx1 expression is reduced/knockout in TRPV4SMC-/- and Panx1SMC-/-, respectively.

We now provide evidence that Panx1 expression is reduced in our inducible, smooth muscle cell (SMC)-specific Panx1-/- mice (Supplemental Figure 1A). We previously provided evidence that the expression of TRPV4 channel in SMCs is reduced in SMC-specific TRPV4-/- mice1.

20. The immunofluorescent images in Figures 1A, 1C, 2C and 3A should be better described in the main text.

The immunofluorescent images in figures 1A and 3A have been described in more detail in the Results section (lines 108-109).

21. page 5, line 111 – I'm a bit puzzled why there was no right hypertrophy in your models with elevated PAP? Was it just that the duration of elevated PAP was insufficient to cause right heart hypertrophy? Please discuss.

The observation that there is no significant right ventricular hypertrophy in our genetic Panx1 and P2Y2R mouse models is in congruence with previous findings from our laboratory1. We agree with the reviewer that this could be due to a short duration of elevated PAP. All the experiments were performed two weeks after the last tamoxifen injection. While we get a PAP phenotype under these conditions, the duration of elevated PAP may not be sufficient to cause ventricular hypertrophy. This possibility has now been discussed (lines 286-290).

22. page 6, lines 125-127 – Were currents different between control and EC Panx1-/- with 30 nM GSK101 in your patch clamp experiments? Why were higher concentrations of GSK101 used in the patch clamp experiments?

We previously showed that 100 nmol/L GSK101 results in near-maximal activation of TRPV4 channel currents in pulmonary artery ECs1. We used two concentrations of GSK101 for our patch-clamp experiments: 10 nmol/L to get a low-level activation of TRPV4 channels; and 100 nmol/L to get near-maximal activation of TRPV4 channels1. 100 nmol/L GSK101 was used to determine the number of functional channels. We postulated that at lower levels of activity (10 nmol/L), the difference in TRPV4 currents due to genetic ablation of Panx1/P2Y2R will be more discernible. At a high concentration (100 nmol/L), the direct channel agonist will surpass the channel regulation by Panx1–ATP–P2Y2R signaling and result in maximal channel activation regardless of the absence of the regulatory proteins. Consistent with this, we previously showed that genetic deletion of endothelial Cav-1 results in reduced TRPV4 currents at 10 nmol/L GSK101 but not at 100 nmol/L GSK1011.

23. page 6, lines 129-131 – Did knock out of EC TRPV4-/- or EC Panx1-/- cause PAs to develop myogenic tone?

We performed PA pressure myography experiments to carefully test if endothelial knockout of Panx1, TRPV4, or P2Y2R results in increased myogenic constriction at 15 mm Hg. We show that PAs from endothelial Panx1, TRPV4, and P2Y2R knockout mice show higher myogenic constriction compared to PAs from the respective control mice (Figure 2B, 2D, and 3G). In combination with the data on U46619-induced constriction, these results support the concept that endothelial Panx1–P2Y2R–Cav-1–TRPV4 signaling axis lowers myogenic and agonist-induced constriction of PAs.

24. page 6 line 134 and onward – in your ex vivo ca2+ imaging experiments, what is the stimulus that is leading to Panx1 activity and release of ATP? It would seem important to identify the stimulus. Also, does luminal apyrase in pressure myograph experiments have the same effect as Panx1 knockout? How about P2Y2R-/- in pressure myography experiments? These experiments also would seem to be important to close the loop.

Our ATP measurements in isolated PAs (Figure 1B) show that ATP is being effluxed from EC Panx1 under resting/baseline conditions. We postulate that there is a similar basal ATP efflux through Panx1 in our imaging experiments, although the exact stimulus is not known. It is possible that Panx1 is activated by intracellular ca2+ or changes in membrane potential or membrane stretch. It is important to note that TRPV4 channels do not appear to be altering ATP efflux through Panx1 under basal conditions (Figure 1B).

Regarding physiological activators of Panx1, flow/shear stress has been shown to activate ATP efflux through Panx14. PAs are a high-flow vascular bed. However, flow/shear stress-activated signaling in PAs remains entirely unknown. In the revised manuscript, we provide evidence that flow/shear stress increases luminal ATP levels in PAs through endothelial Panx1 activation. Specifically, increased flow/shear stress elevated luminal ATP levels in pressurized PAs, and this effect was absent in PAs from endothelial Panx1-/- mice (Figure 2G). The shear stress is negligible in our calcium experiments (< 0.01 dynes/cm2). It is not clear if this level of shear stress can activate Panx1.

Furthermore, we conducted pressure myography experiments where PAs were treated with luminal apyrase (10 U/mL), followed by U46619. We observed that apyrase pretreatment increased the contractility of PAs from control mice (Supplemental Figure 2B) to the level of PA from endothelial Panx1-/- mice in absence of apyrase (Figure 2E), further supporting the dilatory effect of ATP effluxed through Panx1.

We have also added new experiments in PAs from endothelial P2Y2R-/- mice. First, we show that the myogenic constriction is increased in PAs from endothelial P2Y2R-/- mice compared to the control mice (Figure 3G). Next, we demonstrate that constriction to U46619 is significantly higher in PAs from endothelial P2Y2R-/- mice compared to the control mice (Figure 3H). These data are in agreement with the results that RVSP is elevated in endothelial P2Y2R-/- mice compared to the control mice (Figure 3E).

25. page 11, lines 236-239 – How do you reconcile the lack of effect of global TRPV4 knockout on PAP with your finding that EC TRPV4-/- increases PAP? Shoudln't global do the same thing? This should be better discussed.

Cell-specific knockout mice offer a significant advantage over global knockout mice with respect to discerning cell-type specific phenotype. For an endothelium-specific effect, the phenotype is expected to be better in an inducible, endothelium-specific knockout mouse than a global knockout mouse. Indeed, global TRPV4 knockout mice showed no systemic blood pressure or pulmonary arterial pressure phenotype8, 9. However, inducible, endothelium-specific knockout mice had elevated systemic blood pressure and pulmonary arterial pressure1. Lack of a phenotype in global knockout mice could be due to the deletion of TRPV4 channels from multiple cell types or compensatory mechanisms that have developed over time. These issues with the use of non-inducible, global knockout mice are well-documented in the literature. These possibilities have now been discussed (lines 275-281).

26. page 16, lines 351-353 – You did not cannulate the pressure catheter – you cannulated the external jugular vein for access to the right ventricle – please revise to clarify.

We thank the reviewer for this correction. This mistake has now been fixed (line 407-408).

27. Table 2 – How did you determine the specificity of the antibodies used?

The specificity of TRPV4, Panx1, P2Y2R, and Cav-1 antibodies was determined using PAs from the knockout mice as shown in Figure 1A, 3A, and Daneva et al., PNAS, 20211. The specificity of the PKC antibody was tested using a competing peptide, as described earlier7. The negative controls for PLA experiments are now provided in the form of loss of PLA signal in endothelial Cav-1-/- mice for the following pairs: (1) Cav1:TRPV4; (2) Cav-1:Panx1; and (3) Cav-1:P2Y2R. We have also provided negative controls for TRPV4:P2Y2R and Panx1:P2Y2R pairs as loss of signal in PAs from endothelial P2Y2R-/- mice the PLA experiments (Supplemental Figure 4).

28. Figure legends – please provide exact n-values for each panel and also please provide exact p-values for all statistical tests.

We have now added n-values and p-values for each panel in the figure legends.

1. Daneva Z, Marziano C, Ottolini M, Chen YL, Baker TM, Kuppusamy M, Zhang A, Ta HQ, Reagan CE, Mihalek AD, Kasetti RB, Shen Y, Isakson BE, Minshall RD, Zode GS, Goncharova EA, Laubach VE and Sonkusare SK. Caveolar peroxynitrite formation impairs endothelial TRPV4 channels and elevates pulmonary arterial pressure in pulmonary hypertension. Proceedings of the National Academy of Sciences of the United States of America. 2021;118.

2. Marziano C, Hong K, Cope EL, Kotlikoff MI, Isakson BE and Sonkusare SK. Nitric Oxide-Dependent Feedback Loop Regulates Transient Receptor Potential Vanilloid 4 (TRPV4) Channel Cooperativity and Endothelial Function in Small Pulmonary Arteries. J Am Heart Assoc. 2017;6.

3. Good ME, Eucker SA, Li J, Bacon HM, Lang SM, Butcher JT, Johnson TJ, Gaykema RP, Patel MK, Zuo Z and Isakson BE. Endothelial cell Pannexin1 modulates severity of ischemic stroke by regulating cerebral inflammation and myogenic tone. JCI Insight. 2018;3.

4. Wang S, Chennupati R, Kaur H, Iring A, Wettschureck N and Offermanns S. Endothelial cation channel PIEZO1 controls blood pressure by mediating flow-induced ATP release. J Clin Invest. 2016;126:4527-4536.

5. Tajada S, Moreno CM, O'Dwyer S, Woods S, Sato D, Navedo MF and Santana LF. Distance constraints on activation of TRPV4 channels by AKAP150-bound PKCalpha in arterial myocytes. J Gen Physiol. 2017;149:639-659.

6. Sonkusare SK, Bonev AD, Ledoux J, Liedtke W, Kotlikoff MI, Heppner TJ, Hill-Eubanks DC and Nelson MT. Elementary Ca2+ signals through endothelial TRPV4 channels regulate vascular function. Science. 2012;336:597-601.

7. Ottolini M, Hong K, Cope EL, Daneva Z, DeLalio LJ, Sokolowski JD, Marziano C, Nguyen NY, Altschmied J, Haendeler J, Johnstone SR, Kalani MY, Park MS, Patel RP, Liedtke W, Isakson BE and Sonkusare SK. Local Peroxynitrite Impairs Endothelial TRPV4 Channels and Elevates Blood Pressure in Obesity. Circulation. 2020.

8. Xia Y, Fu Z, Hu J, Huang C, Paudel O, Cai S, Liedtke W and Sham JS. TRPV4 channel contributes to serotonin-induced pulmonary vasoconstriction and the enhanced vascular reactivity in chronic hypoxic pulmonary hypertension. Am J Physiol Cell Physiol. 2013;305:C704-15.

9. Yang XR, Lin AH, Hughes JM, Flavahan NA, Cao YN, Liedtke W and Sham JS. Upregulation of osmo-mechanosensitive TRPV4 channel facilitates chronic hypoxia-induced myogenic tone and pulmonary hypertension. Am J Physiol Lung Cell Mol Physiol. 2012;302:L555-68.

https://doi.org/10.7554/eLife.67777.sa2

Article and author information

Author details

  1. Zdravka Daneva

    Robert M. Berne Cardiovascular Research Center, University of Virginia, Charlottesville, United States
    Contribution
    Data curation, Formal analysis, Investigation, Methodology, Validation, Writing – original draft, Writing – review and editing
    Competing interests
    none
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-1141-9697
  2. Matteo Ottolini

    1. Robert M. Berne Cardiovascular Research Center, University of Virginia, Charlottesville, United States
    2. Department of Pharmacology, University of Virginia, Charlottesville, United States
    Contribution
    Data curation, Formal analysis, Investigation, Methodology, Visualization
    Competing interests
    None
  3. Yen Lin Chen

    Robert M. Berne Cardiovascular Research Center, University of Virginia, Charlottesville, United States
    Contribution
    Data curation, Investigation, Methodology, Validation, Visualization
    Competing interests
    none
  4. Eliska Klimentova

    Robert M. Berne Cardiovascular Research Center, University of Virginia, Charlottesville, United States
    Contribution
    Investigation, Methodology, Validation
    Competing interests
    none
  5. Maniselvan Kuppusamy

    Robert M. Berne Cardiovascular Research Center, University of Virginia, Charlottesville, United States
    Contribution
    Data curation, Formal analysis, Investigation
    Competing interests
    none
  6. Soham A Shah

    Department of Biomedical Engineering, University of Virginia, Charlottesville, United States
    Contribution
    Investigation, Methodology
    Competing interests
    none
  7. Richard D Minshall

    Department of Anesthesiology, Department of Pharmacology, University of Illinois, Chicago, United States
    Contribution
    Methodology, Resources
    Competing interests
    none
  8. Cheikh I Seye

    Department of Biochemistry, University of Missouri-Columbia, Columbia, United States
    Contribution
    Methodology, Resources
    Competing interests
    none
  9. Victor E Laubach

    Department of Surgery, University of Virginia, Charlottesville, United States
    Contribution
    Funding acquisition, Resources, Writing – review and editing
    Competing interests
    none
  10. Brant E Isakson

    Department of Molecular Physiology and Biological Physics, University of Virginia, Charlottesville, United States
    Contribution
    Methodology, Resources
    Competing interests
    none
  11. Swapnil K Sonkusare

    1. Robert M. Berne Cardiovascular Research Center, University of Virginia, Charlottesville, United States
    2. Department of Molecular Physiology and Biological Physics, University of Virginia, Charlottesville, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration, Resources, Supervision, Validation, Visualization, Writing – original draft, Writing – review and editing
    For correspondence
    swapnil.sonkusare@virginia.edu
    Competing interests
    none
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9587-9342

Funding

National Institutes of Health (HL146914)

  • Swapnil K Sonkusare

National Institutes of Health (HL142808)

  • Swapnil K Sonkusare

National Institutes of Health (HL157407)

  • Victor E Laubach
  • Swapnil K Sonkusare

National Institutes of Health (P01HL120840)

  • Brant E Isakson

National Institutes of Health (HL137112)

  • Brant E Isakson

National Institutes of Health (R01HL133293)

  • Victor E Laubach

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

The mouse strain Cdh5-optoα1AR was developed by CHROMus, which is supported by the National Heart Lung Blood Institute of the National Institute of Health under award number R24HL120847. This work was supported by grants from the National Institutes of Health to SKS (R01HL142808, R01HL146914, R01HL157407), BEI (P01HL120840, HL137112), and VEL (R01HL133293, R01HL157407).

Ethics

All animal protocols were approved by the University of Virginia Animal Care and Use Committee (protocols 4100 and 4120). This study was performed in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. For surgical procedures, every effort was made to minimize suffering.

Senior Editor

  1. Richard W Aldrich, The University of Texas at Austin, United States

Reviewing Editor

  1. Mark T Nelson, University of Vermont, United States

Reviewer

  1. William Jackson, Michigan State University, United States

Publication history

  1. Received: February 22, 2021
  2. Preprint posted: March 9, 2021 (view preprint)
  3. Accepted: September 6, 2021
  4. Accepted Manuscript published: September 7, 2021 (version 1)
  5. Version of Record published: September 17, 2021 (version 2)

Copyright

© 2021, Daneva et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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