Zinc regulates vascular endothelial cell activity through zinc-sensing receptor ZnR/GPR39

Abstract

Zn²⁺ is an essential element for cell survival/growth, and its deficiency is linked to many disorders. Extracellular Zn²⁺ concentration changes participate in modulating fundamental cellular processes such as proliferation, secretion, ion transport, and cell signal transduction in a mechanism that is not well understood. Here, we hypothesize that the Zn²⁺-sensing receptor ZnR/G protein-coupled receptor 39 (GPR39), found in tissues where dynamic Zn²⁺ homeostasis takes place, enables extracellular Zn²⁺ to trigger intracellular signaling pathways regulating key cell functions in vascular cells. Thus, we investigated how extracellular Zn²⁺ regulates cell viability, proliferation, motility, angiogenesis, vascular tone, and inflammation through ZnR/GPR39 in endothelial cells. Knockdown of GPR39 through siRNA largely abolished Zn²⁺-triggered cellular activity changes, Ca²⁺ responses, as well as the downstream activation of Gαq-PLC pathways. Extracellular Zn²⁺ promoted vascular cell survival/growth through activation of cAMP and Akt as well as overexpressing of platelet-derived growth factor-α receptor and vascular endothelial growth factor A. It also enhanced cell adhesion and mobility, endothelial tubule formation, and cytoskeletal reorganization. Such effects from extracellular Zn²⁺ were not observed in GPR39^−/− endothelial cells. Zn²⁺ also regulated inflammation-related key molecules such as heme oxygenase-1, selectin L, IL-10, and platelet endothelial cell adhesion molecule 1, as well as vascular tone-related prostaglandin I2 synthase and nitric oxide synthase-3. In sum, extracellular Zn²⁺ regulates endothelial cell activity in a ZnR/GPR39-dependent manner and through the downstream G_αq-PLC pathways. Thus, ZnR/GPR39 may be a therapeutic target for regulating endothelial activity.

INTRODUCTION

Zn²⁺ is an essential supplementary nutrient for health and is involved in structural and regulatory cellular functions (11, 24). It interacts with Zn-finger domains and acts as a cofactor for hundreds of enzymes (5, 50). It also binds to numerous membrane receptors, transporters, and channels, thereby regulating their activity (20). Zn²⁺ deficiency is linked to many disorders involving different organs and tissues, such as digestive, immune, neuronal, and vascular systems (43, 50, 53). Severe lack of Zn²⁺ may lead to attenuation of growth and sexual development (50). However, an excess of extracellular Zn²⁺ is toxic; for example, a massive release of synaptic Zn²⁺ induces neuronal cell death in brain ischemia (3, 7, 22). Moreover, dramatic changes in intracellular and extracellular Zn²⁺ concentrations occur during various pathophysiological syndromes such as myocardial infarction, hepatic renal failure, and neoplastic processes (52). Despite the important role of Zn²⁺ in cellular function and the fluctuations in Zn²⁺ concentrations, little is known about the cellular signaling mechanisms regulated by Zn²⁺ and its effect on cell functions and fate in endothelial cells (26, 28).

In vertebrate animals and human beings, endothelial cells are the main vascular cells for blood vessels (33). These cells are involved in many aspects of vascular biology; among them, major functions of vascular cells include formation and repair of blood vessels (angiogenesis) (56), vasoconstriction and vasodilation (vascular tone) (32), and inflammation (35, 49), as well as blood clotting and barrier functions (54), etc. Although Zn²⁺ plays an essential role in endothelial cell health and functions, the link between extracellular Zn²⁺ and cellular signaling pathways involved in these vascular functions is not well understood.

Recently, Zn²⁺ has emerged as an important signaling molecule, activating intracellular pathways and regulating cell fate in various cell types like epithelium and colonocyte. A Zn²⁺-sensing receptor (ZnR), also known as the G protein-coupled receptor GPR39, was identified, which is a Gq-coupled receptor mediating changes in extracellular Zn²⁺ and major intracellular signaling pathways (2, 9, 10, 13, 42, 44–46). It triggers inositol 1,4,5-trisphosphate-dependent release of intracellular Ca²⁺, leading to activation of mitogen-activated protein (MAP) and phosphoinositide 3 (PI3) kinase pathways. Moreover, GPR39 knockout mice exhibited symptoms of Zn²⁺-deficiency, including depression, accelerated gastric emptying, and increased fecal excretion (30, 31). However, how Zn²⁺ or ZnR/GPR39 signaling regulates endothelial activity is not well understood.

In the present work, we investigated the role of ZnR/GPR39 in Zn²⁺-regulated endothelial activity related to angiogenesis, endothelial permeability, vascular tone, and inflammation. They are endothelial cell viability, proliferation, mobility, and tubule formation as related to angiogenesis and regulation of key molecules involved in vasoconstriction and vasodilation as related to vascular tone, as well as expression and secretion of key molecules involved in inflammation. The findings herein provide evidence for a specific Zn²⁺-sensing mechanism involving cellular Ca²⁺ and PI3 signaling and, hence, controlling key vascular cellular processes.

MATERIALS AND METHODS

Solution preparation.

ZnCl₂ solutions (10 mM) were prepared by dissolving ZnCl₂ (Sigma-Aldrich) into deionized water and adjusting to neutral pH by NaOH. Solutions were filtered by 0.22-μm filter (Thermo Fisher Scientific) and autoclaved (Harvey Sterile Max, Thermo Scientific). ZnCl₂-supplemented medium (pH 7.2) were prepared by diluting ZnCl₂ stock solution with cell culture medium (26–28, 59). A physiologically relevant concentration of 25 µM was used as Zn²⁺ treatment group, as the serum concentration of Zn is in the range of 10–30 µM (14). Typical cell culture medium DMEM (Sigma-Aldrich) has no or only a trace amount of Zn²⁺ (<0.7 pM) so was used as the negative control (Zn²⁺ deficiency control).

Cell culture.

Primary human coronary artery endothelial cells (HCAECs) were ordered from ATCC. Three batches of cells from different healthy donors were used. The components of cell culture medium were basal medium, 5% fetal bovine serum, 1% endothelial cell growth supplement, and 1% penicillin-streptomycin. HCAECs were cultured in a flask (Falcon, BD Bioscience) until ~90% confluence was reached. Then cells were detached by trypsin-EDTA (Life Technologies), centrifuged (Sorvall Biofuge Stratos, Thermo Electron), and resuspended. The cell number was counted with a cell counter (Bio-Rad). Cells at passages 3–5 were used (26–28, 59). For all cell treatment experiments, cells were starved overnight in serum-free medium and then treated with different concentrations of Zn²⁺ in basal DMEM medium without serum.

Mouse GPR39^−/− endothelial cells.

Mouse GPR39^−/− endothelial cells were a gift from Capital Medical University. The endothelial cells were >99% pure, as measured by staining for the von Willebrand factor (endothelial cells) and negative immunostaining for smooth muscle cell α-actin (25). Cells at passages 3–5 from six different batches were used.

Cell viability.

Cells were seeded in a 96-well cell culture plate (BD Biosciences) with 5,000 cells per well for 24 h to allow cell attachment. Medium was replaced by medium supplemented with different solutions and incubated for 24 h. Medium with 10% DMSO (Life Technologies) and medium alone were positive and negative controls, respectively. Another blank reference containing the same concentration of Zn ion solution without cells was used to exclude the interference of the ions. The 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrezolium bromide (MTT, Invitrogen) test was performed according to the manufacturer's protocol. Absorbance was measured at 570 nm, using a microplate reader (SpectraMax, Molecular Devices) (26–28, 59).

Cell proliferation.

A BrdU cell proliferation kit (Cell Signaling) was used for the cell proliferation test. Cells were seeded in a 96-well cell culture plate at 5,000 cells per well. After 24 h, medium was replaced by different solutions and incubated for 24 h to allow cell growth and doubling. The proliferation test was performed according to the manufacturer's protocol. Absorbance was measured at 450 nm. Positive control and negative control were medium without Zn ion supplement and medium without cells, respectively (26–28, 59).

Cell adhesion.

Cells were seeded onto a 24-well plate (Falcon, Corning). The final cell density was 50,000 cells per well. Cells were incubated at 37°C, 5% CO₂, and 95% relative humidity for 5 h with the treatment of Zn²⁺. The time of 5 h should be sufficient for complete attachment of cells on a culture surface without cell division/growth during attachment. Then, the cell medium was removed, and cells were washed three times with Dulbecco’s phosphate-buffered saline(DPBS). Images of adhered cells were taken with a microscope (EVOS FL Cell Imaging System, AMG). The plate was sealed with self-sticking tape (Fisherbrand, Fisher Scientific). Then, the plate was put into a rotor inversely and centrifuged at 500 rpm for 5 min. Cells were washed with DPBS and fixed by 4% paraformaldehyde (Boston BioProducts). The images of the adhered cells were taken with a microscope (EVOS FL Cell Imaging System, AMG) and analyzed with ImageJ (National Institutes of Health, Bethesda, MD). At least 10 different fields were used for calculating adhered cell density and cell retention ratio (26–28, 59).

Cell migration.

Cells were seeded on a 12-well cell culture plate (BD Biosciences). The cytostatic agent hydroxyurea (250 µM) was added into culture medium to inhibit cell division/growth during migration. A straight line in a cell monolayer was created by scratching the surface using a p200 pipette tip (Thermo Scientific). Debris was removed by gently washing three times with DPBS, and cells were incubated with 3 ml of medium supplemented with different ion solutions. At 0, 6, and 24 h, optical images were taken using a phase contrast microscope (Advanced Microscopy). The width of the line at top, middle, and bottom positions was measured in Image-Pro Plus 6.0 (Media Cybernetics). The average cell migration rate was calculated as described before (26–28, 59).

Tubulogenesis assays.

Endothelial tubular differentiation assays were carried out according to a previously published method with some modifications (1). Briefly, cells (3,000 per sample) in phenol red-free medium containing 2% FCS were plated in 24-well plates precoated with 250 μl of growth factor-reduced, phenol red-free Matrigel (BD Biosciences) for 1 h at 37°C. After treatment, cells were incubated in a humidified incubator at 37°C and 5% CO₂. After 16 h, bright-field images were captured from five random fields of view per sample at ×10 magnification. Tubule length and number of branching points for each condition were quantified using Image Pro-Plus 6.0 software.

RNA extraction.

Cells were seeded in 100-mm culture dishes (BD Technologies) and allowed to attach for 24 h. Then the cells were treated with medium and medium supplemented with ZnCl₂, respectively, for 24 h. Cells were harvested, and total RNA was extracted using an RNeasy Mini Kit (Qiagen) and subsequently quantified using a spectrophotometer (Nanodrop 2000) with OD260/OD280 ratios between 1.9 and 2.1 (26–28, 59).

cDNA synthesis.

A total of 600 ng of RNA was used for reverse transcription using an RT2 First Strand Kit (Qiagen). Reverse transcription was performed in a thermo cycler (T100, Bio-Rad). Then, 91 μl of RNase-free water was added to the 20 μl of cDNA mix and stored at −20°C in a freezer (Puffer Bubbard, Thermo Scientific) (26–28, 59).

Real-time-PCR.

Gene expression analysis was performed in a CFX96 Touch RT-PCR Detection System (Bio-Rad) using the RT2 Profiler PCR array (Qiagen). The array includes 84 functional genes, five housekeeping genes, three reverse-transcription controls, and three positive PCR controls. A total of 25 μl of PCR components mix including cDNA, SYBR Green Mastermix, and RNase-free water was dispensed to the RT2 Profiler PCR Array plate. After initial heat activation (95°C, 10 min), cDNA was amplified as the following parameters: 95°C for 15 s and 60°C for 1 min. After the amplification, melting-curve analysis was performed using the default melting curve program. Only the genes with a single melting peak were chosen for final analysis. Data were analyzed by Bio-Rad CFX Manager 3.1 (Bio-Rad). The 2−ΔΔC_T method was used to calculate gene fold changes. Real-time PCR was also used to estimate the mRNA levels of ZnR/GPR39. The level of target mRNA was normalized to the endogenous control β-actin. Primers and probes were from Dharmacon RNAi Technologies (Thermo Scientific): for GPR39, forward primer CATCTTCCTGAGGCTGA, reverse primer ATGATCCTCCGTCTGGTTG, probe TATGCTGGATGCCCAAC; and for actin, forward primer TGGAGAAAATCTGGCACCAC, reverse primer GGTCTCAAACATGATCTGG, probe ACCGCCAGAAGATGACC (26–28, 59).

Gene silencing.

Cells were seeded in 60-mm cell-culture dishes (density of 7 × 10⁵ cells/plate) or in 96-well plates (1.5 × 10⁴ cells/well) 24 h before transfection and maintained in standard culture medium as described above but without penicillin or streptomycin. Cells were transfected using Lipofectamine 2000 (Invitrogen) according to the manufacturer’s protocol and used 48 h posttransfection. Small interfering RNAs (siRNA) targeting human GPR39 (5′-CCAUGG AGUUCUACAGCAU-3′) were purchased from Santa Cruz Biotechnology. The corresponding scrambled siRNAs were used as controls (61).

Calcium response through imaging.

Cells were incubated with 3 µM Fura 2-acetoxymethyl ester for 30 min in Ringer’s solution with 0.1 BSA. After dye loading, the cells were washed in Ringer’s solution [sodium chloride (8.6 g/l), potassium chloride (0.3 g/l), and calcium chloride dihydrate (0.33 g/l)], and coverslider chambers were mounted into the microscope for imaging. All treatments with Zn²⁺ were applied in the absence of serum to avoid chelation of this ion. Fura 2 was excited at 340 and 380 nm and imaged with a 510-nm filter. Fura 2 fluorescence change was determined by the rate of initial normalized fluorescence change (ΔR/s) as previously described (2).

Western blot analysis.

Cells were washed twice with ice-cold PBS, followed by the addition of 200 μl of cell lysate medium. After collection of the cell lysate, protein concentrations were determined using the Bradford assay (Bio-Rad). Equal amounts of protein for each sample were applied to SDS-PAGE (10% gels). After electrophoresis, proteins were transferred onto nitrocellulose membranes (0.45 μm, Bio-Rad) and the membranes subsequently blocked with 5% (wt/vol) nonfat dried skimmed milk powder-TBST (TBS containing 0.05% Tween 20) for 1 h at room temperature. Primary antibodies against protein of interest and corresponding secondary antibodies conjugated to horseradish peroxidase (HRP) were applied. To confirm equal protein loading, membranes were stripped at 60°C for 30 min with a stripping buffer. The immunoblots were reblocked in 5% (wt/vol) nonfat dried skimmed milk powder-TBST followed by incubation with antibodies that react with β-actin, or total forms of proteins of interest. Bands were visualized using SuperSignal West Pico chemiluminescence detection reagents from Pierce Biotechnology (61).

Akt serine phosphorylation.

Cells were lysed using RIPA buffer after specified treatment. Proteins (20 µg) were analyzed by 4–15% Tris·HCl gel and transferred to nitrocellulose membranes (0.45 µm, Bio-Rad). The membranes were incubated overnight with primary rabbit polyclonal anti-mouse phospho-Akt (Ser⁴⁷³) antibody or rabbit polyclonal anti-mouse Akt antibody diluted in 5% nonfat milk or 5% BSA in TBS and then incubated with the corresponding secondary HRP-conjugated antibodies for 1 h. Immunoreactivity was detected using the ECL detection system (Amersham, Piscataway, NJ) (61).

cAMP direct immunoassay.

The concentration of cAMP was measured following the instructions of the cAMP Direct Immunoassay Kit (Abcam). Briefly, the cultured cells were scraped and dissociated completely and centrifuged at 14,000 g for 10 min to collect the supernatant as the testing sample. After being neutralized and acetylated with acetylating reagent and neutralizing buffer, respectively, 50 μl of standard cAMP (or testing samples were added to a G protein-coated 96-well plate and incubated with 10 μl of cAMP antibody at room temperature for 1 h with gentle agitation. Then 10 μl of cAMP-HRP was added, and the plates were incubated for another hour. The suspension was discarded, and the cells in the wells were washed with 1× cAMP assay buffer five times. The detecting reaction was conducted by incubating the cells with 100 μl of HRP for 1 h and stopped by adding 100 μl of 1 M HCl. Then, the reaction was checked by a microtiter plate reader at 450 nm. The absorbance of the substrate was also detected as background absorbance and subtracted from all standards and samples. The molar concentration of cAMP in cell pellets was determined from standard curves (60).

Immunofluorescence and cytoskeleton staining.

Cells were seeded in a 12-well cell culture plate and treated with medium supplemented with ZnCl₂ for 24 h. An Image-iT Fix-Perm kit (Invitrogen) was used to fix cells. F-actin was stained with Actin Green 488 Ready Probes Reagent (Invitrogen). The cell nucleus was stained with the SlowFade Gold Anti-fade Reagent with DAPI (Invitrogen). The GPR39 was stained with rabbit anti-GPR39 antibody (Abcam) followed by Alexa Fluor 488 mouse anti-rabbit IgG (Invitrogen). Images were taken using an EVOS inverted fluorescent microscope (Advanced Microscopy). Fluorescence intensity of the cells was extracted by using ImageJ (version 1.49 software). Contrast of the representative images was autoadjusted using Image-Pro Plus 6.0 (60, 61).

ELISA analysis of proteins.

Cells were seeded with 4 ml of culture medium into a 60-mm petri dish (Primaria, Corning). After ~90% confluence was reached, cell culture medium was replaced by different treatment solutions. After 24 h, a 4-ml treatment solution was collected and centrifuged at 20,000 g. The supernatants were collected and aliquoted into 500-μl tubes. Then, 200 μl of cell lysis buffer with 1× protease, and phosphatase cocktail (Thermo Fisher Scientific) was added to the petri dish and the dish was put on ice. After a while, cell lysate was collected by cell scraper (Thermo Fisher Scientific) and centrifuged at 20,000 g. The supernatants were also collected and aliquoted. All samples were stored at −80°C until use. All ELISA reagents were bought from R&D Systems. All assays were performed according to their protocol. The absorbances were measured with a microplate reader (Molecular Devices) (26–28, 59).

Transendothelial electrical resistance.

Cells were cultured on inserts with collagen-coated polycarbonate in a transwell. The transwell had membrane filters with 0.4-µm pores (Corning Life Science, Lowell, MA). Transendothelial electrical resistance (TER) was measured using an Endohmeter (World Precision Instruments, Sarasota, FL). TER values are expressed in Ω·cm². The resistance of collagen-coated inserts was subtracted from the resistance obtained in the presence of the endothelial cultures (61).

Permeability of the endothelial barrier.

We used fluorescein-isothiocyanate (FITC)-labeled dextran (40 kDa, Invitrogen) to measure permeability (61). Briefly, the medium in the upper chamber of an established monolayer was replaced with 500 µl of FITC-labeled dextran solution (2 mg/ml) in phenol-red-free DMEM. The lower chamber containing, 1,500 µl of phenol-red-free DMEM was sampled (20 µl) at 5-min intervals for 30 min. At each time point, 20-µl samples were taken from the upper chamber as well. The fluorescence intensity of each sample was determined using a fluorescence multiwell plate reader (PerkinElmer Victor 3) at excitation/emission wavelengths of 490/525 nm. The corresponding FITC-labeled dextran concentrations were determined from the standard curve. The endothelial monolayer permeability to dextran was expressed as a permeability coefficient in cm/s.

Statistical analysis.

All data are presented as means ± SD (n ≥ 5). One-way or two-way analysis of variance (ANOVA) followed by Tukey's post hoc test was used to determine statistically significant differences. An unpaired Student's t-test was also used to compare differences between two groups as appropriate. P < 0.05 was considered statistically significant.

RESULTS

Zn²⁺ stimulates intracellular Ca²⁺ release and activates downstream pathway mediated by ZnR/GPR39.

To determine the role of ZnR/GPR39, cells were transfected with siRNA against GPR39 to knock down its expression. We controlled GPR39 expression in endothelial cells as shown, with RT-PCR (Fig. 1A), Western blotting (Fig. 1B), and immunofluorescent staining (Fig. 1C) using siGPR39 but not scramble control. GPR39 appeared to be located largely on the cell membrane (Fig. 1C). The normal serum Zn²⁺ concentration is ~10–30 µM (14); thus, we used a Zn²⁺ concentration of 25 µM, which is in the vicinity of physiological condition, to stimulate the cells. Extracellular Zn²⁺ induced a significant Ca²⁺ response (Fig. 1D). Silencing of GPR39 was followed by a diminished Ca²⁺ response triggered by Zn²⁺. To further verify that the response was triggered via the G_αq-PLC pathway, as suggested before (2, 57), we used pharmacological inhibitors and monitored the Zn²⁺-dependent Ca²⁺ response. Inhibition of the G_αq using YM254890 (1 µM) blocked the Zn²⁺-dependent Ca²⁺ response almost completely (Fig. 1E). The PLCβ inhibitor u73122 (10 µM) largely abolished the signaling triggered by Zn²⁺, yet to a lower extent than the G_αq inhibitor or due to the fact that the PLC is downstream from the G_αq. Thus, inhibition of the latter suppressed the Ca²⁺ response more efficiently.

Fig. 1.

Zn²⁺ triggers Ca²⁺ response and downstream signaling through ZnR/GPR39 in endothelial cells. A: RT-PCR analysis of GPR39 expression in endothelial cells treated with different siRNA. B: Western blots showing the expression level of GPR39 in endothelial cells with different siRNA treatment. C: representative immunofluorescent images showing GPR39 expression in endothelial cells treated with different siRNA. (Scale bar = 20 µm.) D: rate of Ca²⁺ response from measurement of fluorescence change with Zn²⁺ (25 µM) treatment. E: rate of Ca²⁺ response from measurement of fluorescence change with treatment of Zn²⁺ (25 µM) and pharmacological inhibitors of G_αq and PLC. *P < 0.01 vs. controls at normal conditions in endothelial cells (n = 6).

Zn²⁺ promotes vascular cell viability and proliferation via ZnR/GPR39.

The effect of Zn²⁺ on endothelial survival and growth was tested by measuring cell viability and proliferation. Cell viability analysis using MTT revealed that Zn²⁺ significantly increased the survival rate of endothelial cells, and knockdown of GPR39 using siRNA dramatically counteracted such an effect induced by Zn²⁺ on cell viability (Fig. 2A). A similar result was observed for cell proliferation (Fig. 2B). Next, to further understand the mechanism of Zn²⁺ on cell survival/growth, we examined the intracellular cAMP level and the activation of the Akt pathway because they are the important modulators in cell viability. The level of cAMP was significantly enhanced after Zn²⁺ treatment (Fig. 2C), and Zn²⁺ also induced significant phosphorylation of Akt (Fig. 2D).

Fig. 2.

Zn²⁺ regulates endothelial cell viability and proliferation via ZnR/GPR39. A: cell viability of endothelial cells with Zn²⁺-deficient condition and Zn²⁺ (25 µM) treatment for 24 h. B: cell proliferation of endothelial cells with Zn²⁺-deficient condition and Zn²⁺ (25 µM) treatment for 24 h. C: intracellular cAMP level in endothelial cells with Zn²⁺-deficient condition and Zn²⁺ (25 µM) treatment for 24 h. D: Akt phosphorylation after Zn²⁺ (25 µM) treatment for 24 h in endothelial cells. *P < 0.05 vs. controls at Zn²⁺-deficient conditions in endothelial cells (n = 6).

Zn²⁺ promotes endothelial cell adhesion, mobility, and cytoskeletal reorganization and reduces transendothelial permeability via ZnR/GPR39.

Cell motility plays a pivotal role in various cellular functions and activity, such as division, migration, contraction, and angiogenesis. To elucidate the effect of Zn²⁺ on cell motility, we measured cell adhesion and mobility with and without extracellular Zn²⁺. The adhered cell density of endothelial cells increased ~30% after being treated with Zn²⁺ for 5 h compared with negative controls (Fig. 3A). Transfection with GPR39 siRNA significantly abolished such enhanced adhesion induced by Zn²⁺. Similar results were observed for cell migration rate (Fig. 3B). Zn²⁺ significantly increased cell migration rate, whereas knockdown of GPR39 by siRNA blocked such effects of Zn²⁺ on cell motility. Since cell movement is largely facilitated by cytoskeletal rearrangement, we next quantitatively measured the amount of F-actin stress fibers. Zn²⁺ treatment significantly promoted F-actin polymerization, leading to the emerging of F-actin stress fibers, as shown by representative images of endothelial cells (Fig. 3C). Quantitative analysis of F-actin fluorescence intensity revealed that significantly more F-actin appeared after Zn²⁺ treatment in cells, and siRNA knockdown of GPR39 abolished this phenomenon, as observed (Fig. 3D). Zn²⁺ treatment also significantly reduced transendothelial permeability but not for GRP39 knockdown cells, as shown by transendothelial electrical resistance (TER) (Fig. 3E) and permeability to dextran-40 (Fig. 3F).

Fig. 3.

Zn²⁺ promotes endothelial cell adhesion and mobility and cytoskeletal organization and reduces transendothelial permeability. A: normalized cell adhesion density after 5 h with or without Zn²⁺ (25 µM) and siRNA treatment. B: cell migration rate (µm/h) after 5 h with or without Zn²⁺ (25 µM) and siRNA treatment. C: representative image of F-actin fibers in endothelial cells with or without Zn²⁺. (Scale bar = 5 µm.) D: quantitative measurement of F-actin fluorescence density in endothelial cells. E: transendothelial electrical resistance (TER) with or without Zn²⁺ (25 µM) treatment. F: transendothelial permeability with or without Zn²⁺ (25 µM) treatment. *P < 0.05 vs. controls at Zn²⁺-deficient conditions in endothelial cells (n = 6).

Zn²⁺ promotes angiogenesis via ZnR/GPR39.

To explore the role of Zn²⁺ in angiogenesis, we examined endothelial tubule formation on growth factor-reduced Matrigel. Zn²⁺ treatment induced significantly more endothelial tubular differentiation compared with control groups without Zn²⁺ or with siGPR39 (Fig. 4A). Both endothelial cord length and branches with Zn²⁺ treatment were 30–40% more than negative controls and groups with siGPR39 (Fig. 4, B and C).

Fig. 4.

Zn²⁺ promotes vascular tubule formation. A: representative images showing tubule formation with or without Zn²⁺ (25 µM) and siRNA treatment. (Scale bar = 100 µm.) B: quantitative measurements of tubule length. C: quantitative measurements of tubule branching points. *P < 0.05 vs. controls at Zn²⁺-deficient conditions in endothelial cells (n = 6).

Zn²⁺ has no effect on endothelial activity in GPR39 knockout cells.

To further validate the role of GPR39 in extracellular Zn²⁺ sensing and binding, we investigated the effects of Zn²⁺ in mouse GPR39^−/− endothelial cells. Immunofluorescent staining of GPR39 verified the absence of GPR39 in the knockout GPR39^−/− cells (Fig. 5A). Addition of extracellular Zn²⁺ did not induce significant changes of Ca²⁺ signaling, cell survival/growth, cellular cAMP level, or cell adhesion and mobility in GPR39^−/− cells (Fig. 5, B–D). Similarly, no significant changes of transendothelial permeability or angiogenesis were observed after Zn²⁺ treatment in GPR39^−/− cells (Fig. 5, E and F).

Fig. 5.

Effects of Zn²⁺ on endothelial cell activity with knockout of GPR39. A: representative images of GPR39 staining and nuclei in wild-type (WT) and GPR39 knockout endothelial cells (GPR39^−/−). (Scale bar = 20 µm.) B: rate of Ca²⁺ response from measurement of fluorescence change with Zn²⁺ (25 µM) treatment in WT and GPR39^−/− cells. C: cell viability and cAMP activity of WT and GPR39^−/− cells with Zn²⁺ (25 µM) treatment. D: cell adhesion density and mobility of WT and GPR39^−/− cells with Zn²⁺ (25 µM) treatment. E: TER and permeability of WT and GPR39^−/− monolayer with Zn²⁺ (25 µM) treatment. F: vascular tubule formation of WT and GPR39^−/− cells with Zn²⁺ (25 µM) treatment. (Scale bar = 100 µm.) *P < 0.05 vs. GPR39^−/− endothelial cells (n = 6).

Effects of Zn²⁺ on expression of key molecules related to typical endothelial activity.

To gain more information on molecular mechanisms behind the observed effects of Zn²⁺ on endothelial cells, we next studied the expression of key molecules related to typical endothelial activity at both gene and protein levels, using RT-PCR and ELISA, respectively.

To further understand the beneficial effect of Zn²⁺ on vascular cell survival and growth, we examined the related expression of platelet-derived growth factor-α receptor (PDGFRA) and vascular endothelial growth factor A (VEGFA). PDGFRA is a cell surface tyrosine kinase receptor for members of the platelet-derived growth factor family, and VEGFA is the main, dominant inducer to the growth of vascular endothelial cells. Extracellular Zn²⁺ significantly enhanced the expression of these genes and proteins involved in cell survival, apoptosis, and proliferation (Fig. 6A). Inhibition of ZnR/GPR39 expression abolished the enhanced expression of these molecules induced by Zn²⁺.

Fig. 6.

Effects of Zn²⁺ on genes and proteins related to vascular cell survival/growth and inflammation and vascular tone. A: mRNA and protein expression levels of PDGFRA and VEGFA in endothelial cells with treatment of Zn²⁺ (25 µM) and siGPR39. B: mRNA and protein expression levels of HMOX1, IL-10, SELL, and PECAM1 in endothelial cells with treatment of Zn²⁺ (25 µM) and siGPR39. C: mRNA and protein expression levels of PTGIS and NOS3 in endothelial cells with treatment of Zn²⁺ (25 µM) and siGPR39. *P < 0.01 vs. controls at Zn²⁺-deficient conditions for mRNA levels (n = 6). #P < 0.05 vs. controls at Zn²⁺-deficient conditions for protein levels (n = 6).

It is known that Zn²⁺ may participate in cell signaling regulation of vascular inflammation (28, 29). We thus studied the gene and protein levels of related genes after treatment of extracellular Zn²⁺. We found that both genes and proteins of heme oxygenase-1 (HMOX1) and IL-10 were significantly upregulated, whereas those of L-selectin (SELL) and platelet endothelial cell adhesion molecule (PECAM1) were significantly downregulated (Fig. 6B). Moreover, no significant regulations of intercellular adhesion molecule 1 (ICAM1) and C-C motif chemokine ligand 2 (CCL2) at the protein levels were observed despite the significant downregulation of the corresponding genes (data not shown). The levels of these inflammatory molecules were restored to normal if ZnR/GPR39 was silenced.

Another important function of vascular cells is vasoconstriction and vasodilation. To examine the role of Zn²⁺ in regulation of such vascular activity, we measured related gene and protein expression. Prostaglandin I2 synthase (PTGIS) and nitric oxide synthase-3 (NOS3) were significantly upregulated at both mRNA and protein levels (Fig. 6C). Blockage of Zn²⁺ signaling by knockdown of its receptor ZnR/GPR39 inhibited such effects induced by Zn²⁺.

DISCUSSION

Zn²⁺ is essential for cell survival and functions in many cells. Changes in Zn²⁺ concentration have a fundamental effect on numerous pathophysiological processes. The sensing and signaling of these changes may have an important homeostatic role. Moreover, many studies have underscored the importance of extracellular Zn²⁺ in cellular signaling associated with survival and proliferation (29). For instance, Zn²⁺ was linked to regulation of the mitogen-activated protein kinase (MAPK) and PI3 pathways. The specific receptor for extracellular Zn²⁺, ZnR/GPR39, is functional in various cell types, such as colonocytes, keratinocytes, neurons, pancreatic β cells, and salivary glands (2, 9, 10, 13, 42, 44–46). Here, we have shown that ZnR/GPR39 is essential for mediating Zn²⁺-dependent cellular signaling pathways leading to survival, proliferation, angiogenesis, inflammation, vascular tone, and other vascular activity. Silencing or knockout of ZnR/GPR39 diminished the Zn²⁺-dependent effect significantly but not completely, suggesting the possible existence of other putative Zn²⁺ receptor, e.g., transient receptor potential ion (melastatin) channel (TRPM)6/7 in vascular cells (21, 55, 58). TRPM6/7 is a ubiquitous membrane channel for cations including Ca²⁺, Mg²⁺, and Zn²⁺, etc. Although it is suggested that extracellular Zn²⁺ levels are low, Zn²⁺ released from the liver or in digested food may be a source for Zn²⁺ signaling (23). Moreover, Zn²⁺ is secreted by healthy epithelial cells with a robust release during epithelial injury (42). In addition, accumulation of Zn²⁺ happens in colonocytes triggered by oxidative stress (8). Interestingly, numerous Zn²⁺-containing vesicles stacked at the apical side of the cells, and this Zn²⁺ is required for cell survival (39).

A growing body of evidence suggests that extracellular Zn²⁺ activates signaling transduction pathways that induce cell survival and proliferation. For example, Zn²⁺ upregulates the PI 3-kinase pathway, leading to activation of Akt in fibroblasts (18). It also induces transactivation of the epidermal growth factor receptor by Src in epithelial cells (40, 47). In addition, extracellular Zn²⁺ triggers the activation of extracellular signal-elated kinase (ERK)1/2, which, in turn, activates p21 and cyclin D1 in colonocytes (18). On the basis of the results from our and others' literature, we propose a mechanistic Zn²⁺ signaling pathway in endothelial cells (Fig. 7). As a natural ligand, extracellular Zn²⁺ activates the promiscuous ZnR/GPR39 and thereby triggers G_αq signaling pathways. Then the G_αq would subsequently activate the PLC, Akt, PI3, and MAPK signaling pathways. At the same time, it may induce an intracellular Ca²⁺ response. The elevated Ca²⁺ level will next trigger many other Ca²⁺-dependent signaling cascades, including ERK1/2 and Akt. Taken together, all these pathways would lead to the differential regulation of genes related to vascular functions, such as survival/growth, angiogenesis, inflammation, vascular tone, coagulation, etc.

Fig. 7.

Proposed schematic pathway showing ZnR/GPR39-mediated Zn²⁺ signaling. The model suggests that ZnR/GPR39 is the major link between changes in extracellular Zn and physiological cell function in vascular cells, such as survival and proliferation, inflammation, and vascular tone.

To further explain the observed cellular function changes triggered by Zn²⁺ signaling, we evaluated the vascular functional gene profile as well as the corresponding protein expression. The enhanced cell viability and proliferation induced by Zn²⁺ could be explained by the increased levels of VEGFA and PDGFRA. VEGFA is a glycosylated mitogen that specifically acts on endothelial cells and has various effects, including mediating increased vascular permeability, inducing angiogenesis, vasculogenesis and endothelial cell growth, promoting cell migration, and inhibiting apoptosis (4). PDGFRA is a member of the platelet-derived growth factor family, and a mitogenic factor for cells of mesenchymal origin. Its deficiency leads to defects in oligodendrocytes, alveolar smooth muscle cells, and Leydig cells (19). Next, we examined the effect of Zn²⁺ on endothelial cell mobility including adhesion and migration. The actin cytoskeleton plays important roles in forming and maintaining the shape and structure of cells (48). The reorganization of cytoskeleton alters cell shape and motility (36). Several cellular processes, such as differentiation, growth, and apoptosis, are arbitrated by cell cytoskeletal organization and cell shape. The significant assembling of F-actin and cytoskeletal reorganization would provide support for the enhanced motility of vascular cells triggered by Zn²⁺.

Another important function of vascular cells is the mediation of inflammatory responses. Our results showed that various inflammatory molecules were upregulated or downregulated. Among them, some typical ones are HMOX1, IL-10, SELL, PECAM1, CCL2, IL-6, COX-2, and IL-1β. The ability of HMOX1 to catabolize free heme and produce carbon monoxide (CO) gives its anti-inflammatory properties by upregulation of IL-10 and IL-1 antagonist IL-1RA expression (34). IL-10 and IL-6 are major anti-inflammatory cytokines, whereas IL-1β is a proinflammatory cytokine (16, 24). The SELL gene encodes L-selectin, which is responsible for the recruitment of circulating leukocytes to the inflammation sites of endothelium (12). PECAM1 protein is a member of the immunoglobulin superfamily and is likely involved in leukocyte transmigration, angiogenesis, and integrin activation (6, 37, 51). CCL2 is a small cytokine that belongs to the C-C chemokine family. CCL2 recruits monocytes, memory T cells, and dendritic cells to the sites of inflammation produced by either tissue injury or infection (15, 38). Because various proinflammatory and anti-inflammatory molecules are differently regulated, it will need further studies to fully understand how Zn²⁺ would orchestrate the vascular inflammatory responses in different pathophysiological conditions.

Vascular tone refers to the degree of constriction experienced by a blood vessel relative to its maximally dilated state. Our results showed that Zn²⁺ also triggered the regulation of genes and proteins involved in vessel constriction and dilation including PTGIS and NOS3. PTGIS encodes a member of the cytochrome P-450 superfamily of enzymes, which catalyze the conversion of prostaglandin H2 to prostacyclin (prostaglandin I2), a potent vasodilator and inhibitor of platelet aggregation. NOS3 has a protective function in the cardiovascular system, which is attributed to NO production. Regulation of vascular tone is one of the best known roles of NO in the cardiovascular system. NO also has antithrombotic effects and antioxidant properties.

GRANTS

This work was supported by the National Heart, Lung, and Blood Institute of the National Institutes of Health (Grant 1R01 HL-140562). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

D.Z., Y.Z., B.M.F., L.T., and Y.-X.Q. conceived and designed research; D.Z. performed experiments; D.Z. analyzed data; D.Z., Y.S., and Y.-X.Q. interpreted results of experiments; D.Z. and Y.S. prepared figures; D.Z. drafted manuscript; D.Z., Y.S., and L.T. edited and revised manuscript; D.Z., Y.Z., B.M.F., L.T., and Y.-X.Q. approved final version of manuscript.