Temporally resolved cAMP monitoring in endothelial cells uncovers a thrombin-induced [cAMP] elevation mediated via the Ca2+-dependent production of prostacyclin

Authors

  • R. C. Werthmann,

    1. Institute of Pharmacology and Toxicology, University of Würzburg, Versbacherstrasse 9, 97078 Würzburg, Germany
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  • M. J. Lohse,

    1. Institute of Pharmacology and Toxicology, University of Würzburg, Versbacherstrasse 9, 97078 Würzburg, Germany
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  • M. Bünemann

    1. Institute of Pharmacology and Toxicology, University of Würzburg, Versbacherstrasse 9, 97078 Würzburg, Germany
    2. Institute of Pharmacology and Clinical Pharmacy, University of Marburg, Karl-von-Frisch-Strasse 1, 35032 Marburg, Germany
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Corresponding author M. Bünemann: Department of Pharmacology and Clinical Pharmacy, University of Marburg, Karl-von-Frisch-Strasse 1, 35032 Marburg, Germany. Email: moritz.buenemann@staff.uni-marburg.de

Non-technical summary

Endothelial cells form the innermost layer of blood vessels and build a barrier between blood and tissue. The permeability of this barrier is antagonistically controlled by the intracellular signalling molecules Ca2+ and cAMP. A rise in Ca2+ concentration increases permeability, whereas increased cAMP levels strengthen the endothelial cell barrier. In this study we investigated the impact of the coagulation factor thrombin that is known to increase Ca2+ concentrations and endothelial permeability, on cAMP levels. Surprisingly we detected that thrombin also led to a delayed and slow increase of cAMP concentrations. We discovered that this increase is due to the production of prostacyclin and a subsequent stimulation of endothelial prostacyclin receptors that finally induce cAMP production. This thrombin-mediated increase of cAMP levels might constitute a negative feedback control to protect endothelial barrier function despite a rise of Ca2+ concentrations.

Abstract

The barrier function of the endothelium is controlled by the second messengers Ca2+ and cAMP that differentially regulate the permeability of endothelial cells. The Ca2+-elevating agent thrombin has been demonstrated to increase endothelial permeability and to decrease cAMP levels as detected via enzyme immunoassays. To study the effects of thrombin on cAMP with high temporal resolution, we utilised the FRET-based cAMP sensor Epac1-camps in single intact human umbilical vein endothelial cells (HUVECs). In these cells, thrombin induced a delayed increase in [cAMP], initiating after about 40 s, with maximum cAMP levels after 130 s of thrombin application. This increase of cAMP levels was Ca2+-dependent, but did not require calmodulin (CaM). Pharmacological approaches revealed that phospholipase A2 (PLA2) activity and cyclooxygenase (COX)-mediated synthesis of prostaglandins was required for the thrombin-induced elevation of [cAMP]. Furthermore, preincubation of HUVECs with a prostacyclin-receptor antagonist significantly reduced the thrombin-induced increase in [cAMP]. We conclude that thrombin causes the synthesis of prostacyclin in endothelial cells and that the subsequent stimulation of Gs-coupled prostacyclin receptors then results in an increase in [cAMP].

Abbreviations 
AC

adenylyl cyclase

CaM

calmodulin

CFP

cyan fluorescent protein

cPLA2

Ca2+-dependent PLA2

COX

cyclooxygenase

Epac

exchange protein directly activated by cAMP

F CFP

CFP fluorescence

F YFP

YFP fluorescence

FRET

fluorescence resonance energy transfer

HDMEC

human dermal microvascular endothelial cell

HUVEC

human umbilical vein endothelial cell

iPLA2

Ca2+-independent PLA2

MAFP

5,8,11,14-eicosatetraenylmethylester phosphonofluoridic acid

PAR-1

protease activating receptor 1

PDE

phosphodiesterase

PGF

prostaglandin F

PKA

protein kinase A

PLA2

phospholipase A2

PMVEC

pulmonary microvascular endothelial cell

R

ratiometric FRET signal

YFP

yellow fluorescent protein

Introduction

The endothelial monolayer functions as a semi-permeable barrier between blood and interstitial tissues. This barrier function is controlled by the second messengers Ca2+ and cAMP, which differentially regulate the permeability of endothelial cells. While Ca2+ increases the permeability by inducing cell contraction, cAMP enhances stability of tight and adherens junctions and, thereby, supports the barrier function (Michel & Curry, 1999; Mehta & Malik, 2006). However, Ca2+ and cAMP signals are not independent, but rather are subject to crosstalk. cAMP signals can be regulated by Ca2+ via several pathways: first, via the Ca2+-dependent phosphodiesterase 1 (PDE1) that is activated by Ca2+ and calmodulin (CaM; Goraya & Cooper, 2005); and second, via adenylyl cyclases (ACs) that are either CaM-dependently activated (AC1, AC8) or inhibited (AC5, AC6) via submicromolar Ca2+ concentrations (Willoughby & Cooper, 2007; Sadana & Dessauer, 2009).

Thrombin, a coagulation factor that activates the protease activating receptor 1 (PAR1), has been reported to enhance endothelial permeability (Lum et al. 1992; Tiruppathi et al. 1992; Cioffi et al. 2002; Baumer et al. 2009). This is caused by the activation of the Gq-signalling cascade and a subsequent increase in intracellular [Ca2+], and by the activation of Rho-GTPase, both events finally promoting actin–myosin interaction and cellular contraction (Vandenbroucke et al. 2008). It has also been reported that the thrombin-mediated increase in endothelial permeability is also induced or maintained by the sustained reduction of cAMP levels that was detected in several studies via enzyme immunoassays (Cioffi et al. 2002; Baumer et al. 2008). For these assays, endothelial cells were incubated with thrombin and PDE inhibitors for several minutes (≥5 min) before cells were disrupted for [cAMP] determination. However, as thrombin-induced Ca2+ signals are highly dynamic and the Ca2+-mediated regulation of [cAMP] is complex, more detailed insights require monitoring thrombin-induced cAMP regulation with much better temporal resolution than can be achieved with biochemical techniques.

Utilising the fluorescence resonance energy transfer (FRET)-based cAMP sensor Epac1-camps (Nikolaev et al. 2004), we recently reported that in human umbilical vein endothelial cells (HUVECs) thrombin induced a transient decrease of cAMP levels that had been elevated by stimulation of β-adrenergic receptors (Werthmann et al. 2009). This effect of thrombin was attributed to the Ca2+-mediated inhibition of AC6. However, the thrombin-induced decrease of [cAMP] was followed by an increase in [cAMP] that was also observed in the absence of a prior β-adrenergic-mediated increase in [cAMP].

In the present study we focused on the molecular mechanism underlying this slowly developing [cAMP] increase caused by exposure of endothelial cells to thrombin.

As the thrombin-activated PAR1 is reported to couple to Gq, Gi and G12/13 but not to Gs (Coughlin, 2000; Bogatcheva et al. 2001; Macfarlane et al. 2001), the question arises how thrombin might cause this increase in [cAMP]. We evaluated two possible mechanisms: on the one hand, the activation of Ca2+-dependent ACs (Simpson et al. 2006; Willoughby & Cooper, 2007) and on the other hand, the thrombin-induced prostacyclin synthesis and subsequent activation of Gs-coupled prostacyclin receptor (Weksler et al. 1978; Jaffe et al. 1987). The results presented here strongly support the conclusion that thrombin elevates cAMP in endothelial cells via a PLA2–COX–prostacyclin pathway.

Methods

Cell culture and cell transfection

Human umbilical vein endothelial cells (HUVECs; Lonza, Cologne, Germany) were cultured in complete medium (EBM-2; Lonza) and grown in the presence of 5% CO2 at 37°C. HUVECs were transfected with plasmid DNA using Amaxa Nucleofector Technology according to the manufacturer's instructions (Basic Nucleofector Kit for Primary Endothelial Cells; Lonza). HUVECs (5 × 105) were transfected with a maximum of 3 μg DNA. Transfection of HUVECs with the plasmid encoding the cAMP sensor Epac1-camps (Nikolaev et al. 2004) via electroporation yielded transfection efficiencies of over 50%. For FRET measurements HUVECs were seeded on fibronectin-coated glass coverslips (Human Plasma Fibronectin Purified Protein; Millipore, Schwalbach, Germany) and FRET experiments were done 24 h after transfection.

Reagents

Adenosine deaminase (0.5 U ml−1; Roche, Palo Alto, CA, USA) was added to the medium 30 min before FRET experiments. To establish a concentration–response curve for the cAMP probe Epac1-camps, transfected HUVECs were stimulated with increasing concentrations of isoproterenol (isoprenaline; Sigma-Aldrich, St Louis, MO, USA). Thrombin from human plasma (Sigma-Aldrich) was used at 10 U ml−1, 0.5 U ml−1 or at the indicated concentrations, respectively. For chelation of Ca2+, HUVECs were incubated with BAPTA-AM (10 μm; Sigma-Aldrich) for 30 min before FRET measurements and BAPTA-AM was present in the external solution during FRET experiments. The Ca2+ ionophore A23187 (1 μm), ATP (100 μm) and histamine (100 μm; Sigma-Aldrich) were added to increase intracellular [Ca2+]. To inhibit cytosolic isoforms of phospholipase A2 (PLA2), MAFP (100 μm; 5,8,11,14-eicosatetraenylmethylester phosphonofluoridic acid; Tocris, Bristol, UK) was added to the medium 10 min before experiments. Indomethacin (100 μm; Sigma-Aldrich) was added 1 h and acetylsalicylic acid (1 mm; Sigma-Aldrich) 30 min before FRET measurements to the cells for the inhibition of cyclooxygenases (COX). Carbaprostacyclin (Cayman Chemical, Ann Arbor, MI, USA) was used at 10 μm to stimulate prostacyclin receptors. The prostacyclin receptor antagonist CAY10441 (10 μm; Cayman Chemical) was added to the external solution during the FRET measurements before thrombin stimulation.

FRET measurements

Fluorescence microscopy and FRET measurements were done as described previously (Nikolaev et al. 2005; Hein et al. 2006). Confluent HUVECs, that were transfected with Epac1-camps and grown on fibronectin-coated glass coverslips, were placed on an Axiovert 200 inverted microscope (Zeiss, Jena, Germany) using a 63× oil immersion objective, a dual-emission photometric system and a polychrome IV light source (both TILL Photonics, Gräfelfing, Germany). Illumination time was 50 ms at a frequency of 1 Hz. Excitation wavelength was set to 436 ± 10 nm (beam splitter Dichroic long pass (DCLP) 460 nm), and emission of single whole cells was recorded at 535 ± 15 nm and 480 ± 20 nm (beam splitter DCLP 505 nm). The FRET-based sensor Epac1-camps consists of a single cAMP-binding domain derived from exchange protein directly activated by cAMP 1 (Epac1), which is flanked by a yellow fluorescent protein (YFP) and a cyan fluorescent protein (CFP) on either side. In the absence of cAMP this sensor exhibits strong FRET. If cAMP is bound to the binding domain, FRET is highly attenuated (Nikolaev et al. 2004). Upon excitation of CFP in intact single cells, CFP and YFP fluorescence intensities and ratiometric FRET were recorded as a read-out for cAMP concentrations. Ratiometric FRET (R) was calculated as a ratio of yellow fluorescent protein emission (FYFP) over cyan fluorescent protein emission (FCFP), where FYFP was the emission at 535 nm, corrected for direct excitation of YFP at 436 nm and bleedthrough of CFP emission into the YFP channel. Additionally ratiometric FRET of all experiments was normalised to the FRET ratio at the timepoint 0 s (R0). Cells were continuously superfused with external buffer (141 mm NaCl, 5.4 mm KCl, 1 mm MgCl2, 10 mm Hepes at pH 7.3) with or without Ca2+ (2 mm CaCl2 or 5 mm EGTA). These reagents were applied from AppliChem (Darmstadt, Germany). Thrombin was freshly prepared and applied using a rapid superfusion system (ALA Scientific Instruments, Westbury, NY, USA). The same experimental conditions were applied for FRET experiments with the cAMP-independent probe XProt (Willemse et al. 2007) or the cAMP probe based on the regulatory and catalytic subunits of PKA (RII-CFP and C-YFP; Zaccolo & Pozzan, 2002).

Ca2+ measurements

Changes in cytosolic Ca2+ levels were monitored in HUVECs loaded with the fluorescent Ca2+-sensitive dye Fluo-4 AM (Molecular Probes, Invitrogen, Eugene, OR, USA). HUVECs were incubated with 2 μm Fluo-4 AM for 20 min at 37°C and washed with external buffer before fluorescence measurements. Fluo-4 fluorescence was recorded with the same photometric microscope setup as described for FRET measurements. Illumination time was 50 ms at a frequency of 1 Hz. Excitation wavelength was set to 490 nm (exciter ET470/40×, dichroic T495LP), and emission of single whole cells was recorded at 535 ± 15 nm (DCLP 505 nm). The fluorescence intensity of Fluo-4 increases upon Ca2+ binding.

6-Keto prostaglandin F detection

The thrombin-induced release of prostacyclin was determined via an enzyme immunoassay for the quantification of 6-keto prostaglandin F (6-keto PGF; Cayman Chemical), the stable hydrolysis product of prostacyclin. 6-Keto PGF in the media of control and thrombin-stimulated HUVECs was measured 2 min after thrombin stimulation (10 U ml−1) following the manufacturer's protocol.

Data analysis and statistics

Fluorescence intensities were acquired using CLAMPEX 9.0 (Axon Instruments, Foster City, CA). Data were processed using Origin 6.1 (Northampton, MA, USA) and statistical analyses were performed using Prism 4.0 (San Diego, CA, USA). Values are given as mean ±s.e.m. For comparison of two individual groups Student's unpaired t tests were performed and differences were considered significant when P < 0.05 (*) or P < 0.01 (**; two-tailed).

Results

Stimulation of HUVECs with thrombin causes an increase of cAMP levels

As the fine-tuned regulation of Ca2+ and cAMP signals is highly relevant for the maintenance or disruption of the endothelial barrier, we studied the temporal pattern of thrombin-induced Ca2+ and cAMP signals in single living endothelial cells. In order to achieve high temporal resolution, we photometrically detected Ca2+ signals by using the fluorescence dye Fluo-4 and cAMP concentrations by monitoring FRET of the cAMP-sensing probe Epac1-camps (see Methods; Nikolaev et al. 2004; Werthmann et al. 2009).

The stimulation of single Epac1-camps-transfected HUVECs with thrombin (10 U ml−1) led to a decrease in FYFP and an increase in FCFP, resulting in a decrease of the ratiometric FRET signal (R) that was normalised to its value at the timepoint 0 s (R0). Thrombin stimulation (10 U ml−1) decreased the ratiometric FRET signal by approximately 50% compared to the maximal changes of the ratiometric FRET signal following a stimulation of cells with isoproterenol (1 μm; representative experiment, Fig. 1A). Stimulation of cells with 1 μm isoproterenol caused maximal changes of the ratiometric FRET signal due to saturation of Epac1-camps (Werthmann et al. 2009). The decrease of the ratiometric FRET signal (R/R0) is consistent with an increase in [cAMP]. The plot of averaged ratiometric FRET signals of single experiments (mean ±s.e.m.) revealed a delayed decrease of the ratiometric FRET signal, that started about 40 s after thrombin application (10 U ml−1) and reached equilibrium after about 150 s (Fig. 1B). To test the potential of thrombin to stimulate cAMP production in HUVECs, concentration–response curves for thrombin and isoproterenol were established with Epac1-camps-transfected cells (Fig. 1C and Supplemental Fig. 1). The concentration–response curve for isoproterenol revealed an EC50 value of 8.1 ± 1.2 nm (Supplemental Fig. 1). Compared to the maximal increase in [cAMP] after isoproterenol stimulation, thrombin application (10–100 U ml−1) only induced about half-maximal cAMP responses (Fig. 1A and C).

Figure 1.

Thrombin stimulation of HUVECs induced an increase of cAMP levels
A, upon CFP excitation, YFP and CFP fluorescence were recorded in single HUVECs expressing the FRET-based cAMP sensor Epac1-camps. Stimulation of cells with thrombin (10 U ml−1) resulted in a decrease in YFP emission (FYFP, yellow trace) and a concomitant increase in CFP emission (FCFP, blue trace). The ratiometric FRET signal (R) was normalised to R0 (R/R0), corresponding to the FRET ratio at timepoint 0 s. Stimulation of cells with isoproterenol (1 μm) led to a maximal decrease of the ratiometric FRET signal. The decrease of the ratiometric FRET signal is consistent with an increase in [cAMP] (representative experiment shown). B, the plot of averaged FRET experiments of Epac1-camps-transfected HUVECs (n = 21) revealed a delayed decrease of the ratiometric FRET signal (R/R0) about 40 s after thrombin application, that is reaching equilibrium after about 150 s. C, Epac1-camps-transfected HUVECs were stimulated with increasing concentrations of thrombin. The establishment of a concentration–response curve revealed an EC50 value of 0.95 ± 1.06 U ml−1 thrombin (n= 5). The stimulation of cells with a maximal effective thrombin concentration (100 U ml−1) only led to half-maximal changes of the ratiometric FRET signal (ΔR) compared to ΔR following stimulation of HUVECs with a maximal effective isoproterenol concentration (ΔRiso max).

In order to test whether this thrombin-induced decrease of the ratiometric FRET signal was due to changes in cAMP or rather reflected unspecific alterations in FRET such as those induced by changes in intracellular ATP (Willemse et al. 2007), we measured thrombin-induced changes of FRET in HUVECs expressing the FRET-based, but cAMP-independent XProt sensor (Willemse et al. 2007). These experiments revealed a markedly retarded, thrombin-induced (10 U ml−1) decline of the mean ratiometric FRET signal (Fig. 2A) that was not due to simple bleaching of the fluorophores (YFP and CFP; not shown). This thrombin-induced decline of the ratiometric FRET signal was considered cAMP-unspecific. So, all further experiments with Epac1-camps-transfected HUVECs that were stimulated with thrombin (10 U ml−1), were corrected for the mean ratiometric FRET signal of XProt-transfected cells and herein after referred to as R/R0*. The thrombin-induced decrease of the ratiometric FRET signal in Epac1-camps-transfected HUVECs (Fig. 1B), background-corrected for the mean ratiometric FRET signal in XProt-transfected HUVECs (Fig. 2A), still revealed a clear decrease of the FRET ratio (R/R0*), i.e. an increase in [cAMP], starting about 40 s and reaching equilibrium about 130 s after thrombin stimulation (Fig. 2B).

Figure 2.

Thrombin led to a cAMP-specific change of the ratiometric FRET signal
A, instead of Epac1-camps, HUVECs were transfected with the FRET-based XProt-protein that has no cAMP-binding domain. Thrombin stimulation (10 U ml−1) of XProt-transfected cells resulted in a slow and steady decrease of the averaged ratiometric FRET signal (n= 19). B, the plot of averaged FRET experiments (Fig. 1B) of Epac1-camps-transfected HUVECs was corrected for the mean XProt-signal (Fig. 2A). This background-corrected ratiometric FRET signal (R/R0*) is reaching equilibrium about 130 s after thrombin application. C, in HUVECs that were transfected with a PKA-based FRET probe for cAMP (RII-CFP and C-YFP), thrombin stimulation led to a transient decrease of the mean ratiometric FRET signal (R/R0; n= 6) that is consistent with a transient increase in [cAMP].

The thrombin-induced increase in [cAMP] was further proven by using a FRET-based cAMP probe that is based on the dissociation of protein kinase A (PKA) regulatory (RII) and catalytic (C) subunits (Zaccolo & Pozzan, 2002). Binding of cAMP to the regulatory subunits leads to subunit dissociation and therefore results in a loss of FRET. Thrombin stimulation (10 U ml−1) of HUVECs, that were transfected with RII-CFP and C-YFP, led to a transient decrease of the ratiometric FRET signal (Fig. 2C), that is consistent with a transient increase in [cAMP].

The thrombin-induced increase in [cAMP] is Ca2+-dependent

As the thrombin receptor PAR1 is not described as activating the Gs protein, which would directly activate ACs to produce cAMP, we tested if the thrombin-induced increase in [cAMP] was Ca2+-dependent. First, we investigated the temporal pattern of thrombin-induced Ca2+-signals. Therefore, HUVECs were loaded with the Ca2+-sensitive dye Fluo-4 AM. Thrombin stimulation (10 U ml−1) of single HUVECs led to a fast increase in Fluo4 emission (FFluo4) reflecting an increase of intracellular [Ca2+]. The detection of the mean Fluo-4 emission of single experiments revealed a maximum Ca2+ signal about 12 s after thrombin stimulation (Fig. 3A). To test the Ca2+-dependency of the thrombin-induced increase in [cAMP], FRET assays were performed with Epac1-camps-expressing HUVECs that were either preincubated with BAPTA-AM (10 μm) for complexation of intracellular Ca2+ or exposed to a nominally Ca2+-free external solution that contained EGTA. The preincubation of HUVECs with BAPTA-AM (10 μm) significantly attenuated the decrease of the averaged ratiometric FRET signal (Fig. 3B; grey curve) after thrombin stimulation (10 U ml−1) compared to control cells (black curve; **P < 0.01). The thrombin-induced changes of the ratiometric FRET signal (Δ(R/R0*): minimal FRET ratio after thrombin stimulation minus FRET ratio at the timepoint 0 s) were almost abolished in cells pretreated with BAPTA-AM (**P < 0.01) and significantly reduced in cells superfused with a Ca2+-free solution (*P < 0.05; Fig. 3C). Further, stimulation of Epac1-camps-transfected HUVECs with either the Ca2+ ionophore A23187 (1 μm; Fig. 3D), or with ATP (100 μm; Fig. 3E) and histamine (100 μm; Fig. 3F), that elevate intracellular [Ca2+] via Gq protein activation, led to a delayed increase in [cAMP] comparable to the thrombin-induced [cAMP] increase (Fig. 3B).

Figure 3.

The thrombin-induced increase in [cAMP] is Ca2+ dependent
A, HUVECs were loaded with Fluo-4 for measurements of Ca2+ signals. After thrombin application (10 U ml−1) Ca2+ binding to Fluo-4 led to a transient increase in Fluo4-emission intensity induced by excitation with 490 nm. The fluorescence intensity of single experiments was normalised to the time point of agonist application and averaged in order to plot the mean ±s.e.m. (n= 14) over the course of the experiment. The averaged fluorescence signal peaked 12 s after thrombin stimulation, reflecting maximal Ca2+ concentrations. B, to prove the Ca2+ dependency of the thrombin-mediated increase in [cAMP], HUVECs were incubated with BAPTA-AM for complexation of Ca2+. Incubation of cells with BAPTA-AM (10 μm; n= 13; grey curve) led to a significantly diminished decrease of the averaged ratiometric FRET signal (R/R0*) compared to control cells (n= 11; black curve; **P < 0.01). C, for the comparison of thrombin-induced changes of the ratiometric FRET signal under different conditions, Δ(R/R0*) (minimal FRET ratio after thrombin stimulation minus FRET ratio at the timepoint 0 s) was calculated of single FRET experiments. The complexation of intracellular Ca2+ by incubating HUVECs with 10 μm BAPTA-AM resulted in a significantly diminished Δ(R/R0*) (black column; n= 13) compared to control cells (n= 11; **P < 0.01) following thrombin stimulation (10 U ml−1; see Fig. 3B). Δ(R/R0*) was also significantly reduced in HUVECs that were measured in EGTA-buffered external solution without Ca2+ (grey column; n= 16) compared to control cells (*P < 0.05). D–F, stimulation of HUVECs with the Ca2+ ionophore A23187 (1 μm; D) or the Ca2+-elevating agonists ATP (100 μm; E) and histamine (100 μm; F) also led to a delayed decrease of the ratiometric FRET signal (R/R0) depicting an increase in [cAMP].

Thus, we conclude that the thrombin-induced decrease of the ratiometric FRET signal that indicates an increase in [cAMP] was dependent on intracellular Ca2+.

To test a possible involvement of Ca2+–calmodulin (CaM)-activated ACs, we expressed the dominant negative CaM mutant CaM12 (Simpson et al. 2006), which failed to have any detectable effect on the thrombin-induced decline of FRET in Epac1-camps-transfected HUVECs (Supplemental Fig. 2). In control experiments we verified that cAMP signals induced via exogenously expressed CaM-activated AC8 were robustly inhibited by CaM12 as reflected by Epac1-camps FRET signals (Supplemental Fig. 2). The observed difference between AC8-transfected HUVECs and control cells regarding the differential sensitivity to the CaM12 mutant suggest that the direct CaM- and Ca2+-dependent activation of endogenous ACs is not the underlying mechanism for the rise in [cAMP] after thrombin stimulation of HUVECs.

The thrombin-induced activation of phospholipase A2 (PLA2) is involved in the increase in [cAMP]

Based on published evidence that thrombin can stimulate prostacyclin synthesis in endothelial cells and that endothelial cells express the Gs-coupled prostacyclin receptor (Weksler et al. 1978; Bundey & Insel, 2006; Wheeler-Jones, 2008), we explored next whether thrombin might induce the activation of ACs indirectly via the eicosanoid pathway. Therefore, we investigated the potential role of thrombin-induced synthesis of prostacyclin in the thrombin-mediated increase in [cAMP] in HUVECs. Prostacyclin synthesis is thought to be initiated by the PLA2-mediated release of arachidonic acid from cell membrane phospholipids, which is then converted via cyclooxygenases (COX) to prostaglandin H2 and finally via prostacyclin synthase to prostacyclin (Jaffe et al. 1987; Wheeler-Jones, 2008).

To test the potential involvement of the Ca2+-dependent cytosolic PLA2 (cPLA2), HUVECs were incubated with MAFP (100 μm; 10 min), which inhibits all cytosolic PLA2s (cPLA2 and Ca2+-independent iPLA2; Lio et al. 1996). Mean ratiometric FRET signals were determined in control (black curve; Fig. 4) and MAFP-treated cells (grey curve) following thrombin stimulation (10 U ml−1). The decrease of the averaged ratiometric FRET signal was significantly reduced in MAFP-treated cells compared to control cells (**P < 0.01). This suggests that PLA2 activity is involved in the thrombin-induced increase in [cAMP] in HUVECs.

Figure 4.

Inhibition of the cytosolic PLA2 by MAFP significantly reduced the thrombin-induced increase in [cAMP]
Incubation of cells with MAFP (100 μm) resulted in a significantly diminished decrease of the ratiometric FRET signal (R/R0*; n= 18; grey curve) compared to control cells (n= 18; black curve; **P < 0.01) following thrombin stimulation (10 U ml−1) as demonstrated by the plot of averaged single cell experiments.

The thrombin-induced increase in [cAMP] is dependent on the activity of cyclooxygenases

Next, we examined the involvement of the cyclooxygenases COX1 and COX2, which catalyse the synthesis of prostaglandin H2, in the thrombin-induced increase in [cAMP]. Therefore, we incubated HUVECs with the COX inhibitors indomethacin (100 μm; 30 min) or acetylsalicylic acid (1 mm; 30 min). The averaged ratiometric FRET signals revealed that indomethacin markedly reduced (grey curve; Fig. 5A) and acetylsalicylic acid completely abolished (grey curve; Fig. 5B) the thrombin-induced (10 U ml−1) decline of the FRET signal in indomethacin- and acetylsalicylic acid-treated cells compared to control experiments (black curve; **P < 0.01; Fig. 5A and B). This implies the involvement of COX in the observed thrombin-mediated effect on cAMP levels.

Figure 5.

Inhibition of COX significantly diminished the increase in [cAMP] following thrombin stimulation
A, incubation of HUVECs with the COX inhibitor indomethacin (100 μm) led to a significantly diminished decrease of the mean ratiometric FRET signal (R/R0*; n= 16; grey curve) after thrombin stimulation (10 U ml−1) compared to control cells (n= 26; black curve; **P < 0.01). B, incubation of HUVECs with the COX inhibitor acetylsalicylic acid (1 mm) completely abolished the thrombin-induced (10 U ml−1) decrease of the mean ratiometric FRET signal (R/R0*; n= 12; grey curve), that is depicted in control cells (n= 26; black curve; **P < 0.01).

The thrombin-induced increase in [cAMP] is caused by the activation of prostacyclin receptors

The prostacyclin receptor has been shown to be the predominant Gs-coupled prostaglandin receptor in HUVECs (Bundey & Insel, 2006). To investigate the relevance of the prostacyclin receptor concerning the thrombin-mediated increase in [cAMP], we stimulated Epac1-camps-transfected HUVECs directly with the stable prostacyclin-analogue carbaprostacyclin. In contrast to thrombin, which was added via the superfusion system, carbaprostacyclin (10 μm) was added with a pipette to avoid retention of the hydrophobic carbaprostacyclin by the tubing. The application of carbaprostacyclin resulted in a fast decrease of the mean ratiometric FRET signal that started after approximately 5 s (Fig. 6A). This confirmed the expression of functional Gs-coupled prostacyclin receptors in HUVECs.

Figure 6.

The thrombin-induced increase of [cAMP] involves the release of prostacyclin and the activation of prostacyclin receptors
A, direct stimulation of endogenous prostacyclin receptors with carbaprostacyclin (10 μm) caused a rapid decrease of the mean ratiometric FRET signal (R/R0; n= 8). B, preincubation of HUVECs with the prostacyclin receptor antagonist CAY10441 (10 μm) resulted in a significantly reduced thrombin-induced (10 U ml−1) decrease of the ratiometric FRET signal (R/R0*; n= 16; grey curve) compared to control cells (n= 14; black curve; *P < 0.05). C, [6-keto PGF], the stable hydrolysis product of prostacyclin, was determined in cell media of thrombin-stimulated and control HUVECs via an enzyme immunoassay. Thrombin stimulation (10 U ml−1) resulted in a significantly increased [6-keto PGF] compared to control cells (n= 4; **P < 0.01).

We then tested the effect of the prostacyclin receptor antagonist CAY10441 on the thrombin-mediated increase in [cAMP]. To do so, FRET measurements were done in buffer containing CAY10441 (10 μm). CAY10441 significantly diminished the thrombin-induced decrease of the mean ratiometric FRET signal (grey curve) in HUVECs compared to control cells (black curve; *P < 0.05; Fig. 6B). Thus, the thrombin-induced increase in [cAMP] was significantly reduced by the prostacyclin receptor antagonist. This led us to conclude that the activation of CAY10441-sensitive prostacyclin receptors was involved in the thrombin-induced rise in [cAMP]. Further, we measured 6-keto PGF, the stable hydrolysis product of prostacyclin, in the cell media of thrombin-stimulated versus unstimulated control cells via an enzyme immunoassay (Bundey & Insel, 2006). Thrombin stimulation (10 U ml−1) of HUVECs led to an ∼8-fold increased [6-keto PGF] in the cell media compared to unstimulated control cells (**P < 0.01; Fig. 6C).

In summary, we observed a delayed thrombin-induced increase of cAMP levels in HUVECs. Similar increases were induced by stimulation of other Gq-coupled receptors, for example with ATP or histamine or by direct elevation of [Ca2+] with A23187. This was dependent on intracellular Ca2+, on PLA2 and COX activity and prostacyclin production, as well as activation of endothelial prostacyclin receptors. These observations suggest that thrombin elevated [cAMP] indirectly, via stimulation of prostacyclin production and subsequent activation of Gs-coupled prostacyclin receptors.

Discussion

The barrier function of endothelial cells is primarily controlled by cAMP and Ca2+ signals. Whereas Ca2+ signals promote contraction of the cells and thereby increase endothelial permeability, cAMP protects the barrier function by strengthening adhesion of the cells. The coagulation factor thrombin was demonstrated to increase the permeability of endothelial cells via a Gq-mediated increase in [Ca2+] and via activation of the Rho-GTPase (Michel & Curry, 1999; Mehta & Malik, 2006). Moreover, several studies reported a decrease of cAMP levels upon exposure of endothelial cells to thrombin (Cioffi et al. 2002; Baumer et al. 2008). Performing real-time monitoring of cAMP levels in single living cells, we recently found a more complex pattern of thrombin-induced cAMP regulation: thrombin inhibited AC6 in a Ca2+-dependent fashion, an effect only detectable when AC6 was stimulated, for instance, via adrenergic receptors (Werthmann et al. 2009). This thrombin-induced decline in [cAMP] appeared to be transient because of the subsequent occurrence of a slowly developing increase in [cAMP]. Utilising the FRET-based cAMP sensor Epac1-camps in HUVECs, we observed thrombin-induced increases in [cAMP] initiating after about 40 s and reaching maximal levels after 130 s stimulation of cells with thrombin (Fig. 2B). Similar [cAMP] increases were detected in HUVECs that were stimulated with other Ca2+-elevating agonists like ATP and histamine (Fig. 3E and F). Particularly for FRET-based studies that deal with slow alterations of the FRET signal, it is important to control the specificity of the signal. Using the XProt probe, representing a CFP–YFP fusion protein that does not react to changes in [cAMP], we also observed thrombin-induced changes in the FRET signal, possibly due to alterations in the ATP content of the cell (Fig. 2A; Willemse et al. 2007). Notably, these decreases in FRET occurred much slower and smaller in amplitude compared to thrombin-induced alterations in Epac1-camps FRET (Fig. 1B). As this change of the ratiometric FRET signal is cAMP-independent, all FRET experiments of Epac1-camps-transfected HUVECs that were stimulated with thrombin (10 U ml−1), were corrected for the mean XProt signal. This corrected, thrombin-induced decrease of the ratiometric FRET signal still reflects a robust increase in [cAMP] (Fig. 2B). Using a PKA-based FRET probe for cAMP (Zaccolo & Pozzan, 2002) we further demonstrated that the thrombin-induced increase in [cAMP] led to the activation and subsequent release of the catalytic subunits of PKA and therefore is potentially relevant for the physiology of endothelial cells. Contrary to Epac1-camps-transfected cells, the thrombin-induced increase in [cAMP] was transient in cells that were transfected with the PKA-based cAMP probe (RII-CFP and C-YFP; Fig. 2C). We hypothesise that this is due to a PKA-induced activation of PDEs (PDE3 and PDE4; Maurice et al. 2003) due to the exogenous overexpression of functional PKA subunits.

This thrombin-induced effect was dependent on elevations of intracellular [Ca2+]. The involvement of Ca2+ is supported by several experimental results: BAPTA-mediated complexation of intracellular Ca2+ significantly reduced the thrombin-induced increase in [cAMP] (Fig. 3B and C). Similarly, removal of external Ca2+ by EGTA significantly attenuated the thrombin-induced elevation of [cAMP] (Fig. 3C). These observations correlate well with results obtained by Lückhoff et al. (1988): in this study a bradykinin-evoked release of prostacyclin was highly but not entirely dependent on both intracellular and extracellular [Ca2+]. Furthermore, a fast increase in [Ca2+] with a maximum value about 12 s after thrombin application preceded the increase in [cAMP] (Fig. 3A), and other Ca2+-elevating agonists like the Ca2+ ionophore A23187, ATP or histamine also resulted in a [cAMP] increase in HUVECs (Fig. 3D–F).

HUVECs predominantly express the Ca2+-inhibitable AC6 besides the Ca2+-independent isoforms AC3 and AC4 (Bundey & Insel, 2003). Neither of the two isoforms (AC1 and AC8) described to be directly stimulated by Ca2+ via CaM (Tang et al. 1991; Cali et al. 1994) have been reported to be expressed in significant amounts in HUVECs or other endothelial cells. Furthermore, the thrombin-induced stimulation of cAMP detected in our single cell FRET assays was not inhibited by a dominant negative CaM mutant (CaM12; Simpson et al. 2006), whereas under similar conditions, CaM12 attenuated thrombin-mediated stimulation of exogenously expressed AC8 (Supplemental Fig. 2). Based on these data it is highly unlikely that endogenous Ca2+-stimulated ACs were major mediators of the observed thrombin-induced elevations in cAMP.

The delayed and slow thrombin-induced increase in [cAMP] pointed more towards an indirect activation of ACs. As many years ago thrombin was described to induce the synthesis of prostacyclin (Weksler et al. 1978; Jaffe et al. 1987), we asked whether thrombin mediates [cAMP] elevations via Ca2+-dependent prostacyclin production and subsequent activation of Gs-coupled prostacyclin receptors. Arachidonic acid, which is a precursor of prostacyclin, has been demonstrated to be produced by Ca2+-dependent activation of cytosolic PLA2 (Clark et al. 1991). Incubation of HUVECs with MAFP, an inhibitor of the Ca2+-dependent cytosolic PLA2 (cPLA2) and of the Ca2+-independent cytosolic PLA2 (iPLA2; Lio et al. 1996), resulted in a significant reduction of the thrombin-induced increase in [cAMP] (Fig. 4). Furthermore, the indomethacin- or acetylsalicylic acid-mediated inhibition of COX that catalyses the formation of prostaglandin H2 from arachidonic acid, attenuated (indomethacin; Fig. 5A) or even abolished (acetylsalicylic acid; Fig. 5B) the thrombin-induced [cAMP] increase compared to control cells.

As the activity of PLA2 and COX significantly contributed to the thrombin-induced increase in [cAMP] and endothelial cells express high levels of the prostacyclin synthase (Wheeler-Jones, 2008), we propose that prostacyclin production is critically important for the thrombin-mediated increase in [cAMP]. Utilising an enzyme immunoassay we detected a significant increase in [6-keto PGF], the stable hydrolysis product of prostacyclin, in the cell media of thrombin-stimulated HUVECs compared to control cells (Fig. 6C). Prostacyclin receptors represent the dominant Gs-coupled prostaglandin receptor in HUVECs as revealed by real-time PCR (Bundey & Insel, 2006). In line with the hypothesis that Ca2+-induced prostacyclin generation will induce cAMP production in HUVECs via prostacyclin receptors, preincubation of HUVECs with the prostacyclin receptor antagonist CAY10441 (synonymous to RO1138452; Jones et al. 2006) resulted in a significantly reduced increase in [cAMP] compared to control cells (Fig. 6B). As a positive control, we showed that direct stimulation of receptors with the stable prostacyclin analogue, carbaprostacyclin, caused a fast increase in [cAMP] initiating 5 s after carbaprostacyclin application (Fig. 6A). As the thrombin-induced increase in [cAMP] begins after a delay of about 40 s, the thrombin-induced prostacyclin synthesis apparently requires about 35 s. The maximum cAMP levels after about 130 s of thrombin application are in agreement with studies by Weksler et al. (1978), who observed a maximum prostacyclin synthesis about 2 min after thrombin stimulation in HUVECs. We conclude that thrombin causes a Ca2+-dependent activation of PLA2 and a COX- and prostacyclin synthase-mediated synthesis of prostacyclin, which subsequently stimulates Gs-coupled prostacyclin receptors, finally resulting in the synthesis of cAMP via activated ACs. Moreover, we claim that prostacyclin production and the stimulation of prostacyclin receptors can also occur in one and the same cell as we observed the thrombin-induced increase in [cAMP] also in single isolated cells.

However, this thrombin-induced increase in [cAMP] was somewhat surprising as thrombin has been demonstrated to decrease cAMP levels in enzyme immunoassays (Cioffi et al. 2002; Baumer et al. 2008). Cioffi et al. (2002) observed a 50% decrease of total [cAMP] in pulmonary microvascular endothelial cells (PMVECs) after incubation of cells with the PDE4 inhibitor rolipram (10 μm) for 10 min and with thrombin (100 U ml−1) for 5 min. Baumer et al. (2008) detected a 40% decrease of total [cAMP] in human dermal microvascular endothelial cells (HDMECs) only after incubation of cells with thrombin (10 U ml−1) for 15 min whereas no significant change in [cAMP] was observed after a 5 min incubation. However, one cannot directly compare the results of enzyme immuno-based and FRET-based cAMP assays. As conventional cAMP assays like the enzyme immuno-based assay measure total [cAMP] of cell lysates at a certain time point, it is not feasible to accurately measure small changes of [cAMP] with high temporal resolution like in FRET-based real-time measurements of [cAMP] in single living cells. Furthermore, thrombin will regulate [cAMP] strongly dependent on the physiological context: when AC6 is stimulated, thrombin will cause a decrease in [cAMP] due to Ca2+-mediated inhibition of AC6 (Werthmann et al. 2009), whereas in the absence of AC6 stimulation, cAMP levels will increase over the time course of a few minutes as shown in this study.

Thrombin is well known to increase the endothelial permeability within minutes (Lum et al. 1992; Tiruppathi et al. 1992) and rises in [cAMP] at the cell membrane are described to antagonize this increase in permeability (Carson et al. 1989; Minnear et al. 1989; Stelzner et al. 1989; Garcia et al. 1995). Comparably, the adenoviral transfer of AC6 in HUVECs resulted in a reduced thrombin-induced increase in endothelial permeability in HUVECs compared to control cells, which was demonstrated to be due to thrombin-induced synthesis of prostacyclin and the activation of prostacyclin receptors (Bundey & Insel, 2006). Thus, the thrombin-mediated prostacyclin synthesis and a subsequent increase of cAMP levels might constitute a negative feedback control to protect endothelial barrier function.

Another explanation for the thrombin-induced increase in permeability despite increased cAMP levels might be compartmentation of cAMP within cells. With the FRET-based probe Epac1-camps we detected global cAMP levels of single cells, but cytoplasmic cAMP was demonstrated to increase endothelial permeability in contrast to cAMP that is synthesised at the cell membrane (Sayner et al. 2006). A cytoplasmic cAMP pool might be generated via intracellular signalling by internalised prostacyclin receptors along with ACs, as it was described for the thyroid-stimulating hormone receptor (Calebiro et al. 2009). Thus, spatial imaging of cAMP will be an important goal for the future to learn more about cAMP compartmentation and its consequences for vascular permeability.

In summary, we detected for the first time a delayed and slow, thrombin-induced increase in [cAMP] in living endothelial cells using the FRET-based cAMP sensor Epac1-camps. This increase in [cAMP] was caused by the Ca2+-dependent activation of PLA2. Pharmacological interference of cyclooxygenases and prostacyclin receptors confirmed that the synthesis of prostacyclin and subsequent stimulation of Gs-coupled prostacyclin receptors caused the thrombin-induced increase in [cAMP].

Appendix

Author contributions

R.C.W. designed and performed the experiments, their analyses and wrote the manuscript. M.J.L. significantly contributed to the study design and revised the manuscript. M.B. conceived the experiments and wrote the manuscript. All parts of this study were conducted at the Department of Pharmacology and Toxicology, University of Würzburg. All authors approved the final version of the manuscript.

Acknowledgements

We are grateful to the following scientists for the kind donation of the following plasmids: Philip Bentley (College of Medicine, Pennsylvania State University, USA), CaM12; John Krupinski (University of Manchester, UK), AC8; Jack Fransen (Department of Cell Biology, Radboud University Nijmegen Medical Centre, Netherlands), XProt; Manuela Zaccolo (University of Glasgow, UK), RII-CFP and C-YFP. This work was supported by the Deutsche Forschungsgemeinschaft (SFB 688, TP B6). The authors declare no conflict of interests.

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