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  • Research article
  • Open Access
  • Open Peer Review

AT1-receptor response to non-saturating Ang-II concentrations is amplified by calcium channel blockers

  • 2,
  • 2,
  • 3,
  • 1, 2,
  • 1Email author and
  • 1
BMC Cardiovascular DisordersBMC series – open, inclusive and trusted201717:126

https://doi.org/10.1186/s12872-017-0562-x

©  The Author(s). 2017

  • Received: 19 December 2016
  • Accepted: 9 May 2017
  • Published:
Open Peer Review reports

Abstract

Background

Blockers of angiotensin II type 1 receptor (AT1R) and the voltage gated calcium channel 1.2 (CaV1.2) are commonly used for treatment of hypertension. Yet there is little information about the effect of physiological concentrations of angiotensin II (AngII) on AT1R signaling and whether there is a reciprocal regulation of AT1R signaling by CaV1.2.

Methods

To elucidate these questions, we have studied the Ca2+ signaling response to physiological and pharmacological AngII doses in HEK293a cells, vascular smooth muscle cells and cardiomyocytes using a Ca2+ sensitive dye as the principal sensor. Intra-cellular calcium recordings were performed in presence and absence of CaV1.2 blockers. Semi-quantitative imaging methods were used to assess the plasma membrane expression of AT1R and G-protein activation.

Results

Repeated exposure to pharmacological (100 nM) concentrations of AngII caused, as expected, a down-regulation of the Ca2+ response. In contrast, repeated exposure to physiological (1 nM) AngII concentration resulted in an enhancement of the Ca2+ response. The up-regulation of the Ca2+ response to repeated 1 nM AngII doses and the down-regulation of the Ca2+ response to repeated 100 nM Angll doses were not accompanied by a parallel change of the AT1R plasma membrane expression. The Ca2+ response to 1 nM of AngII was amplified in the presence of therapeutic concentrations of the CaV1.2 blockers, nifedipine and verapamil, in vascular smooth muscle cells, cardiomyocytes and HEK293a cells. Amplification of the AT1R response was also observed following inhibition of the calcium permeable transient receptor potential cation channels, suggesting that the activity of AT1R is sensitive to calcium influx.

Conclusions

Our findings have implications for the understanding of hyperactivity of the angiotensin system and for use of Ca2+ channel blockers as mono-therapy in hypertension.

Keywords

  • Calcium
  • AT1R
  • Cell imaging
  • VGCC
  • hypertension

Background

Angiotensin II (AngII) is an important modulator of the sympathetic nervous system, cardiac function, blood pressure and salt excretion. The main receptor of AngII in the cardiovascular system is the angiotensin II type 1 receptor (AT1R), which is a Gαq-protein coupled receptor (GqPCR). Binding of AngII to AT1R results in activation of phospholipase C, release of inositol 1,4,5-triphosphate (IP3) and Ca2+ mobilization from intracellular stores. It is well documented that AngII activation of AT1R can be followed by desensitization [1, 2]. The question whether AT1R desensitization has clinical implications remains to be resolved, since the majority of cell signaling studies on AT1R have been carried out using concentrations of AngII that are at least three orders of magnitude higher than circulating levels [2–6]. Angiotensin receptor blockers and angiotensin converting enzyme (ACE) inhibitors are, along with voltage gated calcium channel (VGCC) blockers, among the most commonly used antihypertensive drugs. These drugs are used alone or in combination [7]. Several lines of evidence suggest that activation of AT1R can increase the activity of VGCC [8–10]. Little is however known about the effect of VGCC blockers on the activity of AT1R. This is a highly relevant question, since VGCC blockers are sometimes given as mono-therapy.

In the current study, we have compared the AT1R signaling pattern in response to repeated application of physiological and pharmacological concentrations of AngII, using a Ca2+ sensitive dye as the principal sensor. The effects of physiological concentrations of AngII on the AT1R signal were then examined in the presence of the VGCC inhibitors nifedipine and verapamil in therapeutically relevant concentrations. Since there is emerging evidence that some G-protein coupled receptors (GPCR) may be calcium sensitive [11–13], we also determined the effect of physiological concentrations of AngII on the AT1R signal in the presence of inhibitors of transient receptor potential cation channels (TRPC), another pathway for Ca2+ entry into the cell. The majority of experiments have been performed using a human embryonic kidney cell line 293a (HEK). In order to validate the physiological significance of our findings, key protocols were also performed using rat cardiomyocytes in primary culture.

Methods

Cells

Primary rat ventricular cardiomyocytes (RVCM) were obtained from 3 to 5 day old Sprague Dawley (Scanbur, Sollentuna, Sweden) and cultured on 18 mm diameter coverslips for 5 days previously described [14] using a modified growth medium. Growth medium was either a 2:1 mixture of DMEM/F-12:PC-1, supplemented with 2.5% FBS and 0.05 pM of AngII or DMEM for primary cell isolation (Gibco), 1:1000 Cardiomyocyte Growth Supplement (Pierce), 10% FBS and 50 pM AngII. Rats were euthanized by rapid decapitation and the heart removed for generation of cardiomyocyte cultures. Quality of culture was determined using a cardiomyocyte characterization kit (Chemicon). Cardiomyocytes were cultured for 5 days prior to experiment and contracting clusters, seen with transmission light, were selected for recording. Expression of cardiomyocyte markers were confirmed using Troponin I (Chemicon) and Desmin (Chemicon) antibody staining according to manufacturer’s protocol. Rat aortic smooth muscle cells (ASMC, catalog number R6110, 3H Biomedical, ScienCell) were cultured according to manufacturer’s instruction. Briefly, cells were thawed and plated on poly-L-lysine coated coverslips in complete smooth muscle cell medium (SMCM, ScienCell) including 2% FBS and supplemented with 0.05 pM of AngII. Cells were cultured for three to 5 days before experiments, cells from initial plating and first passage were used. Cells were confirmed to express anti-α-smooth muscle actin by immunostaining (antibody from Abcam). Human Embryonic Kidney 293a (HEK, catalog number R70507, Thermo Fisher Scientific) cells were plated on 18 mm diameter coverslips 2 days prior to transfection with Exgen500 (Fermentas), experiments were performed 24 h after transfection. Manufacturers protocols were used for transfection using 1 μg DNA per transfection. HEK cells between passage 3 and 20 was used in experiments. RT-PCR was used to confirm RNA expression of CaV1.2 (Fig 2a).

All experiments were performed according to Karolinska Institutet regulations concerning care and use of laboratory animals, and were approved by the Stockholm North ethical evaluation board for animal research.

Reagents and constructs

All chemicals and drugs were obtained from Sigma-Aldrich, if not otherwise stated. Bradykinin was purchased from Abcam. 3,5-bis(trifluoromethyl)pyrazole (BTP2) a TRPC channel blocker was obtained from Santa Cruz. Rat angiotensin II type 1 receptor, AT1R, with Venus fused at C-terminus was used for Ca2+ experiments [15]. The HA-tagged AT1R was made by insertion of PvuI restriction site through mutagenesis in the first extracellular loop between Pro331 and Phe332. Agilent QuikChangeII Site-Directed Mutagenesis kit together with the following primers were used: sense 5′ – GGTGATTGCCGAACGATCGGGGCCAGCGGTAC and antisense 5′ GTACCGCTGGCCCCGATCGTTCGGCAATCACC. The product was digested with PvuI (Thermo Scientific), and RSYPYDVPDYARS (Hemaglutinin flanked by PvuI sites) was inserted through ligation (Phusion Hot Start II, ThermoFisher). Structure of the construct was verified by using BigDye® Terminator V3.1 Cycle Sequencing Kit (Applied Biosystems). pGβ-2A-YFP-Gγ2-IRES-GαqmTq [16] was kindly provided by the lab of Th. W. J. Gadella and used for Gαq-Gβγ FRET (Förster Resonance Energy Transfer) measurements.

Membrane recruitment

Control and cells exposed to AngII were fixed using 2% paraformaldehyde for 10 min and stained using 1:1000 dilution of Alexa-594 fluorescence conjugated hemagglutinin (HA) antibody (Invitrogen). Cells were imaged using a Zeiss LSM780 in two channels for Venus and Alexa-594 fluorescence. Mean membrane and cytosol fluorescence intensities were calculated. Membrane fraction is reported as normalized to control ratio between mean membrane to cytosolic intensities.

PCR

Approximately 107 HEK cells were used for RNA extraction using manufacturer’s protocols (RNeasy Plus, Qiagen). cDNA was obtained using iScript cDNA kit (BioRad) following manufacturers protocols. Primers for CaV1.1, CaV1.2 and CaV1.4 were designed and used for PCR amplification (Phusion High-Fidelity DNA polymerase, Thermo Scientific). PCR amplification was run using the manufacturer’s protocols with the following program changes: 40 cycles with 10s extension time. Annealing temperature was 56 °C for CaV1.4, 58 °C for CaV1.1 and CaV1.2. The product was run on a 2% Agarose gel (1:10,000 GelRed, Biotium) and imaged on a Li-Cor Oddyssey system. The following primer pairs were used at 0.5 μM:

CaV11_4689F CAAGAGGAGTATTATGGCTATCGG,

CaV11_5252R GGTCAGCAGTCCCTTCAGCA,

CaV12_6568F CTGCGACATGACCATAGAGG,

CaV12_7188R CAGCGAGGCTCGTACAGA,

CaV14_5045F CAAAGCCACGATGGTCTCCC and.

CaV14_5545R TTTCGCCCACGATGATAGG (Cybergene).

[Ca2+]i measurements

Intracellular Ca2+ concentration ([Ca2+]i) measurements carried out in HEK cells were performed using the fluorescent Ca2+ indicator Fura Red (Invitrogen). Cells were loaded with Fura Red using a buffer consisting of 5 μM Fura Red and 1 μl of 20% Pluronic F-127 (Invitrogen) diluted in KREBs (110 mM NaCl, 4 mM KCl, 1 mM NaH2PO4xH2O, 25 mM NaHCO3, 1.5 mM CaCl2x2H2O, 1.25 mM MgCl2x6H20, 10 mM Glucose and 20 mM HEPES, pH 7.40, sterile filtered). Cells grown on #1.5 18 mm diameter coverslips (Warner Instruments) were rinsed with KREBs buffer and loaded for 45 min with the loading buffer at 37 °C with 5% CO2. After 45 min cells were washed again with KREBs prior to being mounted in a perfusion chamber (Chamlide). Through the use of heated optical table and in-line perfusion feed heater the cells were kept at 37 °C throughout experiments. Loading protocol was evaluated by testing different loading times, both at room temperature and in incubator. The minimal dye concentration usable was identified through Ca2+ calibration of serial dilution of dye.

CaV1.2 inhibitors, nifedipine and verapamil, were both mixed fresh for each experimental day. Solutions were kept in fridge and protected from light. We chose to use a high concentration of nifedipine, 100 μM, to compensate for loss of active drugs during the longer experiments. Degradation was unavoidable within each experiment with cells and solutions kept at 37 °C. UV and 488 nm laser excitation also contributed to some drug degradation throughout experiments.

[Ca2+] calibration was done by the following protocol, first a 3 min long washout of the KREBs buffer using a 0 [Ca2+] containing buffer (110 mM NaCl, 4 mM KCl, 1 mM NaH2PO4xH2O, 25 mM NaHCO3, 1.25 mM MgCl2x6H20, 250 μM EGTA, 10 mM glucose and 20 mM HEPES, pH 7.40, sterile filtered) followed by 1 μM ionomycin in 0 [Ca2+] buffer for 4 min. Calibration ended with 7 min of 1.5 mM [Ca2+] KREBs buffer perfusion.

The strongest AT1R-expressing cells were excluded from the analysis. Cells not responding to the first stimuli were also excluded. Cells were individually selected and the mean Ca2+ dye intensity was calculated and calibrated using the data from the end of each experiment. The mean intensity ratio, R, between the 488 nm excitation image and the 405 nm excitation image for each cell was calculated for each timepoint. Rmin and Rmax are the minimum and maximum intensity ratios obtained during calibration. Calibrated [Ca2+]i, was calculated with kd provided by Invitrogen as:
$$ {\left[{Ca}^{2+}\right]}_i={k}_d\frac{R-{R}_{min}}{R_{max}- R} $$

Ca2+ measurements of RVCM were performed using the fluorescent Ca2+ indicator Oregon Green BAPTA-1 (Invitrogen) with the same protocol as described for Fura Red. Oregon Green BAPTA1 was chosen as its weaker binding affinity compared to Fura Red made it insensitive to background Ca2+ sparklets in RVCM. Calibration was not possible for the RVCM Ca2+ experiments, thus the non-calibrated Ca2+ response for each cell was obtained by the mean intensity within each selected cell.

Nifedipine effects on bradykinin activation of bradykinin B2 receptors were evaluated through comparing signaling strength with or without pretreatment of 100 μM nifedipine on 100 nM bradykinin (Abcam) treatment. Experiments were performed on Oregon Green Bapta 1 loaded HEK cells. Cells were loaded in the same manner as for Fura Red but with 5 μM Oregon Green BAPTA-1. All other parameters were kept identical as for the 1 nM AngII experiments described above. Each experiment was calibrated as per the protocol described above.

Peak amplitudes were in all experiments calculated as peak above baseline. Baseline was defined as the mean intensity of a stable part of the trace prior to the peak. This calculation reflects the cytosolic increase in concentration rather than the absolute concentration.

Gαq dissociation

HEK cells expressing AT1R and Gαq-Gβγ FRET sensor were exposed to 1 nM AngII with or without 3 min pretreatment of 100 μM nifedipine. FRET was measured using a Zeiss LSM 510 LIVE microscope with a 40×/1.2 water immersion objective and excitation of 405 nm. YFP signal was corrected for mTurquoise bleedthrough and used as FRET intensity. Transfecting HEK cells with GαqmTq allowed for determination of donor bleed through into FRET channel. Bleed through coefficient, b, was calculated as ImTq/IFRET. In experiments, FRET channel intensities were then corrected as: FRETcorrected = IFRET – b*ImTq. FRET ratios were finally calculated as ImTurquoise / IFRETcorrected. Increase in FRET ratio corresponds with increased Gq-protein dissociation.

Statistical analysis

Statistical analysis was performed using Matlab, one way Anova analysis and subsequent multiple comparison tests based on studentized range distribution. Quantified data is shown as mean value +/− SEM.

Results

AT1R desensitization is dose-dependent

The majority of studies on AT1R signaling have used AngII concentrations that are at least three orders of magnitude higher than the circulatory levels [1, 2, 17, 18], which range between 20 and 100 pM [5]. Repeated exposure to non-physiological AngII concentrations have consistently been shown to result in desensitization of the signaling response. Rat renal interstitial fluid has been reported to reach 3-5 nM AngII under pathological conditions [19–21]. Here we have compared the effect of doses of 1 nM and 100 nM AngII on the calcium signal from HEK cells expressing Venus tagged AT1R and loaded with the Ca2+ sensitive dye Fura Red. Cells were exposed to three consecutive AngII doses, each 2 min long. The time delay between the 1st and 2nd exposure was 5 min, a common timescale for studies of desensitization [6, 17]. The 3rd exposure to AngII was performed 30 min after the 2nd exposure; a delay selected to test recovery of receptor signaling. Representative [Ca2+]i responses to serial AngII exposure are shown in Fig. 1a. The 1st exposure to 100 nM AngII resulted in a seven-fold stronger [Ca2+]i, response relative the 1st exposure to 1 nM AngII (Fig 1b). The 2nd exposure to 100 nM AngII shows a significant desensitization. In contrast, no desensitization was observed following the 2nd exposure to 1 nM AngII. A partial recovery of the [Ca2+]i response to 100 nM AngII was observed following the 3rd exposure. The [Ca2+]i response to the 3rd exposure to 1 nM AngII was significantly stronger than the [Ca2+]i response following the 1st exposure (Fig 1b). Application of cyclopiazonic acid, an inhibitor of the sarcoendoplasmic reticulum calcium transport ATPase, completely abolished the [Ca2+]i response to AngII, indicating that the increase in [Ca2+]i is mainly due to release of Ca2+ from intracellular stores (data not shown).
Fig. 1
Fig. 1

Physiological and pharmacological AngII results in significantly different Ca2+ signaling pattern. a Representative [Ca2+]i traces from Fura Red loaded HEK cells treated with repeated 1 nM (left) or 100 nM (right) AngII for two minutes at times indicated by the arrows, with 5 min recovery after the first exposure and 30 min after second exposure. b Quantification of peak [Ca2+]i response to repeated AngII treatment 1 nM (open bars) and 100 nM (grey bars) as shown in A. Responses are calculated as peak signal above baseline. *: p ≤ 0.05 compared to 1st AngII exposure, **: p ≤ 0.01 compared to 1st AngII exposure. n = 4 - 8 experiments. c The cartoon shows the principle used to detect AT1R in plasma membrane. Signal from Anti-HA-Alexa594 antibody show plasma membrane AT1R, Venus signal show the total abundance of AT1R. Bars show relative change of plasma membrane AT1R abundance 3 min and 30 min after exposure to AngII, 1 nM AngII open bars and 100 nM grey bars. Data is normalized to untreated control membrane fraction. n = 5 experiments

Next we examined to which extent the dose-dependent [Ca2+]i response to AngII could be attributed to effects on AT1R plasma-membrane expression. For this purpose, we expressed AT1R with an extracellular hemagglutinin (HA) tag in addition to a Venus tag at the intracellular C-terminus of HEK cells. Cells were fixed and labeled with an Alexa-594 conjugated anti-HA antibody 3 or 30 min after exposure to AngII. The AT1R fraction inserted in the plasma membrane was calculated as the ratio between the fluorescence intensity of Alexa-594 (plasma membrane AT1R) and Venus (total AT1R) (Fig 1c). The effect of repeated exposure to 1 and 100 nM AngII on the fraction of AT1R in the plasma membrane were subtle and did not parallel the changes in Ca2+ response.

AT1R signaling is upregulated by nifedipine and verapamil

Drugs based on either the nifedipine or verapamil structure, are inhibitors of the L-type VGCC, CaV1.2 subtype [22], and are commonly used in the treatment of hypertension. Since there is emerging evidence of a cross-talk between the CaV1.2 and AT1R [8, 23, 24], we tested if nifedipine and verapamil would affect the response to AngII in AT1R expressing HEK cells. CaV1.2 is widely expressed, and the RNA expression of CaV1.2 in the HEK cells was confirmed using RT-PCR (Fig 2a). HEK cells were pretreated for 5 min with 1 μM nifedipine or 2 μM verapamil and then exposed to 1 nM AngII for 2 min (Fig. 2b-c). The concentration of nifedipine [25] and verapamil [26] were selected based on human plasma levels. Unexpectedly, we found that in the presence of either drug, there was a robust and highly significant increase of the calcium response to the initial dose of 1 nM AngII in both nifedipine and verapamil treated cells (Fig. 2b and c) To ensure that complete inhibition of CaV1.2 was achieved, as nifedipine is sensitive to heat and light (supplier’s information), we also tested a higher dose, 100 μM nifedipine (Fig 2b). No further increase in response was observed. To eliminate possible errors from drug degradation during the one-hour long protocol we elected to continue with the higher dose of nifedipine.
Fig. 2
Fig. 2

CaV1.2 inhibition up-regulates physiological AT1R signaling. a Gel image shows a typical RT-PCR of HEK mRNA using primers for CaV1.1(left), CaV1.2 (center) and CaV1.4 (right). As a negative control, each reaction was done omitting the cDNA. Expected band size is, for CaV1.1564 bp, CaV1.2621 bp and CaV1.4501 bp. Gene Ruler 1 kb DNA ladder was used as a size marker. b Quantification of peak [Ca2+]i response to 1 nM AngII exposure alone or in presence of 1 μM or 100 μM nifedipine in Fura Red loaded HEK cells. n = 4 - 6 experiments. **: p ≤ 0.01 compared to 1 nM AngII exposure alone. c Quantification of peak [Ca2+]i response to 1 nM AngII exposure alone or in presence of 2 μM verapamil in Fura Red loaded HEK cells. n = 10 experiments. *: p ≤ 0.05 compared to 1 nM AngII exposure alone. d Representative [Ca2+]i traces from Fura Red loaded HEK cells treated with repeated 1 nM AngII for two minutes at times indicated by the arrows in the presence of 100 μM nifedipine. e Quantification of peak [Ca2+]i response to repeated 1 nM AngII treatment as shown in D. Peaks are calculated as peak signal above baseline. Open bars show 1 nM AngII alone and grey bars show 1 nM AngII in presence of 100 μM nifedipine. n = 4 - 6 experiments. **: p ≤ 0.01 compared to vehicle

The effect of nifedipine on the [Ca2+]i response to 1 nM AngII was studied using the same three-pulse protocol as described above with the addition of a pretreatment with 100 μM nifedipine (Fig. 2d and e). There was a subsequent reduction of the [Ca2+]i response to the 2nd and 3rd dose of 1 nM AngII, suggesting a certain desensitization. There was no increase in the relative distribution of AT1R inserted in the plasma membrane in nifedipine pretreated cells compared to control cells (data not shown).

We next asked if the nifedipine potentiation of the response to 1 nM AngII could be attributed to increased G-protein activity. To address this question we examined the dissociation of Gαq and Gβγ subunits induced by AT1R activation using a Gαq-Gβγ FRET sensor [16] (Fig 3a). Increased FRET ratio indicates increased Gq-protein dissociation. The FRET ratio between GαqmTq (donor) and YFP-Gγ2 (acceptor) in response to 1 nM AngII was studied in the presence or absence of nifedipine. No significant difference in FRET ratio was observed between nifedipine treated and control cells (Fig. 3b and c). This suggests that up-regulation of AT1R in nifedipine pretreated cells is not due to increased G-protein activation.
Fig. 3
Fig. 3

G-protein activation is not affected by nifedipine. a The cartoon show the principle of the FRET based assay for detection of G-protein activity. Activity is detected as a decreased FRET signal (increased FRET ratio) when Gαq dissociate from the βγ complex. b Representative FRET ratio traces of HEK cells expressing pGβ-2A-YFP-Gγ2 in response to 2 min exposure to (at arrow) 1 nM AngII in absence (left) or presence of 100 μM nifedipine (right). c Quantification of peak FRET ratio, in response to 1 nM Ang II alone (open bar) and in presence of nifedipine (grey bar) in HEK cells. n = 9 - 11 experiments

AT1R signaling is upregulated by inhibition of TRPC channels

We examined whether the potentiation of the AngII signal observed following CaV1.2 inhibition may also occur following the inhibition of other calcium entry pathways. The TRPC channels are ubiquitously expressed ion channels with high calcium permeability. It was recently reported that the response to AngII was enhanced in arterial myocytes from TRP deficient mice [27]. Here we used BTP2 to inhibit the activity of TRPC channels. BTP2 blocks the calcium influx via TRPC3 and TRPC5 (but not transient receptor potential cation channel subfamily V6) with an IC50 of ~0.1-0.3 μM, and is generally considered to have a many-fold greater potency than other available BTP2 inhibitors [28, 29]. HEK cells were first exposed to 1 nM AngII. This was followed by 5 min treatment with either vehicle or 10 μM BTP2. The cells were then exposed to a second dose of 1 nM AngII (Fig 4a). The relative change in response between first and second AngII exposure was calculated, and showed a significant potentiation of the AngII following BTP2 treatment (Fig 4b).
Fig. 4
Fig. 4

TRPC inhibition up-regulates physiological AT1R signaling. a Representative [Ca2+]i traces from Fura Red loaded HEK cells exposed to 1 nM Ang II (at arrows) followed by 5 min treatment with either vehicle (left) or 10 μM BTP2 (right). b Relative change in response between first and second AngII stimulation for vehicle (open bar) and BTP2 (grey bar). n = 4 - 5 experiments, **: p ≤ 0.01 compared to vehicle

To test whether the sensitivity for calcium influx observed for the AT1R cells is a general phenomenon for GqPCR, we also tested if nifedipine exposure would potentiate [Ca2+]i signaling from the bradykinin receptor B2 (B2 receptors), which are constitutively expressed in HEK cells. Exposure of cells to 100 nM bradykinin resulted in similar [Ca2+]i response as for 100 nM AngII while in the presence of nifedipine a weaker [Ca2+]i response was observed suggesting that not all GqPCRs are potentiated by nifedipine.

AT1R signaling in cardiovascular cells is upregulated by VGCC inhibition

The finding that AT1R signaling is enhanced following inhibition of Ca2+ influx may have implications for AngII regulation of cardiovascular function, since AT1R inhibitors are commonly used to treat cardiovascular disease [30, 31] and CaV1.2 is abundantly expressed throughout arterial and cardiac tissue [32]. We therefore tested if our findings in HEK cells could be reproduced in rat derived primary cultures of both aortic smooth muscle cells (ASMC) and ventricular cardiomyocytes (RVCM).

ASMC at three to 5 days in vitro expressing alpha-smooth muscle actin were used for these experiments. Cells were loaded with the Ca2+ sensitive dye Oregon Green BAPTA1 and exposed to two consecutive physiological doses of AngII thirty minutes apart. Typical [Ca2+]i response to 1 nM AngII exposures are shown in Fig. 5a (upper trace). A weaker response was observed at the 2nd exposure, in contrast to what had been found in HEK cells (Fig. 1a-b). We next tested if blocking of CaV1.2 by nifedipine or verapamil would affect the AngII response in ASMC. Nifedipine and verapamil doses were selected based on blood plasma levels reported in patients [33, 34]. ASMC were exposed to 1 nM AngII for 2 min and then allowed 15 min recovery followed by 15 min treatment with vehicle, nifedipine or verapamil and finally exposed to 1 nM AngII for 2 min. Example traces, of the [Ca2+]i response in presence of nifedipine and verapamil are shown in Fig. 5a (middle and lower trace). The ratio between the second and first AngII response was calculated. Both nifedipine and verapamil treatment were found to strongly (2-fold) enhance the [Ca2+]i response to AngII (Fig 5b) in ASMC.
Fig. 5
Fig. 5

CaV1.2 regulates AT1R signaling in rat aortic smooth muscle cells and ventricular cardiomyocytes. (a) Representative [Ca2+]i traces of Oregon Green Bapta 1 loaded rat aortic smooth muscle cells treated with two minutes of 1 nM AngII at times indicated by arrows. Smooth muscle cells were co-treated from 15 min after first AngII exposure throughout the experiment with vehicle (upper), 0.2 μM nifedipine (middle) and 0.9 μM verapamil (lower). (b) Relative change between first and second response to AngII stimulation for vehicle (open bar), 0.2 μM nifedipine (dark grey bar) and 0.9 μM verapamil (light grey bar). Peaks are calculated as signal above baseline. n = 6 - 7 individual experiments, *: p ≤ 0.05 compared to vehicle. (c) Representative [Ca2+]i traces of Oregon Green Bapta 1 loaded rat ventricular myocytes treated with two minutes of 1 nM AngII at times indicated by arrows. Myocytes were co-treated from 15 min after first AngII exposure throughout the experiment with vehicle (upper), 0.2 μM nifedipine (middle) and 0.9 μM verapamil (lower). (d) Relative change between first and second response to AngII stimulation for vehicle (open bar), 0.2 μM nifedipine (dark grey bar) and 0.9 μM verapamil (light grey bar). Peaks are calculated as signal above baseline. n = 6 - 10 individual experiments, *: p ≤ 0.05 compared to vehicle, #: p ≤ 0.05 compared to nifedipine

The same protocol was used to study the effects of VGCC inhibition on AT1R signaling in RVCM. RVCM at 5 days in vitro expressing the cardiomyocyte marker Troponin I and displaying contractile activity were used for these experiments. A representative [Ca2+]i response trace to 1 nM AngII exposure is shown in Fig. 5c (upper trace). A stronger response was observed at the 2nd exposure, similar to what had been found in HEK cells (see Fig. 1a-b) but in contrast with ASMC response (see Fig 5b). Blocking of CaV1.2 by nifedipine (Fig. 5c middle trace) or verapamil (Fig. 5c bottom trace) were both found to strongly enhance the response to AngII (Fig 5d). Notably, the enhancement by verapamil was significantly stronger than for nifedipine treatment in RVCM but not in ASMC (Fig 5b).

Discussion

Blockers of the AT1 receptor and calcium channel CaV1.2 belong to the most commonly used drugs for treatment of hypertension and cardiovascular disease. Knowledge about AT1R signaling is to a large extent based on studies performed with AngII concentrations that exceed physiological and pathophysiological concentrations by several orders of magnitude. There are few studies, if any, that have examined whether CaV1.2 may control the strength of the AT1R signal. Here we show that the desensitization of AT1R observed following pharmacological concentrations of AngII, does not occur with concentrations of AngII in the physiological range and that acute exposure to therapeutical concentrations of CaV1.2 inhibitors amplify AT1R signaling.

AT1R is the primary receptor mediating the biological functions of AngII and is abundantly expressed in the cardiovascular and renal systems. AT1R signals through release of Ca2+ from the intracellular stores via activation of phospholipase C, IP3 and the IP3 receptor [2]. Both IP3 production and Ca2+ mobilization are common read-outs of AT1R signaling [2, 6, 35, 36]. By performing real-time recordings of [Ca2+]i, the dynamics of AT1R signaling can be monitored continuously in live cell systems, whilst most IP3 protocols rely on extraction of IP3 at individual time-points. A concern when using Ca2+ sensitive dyes is the possibility of toxic effects of the dye and of phototoxicity during prolonged recordings. Our protocols were optimized to ensure minimal toxic effects of loading, minimized pixel dwell time and laser power. End point calibrations were performed to ensure that separate measurements are comparable across different systems.

The majority of previous studies on AT1R signaling have used AngII concentrations that are several orders of magnitude higher than the circulating and/or tissue levels of AngII [19, 21, 37]. Circulating levels of AngII in healthy humans have been reported to be in the order of 20 pM [4, 5], Patients with heart failure often have several-fold increased levels of AngII [38]. The primary cardiomyocyte cells used in this study were derived from rats. The circulating levels of AngII in rats have been reported to be approximately ten times higher than in humans [19, 37]. Local levels of AngII in tissue can be even higher, depending on local production and metabolism, reaching up to 3-5 nM in rat renal interstitial fluid [19–21].

Homologous desensitization of GPCRs is a well-known phenomenon, and numerous studies of desensitization have been carried out on AT1R [1, 2]. Most of these studies have been performed with concentrations of AngII in the range of 10-100 nM. In our study desensitization was only observed after exposure to the high concentration of AngII, while repeated exposure to 1 nM AngII did not result in desensitization. This lack of desensitization to low concentrations of AngII might explain why moderately increased endogenous AngII levels can have a deleterious long-term effect on cardiovascular and renal function.

Voltage-gated calcium channels serve an important role for the control of [Ca2+]i homeostasis in almost all mammalian cells and play a particularly dynamic role in excitable cells. The L-type VGCC 2, CaV1.2, is a common target for anti-hypertensive treatment. It was previously reported from several groups that AngII may modify the activity of the CaV1.2 [8–10, 39]. The results are not unambiguous, but the majority of studies have shown that AngII increases the CaV1.2 activity. Here we showed that pre-incubation of both HEK cells and cardiomyocytes with inhibitors of CaV1.2 activity results in a robust amplification of AngII triggered [Ca2+]i signal. Notably, this amplifying effect of the CaV1.2 inhibitors was observed with concentrations that are comparable to those used to treat hypertension [25, 26, 33, 34]. The amplification of the AngII signal that we observed in both nifedipine and verapamil treated cells may not be specific for CaV1.2, since it was also found following inhibition of the TRPC channels, which represent another ubiquitous Ca2+ influx pathway. We hypothesize that the AT1R complex is sensitive to changes in intracellular Ca2+ concentration, and that increased Ca2+ influx might have a desensitizing effect on the AT1R complex. In support of this hypothesis, it has been shown by other groups that signaling from another G-protein coupled receptor, the dopamine D2 receptors, is controlled by intracellular Ca2+ concentration [11, 12]. A recent study showed that increased calcium influx through either L-type voltage gated calcium channels or from intracellular stores result in D2 receptor desensitization [13]. The sensitivity of AT1R (current study) and D2 receptors [13] to Ca2+ influx was not shared by the bradykinin receptor, and is thus not a common phenomenon for all GPCRs. The molecular mechanism by which calcium influx modulates the strength of AT1R signaling remains elusive. We found no evidence for a recruitment of receptors to the plasma membrane or altered Gq-protein activity.

Conclusions

In conclusion, this study provides novel and clinically relevant information about how inhibition of CaV1.2 upregulate AT1R signaling. Notably these effects were observed with physiological and therapeutically relevant concentration of AngII, nifedipine, verapamil. Calcium channel blockers are commonly used to treat hypertension, either in combination with other drugs such as inhibitors of AT1R or ACE, or as mono-therapy. When calcium channels blockers are used as mono-therapy, the possibility for stimulation of AT1R signaling should be taken into account. Notably, a recent meta-analysis, based on more than 40 studies, has shown that combination therapy that targets both the angiotensin system and the voltage gated calcium channels, will provide a better blood pressure control than mono-therapy that acts at one of these targets [40].

Abbreviations

ACE: 

Angiotensin Converting Enzyme

AngII: 

Angiotensin type II

AT1R: 

Angiotensin II type 1 receptor

BTP2: 

3,5-bis(trifluoromethyl)pyrazole

[Ca2+]i : 

Intracellular Ca2+ concentration

CaV1.2: 

Voltage gated calcium channel subtype 1.2

DMEM: 

Dulbecco’s Modified Eagle Medium

FRET: 

Förster Resonance Energy Transfer

Gαq : 

G-protein subunit αq

GPCR: 

G-protein coupled receptor

GqPCR: 

Gαq-protein coupled receptor

Gβγ : 

G-protein subunit βγ

HA: 

Hemagglutinin

HEK: 

Human Embryonic Kidney cell line 293a

IP3 : 

Inositol 1,4,5-triphosphate

RVCM: 

primary rat ventricular cardiomyocytes

TRPC: 

transient receptor potential cation channels

VGCC: 

Voltage Gated Calcium Channel

ASMC: 

Aortic smooth muscle cells

Declarations

Acknowledgements

The authors wish to thank Allen M. Samarel for valuable advice on the cardiomyocyte culture protocol, and W. J. Gadella for kindly providing the pGβ-2A-YFP-Gγ2-IRES-GαqmTq construct.

Funding

This study has been supported by the Swedish Heart Lung foundation (AA), the Swedish Research Council (AA, HB), the Magnus Bergvalls Foundation (LS), the Söderberg Foundation (AA), Stiftelsen Tornspiran (LS) and the Märta and Gunnar V Philipson Foundation (HB, LS).

Availability of data and materials

The data generated and analyzed within the current study are available from the lead author (KB) and corresponding author (AA) on reasonable request.

Authors’ contributions

The study was conceived by AA, HB and LS. Experimental design was made by KB, KK, HB, AA and LS. Experiments were performed by KB, KK and LS. Plasmid constructs were designed and generated by AB and KB. Data analysis was performed by KB, KK and HB. The manuscript was drafted by KB, HB, AA and LS. All authors read and approved the final manuscript.

Authors’ information

Not applicable.

Competing interests

The authors declare that they have no competing interests.

Consent for publication

Not applicable.

Ethics approval

All experiments were performed according to Karolinska Institutet regulations concerning care and use of laboratory animals, and were approved by the Stockholm North ethical evaluation board for animal research.

Publisher’s Note

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Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.

Authors’ Affiliations

(1)
Science for Life Laboratory, Department of Women’s and Children’s Health, Karolinska Institutet, PO Box 1031, 17121 Solna, Sweden
(2)
Science for Life Laboratory, Department of Applied Physics, Royal Institute of Technology, Stockholm, Sweden
(3)
Institute of Chemical Biology and Fundamental Medicine, 630090 Novosibirsk, Russia

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