Therefore, in the present study, the effects of A2BAR stimulation on EMT was investigated in human lung epithelial cells. Particularly, the role of this adenosine receptor subtype in directly modulating the epithelial-mesenchymal markers under physiological or inflammatory conditions (TGF-β1) was analyzed. Furthermore, the intracellular pathways activated by A2BAR, cAMP/PKA, and MAPK/ERK, involved in EMT, were explored.

### MATERIALS AND METHODS

### Material and Reagents

Human type II alveolar epithelial cells (A549, American Type Culture Collection, CCL-195), were kindly provided

by Dr. R. Danesi, University of Pisa, Pisa, Italy. The chemicals 2-[6-amino-3,5-dicyano-4-[4-(cyclopropylmethoxy) phenyl]pyridin-2-ylsulfanyl] acetamide (BAY 60-6583) and N- (4-acetylphenyl)-2-[4-(2,3,6,7-tetrahydro-2,6-dioxo-1,3-dipropyl-1H-purin-8-yl)phenoxy]-acetamide (MRS1706) were purchased from Tocris Bioscience (Bristol, United Kingdom). The RNeasy minikit was obtained from Qiagen. The Script cDNA synthesis kit was furnished by Bio-Rad s.r.l. Fluocycle II SYBR was from Euroclone (Milan, Italy). TGF-β1 was purchased from Sigma–Aldrich. All other reagents were obtained from standard commercial sources and were of the highest commercially available grade

### Cell Culture

Human type II alveolar epithelial cells (A549) were maintained in DMEM-F12 containing 10% FBS, 2 mM L-glutamine, 100 U/ml penicillin and 100 µg/ml streptomycin at 37◦C in a humidified 5% CO2 atmosphere. For the different experiments, cultures of cells were maintained in serum-free DMEM for 8 h prior to stimulation with cytokines or other reagents.

## RNA Extraction and Real-Time RT-PCR Analysis

A549 cells were untreated or treated with DMSO (CTRL) or BAY 60-6583 (100 nM) in the absence or presence of TGF-β1 (10 ng/ml) and MRS 1706 (1 µM) for the indicated time. At the end of treatments, cells were collected, and total RNA was extracted using Rneasy R Mini Kit (Qiagen, Hilden, Germany) as previously reported (Da Pozzo et al., 2014); residual DNA was removed using RNase free DNAse set (#79254, Qiagen, Hilden, Germany). The RNA purity was checked measuring the A260/280 ratio. cDNA synthesis was performed with 400 ng of RNA using i-Script cDNA synthesis kit (BioRad, Hercules, CA, United States) following manufacturer's instructions. Realtime RT-PCR reactions consisted of 25 µL Fluocycle R II SYBR R (Euroclone, Milan, Italy), 1.0 µL of both 10 µM forward and reverse primers, 2.5 µL cDNA (100 ng), and 20.5 µL of H2O. All reactions were performed for 40 cycles using the following temperature profiles: 98◦C for 30 s (initial denaturation); 55◦C for 30 s (annealing); and 72◦C for 3 s (extension). The primer sequences were listed in **Table 1**. When possible, the primers used were designed to span intron/exon boundaries and the β-actin was used as the housekeeping gene. The mRNA levels for each sample were normalized against β-actin mRNA levels, and the relative expression was calculated by using a Ct value. PCR specificity was determined by both the melting curve analysis and gel electrophoresis.

## Western Blotting Analysis

A549 cells (3.5 × 104 cell/cm2 ) were maintained untreated or treated with DMSO (CTRL) or BAY 60-6583 (100 nM) in the absence or presence of TGF-β1 (10 ng/ml) and MRS 1706 (1 µM) for 48 h. At the end 200 µl RIPA buffer were added for 60 min at 4 ◦C to lyse the cells. Fifty microgram of total proteins was diluted in Laemmli solution, resolved by SDS-PAGE (7.5%), transferred to PVDF membranes and probed overnight at 4◦C with primary anti-A1AR (diluted 1:200, sc-28995; Santa Cruz Biotechnology), anti-A2AAR (diluted 1:200, sc-13937; Santa Cruz Biotechnology), anti-A2BAR (diluted 1:150, sc-28996; Santa Cruz Biotechnology), anti-A3AR (diluted 1:200, sc-13938; Santa Cruz Biotechnology), anti-E-cadherin antibody (diluted 1:200; sc-7870; Santa Cruz Biotechnology), anti-N-cadherin antibody (diluted 1:200; sc-7939; Santa Cruz Biotechnology) anti-Vimentin (diluted 1:1000, #5741; Cell Signaling Technology) or β-actin antibody (diluted 1:1000; MAB1501, Merck KGaA, Darmstadt, Germany). The primary antibody was detected using appropriate secondary antibody. The peroxidase was detected using a chemiluminescent substrate (ECL, Perkin Elmer), and the images were acquired by photographic film or by LAS4010 (GE Health Care Europe, Uppsala, Sweden). Immunoreactive bands were quantified performing a densitometric analysis with Image J Software (version 1.41; Bethesda, MD, United States).

### Immunofluorescence Analysis

A549 cells were seeded at 3.5 × 104 cell/cm2 in chamber slide (BD Biosciences, San Jose, CA, United States). After treatment, cells were fixed in 2% paraformaldheyde in 0.1 M phosphate buffer, washed three times with PBS, rinsed, and blocked for 45 min with PBS containing 0.1% Triton-X 100 and 1% BSA. After washing, cells were incubated overnight at 4◦C with mouse monoclonal anti-β-actin antibody (diluted 1:500; MAB1501, Merck KGaA, Darmstadt, Germany) or a rabbit polyclonal anti-A2BAR (diluted 1:100, sc-28996; Santa Cruz Biotechnology) primary antibodies diluted in PBS containing 0.03% Triton-X 100 and 1% BSA overnight at 4◦C. After washing, to visualize the staining, cells were incubated with Alexa Fluor 488- and Alexa Fluor 568-labeled goat anti-mouse (1:500) or anti-rabbit (1:500) antibodies for 2 h at room temperature. Then slides were covered with Vectashield conjugated with DAPI (Vector Laboratories, Burlingame, CA, United States). Images were obtained with a Nikon Ni-E microscope, using a 20× objective with 1.45 NA and a recommended pinhole size of less than 1.0 mm, and equipped with digital camera Nikon Mod.DS-Ri2. The images were processed with ImageJ software.

## Cell Viability Assay

A549 cells were seeded at a density of 3 × 103 cells/well in a black, clear bottom PerkinElmer 96-well CellCarrierTM microplate (#6005550). After 24 h, the cells were starved for 8 h with non-complete medium; then, cells were treated with different concentrations of BAY 60-6583 (0.5 nM–1 µM) or TGFβ1 (5–20 ng/ml) alone or in combination for 48 or 72 h. Following the treatment period, cells were imaged with the EnSightTM multimode plate reader equipped with well-imaging module, and Kaleido Data Acquisition and Analysis Software. Brightfield images were taken before and 48 or 72 h after treatments to allow for a Cell Count comparison between both time points. The cell count was determined using the pre-defined Brightfield Cell Count algorithm provided by the Kaleido software. Cell with a mean area ≥100 µm2 were counted. Data were normalized

### TABLE 1 | Primers used for real-time RT-PCR.

fphar-09-00054 January 29, 2018 Time: 17:4 # 4

to the cell count at t0 and expressed as the number of cell per well.

### cAMP Quantification

A549 cells were seeded at a density of 3 × 103 cells/well in a black, clear bottom PerkinElmer 96-well CellCarrierTM microplate (#6005550). After 24 h, the cells were starved for 8 h with non-complete medium; then, cells were treated with different concentrations of BAY 60-6583 (0.1 nM–1 µM) in the absence or presence of TGF-β1 (10 ng/ml) in 100 µl of non-complete medium for 48 h. Cells were imaged with the EnSightTM multimode plate reader. The desensitization experiments were performed incubating the cells with BAY 60-6583 (100 nM) in the presence or absence of TGF-β1 (10 ng/ml) for 48 h. At the end the medium was changed and cells were treated with different concentrations of BAY 60- 6583 (0.1 nM–1 µM) for 15 min. Then, cells were lysed with 50 µl of 1X lysis buffer (PerkinElmer, #AL003C) supplemented with protease inhibitors and phosphatase inhibitors for 10 min with gentle shaking. Lysates were either tested immediately in AlphaLISA assays or frozen at –80◦C for later testing. cAMP levels were quantified using cAMP AlphaLISA kit (PerkinElmer, #AL312) following manufacturer's instructions. Standard curves for each AlphaLISA immunoassay were performed in the same diluent as the samples being tested (1X lysis buffer with supplements) using the recombinant standards provided in each kit. Curves were plotted with a sigmoidal concentration-response curve with variable slope. Quantitation of protein levels in cellular assays were interpolated off their respective standard curves.

## MAPK (Mitogen-Activated Phosphorylation Kinase) Assays

A549 cells were seeded at a density of 3 × 103 cells/well in 96 multi-well plate. After 24 h, the cells were starved for 8 h with non-complete medium; then, cells were treated with BAY 60- 6583 (100 nM) in the absence or presence of TGF-β1 (10 ng/ml) and MRS1706 (1 µM) for 5 min, 30 min, 6, 24, and 48 h. In some experiments, before incubation with BAY 60-6583 or TGF-β1, cells were pre-treated for 30 min with PD98059 (1 µM) (MEK, inhibitor). At the end of treatments, cells were fixed with 4% formaldehyde to preserve activation of specific protein modification. Levels of total and phosphorylated extracellular signal-regulated kinases (ERK1/2) were determined by ELISA assays, as previously reported (Giacomelli et al., 2015). Briefly, the cells were washed three times with wash buffer (0.1% Triton X-100 in PBS) and 100 µl of quenching buffer (1% H2O2; 0.1% sodium azide in wash buffer) was added and incubation was protracted for other 20 min. The cells were washed with PBS twice, and then 100 µl of blocking solution (1% BSA; 0.1% Triton X-100 in PBS) was added for 60 min. After blocking, cells were washed three times with wash buffer and the specific primary antibodies (anti-phospho ERK1/2, 1:500, sc-7383 Santa Cruz Biotechnology; anti-ERK1/2, 1:500, #4695 Cell Signaling Technology) were added on at 4◦C. Subsequent incubation with secondary HRP-conjugated antibodies and developing solution allowed a colorimetric quantification of total and phosphorylated levels. Blanks were obtained by treating cells in the absence of the primary antibody. The relative number of cells in each well was then determined using Crystal Violet solution. The results were calculated by subtracting the mean background from the values

obtained from each test condition; values were normalized to the number of cells in each well, and were expressed as the percentage of untreated cells (basal).

## Quantification of E-cad and N-cad Proteins

A549 cells were seeded at a density of 3 × 103 cells/well in a black, clear bottom PerkinElmer 96-well CellCarrierTM microplate (#6005550). After 24 h, the cells were starved for 8 h with non-complete medium; then, cells were treated with different concentrations of BAY 60-6583 (0.1 nM–1 µM) in the absence or presence of TGF-β1 (10 ng/ml) in 100 µl of non-complete medium for 48 h. In some experiments, before incubation with BAY 60-6583 or TGF-β1, cells were pre-treated for 30 min with PD98059 (1 µM, MEK, inhibitor), H89 (100 nM, PKA inhibitor) or 8-Br-cAMP (100 nM–1 µM, cAMP analog & PKA activator). Then, cells were imaged with the EnSightTM multimode plate reader. After imaging, cells were lysed with 50 µl of 1X lysis buffer (PerkinElmer, #AL003C) supplemented with protease inhibitors and phosphatase inhibitors for 10 min with gentle shaking. E-cad were quantified using E-cadherin AlphaLISA kit (PerkinElmer, #AL370), and N-cad were quantified using N-cadherin AlphaLISA kit (PerkinElmer, #AL379) following the manufacturer's instructions. Standard curves for each AlphaLISA immunoassay were performed in the same diluents as the samples being tested (1X lysis buffer with supplements), using the recombinant standards provided in each kit. Curves were plotted with a sigmoidal concentration-response curve with variable slope. Quantitation of protein levels in cellular assays were interpolated off their respective standard curves.

### Statistical Analysis

The Graph-Pad Prism program (GraphPad Software Inc., San Diego, CA, United States) was used for data analysis and graphic presentation. All data are the mean ± SEM of at least three different experiments. Statistical analysis was performed by one-way analysis of variance (ANOVA) with Dunnett post hoc analysis to compare the data to the control, or two-way ANOVA with Bonferroni correction and two-sided tests for multiple comparisons. EC50 values were reported as mean of the values obtained in at least three independent experiments performed in duplicate ± SEM. P ≤ 0.05 was considered statistically significant.

## RESULTS

### Adenosine Receptor Expression in Human Epithelial Lung Cells and Its Modulation by TGF-β1

The A549 human alveolar epithelial cells have been widely used to study the fibrotic process in the lung and related EMT mechanism (Kim et al., 2007; Ji et al., 2016). Furthermore, these cells were maintained in serum-free medium to increase the epithelial phenotype (Dong et al., 2014).

First, the expression of the AR subtypes in A549 cells was evaluated after incubation in serum-free medium for 48 h (**Figure 1A**). All the ARs were expressed under this condition, and the A2BAR subtype was the most expressed with a fold change of approximately 200 (**Figure 1A**).

TGF-β1 has been reported to affect the expression of several proteins (Zhang, 2017); thus, the effects of cytokine treatment on AR expression were evaluated (**Figures 1B–D**). Challenging the A549 cells for 48 h with increasing concentrations of TGF-β1 (5–20 ng/ml) modified the expression of the different adenosine subtypes. Particularly, A1AR expression was significantly increased when a high concentration of TGF-β1 was used; conversely, A2A and A3 receptor expression was slightly decreased when increased concentration of the cytokine was applied. Regarding A2BAR expression, low concentrations of TGF-β1 (5 or 10 ng/ml) were not sufficient to modify the receptor expression; by contrast, a higher concentration of the cytokine significantly decreased its expression. In this light, further experiments were performed using a maximum concentration of the cytokine of 10 ng/ml for 48 h to minimize the change in the receptor expression.

## TGF-β1 Effects on A2BAR Functionality and Lung Cell Growth

A2BAR stimulation or blockade has been correlated to different levels of cell proliferation depending on the cell type and culture condition (Phosri et al., 2017; Zhou et al., 2017). Thus, the effects of A2BAR stimulation in the absence or presence of TGF-β1 were evaluated in the lung cells (**Figure 2**). The selective agonist BAY 60-6583 was used to stimulate selectively the A2BAR subtype. The compound could slightly increase the cell proliferation when used at low concentration (50–100 nM) for 48 or 72 h treatment (**Figure 2A**), conversely, it produced a decrease in proliferation when used at a high concentration (1 µM), with a significant effect after 72 h of treatment (13970 ± 488 n◦ of cells CTRL, 11038 ± 816 n◦ of cells BAY; P ≤ 0.05). TGF-β1 alone presented a hormetic concentrationresponse course: the lower concentration possessed positive effects on cell proliferation that are lost at higher concentrations (**Figure 2B**). The combined treatment with BAY 60-6583 and TGF-β1 for 48 h did not significantly affect the cell proliferation (**Figure 2C**). These data confirmed the diverse role of A2BAR in the cell proliferative mechanism. Based on such results, further experiments were then performed using BAY 60-6583 at a concentration (100 nM) not affecting significantly the A549 cell proliferation.

Next, the effect of cytokine treatment on A2BAR functionality was evaluated by measuring the agonist-mediated cAMP production (**Figure 2D**). The potency of BAY 60-6583 was in the nanomolar range (EC50 = 17.2 ± 2.5 nM), as derived by agonist concentration-response curves, it is a value comparable to that described for the same agonist in transfected cells (**Figure 2D**) (Trincavelli et al., 2014). The presence of TGF-β1 did not significantly affect the agonist potency (EC50 = 10.0 ± 1.8 nM), even if the maximal effect of BAY in the production of cAMP was slightly increased (Emax = 4.3 pmol – TGF-β1; Emax = 5.8 pmol + TGF-β1, P = 0.0565) even if it was

∗P ≤ 0.05, ∗∗∗P ≤ 0.001 vs. the CTRL.

not statistically significant. These results are in accordance with the effect of other cytokines on A2BAR functionality (Daniele et al., 2017); in fact, TNF-α has been demonstrated to increase A2BAR coupling to the Gs protein (Daniele et al., 2017).

In parallel, the BAY-induced ERK phosphorylation was evaluated in order to further assess the A2BAR functionality (**Figure 2E**). The agonist was able to produce a concentrationdependent increase of pERK with a potency in the submicromolar range of (EC50 = 114.8 ± 24.5 nM).

60-6583, and cAMP production was quantified. (E) A549 cells were treated with different concentrations of BAY 60-6583 for 5 min and the levels of pERK were quantified. The data were expressed as pmol of cAMP and are the mean values ± SEM of three independent experiments, each performed in duplicate.

## Effects of A2BAR Stimulation on EMT Markers in the Absence or Presence of TGF-β1

Typically, the EMT process is triggered and maintained by the action of different cytokines and extracellular stimuli that, through the activation of specific intracellular pathways, modify the expression of several proteins (Nieto et al., 2016). A549 cells were treated for 48 h with the A2BAR agonist alone or in the presence of TGF-β1 (10 ng/ml), which has been reported to induce EMT (Kawata et al., 2012). First, a morphological analysis of the cells was performed (**Figures 3A,B**). TGF-β1 treatment induced a change in cell morphology: most of the A549 cells, which normally display an oval shape, showed an elongated shape with a fibroblast-like appearance. Surprisingly, cells challenged with the A2BAR agonist lead to a partial morphological change, with only some cells presenting an elongated shape, suggesting the induction of partial EMT.

The morphological changes, which are characteristic of cells undergoing EMT, are accompanied by a shift in the expression of epithelial genes to a mesenchymal gene repertoire (Nieto et al., 2016). Accordingly, challenging cells with TGF-β1 modified the expression of EMT markers, leading to a significant increase in the expression of the mesenchymal markers, Vimentin and N-cadherin (128.8 ± 6.5, P ≤ 0.01, and 162.7 ± 3.8% P ≤ 0.001 vs. CTRL, respectively; **Figures 3C,D**) and a concomitant decrease in the expression of the epithelial marker E-cadherin (6.7 ± 2.5% P ≤ 0.001 vs. CTRL; **Figures 3C,D**). The data were confirmed at the gene expression level by real-time RT-PCR analysis of E-cadherin (CDH1), Vimentin, and N-cadherin (CHD2) expression (**Figure 3E**).

A2BAR stimulation was able to slightly decrease the expression of E-cad protein (**Figures 3C,D**) and increase the expression of a mesenchymal marker (N-cad, 119.8 ± 0.6% vs. CTRL; P ≤ 0.05). These effects are in accordance with the effects elicited by BAY on epithelial/mesenchymal gene transcription. Conversely, A2BAR activation in the presence of TGF-β1 produced an opposite effect, significantly counteracting the induced EMT (**Figures 3C–E**). These effects were almost completely counteracted by concomitant treatment with the A2BAR selective inverse agonist MRS 1706 (1 µM), demonstrating that they were mediated specifically by A2BAR. The treatment with MRS 1706 alone was not able to significantly modify the expression of mesenchymal and epithelial markers in the absence or presence of TGF-β1 demonstrating that the adenosine in the medium was not sufficient to induce a basal A2BAR activation.

## Effects of A2BAR Modulation on EMT Major Transcription Factors (EMT-TFs)

The modulation of classical EMT centers on the transcriptional control of different transcription factors: Snail (SNAI1), Slug (SNAI2), ZEB1 and TWIST (Nieto et al., 2016). To gain insight into the mechanisms by which A2BAR affects E-cadherin deficiency alone and in TGF-β1-induced A549 cells, the expression levels of the EMT-TFs were examined using real-time RT-qPCR (**Figure 4**).

A549 cells expressed Snail and Slug at low levels (Supplementary Figure 1), in accordance with the high expression of E-cad (**Figure 2**). Treatment with TGF-β1 for 48 h changed the expression pattern of these genes (**Figure 4A**), with a significant increase in ZEB1 (1.57 ± 0.13-fold; P ≤ 0.05), Slug (2.63 ± 0.25-fold; P ≤ 0.001), in accordance with the literature data (Ji et al., 2016). Next, the effects of A2BAR stimulation (BAY 60-6583) or blockade (MRS 1706) were evaluated (**Figure 4B**). The MRS alone could not modify the expression of EMT-TFs, highlighting that the endogenous activation of the receptor was not sufficient to modify the related-EMT gene expression. Conversely, A2BAR activation modified the gene repertoire, producing a significant increase in the Slug expression (1.65 ± 0.16-fold; P ≤ 0.05). These data are in accordance with the effects of A2BAR activation on epithelial/mesenchymal markers (**Figure 3**).

Next, the role of A2BAR stimulation on EMT-TF was evaluated in the presence of TGF-β1 (**Figure 4C**). As expected, the A2BAR activation counteracted the effects of cytokine treatment, with a significant decrease in ZEB1 and Slug expression. When the A2BAR inverse agonist was applied, the TGF-β1-induced decrease in gene expression was completely counteracted, demonstrating that these effects were mediated by A2BAR.

## The Balance of cAMP Production and MAPK/ERK Activation Orchestrates the A2BAR Effects on EMT

A2BAR couples to Gαs proteins, leading to an increase in intracellular cAMP, and to the Gq-PLC pathway, which induces the activation of the MAPK/ERK pathway (Schulte and Fredholm, 2003; Sun and Huang, 2016). Thus, activation of these two intracellular signaling pathways were investigated following A2BAR stimulation, in the absence or presence of TGFβ1 (**Figure 5**). MRS 1706 alone did not affect the activation of the two pathways, demonstrating that the adenosine levels in our cellular model were not sufficient to activate the receptor. BAY was able to increase the intracellular cAMP concentration (**Figure 5A**), and significantly enhance ERK1/2 phosphorylation (178.9 ± 10.5% vs. CTRL, P ≤ 0.05; **Figure 5B**), producing transient activation of these kinases (**Figure 5C**). The effects evoked by A2BAR stimulation were almost completely reversed by the A2BAR inverse agonist MRS1706, demonstrating the specific involvement of the A2BAR subtype. As expected, TGF-β1 did not produce a significant increase in the cAMP levels (**Figure 5A**); conversely, it was able to significantly increase ERK1/2 phosphorylation (341.7 ± 16.2% vs. CTRL, P ≤ 0.001; **Figure 5B**). Both BAY and TGF-β1 induced transient activation of ERK1/2 but with different kinetics; whereas ERK phosphorylation induced by BAY peaked within 30 min and then returned to the basal value within 6 h, TGF-β1 caused a more sustained ERK activation up to 48 h of cell treatment (48 h, 155.2 ± 7.8%, TGF-β1; 85.3 ± 8.8%, BAY; P ≤ 0.05; **Figure 5C**). These data highlighted a difference in the final reorganization of the intracellular pathways activated by the cytokine and the receptor.

FIGURE 3 | A2BAR stimulation affects EMT markers and decreases TGF-β1-induced EMT. A549 cells were treated with BAY 60-6583 (100 nM) in the absence or presence of TGF-β1 (10 ng/ml) for 48 h. When indicated, the A2BAR inverse agonist, MRS 1706 (1 µM), was applied. (A) At the end of the incubation times, representative images were taken. Scale bar = 100 µm. (B) Cells treated as above were fixed and stained with anti-β actin and visualized with goat anti-mouse AlexaFluo 488 (green). Nuclei were counterstained with DAPI (blue). (C,D) A549 cells were treated as described above, and the levels of the EMT markers E-cad, Vimentin and N-cad were evaluated by Western blotting. One representative Western Blot is presented (C). The bar graph (D) shows the densitometric analysis of the Western blot performed using the ImageJ program. The data are presented as the means of three different experiments. (E) Real-time RT-PCR analysis of the same EMT markers were performed. The data were expressed as the fold change vs. the CTRL levels, which were set to 1 and are the mean values ± SEM of three different experiments each performed in duplicate. The significance of the differences was determined by one-way ANOVA, followed by Dunnett's post hoc test or two-way ANOVA with Bonferroni correction and two-sided tests for multiple comparisons. ∗P ≤ 0.05, ∗∗P ≤ 0.01, ∗∗∗P ≤ 0.001 vs. the CTRL; #P ≤ 0.05, ###P ≤ 0.001 vs. TGF-β1 alone; ◦◦P ≤ 0.01 vs. TGF-β1 + BAY.

TGF-β1 + BAY.

fphar-09-00054 January 29, 2018 Time: 17:4 # 10

correction and two-sided tests for multiple comparisons. ∗P ≤ 0.05, ∗∗∗P ≤ 0.001 vs. the CTRL; #P ≤ 0.05, ##P ≤ 0.01 vs. TGF-β1 alone; § P ≤ 0.05 vs. the

When A2BAR was activated in the presence of TGF-β1, the receptor activation produced a significant increase in the intracellular cAMP concentration (56.4 ± 1.9 nM, CTRL; 96.6 ± 5.3 nM, TGF-β1 + BAY P ≤ 0.01; **Figure 5A**). This effect was in accordance with the slight increase in A2BAR functionality that was noticed in the presence of the cytokine (**Figure 2D**). Regarding pERK1/2, the co-treatment led to a modest decrease in phosphorylation. The BAY activities on cAMP and pERK1/2 were most completely counteracted by cell pre-incubation with the A2BAR inverse agonist MRS1706. Finally, TGF-β1 and BAY did not affect the amount of total ERK (Supplementary Figure 2).

To elucidate if the different activation of cAMP and ERK pathway in the presence of the cytokine, could be related to A2BAR desensitization, the functionality of the receptor was evaluated after 48 h of cell treatment in the absence or presence of BAY and TGF-β1 alone or in combination (**Figure 5D**). In the basal condition (48 h without treatment), A2BAR maintained its functional response. The cell treatment with BAY (100 nM) for 48 h induced the desensitization of the A2B receptor (219.9 ± 7.7% in Basal condition; 113.3 ± 13.6% in BAY pre-treated cells; P ≤ 0.001). Cytokine alone was not able to significantly interfere on the receptor desensitization. Interestingly, when cells were treated with BAY in the presence of TGF-β1 for 48 h, the desensitization of the receptor was partially impaired (164.2 ± 8.7%; P ≤ 0.01).

### cAMP/PKA Regulation of EMT Markers in Human Epithelial Cells

The role of the cAMP pathway in EMT is controversial, depending on the cell type, the nature of the EMT inducers and maximum levels of the intracellular cAMP obtained (Weng et al., 2015). Thus, the effects of different levels of cAMP on EMT markers in our cellular model were investigated using the cAMP analog Br-cAMP (8-bromoadenosine 30 ,50 -cyclic monophosphate) as the PKA activator (**Figure 6**). Br-cAMP (100 nm–1 µM) induced a concentration-dependent increase in the intracellular cAMP levels (**Figure 6A**). Surprisingly, different

TGF-β1 alone.

concentrations of the activator caused opposite effects on E-cad (**Figure 6B**) and N-cad (**Figure 6C**) expression. A high Br-cAMP concentration increased E-cad and decreased N-cad expression. A low Br-cAMP concentration (100 nM) was sufficient to determine a modest decrease of E-cad (1646 ± 56 ng/ml, CTRL; 1540 ± 93 ng/ml, Br-cAMP; **Figure 6B**) and an increase of N-cad (246 ± 35 ng/ml, CTRL; 270 ± 8 ng/ml, Br-cAMP; **Figure 6C**) even if these variations were not statistically significant. When TGF-β1 and BAY were used simultaneously, the activator produced higher cAMP levels and a concentration-dependent increase in E-cad and decrease in N-cad.

These data demonstrate that, in our cellular model, when the cAMP was increased up to high concentrations, it could negatively affect the EMT process. Conversely, when cAMP levels remained low, it seems to mediate the opposite effects, promoting a slight decrease in epithelial markers and an increase in the mesenchymal markers. This controversial nature of the cAMP analogue on the EMT process is in accordance with the recent data reported by Zuccarini et al. (2017).

## The cAMP/PKA-MAPK/ERK Balance in the A2BAR Modulation of EMT Markers and EMT-TFs

Nevertheless, high levels of cAMP counteract EMT, and ERK phosphorylation mediated by MAPK activation has been demonstrated to promote EMT (Singh et al., 2017). Thus, the ability of A2BAR to modify the amounts of both pERK and of cAMP could be a possible mechanism by which the receptor agonist affects epithelial/mesenchymal markers. In this respect, the involvement of A2BAR in the regulation of EMT markers and transcription factors was investigated in the presence of PKA inhibitor (H89, 100 nM) and a mitogen-activated protein kinase inhibitor (PD98059, 1 µM) (**Figure 7**, Supplementary

### FIGURE 7 | Continued

fphar-09-00054 January 29, 2018 Time: 17:4 # 14

and ERK 1/2 phosphorylation was evaluated after 30 min of treatment. The data were expressed as the percentage versus untreated cells (CTRL) set to 100% ± SEM of at least three different experiments performed in duplicate. Cells treated as above were lysed, and the expression of E-cadherin (C) or N-cadherin (D) were quantified using AlphaLISA kits. The data were expressed as E-cadherin and N-cadherin concentrations (ng/ml) and are the mean values ± SEM of three independent experiments, each performed in duplicate. The significance of the differences was determined by one-way ANOVA, followed by Dunnett's post hoc test or two-way ANOVA with Bonferroni correction and two-sided tests for multiple comparisons. ∗P ≤ 0.05, ∗∗P ≤ 0.01, ∗∗∗P ≤ 0.001 vs. the CTRL; #P ≤ 0.05, ##P ≤ 0.01, ###P ≤ 0.001 vs. BAY alone. (E,F) Cells were treated as above and real-time RT-PCR analysis of EMT markers (E-cad, N-cad and Vimentin) (E), and EMT transcription factors (ZEB1, Snail, Slug, TWIST) (F) was performed. The data were expressed as the fold change vs. the CTRL levels, which were set to 1 and are the mean values ± SEM of three different experiments each performed in duplicate. The significance of the differences was determined by one-way ANOVA, followed by Dunnett's post hoc test or two-way ANOVA with Bonferroni correction and two-sided tests for multiple comparisons. ∗P ≤ 0.05, ∗∗P ≤ 0.01, ∗∗∗P ≤ 0.001 vs. the CTRL; #P ≤ 0.05, ##P ≤ 0.01 vs. BAY alone.

Figure 3). The BAY-mediated increase in the intracellular cAMP levels was not affected by H89 treatment (**Figure 7A**); conversely, PD98059 was able to block BAY-mediated ERK phosphorylation (**Figure 7B**). In this respect, the inhibitors were used to unveil the involvement of the two signaling pathways in A2BAR mediated triggering of EMT.

BAY decreased the protein density of E-cad and increased that of the N-cad one; these effects were lost in the presence of the inverse agonist MRS 1706, in accordance with the abovementioned data (**Figure 3**). The ability of BAY to decrease the expression of E-cad was augmented by pretreatment with the PKA inhibitor (2718 ± 58 ng/ml, CTRL; 2045 ± 157 ng/ml, BAY; P ≤ 0.01; 1483 ± 122 ng/ml, BAY + H89; P ≤ 0.001 vs. CTRL, P ≤ 0.01 vs BAY, **Figure 7C**). Similarly, A2BAR mediated increase in N-cad was enhanced when the cAMP/PKA axis was abrogated (239 ± 20 ng/ml, CTRL; 406 ± 29 ng/ml, BAY; P ≤ 0.05; 558 ± 72 ng/ml, BAY + H89; P ≤ 0.001 vs. CTRL, P ≤ 0.05 vs BAY; **Figure 7D**). These effects highlighted that cAMP, through PKA activation, could partially antagonize A2BAR-mediated EMT induction. When the MEK inhibitor was used, the changes induced by A2BAR stimulation were almost completely counteracted, restoring the basal conditions (E-cad, 2586 ± 90 ng/ml, BAY + PD98059; P ≤ 0.05 vs BAY; N-cad, 260 ± 49 ng/ml, BAY + PD89059; P ≤ 0.05 vs BAY, **Figures 7C,D**). These results are in accordance with the effects of the inhibitor treatments on A2BAR-mediated alteration of the expression of CDH1, CDH2 and vimentin genes (**Figure 7E**).

Finally, the modulation of the EMT-TF gene expression by BAY was evaluated in the presence of H89 and PD98059. Similarly, regarding the EMT markers, the PKA inhibitor slightly exacerbated the A2BAR-mediated effects; conversely, the MEK inhibitor counteracted the BAY-mediated increase in gene expression. The PD98059 activity was more pronounced on Slug expression, causing a significant decrease of gene expression (1.65 ± 0.16-fold, BAY; 1.61 ± 0.20-fold, BAY + H89; 0.93 ± 0.09-fold vs. CTRL, BAY + PD98059; P ≤ 0.05). These data are consistent with the reported role of ERK phosphorylation in the induction of Slug expression (Choi et al., 2007; Joannes et al., 2014).

### DISCUSSION

Adenosine receptors (ARs) have attracted great attention as possible targets to control the EMT process in pathologies such as fibrosis and cancer (Lu and Insel, 2014). However, among the AR subtypes, no data have been reported that correlate A2BAR activation with EMT in epithelial cells. In this respect, herein, we report for the first time the ability of A2BAR activation to promote EMT or contrast the effects of the extracellular inducer, TGFβ1, in human lung epithelial cells. Furthermore, the ability of the receptor activation to modulate two signaling pathways involved in EMT, cAMP/PKA, and MAPK/ERK, was demonstrated as a possible mechanism explaining the different effects mediated by A2BAR stimulation in different extracellular microenvironment (**Figure 8**).

Human carcinoma epithelial lung cells (A549) were maintained under serum deprivation, a condition that has been demonstrated to amplify the epithelial phenotype (Dong et al., 2014). Although all the ARs are expressed in epithelial lung cells, the A2B receptor subtype was demonstrated to be the most represented, as previously reported (Roman et al., 2006). Challenging these cells with TGF-β1 produced a marked change in the phenotype through the simultaneous decrease in epithelial markers (E-cad) and increase in mesenchymal markers (N-cad). In accordance with the literature (Kawata et al., 2012), these data demonstrated the suitability of the model to study the A2BAR involvement in the EMT process.

A2BAR stimulation modified the expression of epithelial/mesenchymal markers with a low but significant amplitude. Surprisingly, the same receptor could counteract the EMT induction mediated by TGF-β1. These opposite effects elicited by A2BAR stimulation in lung cells reflect the well documented debated role of this receptor in the onset and progression of different pathologies. The receptor activation has been reported to protect the lungs and other tissues from inflammation and acute injury (Eckle et al., 2008; Davies et al., 2014; Hoegl et al., 2015). By contrast, other evidence has been reported concerning the A2BAR-mediated pro-fibrotic effects and promotion of cancer development (Karmouty-Quintana et al., 2012; Huerter et al., 2016; Sepúlveda et al., 2016). Furthermore, Zhou et al. (2011) have shown that A2BAR genetic removal slightly affects acute lung injury but reduces lung fibrosis, supporting a pro-fibrotic role for this receptor.

In our study, the A2BAR selective agonist regulated the gene transcription of different EMT-transcription factors (EMT-TFs). EMT is a complex process regulated by the orchestration of several transcription factors. Among these EMT-TFs, ZEB1, Snail and Slug (SNAI2) are the major transcription factors involved in the promotion of the EMT process in A549 cells,

as previously reported (Ji et al., 2016). A2BAR stimulation did not affect significantly the expression of Snail and TWIST, other master genes of EMT. However, A2BAR activation modulate the expression of Slug and, to a lesser extent, that of ZEB1, which are commonly considered the major actors in myofibroblast differentiation and fibrogenesis (Lamouille et al., 2014). Furthermore, Slug expression has been correlated to direct repression of E-cadherin expression (Choi et al., 2007). This evidence was in accordance with the increase in E-cadherin induced by A2BAR stimulation.

The EMT-TFs are regulated by the activation of different intracellular pathways, which in turn regulate the balance between the epithelial and mesenchymal markers. ERK1/2, GSK3β, p38, STAT3, and cAMP are only few of all the proteins involved in the promotion/inhibition of EMT (Nieto et al., 2016). In our cellular model, activation of the cAMP/PKA axis promoted opposite effects depending on the levels of intracellular the cAMP. Low concentrations led to a slight promotion or no effects on epithelial/mesenchymal markers; conversely, high levels of cAMP could counteract EMT progression. In accordance with our results, it has been reported that cAMP/PKA activation could exert opposite effects on EMT (Zhang et al., 2006a,b; Nadella et al., 2008; Shaikh et al., 2012; Lambers et al., 2015) that could be ascribed to the different cell models, type of EMT inducer and maximum levels of intracellular cAMP obtained (Weng et al., 2015).

A2BAR activation increases the intracellular cAMP levels and promotes the phosphorylation of ERK 1/2 (Grant et al., 2001; Zhou et al., 2017). In this respect, we hypothesize that the controversial role of A2BAR in promoting EMT or reversing TGF-β1-induced EMT could be ascribed to the different balance of the intracellular pathways activated. The activation of other GPCRs coupled to different G proteins has been reported to elicit opposite effects on EMT. Zhang et al. (2016) have demonstrated that norepinephrine, which activates Gs protein, induces the epithelial-mesenchymal transition in A549 cells; similarly, the P2X7 stimulation, which activates the PI3K/Akt and ERK1/2 signaling pathways, promotes the mesenchymal phenotype in epithelial renal cells (Zuccarini et al., 2017). A2AARs have been reported to modify the epithelial/mesenchymal phenotype in renal cells by downregulating TGF-β1-induced EMT (Zuccarini et al., 2017) in accordance with the effects elicited by A2BAR activation in our cellular model when the cytokine was present. TGF-β1 activates both Smad signaling and non-Smad signaling including the ERK pathway, critically regulating EMT (Xie et al., 2004). A2BAR stimulation could counteract cytokine-induced EMT, reducing the transition progression. These effects were correlated with a robust increase in cAMP production and with a concomitant decrease in ERK phosphorylation. In this respect, we could speculate that the increase in the cAMP, mediated by A2BAR activation, could counteract the cytokine-mediated effects on EMT. This is in accordance with the ability of other

Gs coupled receptors (i.e., A2AAR) or cAMP inducers (i.e., forskolin, 8-Br-cAMP) to prevent TGF-β1-mediated EMT (Zhang et al., 2006a,b; Zuccarini et al., 2017). It should also be considered that cAMP/PKA has a negative effect on TGF-β induced pERK 1/2, but only when the cytokine downstream pathways are highly expressed (Weng et al., 2015). This mechanism could explain the reduction in ERK1/2 phosphorylation in the presence of the A2BAR agonist.

The modulation of cAMP production and ERK phosphorylation could be affected by internalization or desensitization processes of the receptor. Several GPCRs employ a variety of signaling mechanisms to exert their functions. The production of second messengers, such as cAMP, is the results of the receptor-G protein coupling. However, other signals produced in conjunction with, or even independent of heterotrimeric G protein could be activated. It is well known that the arrestins, a family of four GPCR-binding proteins originally described for their role in the GPCR desensitization, have been found to interact with Src family kinases and to serve as scaffolds for activation of ERK1/2 and c-Jun N-terminal kinase 3 mitogen-activated protein kinases (Ferguson, 2001; Luttrell, 2005; Lee et al., 2008). Thus, the desensitization could represent a mechanism to maintain the activation of G-protein independent pathways. The activation of the A2BAR for long time produced the desensitization of the receptor. This may be another mechanism through which A2BAR mediate the ERK phosphorylation favoring the EMT traits. Conversely, the TGF-β1 was able to reduce the agonist-induced A2BAR desensitization, maintaining the ability of A2BAR to produce cAMP that negatively affected EMT process. These data could better highlight the controversial role of A2BAR in the EMT. In different pathological status in which adenosine raise to micromolar concentration, the extracellular microenvironment and the localization of the receptor could interfere with the levels of the activated intracellular signaling shifting the balance in favor of or against the EMT.

In our cellular model, A2BAR activation was able to induce partial EMT; however, it was not sufficient to promote a complete epithelial-mesenchymal transformation of the lung cells. The involvement of the cAMP/PKA and MAPK/ERK signaling pathways was demonstrated by the use of selective inhibitors. The EMT traits, in the presence of A2BAR agonist was enhanced by PKA inhibition and decreased by the ERK1/2 phosphorylation blockade. Furthermore, the pretreatment with a PKA inhibitor did not enhance receptor effects on TF expression. Conversely, the presence of the MEK inhibitor completely counteracted the A2B-mediated increase in Slug and ZEB1 gene expression, demonstrating the involvement of MAPK/ERK pathway activation in A2BAR-mediated EMT induction.

### REFERENCES

## CONCLUSION

For the first time, these results highlight the possibility that A2BAR is one of the numerous receptors that could be involved in EMT regulation. Although the expression of the other adenosine receptor subtypes are low, in physiological condition the involvement of all the AR in the EMT regulation could not be excluded. A2BAR activation is not enough to trigger a complete transition; it can affect the expression of the epithelial and mesenchymal markers promoting EMT traits. These effects are related to the ability of A2BAR to modify the balance between cAMP/PKA and MAPK/ERK activation. The presence of extracellular cytokines that activate the transition could change the balance and levels of these two intracellular pathways, shifting the effects exerted by the receptor stimulation on EMT trigger and maintenance. Thus, the A2BAR seems to act as an EMT regulator rather than a main actor in the EMT induction. Although the EMT is a complex machinery and further investigation are needed to better elucidate the effects of adenosine and the contribution of the other AR subtypes in physiopathological conditions, herein, for the first time, the A2BAR was shown to be related to the EMT process, highlighting its potential role as pharmacological target in EMT-related pathologies.

### AUTHOR CONTRIBUTIONS

CG designed and performed the biological experiments. CG, SD, and MT. analyzed the data and wrote the manuscript. IP performed the immunofluorescence analysis. CR, LT, and TN contributed to the experimental work. MT, AC, and CM designed the study and played a key role as project supervisor. MT and CM coordinated the project. All the authors contributed to and approved the final manuscript.

## FUNDING

This work was financially supported by the Italian Ministry of Education (Project of National Research Interest PRIN 2015, 2015E8EMCM\_007).

### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fphar. 2018.00054/full#supplementary-material

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through activation of a functional adenosine 2B receptor in renal fibroblasts. Ann. Clin. Lab. Sci. 46, 339–345.

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Giacomelli, Daniele, Romei, Tavanti, Neri, Piano, Celi, Martini and Trincavelli. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Biology of Platelet Purinergic Receptors and Implications for Platelet Heterogeneity

Milka Koupenova1 \* and Katya Ravid2 \*

1 Department of Medicine, Division of Cardiovascular Medicine, University of Massachusetts Medical School, Worcester, MA, United States, 2 Departments of Medicine and Biochemistry and Whitaker Cardiovascular Institute, Boston University School of Medicine, Boston, MA, United States

Platelets are small anucleated cells present only in mammals. Platelets mediate intravascular hemostatic balance, prevent interstitial bleeding, and have a major role in thrombosis. Activation of platelet purinergic receptors is instrumental in initiation of hemostasis and formation of the hemostatic plug, although this activation process becomes problematic in pathological settings of thrombosis. This review briefly outlines the roles and function of currently known platelet purinergic receptors (P1 and P2) in the setting of hemostasis and thrombosis. Additionally, we discuss recent novel studies on purinergic receptor distribution according to heterogeneous platelet size, and the possible implication of this distribution on hemostatic function.

### Edited by:

Kenneth A. Jacobson, National Institutes of Health (NIH), United States

### Reviewed by:

Alvin H. Schmaier, Case Western Reserve University, United States Martyn P. Mahaut-Smith, University of Leicester, United Kingdom

### \*Correspondence:

Milka Koupenova<EMAIL_ADDRESS>Katya Ravid<EMAIL_ADDRESS>
### Specialty section:

This article was submitted to Experimental Pharmacology and Drug Discovery, a section of the journal Frontiers in Pharmacology

> Received: 20 November 2017 Accepted: 12 January 2018 Published: 30 January 2018

### Citation:

Koupenova M and Ravid K (2018) Biology of Platelet Purinergic Receptors and Implications for Platelet Heterogeneity. Front. Pharmacol. 9:37. doi: 10.3389/fphar.2018.00037 Keywords: platelets, purinergic receptors, ATP, ADP, adenosine

## INTRODUCTION

Adenine nucleosides and nucleotides accumulate in the circulation under various pathological conditions and can initiate and mediate the response to cell damage, hypoxia, and inflammation (Koupenova and Ravid, 2013). Additionally, adenosine diphosphate (ADP) and adenosine triphosphate (ATP) are released from dense-granules during platelet activation and propagate platelet–platelet interactions, ultimately leading to three-dimensional plug formation that seals endothelial damage (McNicol and Israels, 1999; Koupenova and Ravid, 2013; Koupenova et al., 2017). Adenosine, in turn, is not known to be present in platelets but is generated in the extracellular space from ATP and ADP by two ectonucleotidases, CD39 (ENTPD1) and CD73 (NT5E) (reviewed in Koupenova and Ravid, 2013), that are present on endothelial and platelet surfaces (Koziak et al., 1999; Castilhos et al., 2016).

Two major classes of receptors mediate the physiological effect of adenosine and adenosine phosphates. These are the P1 and P2 purinergic receptors, classified based on their preference for adenosine (P1) or adenosine phosphates (P2). The P1 purinergic receptors include four G-proteincoupled receptors, two that activate adenylate cyclase [Adora2A (A2aAR) and Adora2b (A2bAR)] and generate cyclic AMP (cAMP), and two that inhibit adenylate cyclase [AdoraA1 (A1AR) and AdoraA3 (A3AR)] and decrease cAMP. The P2 purinergic receptor group is subdivided further into two groups that, in platelets, are represented by receptors activated by ATP (P2X1) and receptors activated by ADP (P2Y1 and P2Y12). P2Y1 and P2Y12 are G-protein-coupled receptors, while P2X1 is a ligand-gated ion channel receptor (Jacobson et al., 2006; Cattaneo, 2007). Adenosine and adenosine phosphates play a central role in regulating platelet behavior during hemostasis and thrombosis, as ADP induces platelet activation, while adenosine, for the most

part, inhibits it. ATP, in turn, can initiate platelet activation through the P2X1 receptor and inhibit ADP-mediated activation when acting as an antagonist on the P2Y receptors (Cattaneo, 2007). The potential explanation as to why platelets have both ATP and ADP receptors is that all of the purinergic receptors are necessary for establishing a balanced regulation of hemostasis or thrombus formation, depending on the intensity of vessel injury. Nucleotides released from damaged cells may occupy platelet P2 receptors and initiate shape change (via P2X receptors) and aggregation (via P2Y12 and P2Y1 receptor). The level of ADPmediated P2Y12 or P2Y1 receptor activation can be controlled by ATP. Conversion of ATP and ADP (by CD39/CD73) to adenosine provides a second level of regulation that controls propagation of the hemostatic plaque by inhibiting activation through the A2 ARs. Deeper damage of the vessel leads to an increase in platelet aggregation response (collagen- or tissue factormediated) in order to prevent leakage into the interstitial tissue. This simplified explanation, however, is complicated by the fact that platelets are a heterogeneous population (as discussed in the last section of this review) and there is differential expression of distinct purinergic receptors based on size and function. In this review, we will focus on the role of purinergic receptors in platelet function and their distribution according to platelet heterogeneity.

### PURINERGIC RECEPTORS IN HEMOSTASIS

Recent in vivo studies have further clarified the complex interactions and agonist distribution in the formation of threedimensional platelet arrangement during hemostasis. According to this newly elucidated model, the hemostatic plug is composed of a core and an outer shell through which platelets are differentially activated. The stringent plug architecture consists of a platelet activation gradient with the most activated platelets in the core of the clot, surrounded by less activated platelets in the outer shell region. Fibrin deposition is localized distinctly at the base of the core in the extravascular space before hemostasis is achieved (Stalker et al., 2013; Tomaiuolo et al., 2017). The inner core of the hemostatic plug is packed tightly with degranulated platelets that are P-selectin positive. The outer shell is composed of loosely packed platelets that do not express P-selectin, and there is little to no fibrin present.

Although stable, the outer shell is porous and permeable to plasma solutes. Consistent with the platelet activation distribution gradient, there is a distinct distribution of platelet agonists throughout the hemostatic plug. The core of the plug contains a high concentration of thrombin (factor IIa) and, as the plug becomes more porous, a gradient of ADP and thromboxane A2 (TxA2) develops (Stalker et al., 2013; Tomaiuolo et al., 2017). The porous outer shell of the thrombus allows for recruitment of leukocytes necessary for injury repair or pathogen elimination. An increase in thrombin leads to PAR4 cleavage, consequently leading to leukocyte recruitment and migration to the damaged endothelium (Kaplan et al., 2015). Leukocyte recruitment, in turn, is limited by binding of thrombin to platelet GP1bα that can reduce platelet activation. Additionally, fibrin deposition in the thrombus physically inhibits leukocyte migration (Kaplan et al., 2015). The distinct distribution of P-selectin expressing platelets in the core vs. P-selectin-negative platelets in the shell suggests a possibility for a specific distribution of different platelet subpopulations throughout the hemostatic plug, according to their function in the interaction with either damaged endothelium or circulating leukocytes.

## PLATELETS AND P2 RECEPTORS

### ATP Receptors in Platelets

The P2X1 receptor is a ligand-gated ion channel receptor (Sun et al., 1998) that is activated by ATP and inhibited by ADP. Binding of ATP to the P2X1 receptor leads to calcium influx into platelets (Rolf et al., 2001; Mahaut-Smith, 2012) which consequently results in a transient change of platelet shape, platelet degranulation, pseudopodia formation, and platelet activation (Rolf et al., 2001; Toth-Zsamboki et al., 2003; Mahaut-Smith, 2012). P2X1 receptor activation by ATP alone does not mediate platelet aggregation; however, it can amplify ADP-mediated aggregation through the platelet-P2Y1 receptor (Jones et al., 2014). Furthermore, during early stages of vessel damage, in the presence of a low concentration of collagen, ATP contributes to increased aggregation through the P2X1 receptor (Oury et al., 2001). Similarly, P2X1 receptors can amplify thrombin-mediated platelet aggregation through protease-activated receptor 1 (PAR1) at low levels of thrombin (Erhardt et al., 2006). Importantly, in the presence of collagen or pathogenic stimuli, endothelial inhibitors (such as prostacyclins) are unable to completely inhibit calcium-mediated platelet aggregation partially due to activation of P2X1 receptor by ATP (Fung et al., 2012).

Intracellularly, P2X1 activation leads to MAPK/ERK2 pathway signaling that contributes to myosin light chain (MLC) phosphorylation and propagation of collagen-mediated platelet secretion (Toth-Zsamboki et al., 2003). During high shear stress, ATP-activated P2X1 also contributes to platelet-induced aggregation by MLC-mediated cytoskeletal rearrangements (Oury et al., 2004). P2X1 activation by ATP can also contribute to platelet secretion of TxA2 and enhance TxA2-mediated platelet aggregation (Huang et al., 2014). Additionally, in cases of a co-stimulatory role with P2Y1 signaling, P2X1 increases the influx of calcium and amplifies the consequent calcium signaling through P2Y1 and other Gαq-coupled platelet receptors (Jones et al., 2014). Therefore, at sites of vascular injury, intensity of the platelet response can be regulated by the availability of various forms of adenosine phosphates.

Murine platelets lacking the P2X1 receptor exhibit decreased collagen-induced aggregation and adhesion (Hechler et al., 2003a). Furthermore, these platelets show diminished thrombus growth on collagen-coated slides, particularly at higher shear stress (Hechler et al., 2003a). Overall, P2X1 activation seems to be important at high shear stress and low agonist concentration, suggesting that ATP contributes to platelet aggregation at the initial stages of platelet attachment to damaged endothelium, particularly in the arteries.

### ADP Receptors in Platelets

fphar-09-00037 January 27, 2018 Time: 15:15 # 3

P2Y12 and P2Y1 receptors are G-protein-coupled receptors activated by ADP and inhibited by ATP. With respect to adenine nucleotide-mediated aggregation in platelets, the ADPactivated P2Y12 receptor is the most important receptor. P2Y12 was first discovered in 2001 (Hollopeter et al., 2001), and it can couple to the Gαi subunit leading to the inhibition of adenylyl cyclase that mediates the conversion of ATP to cAMP (Hollopeter et al., 2001; Cattaneo, 2007; Hechler and Gachet, 2011). Consequently, the overall levels of cAMP can be decreased which may lead to an increase in platelet activation state. One of the mechanisms by which cAMP reduces platelet aggregation involves activation of protein kinase A (PKA) which leads to the phosphorylation and inhibition of inositol 1,4,5 triphosphate (IP3) receptor-mediated increase of calcium from the dense tubular system; reduced levels of cAMP have the opposite effect on IP3 and consequently increase intracellular calcium (Quinton and Dean, 1992; Tertyshnikova and Fein, 1998). In addition to coupling through Gαi and inhibition of adenylyl cyclase, ADP-mediated activation of P2Y12 receptors leads to activation of PI-3 kinase (PI-3K) through Gβγ (Shankar et al., 2004). It is now understood that the P2Y12 receptor contributes to the ADP-mediated calcium response (by P2Y1) in platelets by activating PI-3K, in addition to simultaneously inhibiting adenylyl cyclase, i.e., by reducing cAMP levels (Hardy et al., 2004). Importantly, ADP activation through Gαi-coupled P2Y12 receptors and the consequent RAP1 activation is critical for GPIIb/IIIa integrin-mediated platelet aggregation (Larson et al., 2003; Stefanini and Bergmeier, 2017). The P2Y12, but not P2Y1 receptor is required for sustained activation of RAP1 (Ras-related protein 1). P2Y12 is necessary for the formation of a shear-resistant hemostatic plug which is mediated by locally secreted ADP from platelets (Larson et al., 2003; Stolla et al., 2011b; Stefanini and Bergmeier, 2017). RAP1 activation is also dependent on guanine exchange factor (GEF), proteins RapGEF and RAS guanyl releasing protein 2 (RASGRP2), that are regulated by intracellular calcium signaling and by inhibition from the GTP-ase activating protein Ras GTPase-activating protein 3 (RASA3) (Stolla et al., 2011a; Stefanini and Bergmeier, 2017). ADP-activated P2Y1 receptor, on the other hand, does not affect cAMP levels (Fabre et al., 1999), couples to the Gαq subunit leading to the activation of phospholipase C (PLC) (Cattaneo, 2007; Hechler and Gachet, 2011). In platelets, PLC activation leads to a cascade of molecular signaling events that result in increased levels of cytosolic calcium, particularly from the dense tubular system. Interestingly, the P2Y1 receptor is able to activate Src kinase and consequently establish negative regulation over P2Y12 signaling through PI-3K (Hardy et al., 2004). This suggests that for ADP-mediated calcium signaling in platelets, P2Y12 and P2Y1 receptors cross-communicate to establish a controlled response to vascular damage; the P2Y12 receptor contributes to the absolutely required P2Y1-mediated calcium response by activating PI-3K and reducing cAMP levels, while the P2Y1 receptor inhibits PI-3K by coupling with Src kinase (Hardy et al., 2004). Thus, now it is understood that, in addition to ADP activation of platelets through two different molecular signaling mechanisms in which platelet activation by P2Y1 leads to rapid shape change and reversible aggregation while P2Y12 activation induces sustained platelet aggregation but without a shape change (Cattaneo, 2007; Hechler and Gachet, 2011), there is also a synergy between these receptors at the level of calcium signaling (Hardy et al., 2004). Importantly, other platelet G-protein-coupled receptor signaling through Gαq and contribution to the overall calcium pool can further synergize with ADP receptors and affect platelet function. Detailed reviews of platelet P2 signaling and synergy at calcium regulation can be found elsewhere (Dorsam and Kunapuli, 2004; Cattaneo, 2007; Varga-Szabo et al., 2009; Hechler and Gachet, 2011; Stefanini and Bergmeier, 2017).

It is important to mention that the role of ADP secreted from platelet dense-granules is to amplify the aggregation signal induced by other strong platelet agonists (such as TxA2), assuring stable platelet aggregation. Conversely, coordinated signaling involving activation of the P2Y12 and P2Y1 ADP receptors and integrin GPIIb/IIIa can mediate TxA2 generation (Jin et al., 2002). This multifactorial ADP receptors–TxA2 axis in platelets is important for thrombus growth and stabilization. Interestingly, patients deficient only in P2Y12 receptors have unaffected TxA2 production (Cattaneo et al., 1992, 2000), suggesting an important requirement of the two ADP receptors for proper hemostasis.

Studies utilizing murine platelets deficient in P2Y1 receptor showed that these platelets lack shape change and are prone to decreased aggregation in response to high concentrations of ADP (Fabre et al., 1999; Leon et al., 1999). However, P2Y1-deficient platelets had strongly impaired aggregation at low concentrations of ADP or collagen (Leon et al., 1999), suggesting that the P2Y1 receptor contributes to the overall ADP-mediated platelet aggregation and responses to other agonists. On the other hand, platelet overexpression of the P2Y1 receptor induces platelet hyperactivity (Hechler et al., 2003b). Bleeding time of mice deficient in this receptor is increased and they are protected from collagen- or ADP-induced thromboembolism (Fabre et al., 1999; Leon et al., 1999), suggesting that P2Y1 mediates ADP– platelet aggregation effects in vivo. Studies utilizing a murine model deficient in P2Y12 (Andre et al., 2003) in the context of the mesenteric artery injury model, support the observation that this receptor mediates platelet adhesion and activation, in addition to mediating the growth and stability of the thrombus. Consistently, in vivo antagonism of the P2Y12 receptor in rabbits reduces the size of the initial thrombus in injured mesenteric arterioles. However, in this study utilizing P2Y12 antagonists, AR-C69931 MX or clopidogrel (vs. genetic elimination), the stability of the thrombus was not affected by P2Y12 and the formation of the hemostatic plug was also not affected (van Gestel et al., 2003).

There are various reasons as to why a genetic deletion may lead to a different outcome than pharmacological inhibition in growth and stability of the thrombus. An inhibitor can vary in binding affinity for the receptor, reversibility, availability at specific sites, and off-target effects, as compared to a complete lack of receptor expression. Genetic deletion, on the other hand, can lead to elimination not only of the gene of interest but

can also alter levels of other genes that may be transcriptionally regulated by the particular gene. Finally, since human and murine platelets exhibit differences in content, the differences in pharmacological inhibition vs. deletion can additionally be related to species variability. Regardless, although an antagonist may not completely inhibit the action of P2Y12 or account for other functions of P2 receptors, it is clear that inhibition of P2Y12 can be beneficial in limiting thrombus growth in arterial thrombosis and secondary prevention of ischemia (Andre et al., 2003; van Gestel et al., 2003).

### PLATELETS AND P1 RECEPTORS

### Adenosine Receptors in Platelets

In addition to P2 receptors, platelets express functional P1 purinergic receptors on their surfaces. As mentioned earlier, P1 purinergic receptors are activated by adenosine at various concentrations that may be generated by ATP/ADP hydrolysis. In addition, an increase in levels of nucleotides, and consequent adenosine generation, can occur during inflammation, cell damage during infection, or other cardiovascular pathologies (Fredholm, 2007; Koupenova and Ravid, 2013). Human and murine platelets express functional cAMP-increasing P1 receptors A2aAR and A2bAR (Ledent et al., 1997; Yang et al., 2010), but are not known to have functional cAMP-inhibiting P1 receptors A1AR and A3AR. It is noteworthy that A1AR mRNA has been detected by qRT-PCR (Amisten et al., 2008) but significant expression was not detected by sequencing in more recent studies (Rowley et al., 2011; Clancy et al., 2017). An increased level of intracellular cAMP, in turn, inhibits platelet aggregation/function.

Among the A2 adenosine receptors, A2aAR has the highest affinity for adenosine and is the major adenosine receptor expressed in platelets that mediates inhibition of platelet aggregation. A2aAR is a G-protein-coupled receptor that couples with Gαs and activates adenylyl cyclase (Koupenova et al., 2012). A2aAR mediates platelet function through increasing cAMP levels (Yang et al., 2010), and by inhibiting the rise of thrombin-mediated increase in intracellular calcium levels (Paul et al., 1990). Both of these effects on intracellular second messengers in platelets are presumed to be mediated by the action of adenylyl cyclase (Paul et al., 1990). It is now understood that the inhibitory effect of cAMP on platelet aggregation involves complex regulation of diverging signaling pathways. Intracellularly, cAMP affects calcium signaling by inhibiting PLC-mediated diacylglycerol (DAG) and inositol triphosphate formation, by inhibiting protein kinase C directly or indirectly through reduction of DAG level, or by affecting actin polymerization through phosphorylation of the β-subunit of the glycoprotein receptor GPIb (Fox and Berndt, 1989; Schwarz et al., 2001; Varga-Szabo et al., 2009). In addition to inhibiting platelet aggregation in human blood, activation of A2aAR by specific agonists leads to a reduction in P-selectin expression on the platelet cell surface, as a result of TxA2 or ADP stimulation. This is followed by a greater than 50% reduction in platelet–monocyte aggregate formation (Linden et al., 2008). This suggests that the A2aAR has the potential to mediate the impact of prothrombotic agonists during hemostasis and thrombosis as a fast responder.

Studies utilizing platelets from A2aAR null mice have confirmed that inhibition of platelet aggregation is mediated by this receptor, as platelets without A2aAR have an increased aggregation potential and are not affected by any concentration of the adenosine receptor (non-specific) agonist, 50 -N-ethylcarboxamidoadenosine (NECA) (Ledent et al., 1997). Similarly, inhibition of A2aAR by caffeine, a non-specific adenosine receptor antagonist, also leads to decreased aggregation potential of human platelets (Varani et al., 1999, 2000). However, chronic intake of caffeine leads to a desensitization effect on aggregation, possibly mediated by increased upregulation of A2aAR (or A2bAR) on the platelet surface (Varani et al., 1999, 2000).

Although platelets have a large density of A2aAR, they also contain A2bARs. Similarly to A2aAR, A2bAR is a G-protein receptor that couples to Gαs (Koupenova et al., 2012), but can also couple to Gαq (Feoktistov et al., 1994; Linden et al., 1999; Gao et al., 2017) and, in some cells, to Gαi (Gao et al., 2017). Importantly, in different cells, A2bAR can couple to the same pathways through different G-proteins possibly depending on specific intracellular G-protein signatures (Gao et al., 2017). In cells of the same type, A2bAR can also couple to Gαs or Gαi leading to activation of opposing or synergistic signaling pathways (Gao et al., 2017), suggesting that these adenosine receptors have the ability to balance different signaling mechanisms in different cell types. Contrary to the other P1 receptors, A2bAR is a low affinity receptor that requires the concentration of adenosine to be higher than 10 µM (reviewed in Koupenova and Ravid, 2013). Additionally, A2bAR is an inducible receptor, the expression of which is known to be upregulated under inflammation, stress, or vascular injury (Yang et al., 2006, 2008, 2010; St. Hilaire et al., 2008). Since platelets are anucleate cells, increase in A2bAR expression most likely occurs in the platelet precursors, the megakaryocytes (MK). The low affinity for adenosine, the inducibility of A2bAR expression, and the promiscuous coupling with downstream G-proteins suggest that this receptor may play an instrumental role in platelet function during chronic inflammation and pathophysiological conditions.

Recent sequencing studies show that mRNA expression of A2bAR in human platelets is almost sixfold lower than A2aAR (**Table 1**) (Rowley et al., 2011). Murine platelets lacking A2bAR have reduced cAMP levels and are prone to increased ADP- or collagen-mediated aggregation (Yang et al., 2010). Additionally, A2bAR increases the inhibition of ADP-mediated aggregation of platelets isolated from LPS-injected animals (Yang et al., 2010). Elimination of A2bAR also leads to increased platelet expression of the P2Y1 receptor (Yang et al., 2010), and P2Y1 activation, as mentioned before, leads to an increase in intracellular calcium levels (Cattaneo, 2007; Hechler and Gachet, 2011). This suggests that the absence of A2bAR can have a stimulatory effect on platelet aggregation through simultaneous reduction of cAMP and increase in calcium levels. A2bAR signaling could be particularly important over prolonged periods of inflammation or injury.

The two studies summarized here included healthy human donors and isolated platelets by sorting [large and small platelets (Clancy et al., 2017)] or by centrifugation followed by leukocyte depletion (Rowley et al., 2011). The human cohorts, methods of platelet isolation, and RNA extractions are described in Rowley et al. (2011) and Clancy et al. (2017). Mice sequencing data represent RNA transcripts in platelets from male and female C57BL/6 murine models (Rowley et al., 2011). Of note: The data summarized here are derived from platelets from two human male donors (large and small platelets) and from one male and one female donor (data in "all platelet" column). Data validation is required in various individuals of different ages and sexes in order to establish a broader distribution profile of purinergic receptors. cAMP, cyclic AMP; FPKM, fragments per kilobase of transcript per million mapped reads; RPKM, reads per kilobase of transcript per million mapped reads; Ado, Adenosine; ATP, adenosine triphosphate; ADP, adenosine diphosphate.

### PURINERGIC RECEPTORS AND PLATELET HETEROGENEITY

Circulating platelets are anucleate cell fragments originating from their bone marrow precursor, the MK. Human platelets are not a homogeneous population, but vary in size (2–5 µm) and content. RNA sequencing shows that platelets may contain as many as 9500 different transcripts (Rowley et al., 2011; Bray et al., 2013; Clancy et al., 2017), most of which are prepackaged from the MK. Additionally, platelets uptake transcripts and transcript fragments from circulating cells and the endothelium (Clancy et al., 2017). The heterogeneous size of platelets (Thon and Italiano, 2012) is hypothesized to be a result of platelets losing their contents into the circulation as they age; hence, large platelets are also referred to as "immature" and small platelets are referred to as "mature" (Penington and Streatfield, 1975; Penington et al., 1976a,b). Alternatively, in the late 1970s, it was proposed that platelets vary in density and structure depending on ploidy of the MK from which platelets are originating. Low ploidy MKs produce less hemostatically active platelets while high ploidy MKs produce hemostatically functional platelets (Penington and Streatfield, 1975; Penington et al., 1976a,b). Direct experimental evidence is limited for each of these hypotheses and it is possible that both of these explanations are valid. Studies have shown that large (immature) platelets are highly hemostatically active while small (mature) platelets are less hemostatically functional (Thompson et al., 1984; Guthikonda et al., 2008). It is noteworthy that the predominant size of platelets in a healthy individual lies somewhere in between (Clancy et al., 2017). In patients with acute coronary syndrome (developing early stent thrombosis) there is a baseline platelet size increase predicting platelet reactivity (Huczek et al., 2010). A population study performed with patients after non-cardiac surgery has provided evidence that large platelets may be a novel biomarker for adverse cardiovascular events (Anetsberger et al., 2017).

Sequencing studies have addressed the overall platelet mRNA transcriptome of the entire platelet population (Rowley et al., 2011) and that of small and large platelets sorted from the blood of healthy human donors (Clancy et al., 2017). Of note, in platelets there is a reported mismatch between mRNA and protein expression (Burkhart et al., 2012) that can be partially justified by the ability of platelets to uptake mRNA transcripts (Clancy et al., 2017). However, in cases when both proteins and transcripts are detectable in platelets, there is strong evidence that mRNA and the corresponding protein expression are correlated (Rowley and Weyrich, 2013). Large platelets (sorted as 10% of the entire platelet population) have transcripts associated with classical platelet functions, such as platelet activation/aggregation, hemostasis, and wound healing (Clancy et al., 2017). Small platelets show a distinct and more diverse platelet transcriptome as compared to large platelets, and those transcripts are more involved in platelet-immune cell interactions and apoptosis (Clancy et al., 2017). For instance, small platelets contain distinct transcripts that are associated with activation, proliferation and differentiation of T- and Blymphocytes (Clancy et al., 2017). Consistent with the hemostatic role of large platelets and their contribution to aggregation, transcripts for the P2 purinergic receptors, P2Y12 and P2X1, seem to be present at higher levels in this platelet population (**Table 1**). mRNA from the inhibitory P1 receptor A2aAR is distinctly located in small platelets, similar to P1 A3AR (**Table 1**). The function of platelet-A3AR is unclear. On the other hand, sequencing of the entire platelet population, that includes platelets of all sizes, shows the presence of all four P1 purinergic receptors with the inhibitory A2aAR and A2bAR receptors in the highest proportions (**Table 1**). Interestingly, P2Y1 is detected

only in the mixed, non-sorted platelet population, but not in the large or small platelet subpopulations. There is a possibility that the P2Y1 receptor is present on platelets which compose the rest of the platelet population that was not included in the large/small platelet sorting, or P2Y1 was not detected due to low expression levels in each group or in these particular donors. The two sequencing studies also agree on transcripts for other P2 purinergic receptors such as P2X5, P2Y10, and P2Y13 that have not been previously reported to be present in platelets or to have a functional role. Distinct signatures of purinergic receptors in different platelet subpopulations suggests that the architecture of the hemostatic plug may involve platelets of heterogeneous size and function. Large platelets may be predominantly located at the core while small platelets may be located (together with the large) in the shell (**Figure 1**). Since small platelets have more immune transcripts and are less hemostatically active, their presence may allow for direct interactions with immune cells inside of the shell of the thrombus. The mechanism by which MKs mediate this diverse distribution of distinct transcripts in platelets according to size and function is still controversial.

### CONCLUSION AND FUTURE DIRECTIONS

The distinct distribution of purinergic receptor types across different platelet sizes may provide a new approach to

FIGURE 1 | Distribution of purinergic receptor transcripts with known function in the entire platelet population vs. large and small platelets. Platelets are a heterogeneous population of various sizes. Large platelets are highly hemostatically active while small ones are known to be much less active in hemostasis (Thompson et al., 1984; Guthikonda et al., 2008). RNA sequencing studies shown in Table 1 suggest differential distribution of purinergic receptors across platelets of different sizes, providing a provocative hypothesis related to potential functional differences in hemostasis when activated by adenine nucleosides and nucleotides, depending on platelet size. Future protein studies will be needed to test this contention. P2X1 is a ligand-gated ion channel that requires binding of ATP for influx of calcium; the rest of the P2 and P1 receptors in platelets are G-protein-coupled receptors. Of note, with the sensitivity of the above method of detection, P2Y1 or A2bAR transcripts are not found in the small or large platelets; however, these receptors for ADP or adenosine (respectively) are detected in the entire platelet population, suggesting a differential expression level in different platelet populations, possibly necessary for an extra layer of control over platelet function. These findings further encourage future examination of the receptors at protein and functional levels in different platelet populations. It is not known, however, if all of the purinergic receptors depicted in the top (or bottom) panel can be expressed on the same platelet, if there is differential signature of co-expression or if there is a mix of both of these possibilities. Ado, adenosine; ATP, adenosine triphosphate; ADP, adenosine diphosphate; cAMP, cyclic adenosine monophosphate; AC, adenylate cyclase.

purinergic signaling manipulation in order to establish effective antithrombotic therapies. The platelet mRNA profile may have a broader impact on overall platelet function, and proposes an explanation of previously identified functional differences between small and large platelets, with increased mean platelet volume being historically associated with increased hemostatic potential. It remains unclear, however, if within the sophisticated architecture of the thrombus there is a distinct arrangement of platelets with differential expression of purinergic or immune receptors. Perhaps, big platelets are primarily responsible for the formation of the core of the hemostatic plug since they predominantly contain transcripts related to the hemostatic function of platelets, such as the P2 receptors. Small platelets in turn contain more immune transcripts. These cells may need to communicate with both aggregated platelets and leukocytes and may be located in the porous shell of the hemostatic plaque. In that sense, it would be necessary for small platelets to be less prone to aggregation and hence express only purinergic receptors for shape change (P2X1) and reduction of aggregation (A2ARs). It is also not completely clear if platelet heterogeneity increases as a function of pathological conditions, thereby changing the balance between P2/P1 receptors and limiting the beneficial effect of current therapies.

It would be important to test these receptors at protein and agonist functional levels in the different platelet subpopulations. Additionally, future studies are necessary to elucidate the

### REFERENCES

complex cross-communication of purinergic receptors in the hemostatic plug as a function of platelet heterogeneity and to determine if there is a functional presence of the other ADP/ATP receptors that were detected at mRNA level. These future findings may provide additional pharmacological targets in the management of thrombus growth and stability in patients.

### AUTHOR CONTRIBUTIONS

MK and KR co-wrote and edited this review article.

### FUNDING

This work was supported by AHA Award 16SDG30450001 (to MK) and the Thrombosis and Hemostasis Affinity Research Collaborative and the Evans Center for Interdisciplinary Biomedical Research at Boston University School of Medicine (to KR). KR is an Established Investigator with the American Heart Association.

### ACKNOWLEDGMENTS

We apologize to all whose work was not cited due to limited space.

severe impairment of platelet responses to adenosine diphosphate. Blood 80, 2787–2796.

platelet activity after dual antiplatelet therapy with aspirin and clopidogrel in patients with stable coronary artery disease. J. Am. Coll. Cardiol. 52, 743–749. doi: 10.1016/j.jacc.2008.05.031

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Koupenova and Ravid. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Elevated Pressure Changes the Purinergic System of Microglial Cells

Ana C. Rodrigues-Neves 1,2, Inês D. Aires 1,2, Joana Vindeirinho2,3, Raquel Boia1,2 , Maria H. Madeira1,2, Francisco Q. Gonçalves 2,3, Rodrigo A. Cunha2,3,4, Paulo F. Santos 1,2,5 , António F. Ambrósio1,2,4,6 and Ana R. Santiago1,2,4,6 \*

1 Coimbra Institute for Clinical and Biomedical Research (iCBR), Faculty of Medicine, University of Coimbra, Coimbra, Portugal, 2 CNC.IBILI Consortium, University of Coimbra, Coimbra, Portugal, 3 Center for Neuroscience and Cell Biology, University of Coimbra, Coimbra, Portugal, 4 Faculty of Medicine, University of Coimbra, Coimbra, Portugal, 5 Department of Life Sciences, Faculty of Sciences and Technology, University of Coimbra, Coimbra, Portugal, 6 Association for Innovation and Biomedical Research on Light and Image, Coimbra, Portugal

### Edited by:

Francisco Ciruela, University of Barcelona, Spain

### Reviewed by:

Rodrigo Pacheco, Universidad Andrés Bello, Chile Claire H. Mitchell, University of Pennsylvania, United States

> \*Correspondence: Ana R. Santiago<EMAIL_ADDRESS>
### Specialty section:

This article was submitted to Experimental Pharmacology and Drug Discovery, a section of the journal Frontiers in Pharmacology

> Received: 17 October 2017 Accepted: 05 January 2018 Published: 24 January 2018

### Citation:

Rodrigues-Neves AC, Aires ID, Vindeirinho J, Boia R, Madeira MH, Gonçalves FQ, Cunha RA, Santos PF, Ambrósio AF and Santiago AR (2018) Elevated Pressure Changes the Purinergic System of Microglial Cells. Front. Pharmacol. 9:16. doi: 10.3389/fphar.2018.00016 Glaucoma is the second cause of blindness worldwide and is characterized by the degeneration of retinal ganglion cells (RGCs) and optic nerve atrophy. Increased microglia reactivity is an early event in glaucoma that may precede the loss of RGCs, suggesting that microglia and neuroinflammation are involved in the pathophysiology of this disease. Although global changes of the purinergic system have been reported in experimental and human glaucoma, it is not known if this is due to alterations of the purinergic system of microglial cells, the resident immune cells of the central nervous system. We now studied if elevated hydrostatic pressure (EHP), mimicking ocular hypertension, changed the extracellular levels of ATP and adenosine and the expression, density and activity of enzymes, transporters and receptors defining the purinergic system. The exposure of the murine microglial BV-2 cell line to EHP increased the extracellular levels of ATP and adenosine, increased the density of ecto-nucleoside triphosphate diphosphohydrolase 1 (E-NTPDase1, CD39) and decreased the density of the equilibrative nucleotide transporter 2 as well as the activity of adenosine deaminase. The expression of adenosine A1 receptor also decreased, but the adenosine A3 receptor was not affected. Notably, ATP and adenosine selectively control migration rather than phagocytosis, both bolstered by EHP. The results show that the purinergic system is altered in microglia in conditions of elevated pressure. Understanding the impact of elevated pressure on the purinergic system will help to unravel the mechanisms underlying inflammation and neurodegeneration associated with glaucoma.

Keywords: purinergic system, ATP, adenosine, microglia, elevated pressure, glaucoma

## INTRODUCTION

Glaucoma is a multifactorial degenerative disease characterized by the progressive loss of retinal ganglion cells (RGCs) and optic nerve fibers that leads to vision loss (Casson et al., 2012). Elevated intraocular pressure (IOP) is the most common risk factor of glaucoma and the current treatments are based on lowering the IOP (Cohen and Pasquale, 2014). However, some patients still progress to blindness despite successful control of IOP (Song et al., 2015), suggesting that the control of IOP is not sufficient to prevent RGC loss (Levin et al., 2017).

Microglial cells are the immunocompetent cells of the central nervous system (CNS). Under resting conditions, microglial cells are ramified and constantly monitor their surrounding environment. However, after an insult, microglia change their morphology and become reactive, migrate toward the lesion site, release pro-inflammatory mediators and phagocyte cell debris (Ransohoff and Perry, 2009). Sustained microgliamediated neuroinflammation is associated with several retinal neurodegenerative diseases, namely glaucoma (Madeira et al., 2015a). Exacerbated microglia reactivity has been detected early in experimental models of glaucoma, even before RGC death (Bosco et al., 2011, 2015), suggesting that the modulation of microglia reactivity may attenuate disease progression.

Purine nucleotides are, among others, primordial mediators of cell-to-cell communication (Burnstock and Verkhratsky, 2009; Verkhratsky and Burnstock, 2014). Particularly, purines control several functions of microglia, such as chemotaxis, phagocytosis, and cytokine and chemokine release (reviewed in Di Virgilio et al., 2009). In pathological conditions, neurons, astrocytes and microglia release ATP and adenosine to the extracellular space (Sperlágh and Illes, 2007; Rodrigues et al., 2015). The ratio between the release and the removal, by enzymatic degradation and uptake, determines the extracellular levels of ATP and adenosine. ATP has been postulated to be released by vesicular exocytosis, carrier-mediated release, cytolytic release and through channels and membrane pores (Ballerini et al., 2002; Cook and McCleskey, 2002; Stout et al., 2002; Sperlágh and Illes, 2007; Gajardo-Gomez et al., 2016). Adenosine is considered to be generated by the breakdown of ATP through the actions of NTPDase (CD39) and ecto-5′ -nucleotidase (CD73) (Yegutkin, 2008; Cunha, 2016), but it can also be released on its own (Sperlágh and Illes, 2007). The extracellular levels of adenosine are regulated by equilibrative nucleoside transporters (ENTs), that mediate the bidirectional adenosine transport, or through concentrative transporters (CNTs), that mediate the influx of adenosine (Yegutkin, 2008; Melani et al., 2012). Moreover, adenosine can be metabolized into adenosine monophosphate (AMP) by adenosine kinase (ADK) or into inosine by adenosine deaminase (ADA).

There are indications that the purinergic system is altered in glaucoma, as heralded by the increased levels of ATP detected in the aqueous humor of patients with glaucoma (Zhang et al., 2007; Li et al., 2011) and in experimental models of the disease (Resta et al., 2007; Reigada et al., 2008). Since microglial cells have a role in the pathophysiology of glaucoma and the purinergic system in microglial cells has been demonstrated to play a critical role in controlling inflammation and immune response in several models of neurological diseases (reviewed in Harry, 2013), we now studied the impact of elevated pressure in the set-up of the purinergic system in microglia and also assessed the role of ATP and adenosine in mediating the effects of elevated pressure in microglia phagocytosis and motility.

## MATERIALS AND METHODS

## Microglial Cell Culture

The immortalized murine BV-2 microglia cell line was cultured in Roswell Park Memorial Institute (RPMI) medium, supplemented with 10% fetal bovine serum (FBS) and 1% of antibiotics (penicillin 100 U/ml, streptomycin 100µg/ml) and maintained at 37◦C under a humidified atmosphere with 5% CO2. For experiments, cells were cultured in RPMI with 2% FBS and 1% antibiotics at a density of 6 × 103 cells/cm2 in 6-well-plates or at density of 1 × 104 cells/cm2 in 12-well-plates.

Cells were exposed to elevated hydrostatic pressure (EHP, 70 mmHg above atmospheric pressure) for 4 or 24 h, as previously described (Madeira et al., 2016). Control cultures were kept in a standard incubator at atmospheric pressure.

### ATP Quantification

The extracellular levels of ATP were measured with a luciferinluciferase assay kit, as previously described (Cunha et al., 2000; Madeira et al., 2015b). Briefly, cell culture supernatants were collected and stored at −80◦C until used. Cells were collected and protein concentration was determined by the bicinchoninic acid (BCA) method (Pierce Biotechnology). Supernatants were quickly defrosted and incubated with ATP assay mix (Sigma-Aldrich) in an opaque 96-well plate. ATP levels were measured using a VICTOR multilabel plate reader (Perkin Elmer). The ATP concentration in each sample was determined by interpolation with a standard curve obtained from an ATP stock solution and was normalized to the amount of protein. Results are presented as percentage of control.

## Adenosine Quantification

The extracellular levels of adenosine were quantified using high-performance liquid chromatography (HPLC), as previously described (Vindeirinho et al., 2013). Briefly, cell supernatants were analyzed in Beckman-System Gold HPLC apparatus, with a computer controlled 126 Binary Pump Model using a 166 Variable UV detector (detection at 254 nm) and a Lichrospher 100 RP-18 (5 mm) column from Merck. An isocratic elution with 10 mM phosphate buffer (NaH2PO4; pH 6.0) and 14% methanol was performed with a flow rate of 1.5 ml/min. Adenosine was quantified by considering the retention time and absorption spectra, and then comparing with the standard curve. Results are presented as percentage of control.

(AMP; Sigma-Aldrich) for 40 min at 37◦C. The medium was collected to quantify the free inorganic phosphate, as a measurement of AMP degradation with the Malachite Green Phosphate Assay kit, following the instructions provided by the manufacturer (Cayman Chemicals). Inorganic free phosphate was detected by spectrophotometry (620–630 nm). Results are presented as fold-change of control.

### Measurement of ADA Activity

BV-2 cells were washed twice with ice-cold phosphate-buffered saline (PBS, in mM: 137 NaCl, 2.7 KCl, 10 Na2HPO4, 1.8 KH2PO4, pH 7.4) and lysed in 50 mM Tris-HCl (pH 7.2), supplemented with complete mini protease inhibitor cocktail tablets (Roche). The enzymatic activity of ADA was measured following the instructions provided by the manufacturer (Diazyme). Results were normalized to the amount of protein quantified by the BCA method, and are presented as percentage of control.

### Western Blotting

BV-2 cells were washed twice with ice-cold PBS at 4◦C. Cells were lysed in radioimmunoprecipitation assay buffer [RIPA, in mM: 50 Tris HCl, pH 7.4; 150 NaCl; 5 EDTA; 1% Triton X-100; 0.5% sodium deoxycholate; 0.1% sodium dodecyl sulfate (SDS)] supplemented with complete mini protease inhibitor cocktail tablets and 1 mM of dithiothreitol (DTT) at 4◦C. Samples were denatured by adding 6x concentrated sample buffer (0.5 M Tris, 30% glycerol, 10% SDS, 0.6 M DTT, 0.012% bromophenol blue) and heating for 5 min at 95◦C. When blotting for CD39, cells were lysed in non-reduced conditions.

Samples were separated in SDS-polyacrylamide gel electrophoresis (SDS-PAGE) and transferred onto PVDF membranes (Millipore). The membranes were blocked with 5% milk and then were incubated with the antibodies indicated in **Table 1**. Immunoreactive bands were visualized using Enhanced Chemi-Fluorescence system (ECF; GE Healthcare) on a Storm device (Molecular Dynamics, GE Healthcare) or with Enhance Chemiluminescence system (ECL; Bio-Rad) on a Versadoc (Bio-Rad). Digital quantification of band intensity was performed using Quantity One software (Bio-Rad). The membranes were reprobed for glyceraldehyde 3-phosphate dehydrogenase (GAPDH) to control for similar amounts of protein.

### Immunocytochemistry

Cells were washed with ice-cold PBS and fixed using 4% of paraformaldehyde (PFA) with 4% sucrose for 10 min at room temperature (RT). After fixation, the cells were washed with PBS and permeabilized during 5 min with 1% Triton X-100 in PBS. Then, cells were blocked with 3% of bovine serum albumin (BSA; NZYtech) and 0.2% Tween-20 in PBS for 1 h at RT to prevent the non-specific binding. Afterwards, cells were incubated with the primary antibody diluted in the blocking solution for 90 min at RT (**Table 2**). After washing, the cells were incubated with the secondary antibody in the same solution for 1 h at RT in the dark. Then, cells were rinsed with PBS. Nuclei were stained with 4′ ,6-diamidino-2-phenylindole (DAPI 1:2,000; Molecular Probes, Life Technologies) and F-actin was

stained with phalloidin conjugated to Tetramethylrhodamine B isothiocyanate (TRITC, 1:500; Sigma-Aldrich). Upon rising with PBS, the coverslips were mounted in glycergel mounting medium and observed with a fluorescence microscope (Axio Observer.Z1), using a LD Plan-Neofluar 40x/0.6 Korr Ph2 M27 objective.

Representative images were acquired with a confocal microscope (LSM 710, Zeiss) on an Axio Observer.Z1 microscope using Plan-Apochromat 63x/1.40 Oil Dic M27 objective.

### Boyden Chamber Migration Assay

BV-2 cells were cultured in serum-free medium for 24 h before the experiment. Then, cells were plated in transwell cell culture inserts (8.0µm pore diameter; Merck Millipore) in RPMI with 2% FBS and 1% antibiotics. Cells were incubated with apyrase (30 U/mL) and ADA (1 U/mL) followed by exposure to EHP for 4 h. At the end of the experiment, cells were washed with warm PBS and the cells in upper side of the insert were removed with a cotton swab. After fixation with 4% PFA with 4% sucrose during 10 min, the nuclei were stained with DAPI (1:2,000). The membranes were mounted in glass slides with glycergel mounting medium, and the preparations were observed with a fluorescence microscope (Axio Observer.Z1) using a N-Achroplan 5x/0.15 M27 objective. Five random images per sample were acquired. The number of cells in the bottom side of the insert (the cells that migrated) was counted and the results were expressed as percentage of control.

### Phagocytosis Assay

BV-2 cells were plated in RPMI with 2% FBS and 1% antibiotics and exposed to EHP for 24 h. Cells were incubated with apyrase (30 U/mL) and ADA (1 U/mL) 2 h before the end of the experiment. One hour before the end of the experiment, BV-2 cells were incubated with 0.025% fluorescent beads at 37◦C. Then, cells were washed with warm PBS and fixed with 4% PFA with 4% sucrose. BV-2 cells were stained with TRITCconjugated phalloidin (1:500; Sigma-Aldrich) and nuclei were stained with DAPI (1:2,000). Cells were observed with a confocal microscope (LSM 710, Zeiss) using a Plan-Apochromat 20x/0.8 M27 objective and five random fields were acquired from each condition. The total number of cells in each field, the number of cells with incorporated beads and the number of fluorescent beads phagocytized by each cell were counted. The phagocytic efficiency was calculated:

$$\text{Phosphoytic efficiency } (\%) = \frac{(1 \times \text{x1} + 2 \times \text{x2} + 3 \times \text{x3} + n \times \text{xn})}{\text{total number of cells}}$$

$$\times 100\%$$

where xn represents the number of cells containing n microspheres (n = 1, 2, 3 . . . up to a maximum of 6 points for more than 5 microspheres ingested per each cell).

### Statistical Analysis

Results are presented as mean ± SEM. The number of independent experiments is indicated in each bar. Statistical analysis was performed using GraphPad Prism Version 6 (GraphPad Software). The normality of the data was assessed with Shapiro-Wilk test. Data was analyzed using the nonparametric Kruskall-Wallis test, followed by Dunn's multiple comparison test. Differences were considered significant for p < 0.05.

## RESULTS

Microglial cells are endowed with the machinery of the purinergic system (Sperlágh and Illes, 2007; Castellano et al., 2016). We now assessed how the purinergic system of microglial cells is altered after challenging the microglial cells in a pressure chamber to mimic elevated IOP.

## Elevated Hydrostatic Pressure Increases Extracellular Levels of ATP and Adenosine

BV-2 cells were exposed to elevated pressure for 4 and 24 h and the levels of ATP (**Figure 1A**) and adenosine (**Figure 1B**) were quantified in cell culture medium supernatants. The exposure of microglia to EHP for 4 and 24 h increased the extracellular levels of ATP to 233.1 ± 49.9% (p < 0.01) and 187.9 ± 33.4% of control, respectively, and the adenosine levels to 124.1 ± 9.6% and 131.9 ± 9.6% of the control (p < 0.05), respectively.

## Elevated Hydrostatic Pressure Increases CD39 but Does Not Affect AMP Catabolism

Adenosine can be formed through the hydrolysis of adenine nucleotides [ATP, adenosine di-phosphate (ADP) and AMP] by a cascade of ectonucleotidases, including CD39 and CD73 that are expressed in several cell types, including microglia (Haskó et al., 2005).

Here, we addressed whether EHP could affect the expression of CD39 as well as AMP catabolism, both involved in adenosine formation through ATP hydrolysis. CD73 was not detected in BV-2 cells either by qPCR or Western blot (data not shown). The protein levels of CD39 significantly increased in BV-2 cells exposed to EHP for 4 and 24 h (147.3 ± 23.1% and 128.6 ± 11.0% of the control, respectively; p < 0.05; **Figure 2A**), which is in agreement with the previous proposal that CD39 might be a potential indicator of increased extracellular levels of ATP in retina cells (Lu et al., 2007). However, the dephosphorylation of AMP into adenosine was not altered in BV-2 cells exposed to 4 h EHP (1.0 ± 1 fold-change; **Figure 2B**).

### Elevated Hydrostatic Pressure Impairs the Activity of ADA, but Not the Protein Levels of ADA and ADK

Adenosine can be removed from the extracellular space by degradation into inosine mediated by ADA, while the intracellular the removal of adenosine also occurs by the conversion into AMP mediated by adenosine kinase (Yegutkin, 2008).

The exposure of microglial cells to EHP for 4 and 24 h did not affect the protein levels of ADA and ADK (**Figures 3A,C**). Nevertheless, the activity of ADA (**Figure 3B**) significantly decreased in microglia exposed to EHP for 24 h to 67.9 ± 10.2% of the control (p < 0.05).

## Elevated Hydrostatic Pressure Decreases ENT2 but Does Not Change CNT2 Protein Levels

Extracellular adenosine levels are typically regulated by nucleoside transporters or metabolism (Latini and Pedata, 2001). Hence, we also evaluated the impact of EHP exposure on the expression of nucleoside transporters ENT1, ENT2, and CNT2 in BV-2 cells.

ENT1 was not detected by Western blot (data not shown). The protein levels of ENT2 in BV-2 microglia exposed to EHP for 4 h were similar to the control. However, exposure to EHP for 24 h significantly decreased the protein levels of ENT2 in microglia (46.5 ± 9.6% of the control; p < 0.05; **Figure 4A**). The protein levels of CNT2 were not altered in BV-2 cells exposed to EHP (83.8 ± 13.5% and 105.2 ± 10.0% of the control, for 4 and 24 h, respectively; **Figure 4B**).

## Elevated Hydrostatic Pressure Changes the Expression and Density of A1R but Not A3R

The actions of adenosine are dependent on the activation of the adenosine receptors. We evaluated the impact of EHP on the expression of A1R and A3R receptors in BV-2 microglial cells.

The exposure of BV-2 cells to EHP, for 4 and 24 h, decreased the protein levels of A1R to 64.5 ± 15.6% and 45.9 ± 4.2% (p < 0.05) of the control, respectively, as assessed by Western blot (**Figure 5A**). As we previously reported (Lopes et al., 2003), Western blot for A3R yields three bands but we quantified the more intense 44 kDa that corresponds to the expected molecular weight of A3R. This analysis revealed that the protein levels of A3R (**Figure 5B**) did not change in microglial cells upon exposure

to EHP for 4 and 24 h (97.4 ± 17.2% and 124.0 ± 27.8% of the control, respectively).

The effects of EHP on the immunoreactivity of A1R and A3R were also assessed by immunocytochemistry in microglia (**Figures 5C,D**). In microglial cells exposed to EHP, A1R immunoreactivity decreased when compared with the control, whereas the A3R immunoreactivity was similar to the control, supporting the results obtained by Western blotting.

### ATP and Adenosine Mediate Microglia Migration Induced by Elevated Hydrostatic Pressure

A feature of reactive microglia is their ability to migrate toward the site of injury, where they release pro-inflammatory mediators and phagocyte cell debris (Karlstetter et al., 2010). Although the role of the purinergic system in microglia chemotaxis is well established (Honda et al., 2001; Davalos et al., 2005; Wu et al., 2007), the contribution of ATP and adenosine to the motility of microglia in conditions of elevated pressure it still unknown. Therefore, we evaluated whether EHP could affect microglia migration and if ATP and adenosine affected this process (**Figure 6**). When microglial cells were exposed to EHP for 4 h, the number of microglia that migrated to the bottom side of the transwell significantly increased to 220.5 ± 22.8% of the control (p < 0.05). The treatment of BV-2 cells with 30 U/ml apyrase, to remove the extracellular ATP, prevented the effect of EHP in microglia migration (98.0 ± 10.4% of the control; p < 0.01 compared with EHP). BV-2 cells were also pre-treated with 1 U/ml ADA, to remove the extracellular adenosine. The removal of extracellular adenosine also prevented the migration of microglial cells to the bottom side of the membrane elicited by EHP (115.5 ± 19.2% of the control, p < 0.01 compared with EHP). This strict dependency on extracellular ATP and adenosine for the increased migration of microglia upon EHP is also an important control to exclude the possibility that migration in

EHP conditions might result from cells that were just pushed through the pores.

### ATP and Adenosine Do Not Contribute to the Increase of Phagocytic Efficiency in Microglia Induced by Elevated Hydrostatic Pressure

Increased phagocytic efficiency is a feature of reactive microglia (Kettenmann et al., 2011). The contribution of adenosine and ATP to the phagocytic efficiency of microglia upon exposure to EHP was assessed by incubating microglia with fluorescent microbeads (**Figure 7**). The phagocytic efficiency of microglia exposed to EHP increased by nearly 200% when compared with control cells (from 13 ± 2% in control conditions to 25 ± 3% under EHP; p < 0.01). Incubation with apyrase (30 U/ml) and ADA (1 U/ml) was unable to decrease the phagocytic efficiency in cells under EHP (23 ± 2% and 19 ± 3%, respectively).

## DISCUSSION

The purinergic system is a controller of inflammation (reviewed in Di Virgilio et al., 2009) and microglia-mediated neuroinflammation contributes to the pathogenesis of glaucoma (reviewed in Madeira et al., 2015a). Here, we show for the first time that elevated pressure modifies the purinergic system of BV-2 microglial cells, altering the levels of ATP and adenosine, as well as enzymes responsible for the maintenance of purine levels, which we showed to regulate the EHP-induced microglial cell migration.

EHP increased the extracellular levels of ATP in BV-2 cells, without increasing cell death (data not shown), indicating that

FIGURE 4 | Elevated hydrostatic pressure decreases ENT2 but does not change CNT2 protein levels. Total BV-2 cell extracts were assayed for ENT2 (A) and CNT2 (B) by Western blot. Representative images for ENT2, CNT2, and GAPDH (loading control) are presented above the graphs. Results are expressed as percentage of control. \*p < 0.05, different from control; Kruskal-Wallis test, followed by Dunn's multiple comparison test.

ATP released from these cells occurs by a non-lytic mechanism. Several reports have demonstrated the ability of microglia to release ATP under different conditions (Ferrari et al., 1997; Imura et al., 2013; Shinozaki et al., 2014; George et al., 2015). Indeed, microglia are endowed with the nucleotide transporter (VNUT) that transports cytosolic ATP into vesicles (Sawada et al., 2008) and the genetic deletion of VNUT impairs ATP release from microglia (Shinozaki et al., 2014). Other mechanisms mediate the release of ATP in microglia, namely, the carrier-mediated release (Ballerini et al., 2002), pannexins and/or hemi-channels (Gajardo-Gomez et al., 2016), as well as the cytolytic release (Sperlágh and Illes, 2007).

Increased ATP and adenosine levels have been reported in the aqueous humor of glaucoma patients (Daines et al., 2003; Zhang et al., 2007; Li et al., 2011), implicating the purinergic system in the pathophysiology of glaucoma. Increased ATP levels have been also documented in the retinas of animal models of glaucoma (Lu et al., 2015; Pérez de Lara et al., 2015) and in the vitreous of bovine eyecup preparations subjected to elevated pressure (Reigada et al., 2008). Moreover, we have recently reported that elevated pressure increases extracellular ATP in rat retinal organotypic cultures (Madeira et al., 2015b). In the retinal eyecup and organotypic models it is not possible to identify which cells release ATP, even though several cells are potential sources of extracellular ATP, such as neurons, astrocytes, Müller cells, and microglia (Rodrigues et al., 2015). Astrocytes, for instance, can release ATP when subjected to chronic mechanical strain (Beckel et al., 2014). Under mechanical strain, NLRP3 inflammasome is activated in the retina and astrocytes, due to the upregulation of IL-1β expression, which depends on ATP (released from pannexin hemichannels) and P2X7 receptor stimulation (Albalawi et al., 2017).

The levels of extracellular adenosine were increased in BV-2 microglia exposed to elevated pressure. Extracellular adenosine can arise from the direct release of adenosine into the extracellular space, or by ATP hydrolysis through ecto-enzymes (Yegutkin, 2008; Cunha, 2016), involving the conversion of ATP or ADP into AMP by CD39, followed by AMP dephosphorylation into adenosine by ecto-5′ -nucleotidase (Ecto5′NTase, CD73). Increased expression of NTPDase1 in the posterior eye of rat, mouse, and primate models of glaucoma has been demonstrated (Lu et al., 2015), which is in line with our findings of increased CD39 protein levels in BV-2 cells under elevated pressure. Indeed, NTPDase1 was already proposed as an index for increased ATP levels under pathological conditions (Lu et al., 2007). However, AMP dephosphorylation in microglial cells was not altered by elevated pressure. In addition, we were unable to detect CD73, which is in accordance with a recent study that reported the absence of CD73 expression in the BV-2 cell line (Murphy et al., 2017). Importantly, other enzymes, such as alkaline phosphatase, widely distributed in cell surface, can also degrade ATP, ADP, and AMP into adenosine (Zimmermann, 2006; Yegutkin, 2008; Vardy et al., 2012), and may be responsible for the conversion of extracellular AMP into adenosine in these cells. The half-life of extracellular adenosine may be regulated by extracellular adenosine deaminase (Ecto-ADA) that irreversibly

FIGURE 7 | ATP and adenosine do not contribute to the increase of phagocytic efficiency in microglia induced by elevated hydrostatic pressure. Phagocytosis was assessed using fluorescent microbeads. (A) Representative images of BV-2 cells stained with phalloidin (red) with incorporated beads (green). Nuclei were counterstained with DAPI (blue). Arrowheads show some beads engulfed by microglia. (B) Phagocytic efficiency. Scale bar: 50µm. \*\*p < 0.01, different from control; Kruskal-Wallis test, followed by Dunn's multiple comparison test.

FIGURE 8 | Main alterations in the purinergic system of microglial cells in conditions of elevated pressure. The exposure of microglial cells to elevated pressure changes several players of the purinergic system in microglia, including enzymes, receptors, and transporters. Color code: red means increased levels, expression, or density; blue means decreased expression, density, or activity; and yellow means no changes at elevated pressure when compared with control pressure. Taking into account the lack of CD73 in BV-2 cells, the enzyme responsible for converting AMP into adenosine was not identified; it could be for instance alkaline phosphatase (AP), but we did not assess it experimentally. Adenosine triphosphate (ATP), adenosine monophosphate (AMP), adenosine deaminase (ADA), adenosine kinase (ADK), adenosine A1 receptor (A1R), adenosine A3 receptor (A3R), concentrative nucleoside transporter 2 (CNT2), equilibrative nucleoside transporter 2 (ENT2).

deaminates adenosine into inosine (Regateiro et al., 2013). Although no alterations were detected in the protein levels of ADA, the activity of this enzyme was decreased in cells exposed to elevated pressure, probably as a result of an allosteric modulation of ADA (Levine and Ginsburg, 2013), which we did not experimentally confirmed. This may explain the increase in extracellular adenosine levels. No changes were found in the protein levels of ADK (which converts intracellularly adenosine into AMP). Notably, this enzyme becomes easily saturated with basal concentrations of adenosine (Dunwiddie and Masino, 2001; Latini and Pedata, 2001), which prompts hypothesizing that ADK mostly controls basal levels of adenosine, but does not contribute to these increased levels of adenosine during elevated pressure.

Apart from metabolism, adenosine can also be cleared from the extracellular space by nucleoside transporters (Latini and Pedata, 2001). There are two types of adenosine transporters: equilibrative nucleoside transporters (ENTs) 1-4 and concentrative nucleoside transporters (CNTs) 1–3. Adenosine follows a concentrative gradient through ENTs, while CNTs transport nucleosides against the gradient (Thorn and Jarvis, 1996). ENTs are blocked by S-(4-nitrobenzyl)- 6-thioinosine (NBMPR). In EOC-20 murine microglial cell line and in RAW264.7 macrophage cells, the majority of the adenosine transport is NBMPR-sensitive and insensitive to sodium removal, suggesting that ENTs are the primary transporters functioning in microglia (Carrier et al., 2006), where the presence of ENTs other than 1 and 2 has never been described. The decrease of ENT2 density in microglia caused by exposure to elevated pressure may affect the removal of adenosine from the extracellular space.

Adenosine mediates its actions by means of activation of G protein-coupled receptors named A1R, A2AR, A2BR, and A3R (Fredholm et al., 2001). All adenosine receptors have been described in microglial cells and microglia cell lines (Hammarberg et al., 2003; van Calker and Biber, 2005; Dare et al., 2007; Beckel et al., 2016). It is well documented that under brain noxious conditions, A1Rs are downregulated with a concomitant increase in A2AR density (reviewed in Cunha, 2005). Accordingly, we found that EHP caused A1R downregulation in microglia and in previous reports, we demonstrated the up-regulation of A2AR in microglial cells triggered by elevated pressure (Madeira et al., 2015b, 2016). The downregulation of A1R may result from the exposure of the receptor to an excessive amount of adenosine that results in ligand-induced internalization of the receptor (Ciruela et al., 1997; Coelho et al., 2006).

The purinergic system regulates several functions of microglial cells, such as cell process extension and retraction, migration, proliferation, and phagocytosis (Davalos et al., 2005; Koizumi et al., 2007; Gomes et al., 2013; Matyash et al., 2017). Microglia migration increased under elevated pressure, and this process appears to be mediated by ATP and adenosine. Cell migration can occur by three mechanisms: basal motility (in the absence of a chemical stimulus), chemokinesis (random motility in response to a chemical stimulus), and chemotaxis (migration toward and dependent of a chemical gradient; Wilkinson, 1998; Miller and Stella, 2009). ATP is a known microglial chemotactic agent (Honda et al., 2001; Davalos et al., 2005; Wu et al., 2007) acting on P2Y12 and P2X4 receptors (Honda et al., 2001; Haynes et al., 2006). In addition to chemotaxis, ATP can mediate migration by chemokinesis (Miller and Stella, 2009). Moreover, adenosine may also play a role although it stills remains to be determined which adenosine receptor might bolster microglia chemotaxis: in fact A2ARs are responsible for microglia retraction (Orr et al., 2009), whereas it was proposed that the activation of A3R is required for ADP-induced P2Y12-mediated migration of microglia (Ohsawa et al., 2012).

Phagocytosis is a crucial function of microglia both in the surveillance and in reactive states (Mandrekar et al., 2009; Karlstetter et al., 2014; Scheiblich and Bicker, 2016). Elevated pressure increased microglia phagocytosis, but extracellular ATP and adenosine failed to prevent the increased phagocytosis, indicating that ATP and adenosine do not control this microglial function under elevated pressure conditions. In addition, ATP and adenosine per se did not change the phagocytic efficiency under basal conditions (data not shown). Others reported that the dual activation of P1 and P2 are mandatory for both migration and phagocytic capacity of microglial cells (Bulavina et al., 2013), which is not in total accordance with our results. Therefore, we cannot discard the possibility that the synergistic action of ATP and adenosine could be pivotal in phagocytosis, during elevated pressure conditions.

Our data showed for the first time that elevated pressure impaired the purinergic system of microglial cells (**Figure 8**) and this could interfere with microglial functions under elevated pressure. While it is important to keep in mind that an in vitro model of elevated pressure is an over-simplified model of glaucoma and that the BV-2 microglia cell line has particularities different from endogenous microglia in their native environment, the present results still pave the way to better appreciate the purinergic system in microglial cells as a putative target to be further considered for the management of glaucoma.

## AUTHOR CONTRIBUTIONS

AR-N, IA, and AS conceived and designed the experiments; AR-N, IA, JV, and FG performed the experiments; AR-N, IA, JV, RB, MM, FG, RC, PS, AA, and AS analyzed and interpreted the results; RC, PS, AA, and AS contributed with reagents, materials, analysis tools; AR-N wrote the first draft of the manuscript and all authors have read and approved the final version.

## FUNDING

This work was supported by FCT, Portugal (Project PTDC/BIM-MEC/0913/2012 and Strategic Projects PEst-C/SAU/UI3282/ 2013 and UID/NEU/04539/2013), FEDER-COMPETE (POCI-01-0145-FEDER-007440), Centro 2020 Regional Operational Programme (CENTRO-01-0145-FEDER-000008: BrainHealth 2020).

## ACKNOWLEDGMENTS

The authors acknowledge Prof. Sancha Santos, Faculty of Pharmacy, University of Coimbra, for the help with the HPLC equipment.

## REFERENCES

role for pannexin hemichannels. Neuroscience 157, 396–404. doi: 10.1016/j.neuroscience.2008.08.036

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Rodrigues-Neves, Aires, Vindeirinho, Boia, Madeira, Gonçalves, Cunha, Santos, Ambrósio and Santiago. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Phenotyping of Mice with Heart Specific Overexpression of A2A-Adenosine Receptors: Evidence for Cardioprotective Effects of A2A-Adenosine Receptors

### Edited by:

Francisco Ciruela, University of Barcelona, Spain

### Reviewed by:

Jason N. Peart, Griffith University, Australia Andrew F. James, University of Bristol, United Kingdom

### \*Correspondence:

Joachim Neumann joachim.neumann@medizin. uni-halle.de Ulrich Gergs<EMAIL_ADDRESS>
### Specialty section:

This article was submitted to Experimental Pharmacology and Drug Discovery, a section of the journal Frontiers in Pharmacology

Received: 06 November 2017 Accepted: 05 January 2018 Published: 22 January 2018

### Citation:

Boknik P, Drzewiecki K, Eskandar J, Gergs U, Grote-Wessels S, Fabritz L, Kirchhof P, Müller FU, Stümpel F, Schmitz W, Zimmermann N, Kirchhefer U and Neumann J (2018) Phenotyping of Mice with Heart Specific Overexpression of A2A-Adenosine Receptors: Evidence for Cardioprotective Effects of A2A-Adenosine Receptors. Front. Pharmacol. 9:13. doi: 10.3389/fphar.2018.00013 Peter Boknik1 , Katharina Drzewiecki1 , John Eskandar1 , Ulrich Gergs2 \*, Stephanie Grote-Wessels1 , Larissa Fabritz3 , Paulus Kirchhof3 , Frank U. Müller1 , Frank Stümpel1 , Wilhelm Schmitz1 , Norbert Zimmermann4 , Uwe Kirchhefer1 and Joachim Neumann2 \*

1 Institut für Pharmakologie und Toxikologie, Universitätsklinikum Münster, Westfälische Wilhelms-Universität, Münster, Germany, 2 Institut für Pharmakologie und Toxikologie, Medizinische Fakultät, Martin-Luther-Universität Halle-Wittenberg, Halle, Germany, 3 Centre for Cardiovascular Sciences, University of Birmingham, Birmingham, United Kingdom, 4 Bundesinstitut für Arzneimittel und Medizinprodukte, Bonn, Germany

Background: Adenosine can be produced in the heart and acts on cardiac adenosine receptors. One of these receptors is the A2A-adenosine receptor (A2A-AR).

Methods and Results: To better understand its role in cardiac function, we generated and characterized mice (A2A-TG) which overexpress the human A2A-AR in cardiomyocytes. In isolated atrial preparations from A2A-TG but not from WT, CGS 21680, an A2A-AR agonist, exerted positive inotropic and chronotropic effects. In ventricular preparations from A2A-TG but not WT, CGS 21680 increased the cAMP content and the phosphorylation state of phospholamban and of the inhibitory subunit of troponin in A2A-TG but not WT. Protein expression of phospholamban, SERCA, triadin, and junctin was unchanged in A2A-TG compared to WT. Protein expression of the α-subunit of the stimulatory G-protein was lower in A2A-TG than in WT but expression of the α-subunit of the inhibitory G-protein was higher in A2A-TG than in WT. While basal hemodynamic parameters like left intraventricular pressure and echocardiographic parameters like the systolic diameter of the interventricular septum were higher in A2A-TG than in WT, after β-adrenergic stimulation these differences disappeared. Interestingly, A2A-TG hearts sustained global ischemia better than WT.

Conclusion: We have successfully generated transgenic mice with cardiospecific overexpression of a functional A2A-AR. This receptor is able to increase cardiac function per se and after receptor stimulation. It is speculated that this receptor may be useful to sustain contractility in failing human hearts and upon ischemia and reperfusion.

Keywords: A2A-adenosine receptor, contractility, ischemia, reperfusion, protein phosphorylation

### INTRODUCTION

fphar-09-00013 January 18, 2018 Time: 17:55 # 2

Adenosine elicits multiple effects in the human body. These include effects on cardiac function that have been studied for many years and some effects of adenosine including the downstream signaling mechanisms are similar to those of vagal stimulation. For example, the negative chronotropic effect on the sinus node, the negative dromotropic effect on the AV-node, and the negative inotropic effect in atrial tissue of adenosine (Shryock and Belardinelli, 1997). Extracellular adenosine acts on A1-, A2-, and A3-adenosine receptors (AR). Typically, A1-AR inhibits and A2-AR stimulates adenylyl cyclase activities (Olsson and Pearson, 1990). The A2A-AR was the first of the P1-purinoceptor family to be cloned (Libert et al., 1989).

The negative inotropic effects of adenosine in the rat atrium were mediated by inhibition of adenylyl cyclase activity followed by a decreased cAMP content (Linden et al., 1985). But in guinea pig atrium, no adenosine-induced decrease in cAMP content was found (Böhm et al., 1984). Here, the negative inotropic effect of adenosine was inhibited by pertussis toxin (Böhm et al., 1986) and therefore mediated by a pertussis toxin-sensitive G-protein. Probably, an enhanced conductance of potassium in the sarcolemma was involved (Belardinelli and Isenberg, 1983). However, the activation of protein phosphatases has also been suggested (Gupta et al., 1993a,b).

In contrast, in the ventricle of most species, adenosine is nearly ineffective to influence force of contraction but via activation of A1-adenosine receptors, it can decrease the effects of β-adrenergic stimulation or in other words the inotropic effects of cAMP-elevating agents are decreased by adenosine [for example forskolin or phosphodiesterase inhibitors; rat (Dobson, 1978, 1983) or guinea pig (Böhm et al., 1984; Brückner et al., 1985)]. Adenosine can also provoke relaxation of the vasculature via activation of A2A-AR on smooth muscle cells [human coronaries (Makujina et al., 1992)].

Adenosine is produced in cardiomyocytes and its release there is markedly enhanced during β-adrenergic stimulation, ischemia, or necrosis. In the clinic, adenosine and its precursor ATP can be used to stop supraventricular arrhythmias and therefore, the effects of adenosine in the mammalian heart are of clinical relevance and should be further investigated. It is controversial and may be species dependent (or method dependent) whether A2A-AR is functional (increase in cAMP, increase in contractility) in the mammalian heart: some reported a lack of effect [rat (Shryock et al., 1993), guinea-pig (Boknik et al., 1997), rabbit (Kilpatrick et al., 2002)] whereas other reported a functional response [mouse (Morrison et al., 2006), rat (Monahan et al., 2000)].

It is important to note, that A2A-receptors in human hearts have been detected on the protein level (Marala and Mustafa, 1998). Moreover, work in isolated perfused A2A-KO (constitutive deletion) mouse hearts clearly established that CGS 21680 a classical A2A-receptor agonist was selective: only in WT hearts CGS 21680 (up to 1 µM) increased contractility but not in hearts from A2A-KO mice (Ledent et al., 1997). However, under basal conditions (no CGS 21680 given) contractility of WT and KO were not different (Ledent et al., 1997; Ashton et al., 2017), suggesting that CGS 21680 is an appropriate tool to assess A2A function in mice. Moreover, there is evidence that A2A-receptor stimulation can protect the heart against reperfusion injury [e.g., rat (Cargnoni et al., 1999)].

As part of our long-standing effort to characterize all known adenosine receptors in the mammalian heart, we report here on the generation and phenotyping of a novel transgenic mouse model in which we successfully expressed an inotropically active A2A-AR. We used this model to study reperfusion injury in the isolated perfused heart. A progress report of this work has appeared as abstract (Grote-Wessels et al., 2007).

### MATERIALS AND METHODS

### Isolation of A2A-Receptor cDNA and Generation of Transgenic Mice

The investigation conforms to the Guide for the Care and Use of Laboratory Animals published by the National Research Council (2011). Animals were handled and maintained to according to approved protocols of the animal welfare committee of the University of Münster, Germany.

The PCR generated human A2A-AR cDNA fragment containing a 30 and 50 engineered NotI digestion site was inserted into a mouse cardiac α-myosin heavy chain promoter expression cassette via NotI. The orientation of the cDNA was confirmed by sequencing. The A2A-AR cDNA promoter construct was digested with NruI and purified by a cesium chloride gradient centrifugation for injection into the pronuclei of single-cell fertilized mouse embryos. Generation of transgenic (A2A-TG) mice was performed by standard procedures (mouse strain: FVB/N). One transgenic line overexpressing the A2A-AR under the control of the α-myosin heavy chain (MHC) promoter was established, which was investigated in the present study. Genotypes were identified by PCR analyses of tail-tip DNA using the following primers 50 -acaaagcaggcgatgaag-30 and 50 -acccttaccccacatagacc-30 . For the reverse transcription 4 µg RNA and random primers were used (Transcriptor High Fidelity cDNA Synthesis Kit, Roche Applied Science, Mannheim, Germany). The PCR reaction was performed using the Ampliqon Taq DNA polymerase (Biomol, Hamburg, Germany) according to the manufacturer's instructions. For all experiments, 12–30 weeks old A2A-TG mice and WT littermates of both sexes were used.

### Western Blot Analysis

Homogenates from ventricular tissue samples were prepared in 300 µl of 10 mM NaHCO3 and 100 µl 20% SDS. Crude extracts were incubated at 25◦C for 30 min before centrifugation to remove debris and thereafter, the supernatants (= homogenates) were separated and stored at −20◦C until further use. Western blot analysis was performed as previously described (Gergs et al., 2004). Briefly, aliquots of 100 µg of protein were loaded per lane and finally, bands were detected using enhanced chemifluorescence (ECF, GE Healthcare, Munich, Germany) together with a StormTM PhosphorImager (GE Healthcare, Munich, Germany).  The characteristics and use of these antibodies has been reported repeatedly by our group (Kirchhefer et al., 2002). The antibodies against TnI and phosphor-TnI were obtained from GE Healthcare (Freiburg, Germany), the antibodies against α-subunits of Gi-protein and Gs-protein were purchased from Calbiochem (Darmstadt, Germany).

### Histological Analysis

fphar-09-00013 January 18, 2018 Time: 17:55 # 3