Blood from healthy donors was collected under informed written consent as approved by the ethical committee of Region Hovenstaden, Denmark, under license H-3-2013-054. Peripheral blood mononuclear cells (PBMCs) were isolated from buffy coats from healthy donors by gradient centrifugation in LymphoPrep (#1114740, Axis-Shield). Negative selection was carried out on fresh cells with RosetteSep (#15022, #15023, StemCell) or from frozen PBMCs using EasySep (#19052, #19053, StemCell). Frozen PBMCs were quickly thawed, resuspended in fresh medium, and rested for 2 h at 37°C before use. Cells were kept in RPMI-1640 (#BE12-702F, Lonza) + 10% fetal bovine serum (FBS) (#S0115, Biochrom). Freezing was done in additional 10% FBS and 10% DMSO.

Transfected cells in 24 wells were harvested and cell pellets stored at −80°C before mRNA extraction and gene expression measurements. Primary RosetteSep isolated cells were maintained as 8 × 10⁵/mL with or without the addition of Dynabeads-Human T-Activator CD3/CD28 (#11161D, Gibco, Life Technologies). Cell pellets were collected and quickly frozen days 0–3 at −80°C.

mRNA was extracted by RNeasy Mini Kit (#74106, Qiagen). cDNA synthesis was carried out using TaqMan Reverse Transcription Reagents (#N8080234, Invitrogen, Life Technologies). qPCR gene expression was performed using TaqMan Universal PCR Master Mix (#4369016, Applied Biosystems, Life Technologies) with ACTB (β-actin) and GAPDH as housekeeping genes (list of primers shown in Table 1). Two separate primer/probe sets were used to analyze P2RY11 expression in primary T lymphocytes and transfected cells, as the primer/probe set used for primary cells spanned the 3'-untranslated region of the gene, which was not present in the vector. B2M and EEF1A1 genes were used as housekeeping genes because GAPDH and ACTB are not stable following immune activation (14, 15).

Cell pellets were snap-frozen in dry ice and stored at −80°C until later use. The cells were lysed in RIPA buffer (#856545, RegionH Pharmacy) supplemented with Triton X-100 for a concentration of 1% and cOmplete protease inhibitor was added (#11697498001, Roche Diagnostics). Samples were stored at −20°C until further use. Protein was denatured for 5 min at 95°C in 25% Laemmli buffer [0.25 M Tris-HCl (#839847, RegionH Pharmacy), 40% glycerol (#104092, Merck Millipore), 20% 2-mercaptoethanol (#M-6250, Sigma-Aldrich), 2% sodium dodecyl sulfate (SDS) (#L4390, Sigma-Aldrich), 0.01% bromophenol blue (#B-7021, Sigma-Aldrich)] before loading into RunBlue SDS Precast 4–20% in 12-well 10 × 10 cm gel (#NXG42012, Expedeon) and run in RunBlue SDS running buffer (#NXB50500, Expedeon). Protein was transferred onto an ethanol-activated Hypond-P membrane (#10600057, GE Healthcare Life Sciences) in transfer buffer [Tris-Gly buffer (#843856, RegionH Pharmacy), 20% ethanol, 0.1% SDS (#161-0416, BioRad)]. Blocking solution for unspecific binding was prepared as 2% Amersham ECL Prime Blocking Reagent (#RPN418V, GE Healthcare Life Sciences) in TBST buffer [TBS (#838571, RegionH Pharmacy), 0.1% Tween20], and membranes were blocked for 1 h at room temperature (RT) before incubation in primary rabbit anti-human P2X7 receptor (#ab109246, Abcam) or loading control diluted in TBST buffer with 0.4% blocking solution at 4°C overnight. Loading control was 1:5,000 mouse anti-β-actin (#A5441, Sigma-Aldrich) with a size of 42 kDa. On the following day, membranes were washed three times in TBST before shaking for 1 h at RT in 1:40,000 secondary ELC donkey HRP-linked anti-rabbit IgG (#NA934V, GE Healthcare Life Sciences) or 1:20,000 ELC sheep HRP-linked anti-mouse IgG (#NA931V, GE Healthcare Life Sciences) diluted in TBST. After washing, membranes were developed with Amersham ECL Select Western Blotting Detection Reagent (#RPN2235, GE Healthcare Life Sciences) for 5 min at RT and illuminated with chemiluminescence on LAS-4000 system.

Stripping and re-blotting were done with re-blot strong solution (#2504, Merck Millipore) for 15 min followed by blocking and staining with a primary rabbit anti-human P2Y₁₁ receptor antibody (#ab183096, Abcam) as described above. This antibody has been found suitable for the detection of the P2Y₁₁ receptor in Western blotting (16). Loading control staining with anti-β-actin was done analog to the above following another round of stripping of the membrane.

Peripheral blood mononuclear cells were thawed and rested as described above. 10⁶ cells were placed per tube and washed with phosphate buffered saline (PBS) (#L0615-500, Biowest), 400 g for 5 min at 4°C before staining with Fixable Viability Dye eFlouro 506 (#65-0866, eBioscience) for 20 min at RT. Cells were blocked for unspecific binding in PBS with 10% human serum (#H4522, Sigma-Aldrich) for 15 min on ice. Then, they were washed with PBS and incubated with combinations of extracellular anti-human CD3-BV421 (#563798, BD Bioscience), anti-human CD4-FITC (#11-0047-42, eBioscience), or CD8-PE-Cy7 (#344712, Nordic BioSite) for 30 min at 4°C. The cells were washed in PBS with 0.5% human serum before fixation and stained as previously described (16). Briefly, human P2Y₁₁ receptor staining was performed on unpermeabilized cells with 0.1-µg Lightning Link (#705-0030, Innova Biosciences, UK) APC-conjugated 1:100 rabbit anti-human P2Y₁₁ receptor antibody (#ab140878, Abcam) against the extracellular part for 30 min. We have previously tested this antibody and found it specific for the detection of human P2Y₁₁ over P2Y₁ and P2Y₂ in transfected cells by flow cytometry (16). Flow cytometry was done on FACSverse (BD Biosciences, USA), and the FACSuite software (BD Biosciences, USA) was used for data analysis.

Cells were transfected with siRNA against P2RY11 (#L-005691-00-0005) and non-target control #1 (#D-001810-10-05, SMARTpool, ON-TARGETplus, Thermo Scientific) using 0.125 nmol on 2.0 × 10⁶ cells in an Amaxa Nucleofector (Lonza) as previously described (17). The cells were following transfection cultured in RPMI-1640 containing 2 mM l-glutamine and 100 mg/mL penicillin/streptomycin (Sigma-Aldrich). All cells were supplemented with 10% human serum (The Danish National Bloodbank, Denmark) and stimulated with Dynabeads-Human T-activator CD3/CD28 (#11131D, Thermo Scientific).

Human embryonic kidney (HEK-293) cells were maintained in culture medium: DMEM (#BE12-604F, Lonza) + 10% FBS + 0.5% p/s (#DE17-602E, Lonza) at 37°C, 5% CO₂, and humidified air. GripTite 293 MSR cell line (a generous gift from Søren Gøgsig Faarup Rasmussen, University of Copenhagen, Denmark) is a genetically engineered HEK-293 line expressing the human macrophage scavenger receptor for better surface adherence. GripTite cells were maintained in culture medium supplemented with 1% non-essential amino acid (NEAA) (#M7145, Sigma-Aldrich) and 600 µg/mL antibiotic selection agent Geneticin (#10131-019, Gibco, Life Technologies) at 37°C, 5% CO₂, and humidified air. Cells were passaged two to three times per week with trypsin (#L0930-100, Biowest) and versene (#15040-066, Gibco, Life Technologies).

Vectors used for transfection were pcDNA3.1 (empty vector), eGFP, human P2RX7 in a pcDNA3.1 backbone, human P2RY11 (#EX-Z1416-M02, GeneCopoeia), and human P2RY11-mCherry (EX-Z1416-M56, GeneCopoeia). The C919T mutation was created in the P2RY11-mCherry vector using QuikChange Lightning Site-Directed Mutagenesis Kit (#210518, Agilent Technologies) following the manufacturer’s protocol. Mutation primers were F: 5'-ggcatgaggccccacatcacctggtag-3' and R: 5'-ctaccaggtgatgtggggcctcatgcc-3' (LGC Biosearch Technologies, Denmark). The mutated plasmid construct was sequenced at Eurofins Genomics Sequencing Department, Germany.

We used a reverse transfection protocol where the cells are seeding in wells during transfection or by seeding transfected cells into wells to control for transfection efficiency. Cotransfection of eGFP was included in the control group in some experiments or the application of a vector construct of P2Y₁₁ receptor fused to mCherry to visualize successful transfection. The mCherry tag changed neither the calcium nor the cyclic adenosine monophosphate (cAMP) response of the P2Y₁₁ receptor as shown in Figure S1A in Supplementary Material. Also, the gene expression levels in cells following transfection were measured to compare gene expression levels to rule out group-to-group variation (Figure S1B in Supplementary Material).

Transfection of cells for lactate dehydrogenase (LDH) and pore formation measurements was carried out by allowing the formation of DNA:Lipofectamine complexes in a medium without selection agent for 5 min. DNA:Lipofectamine mixture was dissolved to a final concentration of 0.5% Lipofectamine 2000 Reagent (#11668-019, Invitrogen, Life Technologies) and 1 µg DNA/mL. Cells were resuspended in this solution and seeded as 5 × 10⁴cells/well poly-d-lysine (#P6407, Sigma)-coated transparent or black clear bottom 96-well plate or 2.5 × 10⁵cells/well in poly-d-lysine-coated 24-well plate for gene expression. After a 5-h incubation, the medium was changed to culture medium and incubated overnight at 37°C, 5% CO₂, and humidified air.

Cells for calcium imaging were cultured as 9.5 × 10⁵cells/well in a 6-well plate for 24 h. Lipofectamine and DNA were dissolved in DMEM + 10% FBS + 1% NEAA for a total of 2 µg DNA/well and 0.25% Lipofectamine, respectively, and allowed to transfect overnight. Cells were washed in PBS and loosened with versene and trypsin and cultured overnight as 3 × 10⁵cells/well in a black clear-bottomed poly-d-lysine-coated 96-well plate before further analysis.

Cells were incubated with the synthetic ATP-analog 3'-O-(4-benzoyl)benzoyl-ATP (BzATP, #B6396-25MG, Sigma) dissolved in water, P2X7 receptor antagonist A740003 (#sc-291774, Santa Cruz Biotechnology) dissolved in ethanol, 4,4'-[carbonylbis(imino-3,1-(4-methyl-phenylene)-carbonylimino)]-bis(naphthalene-2,6-disulfonic acid) tetrasodium salt (NF340) dissolved in water (#3830/10, Tocris) or 1% Triton X-100 (#T9284-500 mL, Sigma-Aldrich) for 2 h before LDH measurement. EasySep-isolated lymphocytes at a concentration of 10⁵ cells/well were incubated for 2 h in round-bottomed 96-well plates with various concentrations of the abovementioned compounds before LDH measurement. LDH release was measured using an LDH colorimetric assay (#ab102526, Abcam). Absorbance for the culture medium without cells was subtracted from the values before calculating LDH release. All experiments included a nicotinamide adenine dinucleotide standard curve and a positive LDH control supplied with the kit.

Twenty-four hours after medium change, transfected cells were washed once in dye buffer prepared as Hank’s Balanced Salt Solution (HBSS), (#14170112, Gibco, Life Technology) with 0.2% YO-PRO-1^® Iodide 491/509 (#Y3603, Life Technologies). Plates were read with NOVOStar (BMG LABTECH GmbH) microplate reader for every 2 min at 489/510 for 8 min before manually adding stimulatory compounds. The plate was then read for 64 min every 2 min. Pore formation was assessed by calculating the change in fluorescence intensity as F_max − F_min.

One day after seeding in 96-well plate, the cells were washed with a washing buffer (HBSS (#14065-049, Gibco), 10 mM HEPES (#15630-080, Gibco), 2 mM probenecid (#P8761, Sigma) (pH = 7.4)) and next incubated in a loading buffer (2 µM Fluo-4 AM (#F14201, Molecular Probes, in DMSO), 0.02% pluronic acid F-127 (#P2443, Sigma, in DMSO) in washing buffer) for 30 min. Cells were washed and allowed to rest for 10 min at 37°C in a washing buffer before automatic addition of BzATP. The signal was detected using the NOVOStar microplate reader system in a 2-min time span. For each data point, the maximal increase in the signal above baseline signal before BzATP addition was calculated. The resulting dose–response curves were normalized using baseline as 0% signal and signal at the highest dose as 100%. Next, the curve was fitted using a four-parameter fit of either a sigmoidal dose–response curve (one receptor) or a biphasic sigmoidal dose–response curve (two receptors). The fitted curves were finally used to calculate EC₅₀. All calculations were performed using GraphPad Prism 7.

Transfected cells in 96-well plates were lysed for 10 min in 0.1 M HCl. Debris was removed by centrifugation at ≥600 g 5 min. The supernatant was analyzed with cAMP ELISA colorimetric kit (#ADI-900-066, Enzo Life Sciences) according to manufacturer’s protocol. Briefly, the plate was prepared with 50 µL/well neutralizing reagent. Blue conjugate and yellow antibody were added to each well and incubated for 2 h on a plate shaker. The samples were washed three times in a washing buffer before the addition of substrate and incubation 1 h without shaking. The reaction was stopped and read at 405 nm. For each data point, cAMP concentration in pmol/mL was calculated on the basis of a standard curve. The resulting dose–response curve was fitted using a four-parameter fit of a sigmoidal dose–response curve using GraphPad Prism 7.

All statistical analyses were carried out in GraphPad Prism 7. Gene expression in CD4⁺ and CD8⁺ T lymphocytes was compared in a two-sided t-test. Gene expression changes in CD4⁺ T lymphocytes following in vitro immune activation were analyzed with one-way ANOVA and Dunnett’s post hoc testing against day 0. LDH activity in transfected CD8⁺ T lymphocytes was analyzed with one-way ANOVA and Dunnett’s post hoc testing. The mean values from individual donors or experimental runs were used for statistical analysis using two-way ANOVA with two groups post hoc tested using Sidak’s multiple comparison testing for each concentration. Multiple dose–response curves were compared with two-way ANOVA and Tukey’s multiple comparison post hoc testing.

To study the ATP-induced death of human CD4⁺ and CD8⁺ T lymphocytes, we treated cells isolated from healthy donors with increasing concentrations of the synthetic ATP-analog BzATP. Cell death was estimated by measuring LDH activity in the supernatant. This enzyme leaks from dying cells as a result of membrane permeabilization and can be used as an indirect indicator of plasma membrane integrity and cell death. The results showed that human CD4⁺ T lymphocytes released more LDH than CD8⁺ T lymphocytes (Figure 1A) after 2 h of incubation with BzATP. This difference is similar to what has been observed for CD4⁺ and CD8⁺ T lymphocytes in mice (7, 8). The increase in LDH release induced by BzATP could be completely blocked by the P2X7 receptor antagonist, A740003 (Figure 1B). Cells treated for 24 h showed similar levels of LDH release (Figure S2A in Supplementary Material), suggesting that cell death occurred during the first 2 h after BzATP stimulation. This is consistent with a previous report where P2X7 receptor-mediated cell lysis was observed as early as 15 min after stimulation with ATP in mouse lymphocytes (18).

The higher P2X7 receptor-mediated response in mouse CD4⁺ T lymphocytes compared to CD8⁺ T lymphocytes has been explained by differences in expression levels of the P2X7 receptor (7, 8). Therefore, P2RX7 gene and protein expression in human CD4⁺ and CD8⁺ T lymphocytes were measured. Surprisingly, there was no difference in the expression levels of P2RX7 mRNA between human CD4⁺ and CD8⁺ T lymphocytes (Figure 2A), and the P2X7 receptor protein level was similar or slightly lower in CD4⁺ T lymphocytes compared to that in CD8⁺ T lymphocytes from five different human donors when examined by Western blot (Figure 2B). A similar human P2X7 receptor expression was previously been found in human CD4⁺ and CD8⁺ T lymphocytes with flow cytometry (19, 20). This shows that in human individuals, the greater P2X7 receptor-induced LDH release from CD4⁺ T lymphocytes following BzATP stimulation is not explained by higher P2X7 receptor expression levels.

We, therefore, hypothesized that the differential BzATP response in human CD4⁺ and CD8⁺ T lymphocytes was due to a protective effect of a high P2Y₁₁ receptor expression counteracting P2X7 receptor-mediated cell death in CD8⁺ T lymphocytes. This was based on previous findings that P2Y₁₁ receptor stimulation mitigated the effect of ATP-induced cell death in PBMCs (12) and delayed apoptosis in neutrophils (13). The mRNA expression and protein levels of the P2Y₁₁ receptor were indeed found to be significantly higher in human CD8⁺ compared to that in CD4⁺ T lymphocytes (Figures 3A,B) complying with the hypothesis.

Immune-activated murine CD4⁺ T lymphocytes have lower levels of P2X7 receptor and are more resistant to P2X7 receptor-dependent cell death than their naïve counterparts (21). Therefore, the level of P2RX7 and P2RY11 gene expression in human CD4⁺ T lymphocytes following in vitro immune activation was investigated. Over a 3-day period, both P2RX7 and P2RY11 gene expression levels increased in CD4⁺ T lymphocytes following immune activation. No changes in P2RY11 or P2RX7 gene expression levels were found in CD8⁺ T lymphocytes following immune activation (data not shown). In contrast to murine CD4⁺ T lymphocytes, human CD4⁺ T lymphocytes did not downregulate P2RX7 gene expression upon immune activation. Instead, a higher P2RX7 level was observed (Figure 3C). Immune-activated CD4⁺ T lymphocytes showed a marked decrease in BzATP-induced LDH release compared to naïve cells when tested at day 2 (Figure 3D). Clearly, the reduced BzATP response in the human cells was not caused by a decrease in P2RX7 expression as shown for murine cells. In line with our hypothesis, P2RY11 gene expression was more than 10-fold upregulated in human CD4⁺ T lymphocytes following immune activation (Figure 3C), suggesting that in humans, the lower P2X7 receptor-mediated LDH release from immune-activated CD4⁺ T lymphocytes is caused by an increased P2Y₁₁ receptor expression.

Currently, the most specific P2Y₁₁ antagonist known is NF340 that blocks P2Y₁₁ stimulation by BzATP in the dose range of 0.1–10 µM (22). If the P2Y₁₁ signaling inhibited P2X7 activation, we expected that antagonizing the P2Y₁₁ receptor with NF340 would increase LDH release. NF340 had, however, no effect on BzATP-induced P2X7 activity in primary CD4⁺ and CD8⁺ T lymphocytes when tested in doses ranging from 7.4 to 200 µM (Figure S2B in Supplementary Material) on a baseline of 1,000 µM BzATP. We worried that the high concentrations of BzATP applied to induce significant P2X7 activation would make it impossible to block P2Y₁₁ activation with NF340 or any other pathway-interfering compound. Therefore, to test the effect of P2Y₁₁ signaling on P2X7 cell death, we turned to the use of siRNA.

Transient transfection of CD8⁺ T lymphocytes with P2RY11 siRNA resulted in an almost complete removal of P2RY11 mRNA after 24 and 48 h (Figure 4A), but not in CD4⁺ T lymphocytes (data not shown). This decrease was not equally reflected in protein expression when examined with a flow cytometer. The P2Y₁₁ receptor expression was close to identical between groups (Figures 4B,C). This suggested a low turnover of P2Y₁₁ receptor on the CD8⁺ T lymphocytes. Having already found that immune activation could increase P2RY11 gene expression in CD4⁺ T lymphocytes, we tested if immune activation upregulated P2Y₁₁ receptor protein in the CD8⁺ T lymphocytes. The CD8⁺ T lymphocytes showed two populations with high and low P2Y₁₁ receptor expressions, respectively (see gating strategy in Figure S3 in Supplementary Material). We found that the level of P2Y₁₁ receptor protein was upregulated following immune activation in CD8⁺ T lymphocytes in the P2Y₁₁^high subset (Figure 4C). This was not seen in immune-activated CD8⁺ T lymphocytes when P2RY11 was silenced. Hence, silencing of P2RY11 in human CD8⁺ T lymphocytes decreased P2RY11 gene expression and prevented the cells from upregulating P2Y₁₁ receptor protein expression following immune activation.

When the transfected CD8⁺ T lymphocytes were stimulated with BzATP, the immune-activated P2RY11-silenced CD8⁺ T lymphocytes had a significantly higher LDH release than immune-activated non-silenced cells (Figure 4D). This suggests that following immune activation, the T lymphocytes upregulate P2Y₁₁ receptor, making them more resistant to cell death and that silencing P2RY11 prevents the cells from using this protective mechanism.

Next, we studied the mechanism behind how P2Y₁₁ receptor interfered with P2X7 receptor-driven membrane permeabilization and consecutive cell death in cell cultures overexpressing P2RY11 and P2RX7 separately or in combination. The cells were treated with various doses of BzATP, and P2X7 receptor-induced cell death was measured by LDH release into the supernatant after 2 h stimulation. As expected, cells overexpressing P2RX7 displayed a significant increase in LDH release compared to control cells transfected with an empty vector (Figure 5A). This effect was mediated by the P2X7 receptor, as LDH release was completely inhibited by adding the P2X7 receptor antagonist A740003 (Figure 5B). Interestingly, coexpression of P2RY11 prevented the BzATP-induced LDH release. This complies with our hypothesis that the P2Y₁₁ receptor protects against P2X7 receptor-driven cell death. To rule out that the inhibition of LDH release was caused by a lower P2RX7 expression in the cotransfected cells, the gene expression levels for P2RX7 and P2RY11 were measured in the transfected cells. Coexpression of P2RY11 did not result in a decreased gene expression of P2RX7 (Figure S1C in Supplementary Material).

Membrane permeabilization and LDH release as a response to ATP stimulation are believed to be caused by a pore-forming action of the P2X7 receptor (23). We, therefore, tested the hypothesis that P2RY11 coexpression interfered with membrane permeabilization and LDH release observed in BzATP-treated cells by preventing P2X7 receptor pore formation. Cells overexpressing P2RX7 were stimulated with BzATP, and YO-PRO-1 uptake was quantified as a measure of pore formation (Figure 5C). YO-PRO-1 enters the cell through the P2X7 receptor pore and becomes fluorescent upon DNA binding. A significant P2X7 receptor pore formation was observed at lower concentrations of BzATP than LDH release. The BzATP-induced increase in YO-PRO-1 signal was exclusively caused by dye uptake through the P2X7 receptor pore, as the P2X7 receptor antagonist A740003 completely removed the effect of BzATP (Figure 5D). At concentrations above 500 µM BzATP, the YO-PRO-1 dye signal from P2RX7 transfected cells decreased (data not shown) most likely due to cell detachment following cell death. Coexpression of P2RY11 completely inhibited pore formation in P2RX7 expressing cells (Figure 5C).

By contrast, calcium signaling through P2X7 receptor appeared unaffected as cells expressing both P2RX7 and P2RY11 showed a biphasic response to BzATP with EC₅₀ values equal to their respective single-transfected counterparts (Figure 6). Values from fitted sigmoidal curves were P2RY11 EC₅₀ = 0.29 (0.23–0.36) μM and P2RX7 EC₅₀ = 53 (47–61) μM, respectively, and the values for the biphasic sigmoidal dose–response curve for P2RX7 + P2RY11 were EC₅₀ = 0.56 (0.35–1.05) μM and EC₅₀ = 82 (54–132) μM. This suggested that P2RY11 expression interfered with P2X7 receptor pore formation independent of calcium signaling.

The P2Y₁₁ receptor is a GPCR that signals via both G_s and G_q to activate the adenylyl cyclase and the phosphate inositol pathway (9, 10). We next asked whether the block of pore formation was caused by P2Y₁₁ receptor signaling through G_s and/or G_q pathways. However, as the BzATP concentration needed to activate the P2X7 receptor at the same time saturated the P2Y₁₁ receptor, it was not possible to activate P2X7 receptor while at the same time inhibiting P2Y₁₁ receptor signaling (Figure 6). Instead, we tested a non-signaling P2Y₁₁ receptor mutant in the cotransfection system. Among other amino acids, arginine 307 in the P2Y₁₁ receptor has been predicted to be involved in ATP binding, interacting with the phosphate P_α moiety (24). This has been confirmed experimentally by mutational analysis, revealing inactivation of the receptor as measured by intracellular calcium release (24). Therefore, vectors expressing wild-type P2Y₁₁ receptor and C919T-mutated P2RY11 sequence resulting in the Arg³⁰⁷ being substituted with a tryptophan were tested. The mutated P2Y₁₁ receptor was completely non-signaling as expected with regard to both calcium and cAMP signaling (Figures 7A,B). Coexpression of either P2Y₁₁ or P2Y₁₁ C919T receptors showed identical inhibition of P2X7 receptor YO-PRO-1 dye uptake, demonstrating that the P2Y₁₁ receptor-induced inhibition of P2X7 receptor function is independent of signaling (Figure 7C).