Pharmacological and genetic characterisation of the canine P2X4 receptor

Abstract

Background and Purpose

P2X4 receptors are emerging therapeutic targets for treating chronic pain and cardiovascular disease. Dogs are well‐recognised natural models of human disease, but information regarding P2X4 receptors in dogs is lacking. To aid the development and validation of P2X4 receptor ligands, we have characterised and compared canine and human P2X4 receptors.

Experimental Approach

Genomic DNA was extracted from whole blood samples from 101 randomly selected dogs and sequenced across the P2RX4 gene to identify potential missense variants. Recombinant canine and human P2X4 receptors tagged with Emerald GFP were expressed in 1321N1 and HEK293 cells and analysed by immunoblotting and confocal microscopy. In these cells, receptor pharmacology was characterised using nucleotide‐induced Fura‐2 AM measurements of intracellular Ca²⁺ and known P2X4 receptor antagonists. P2X4 receptor‐mediated inward currents in HEK293 cells were assessed by automated patch clamp.

Key Results

No P2RX4 missense variants were identified in any canine samples. Canine and human P2X4 receptors were localised primarily to lysosomal compartments. ATP was the primary agonist of canine P2X4 receptors with near identical efficacy and potency at human receptors. 2′(3′)‐O‐(4‐benzoylbenzoyl)‐ATP, but not ADP, was a partial agonist with reduced potency for canine P2X4 receptors compared to the human orthologues. Five antagonists inhibited canine P2X4 receptors, with 1‐(2,6‐dibromo‐4‐isopropyl‐phenyl)‐3‐(3‐pyridyl)urea displaying reduced sensitivity and potency at canine P2X4 receptors.

Conclusion and Implications

P2X4 receptors are highly conserved across dog pedigrees and display expression patterns and pharmacological profiles similar to human receptors, supporting validation and use of therapeutic agents for P2X4 receptor‐related disease onset and management in dogs and humans.

Abbreviations

5‐BDBD

5‐(3‐bromophenyl)‐1,3‐dihydro‐2H‐benzofuro[3,2‐e]‐1,4‐diazepin‐2‐one

BX430

1‐(2,6‐dibromo‐4‐isopropyl‐phenyl)‐3‐(3‐pyridyl)urea

BzATP

2′(3′)‐O‐(4‐benzoylbenzoyl)‐ATP

ECS

extracellular Ca²⁺ solution

EmGFP

Emerald GFP

SR

seal resistance

TNP‐ATP

2′,3′‐O‐(2,4,6‐trinitrophenyl)‐ATP

What is already known

• P2X4 receptors in humans and rodents have established roles in nociception and cardiovascular regulation.

• No data regarding functional canine P2X4 receptors have been reported.

What this study adds

• Canine P2RX4 encodes a functional ATP‐gated Ca²⁺ channel with pharmacological similarities to human P2X4 receptors.

• The P2X4‐selective inhibitor BX430 was less potent and sensitive against canine, than human, P2X4 receptors.

What is the clinical significance

• Compounds targeting human P2X4 receptors also target canine P2X4 receptors with similar effectiveness.

• Canine models provide valid preclinical assessment of P2X4 receptor‐targeting compounds as potential therapeutic agents.

1. INTRODUCTION

The https://www.guidetopharmacology.org/GRAC/ObjectDisplayForward?objectId=481 receptor is a member of the ionotropic https://www.guidetopharmacology.org/GRAC/FamilyDisplayForward?familyId=77 and forms homo‐trimeric, https://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=1713‐gated Ca²⁺ channels in macrophages and glial cells (Trang & Salter, 2012; Tsuda, Tozaki‐Saitoh, & Inoue, 2010). Here, it has important roles in signalling the release of https://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=1883 (Ulmann, Hirbec, & Rassendren, 2010) and https://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=4872 (Trang, Beggs, Wan, & Salter, 2009; Ulmann et al., 2008), respectively. Elucidation of the role of P2X4 receptors in these well‐known nociceptive signalling pathways has led to suggestions that P2X4 receptors are an attractive therapeutic target for the treatment of a number of major health issues such as inflammation and chronic pain (Bernier, Ase, & Seguela, 2018), nerve injuries (Su et al., 2019), and alcohol use disorder (Huynh et al., 2016). Functional P2X4 receptors are also present on endothelial cells in humans and mice where it is associated with blood flow‐dependent calcium signalling (Yamamoto et al., 2000, 2006).

To date, two different missense variants in the gene coding P2X4 (P2RX4) have been associated with human disease. The missense Y315C variant (rs28360472) is significantly associated with increased pulse pressure (Stokes et al., 2011). The same variant is associated with increased susceptibility to age‐related macular degeneration when in linkage disequilibrium with a P2RX7 missense G150R variant (rs28360447), which results in a reduction of P2X7 receptor‐mediated phagocytosis by macrophages and microglia (Gu et al., 2013). A second P2RX4 variant (G135S; rs765866317) in a three‐variant haplotype (together with the P2RX7 variants T205M [rs140915863] and N361S [rs201921967]) is associated with multiple sclerosis in a multi‐incident family (Sadovnick et al., 2017). As such, P2X4 receptors have been proposed as possible therapeutic targets for the treatment of multiple sclerosis (Zabala et al., 2018).

Despite the publication of high‐resolution crystal structures of zebrafish P2X4 receptors (Hattori & Gouaux, 2012; Kawate, Michel, Birdsong, & Gouaux, 2009), researchers have struggled to identify potent and selective P2X4 receptor antagonists (Tian et al., 2014). However, in recent years, several commercially available antagonists with greater selectivity for these receptors have appeared, although with disputed effectiveness. Data regarding P2X receptor subtype specificity and species selectivity are incomplete (see Stokes, Layhadi, Bibic, Dhuna, & Fountain, 2017, for a review of current P2X4 receptor antagonists). Overcoming species‐specific pharmacological differences is a huge challenge faced by researchers developing drugs that target ion channels such as P2X4 receptors. Such differences often hinder the translation of observations from preclinical animal models to humans. However, to date, the characterisation of canine P2X4 receptors has not been reported.

To assist with preclinical validation of therapeutic agents for P2X4 receptor‐related disease, we have synthesised canine P2X4 receptors and subcloned them into a mammalian expression vector for functional characterisation, revealing a striking pharmacological similarity to human P2X4 receptors. Recently described antagonists at human P2X4 receptors were further characterised, and their ability to inhibit Ca²⁺ flux through canine and human P2X4 receptors was directly compared.

2. METHODS

2.1. Isolation and sequencing of genomic DNA from dogs

All animal care and experimental procedures complied with institutional (University of Wollongong) guidelines. Informed consent was obtained from all pet owners. This work was approved by the Animal (protocols AE10/01 and AE14/09) and Human (protocol HE10/063) Ethics Committees of the University of Wollongong. Animal studies are reported in compliance with the ARRIVE guidelines (Kilkenny, Browne, Cuthill, Emerson, & Altman, 2010) and with the recommendations made by the British Journal of Pharmacology.

Canine peripheral blood (≤1 ml·kg body weight) was collected into VACUETTE lithium heparin tubes (Greiner Bio‐One, Frickenheisen, Germany) from the jugular vein of 101 non‐sedated or anaesthetised pedigree and non‐pedigree dogs (details in Table S2). Genomic DNA was extracted from these blood samples using the Wizard Genomic DNA Purification Kit (Promega, Madison, USA) according to the manufacturer's instructions and stored at −80°C. Canine genomic DNA was amplified using forward and reverse primer pairs designed to intronic regions flanking between one and three exons of canine P2RX4 as indicated (Table S1). PCR amplification was carried out using MangoTaq DNA polymerase with an initial denaturation at 95°C for 2 min, followed by 35–40 cycles of denaturation at 95°C for 30 s, annealing at 58.8–67°C for 30–60 s, and extension at 72°C for 1 min. After cycling was complete, a final extension was carried out at 72°C for 5 min, and then amplicons were incubated with ExoSAP‐IT at 37°C for 15 min, followed by heat inactivation at 80°C for 15 min. P2RX4 sequences from amplicons were determined using the respective primers listed in Table S1, the Applied Biosystems (Foster City, USA) ABI PRISM BigDye Terminator v3.1 Cycle Sequencing Kit and a 3130xl Genetic Analyser. Resulting sequence chromatograms were compared to the ENSEMBL reference gene transcripts for canine P2RX4 described below.

2.2. Expression constructs

Synthetic canine and human P2RX4 cloned into pUC57 plasmids were purchased from Sigma‐Aldrich (St. Louis, USA) and subcloned into a mammalian pEGFP‐N3 vector with a C‐terminal Emerald GFP (EmGFP) tag connected by a 3x(GGGGS) flexible linker. Plasmid DNA was amplified by transformation into chemically competent DH5α (Thermo Fisher Scientific, Melbourne, Australia) bacterial cells and isolated using the Wizard Plus SV Minipreps DNA Purification kit (Promega, Madison, USA) according to the manufacturer's instructions. Plasmid DNA was sequenced as above using primer pairs listed in Table S1. The canine and human P2RX4 sequences obtained from these vectors were identical to their respective full‐length reference transcripts obtained from ENSEMBL (http://www.ensembl.org; canine P2RX4‐201 ENSCAFT00000013312.3; human P2RX4‐202 ENST00000337233.8).

2.3. Cell lines

1321N1 human astrocytoma (ECACC Cat# 86030402, RRID:CVCL_0110) and HEK293 cells (ECACC Cat# 85120602, RRID:CVCL_0045) were maintained in DMEM/F12 medium containing 15% or 10% FBS, respectively, 2‐mM GlutaMAX, 100 U·ml⁻¹ penicillin, 10 μg·ml⁻¹ streptomycin, and 1% non‐essential amino acids at 37°C/5% CO₂. Human cell line identities were confirmed through STR profiling (Garvan Molecular Genetics, Darlinghurst, Australia) and were routinely assessed for mycoplasma contamination using the MycoAlert Mycoplasma Detection Kit (Lonza, Waverley, Australia). Cells were plated at 3.5 × 10⁵ cells per well in poly‐d‐lysine‐coated six‐well plates (Greiner Bio‐One, Frickenheisen, Germany) and transiently transfected the following day with canine or human P2X4‐EmGFP, or the EmGFP empty vector using Lipofectamine 3000 as per the manufacturer's instructions. Briefly, 0.4 μg of plasmid DNA, 5 μl of Lipofectamine reagent, 4 μl of P3000 reagent, and 250 μl of Opti‐MEM (per well of a six‐well plate transfection) were incubated at room temperature for 15 min prior to adding to cells. After transfection (24–48 hr), cells were plated at 5 × 10⁴ cells per well in black‐walled μClear bottom, poly‐d‐lysine‐coated 96‐well plates (Greiner Bio‐One) for Ca²⁺ flux assays, or at 1 × 10⁵ cells per 18‐mm poly‐d‐lysine‐coated glass coverslip for confocal microscopy, and incubated at 37°C/5% CO₂ overnight.

2.4. Fura‐2 calcium response assay

P2X4 receptor activity was assessed using Fura‐2 measurements of nucleotide‐induced Ca²⁺ influx as described (Ma, Hui, Pelegrin, & Surprenant, 2009). Unless stated otherwise, all assays and compound dilutions were carried out using a standard extracellular Ca²⁺ solution (ECS; 145‐mM NaCl, 2‐mM CaCl₂, 1‐mM MgCl₂, 5‐mM KCl, 13‐mM glucose, and 10‐mM HEPES, pH 7.4). Cells were washed twice with ECS and then preincubated with Fura‐2 AM loading buffer (2.5‐μM Fura‐2 AM/0.2% pluronic acid in ECS) in the dark for 30 min at 37°C. Before recording fluorescence, excess Fura‐2 loading buffer was removed, and cells were washed with ECS and then incubated for a further 20 min to allow complete de‐esterification. Fura‐2 fluorescence emission at 510 nm was recorded every 5 s using a Flexstation3 (Molecular Devices, Sunnyvale, USA) following excitation at 340 and 380 nm (six reads per well, photomultiplier tube setting medium). Baseline recordings were taken for 15 s, and then following the addition of nucleotide or vehicle (up to a final volume of 100 μl per well), recordings were taken for up to 3 min. Where indicated, antagonists, https://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=2373 (IVM), or respective vehicle controls were added to each column of the 96‐well assay plate at regularly spaced 2‐min intervals at 37°C. After 20 min of preincubation, Fura‐2 fluorescence (as described above) was measured for 15 s, and then following the addition of ATP or vehicle (up to a final volume of 100 μl per well), recordings were taken for up to 2 min. Compound additions were carried out using the Flexstation3 built‐in fluidics system. In some experiments, hexokinase (4.5 U·ml⁻¹) was incubated with 1.5‐mM https://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=1712 (or ATP) in ECS for 60 min at 37°C prior to addition of nucleotides. Data were acquired using SoftMax Pro 7 software (Molecular Devices).

2.5. Automatic planar patch clamp electrophysiology

HEK293 cells transfected with canine or human P2X4 were detached from six‐well plates using TrypLE and resuspended in cold external recording solution (140‐mM NaCl, 5‐mM glucose, 4‐mM KCl, 2‐mM CaCl₂, 1‐mM MgCl₂, and 10‐mM HEPES, 298 ± 3 mOsm, pH 7.4 with NaOH) containing 1 U·ml⁻¹ apyrase. Cells were kept in suspension by automatic pipetting at 4°C. Patch clamp measurements were performed on an NPC‐16 Patchliner (Nanion Technologies, Munich, Germany) using multihole medium resistance NPC‐16 chips with an average resistance of 1.1 MΩ. Recordings were made in the whole‐cell configuration with internal solution (60‐mM CsF, 50‐mM CsCl, 20‐mM NaCl, 10‐mM HEPES, and 10‐mM EGTA, 285 ± 3 mOsm, pH 7.2 with CsOH) and external recording solution in absence of apyrase. Seal formation was enhanced by brief treatment with SE seal enhancement solution (80‐mM NaCl, 3‐mM KCl, 10‐mM MgCl₂, 35‐mM CaCl₂, and 10‐mM HEPES, 298 ± 3 mOsm, pH 7.3 with NaOH) until stable seals were obtained and replaced with standard external solution. Solutions were 0.2‐μm membrane filtered. Recordings where seal resistance (SR) was >500 MΩ and access resistance was <3xSR were considered acceptable. Chip and whole‐cell capacitance were fully compensated, and series resistance compensation (70%) was applied via Auto Rs Comp function. Recordings were acquired with PatchMaster (HEKA Elektronik, Lambrecht/Pfalz, Germany) and stored on a computer running PatchControlHT software (Nanion Technologies GmbH, Munich, Germany). A test pulse in control bath solution application (0 ATP) was used to subtract leak currents. Offline analysis was performed using Igor Pro‐6.37 (WaveMetrics Inc.). Solutions were puffed (~1 s) during a 5‐s test pulse from a holding potential (Vh) of −70 mV. The integer of AUC described by the inward currents in response to the ATP puff was used for quantification of agonist‐induced responses, with concentration–response curves plotted as charge density (pC·pF⁻¹) normalised to current at 1‐mM ATP. All recordings were conducted at room temperature.

2.6. Confocal microscopy

The antibody‐based procedures used in this study comply with the recommendations made by the British Journal of Pharmacology (Alexander et al., 2018). Cells on glass coverslips were fixed with 3% paraformaldehyde at 4°C for 15 min and then washed three times with PBS. Cells were permeabilised with 0.1% saponin in 2% BSA/PBS at room temperature for 15 min and then incubated with goat anti‐P2X4 (1:250; Sigma‐Aldrich Cat# SAB2500734, RRID:AB_10604119, Lot# 8830P1) or mouse anti‐LAMP1 (1:100; DSHB Cat# H4A3, RRID:AB_2296838) antibodies in 2% BSA/PBS at room temperature for 2 hr. Cells were washed three times with PBS and then incubated with Alexa Fluor 594‐conjugated donkey anti‐goat (1:200; Abcam Cat# ab150136, RRID:AB_2782994, Lot# GR308670‐1) or PE‐conjugated sheep anti‐mouse (1:200; Chemicon Cat# AQ326H, Lot# 985052005) antibodies in 2% BSA/PBS at room temperature for 60 min. Washed coverslips were mounted onto a glass slide using 50% glycerol in PBS and sealed with nail polish. Cells were visualised on a Leica (Mannheim, Germany) SP5 confocal microscope.

2.7. Western blotting

Cells (1 × 10⁶) were washed three times with ice‐cold PBS and harvested by mechanical scraping and centrifugation (300× g for 5 min). Following incubation in complete lysis buffer (50‐mM BisTris, 750‐mM 6‐aminohexanoic acid, 1‐mM PMSF, 1% n‐dodecyl β‐d‐maltoside, and 1 cOmplete EDTA‐free protease inhibitor cocktail tablet, pH 7.0) at 4°C with gentle agitation for 60 min, cells were sheared 10 times through an 18‐gauge needle and centrifuged at 16,000× g for 15 min at 4°C. Cell supernatants were separated by denaturing SDS‐PAGE under reducing conditions (10‐mM DTT) using Mini‐Protean TGX Stain‐Free gels (4–20%) and then transferred onto a nitrocellulose membrane. The membrane was washed three times in Tris‐buffered saline solution containing Tween‐20 (TBST; 20‐mM Tris, 500‐mM NaCl, 0.1% Tween‐20, pH 7.5) and blocked in blocking buffer (TBST containing 5% skim milk powder) for 60 min at room temperature. The membrane was then incubated overnight with goat anti‐P2X4 (1:500) or rabbit anti‐actin (1:2,000; Sigma‐Aldrich Cat# A2066, RRID:AB_476693) antibodies in blocking buffer at 4°C. The membrane was washed three times for 5 min with TBST and incubated with HRP‐conjugated mouse anti‐goat (Thermo Fisher Scientific Cat# 31400, RRID:AB_228370, Lot# KC1205131) or goat anti‐rabbit IgG (Rockland Cat# 611‐103‐122, RRID:AB_218567, Lot# 21231; both 1:5,000) antibodies in blocking buffer at room temperature for 60 min. The membrane was washed as above and visualised using chemiluminescent substrate and an Amersham Imager 600RGB (GE Healthcare Lifesciences).

2.8. Data and statistical analysis

The data and statistical analysis comply with the recommendations of the British Journal of Pharmacology on experimental design and analysis in pharmacology (Curtis et al., 2018). The relative change in intracellular Ca²⁺ (ΔCa²⁺) data was expressed as the ratio of Fura‐2 fluorescence following excitation at 340 and 380 nm (F _340/380) and then normalised to the mean basal fluorescence according to the formula ΔCa²⁺ = ΔF/F = (F − F _rest)/F _rest, where F is the F _340/380 at a given time and F _rest is the mean fluorescence of the given well prior to the addition of nucleotides (Paredes, Etzler, Watts, & Lechleiter, 2008). Nucleotide‐induced ΔCa²⁺ responses were then calculated by subtracting the ΔCa²⁺ response in absence of the respective nucleotide (measured simultaneously). To minimise potential caveats of high‐throughput fluorescence‐based assays such as large, erroneous ΔCa²⁺ measurements (Heusinkveld & Westerink, 2011), peak nucleotide‐induced ΔCa²⁺ responses were defined by the median of five readings surrounding the peak nucleotide‐induced ΔCa²⁺ response. To control for plate‐to‐plate variation over time and slight variations in P2X4 receptor expression, the peak nucleotide‐induced ΔCa²⁺ response was normalised to the response to 10‐μM ATP alone (per cent of maximum), which was measured for both canine and human P2X4 receptors for each independent experiment. Agonist concentration–response curves were plotted by a non‐linear regression fit to the Hill equation using the least squares method, following a replicates test to confirm adequate fit of the curve. Half‐maximal responses are expressed as their negative logarithm (pEC₅₀/pIC₅₀) ± SEM. Unpaired two‐tailed Student's t test or two‐way ANOVA with Bonferroni post hoc test was used as indicated in the figure and table legends where group size was equal to at least five independent values (with two or more technical replicates per sample to ensure the reliability of single values), there was no significant variance in homogeneity (Bartlett's or F test for equal variance), and the data followed a Gaussian distribution (determined by a D'Agostino–Pearson omnibus K ² test). Any data with less than five independent values were not subject to statistical comparisons. A non‐parametric Mann–Whitney U test was carried out to compare differences when data were normalised to a 100% control with no variance. Curve fitting and statistical comparisons were made using GraphPad Prism 5 (GraphPad Software, San Diego, CA). Colocalisation analysis and calculation of Pearson's coefficient was performed using the JACoP (Just Another Co‐localization Plugin) tool for ImageJ. Differences between P2RX4 variant prevalence were compared using the Fisher's exact test. For determining differences between groups, P < .05 was considered statistically significant throughout the entire manuscript, and statistical analysis was carried out only on independent values, including any outliers. Assays and data analysis were performed unblinded and nonrandomised. However, where possible, plate structure was routinely altered to mitigate positioning bias within data sets.

2.9. Materials, compounds, and solutions

DMEM/F12 medium, ExoSAP‐IT, Fura‐2 AM, GlutaMAX, HRP‐conjugated anti‐goat antibody, Lipofectamine 3000, Opti‐MEM I reduced serum medium, penicillin–streptomycin, ExoSAP‐IT, Subcloning Efficiency™ DH5α™ Competent Cells, TrypLE Express, and trypsin–EDTA were from Thermo Fisher Scientific. ABI PRISM BigDye Terminator v3.1 Cycle Sequencing Kit was from Applied Biosystems (Carlsbad, USA). MangoTaq DNA polymerase was from Bioline (London, UK). HRP‐conjugated anti‐rabbit IgG antibody was from Rockland Immunochemicals (Pottstown, USA). Mini‐Protean TGX Stain‐Free™ gels, Precision Plus Protein™ Dual Colour standards, and nitrocellulose membrane were from Bio‐Rad (Hercules, USA). Alexa Fluor 594‐conjugated donkey anti‐goat IgG antibody was from Abcam (Melbourne, Australia). PE‐conjugated sheep anti‐mouse IgG antibody was from Chemicon (Boronia, Australia). Mouse anti‐LAMP1 antibody (H4A3) was from the Developmental Studies Hybridoma Bank (Iowa City, USA). FBS (heat inactivated) was from Bovogen Biologicals (East Keilor, Australia). Wizard Genomic DNA Purification and Wizard Plus SV Minipreps DNA Purification kits were from Promega (Madison, USA). 6‐Aminohexanoic acid, ADP (sodium salt, cat. no. A2754), apyrase (cat. no. A6535), ATP (disodium salt, cat. no. A7699), 2′(3′)‐O‐(4‐benzoylbenzoyl)‐ATP (https://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=1757; triethylammonium salt, cat. no. B6396), cOmplete™ EDTA‐free protease inhibitor cocktail, n‐dodecyl β‐d‐maltoside, DMSO, (S)‐https://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=202 (HCl, cat. no. SML0474), EGTA, goat anti‐P2X4 antibody (SAB2500734), hexokinase from Saccharomyces cerevisiae (cat. no. H4502), ivermectin (cat. no. 18898), MEM non‐essential amino acid solution, https://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=4790 (maleate salt, cat. no. P1372), PBS, pluronic F‐127, PMSF, poly‐d‐lysine hydrobromide (5 μg·ml⁻¹ working stock), rabbit anti‐actin antibody, and saponin were from Sigma‐Aldrich. 5‐(3‐Bromophenyl)‐1,3‐dihydro‐2H‐benzofuro[3,2‐e]‐1,4‐diazepin‐2‐one (https://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=9541; cat. no. 3579), 1‐(2,6‐dibromo‐4‐isopropyl‐phenyl)‐3‐(3‐pyridyl)urea (https://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=9543; cat. no. 5545), 2′,3′‐O‐(2,4,6‐trinitrophenyl)‐ATP (https://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=4324; tetra(triethylammonium) salt, cat. no. 2464), and https://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=5907 (cat. no. 4890) were from Tocris Bioscience (Bristol, UK). BSA, EGTA, and reagents for preparing buffers were from Sigma‐Aldrich or Amresco (Solon, USA). Primers for sequencing were obtained from GeneWorks (Hindmarsh, Australia) or Integrated DNA Technologies (Coralville, USA).

Compounds were initially made up to stock concentrations in either Milli‐Q H₂O (100‐mM ATP [pH 7.4 with 5‐M NaOH], 30‐mM ADP, 40‐mM BzATP, 10‐mM duloxetine, and 10‐mM TNP‐ATP) or DMSO (10‐mM 5‐BDBD, 30‐mM BX430, 30‐mM paroxetine, 3‐mM IVM, and 30‐mM AR‐C118925). Compounds were stored at −20°C and then diluted in ECS for Ca²⁺ response assays or standard external solution for patch clamp recordings on the day of use, performed unblinded by the experimenter. DMSO controls for each compound (up to 1% in ECS) did not appear to have an effect on ATP‐induced Ca²⁺ responses.

2.10. Nomenclature of targets and ligands

Key protein targets and ligands in this article are hyperlinked to corresponding entries in http://www.guidetopharmacology.org, the common portal for data from the IUPHAR/BPS Guide to PHARMACOLOGY (Harding et al., 2018) and are permanently archived in the Concise Guide to PHARMACOLOGY 2019/2020 (Alexander, Mathie et al., 2019).

3. RESULTS

3.1. Genetic characterisation of canine P2X4 receptors

To determine the presence of exonic variants in canine P2RX4, genomic DNA samples from a randomly selected cohort of 101 dogs were isolated and sequenced using Sanger sequencing. The cohort consisted of 69 pedigree dogs (representing 32 different pedigrees) and 32 non‐pedigree dogs (Table S2). Approximately 85% of the protein‐coding region of P2RX4 for this cohort was sequenced; some regions of various samples could not be amplified or sequenced despite three separate attempts. A query of the NCBI dbSNP, the EMBL‐EBI European Variation Archive (EVA), and the iDOG databases (last accessed March 2019) revealed three reported missense variants: one in the first exon of P2RX4 (no rsID, 26C>A, A9D; EVA database), one in exon 7 (no rsID, 691C>T, R231C; EVA database), and one in exon 10 (rs852294963, 1033C>G, L345V; NCBI dbSNP database). However, these three reported variants were not observed in our cohort, nor were any novel missense variants identified (Table 1). Several synonymous variants are also reported in the iDOG database (Table S3). However, only rs852444127 (798C>T, T266T) was identified in our cohort and then only in heterozygous dosage in two Labrador retrievers and one beagle (Table 1). A previously unreported synonymous variant (15C>T) resulting in an unchanged cysteine at position five (C5C) was found in our cohort in heterozygous dosage only in both pedigree dogs (n = 5) and non‐pedigree dogs (n = 3; Table 1), four of which were of Maltese pedigree. This variant was absent in two other Maltese, whilst exon 1 could not be amplified in a further four Maltese. Both the allele frequency (0.33 and 0.04) and prevalence (66% and 7%) of this variant were significantly different between dogs of Maltese and non‐Maltese pedigree, respectively. Combining these data indicates that the C5C variant may be associated with, but not limited to, dogs of Maltese pedigree.

TABLE 1.

Distribution of canine P2RX4 exon variants in our cohort of dog samples

Variant	Allele change	cDNA position	n	Genotype		Allele frequency	Prevalence
				Wild type (n)	Mutant a (n)
Observed variants b
C5C	tgC > tgT	15	63	55	8	0.063	12.7%
T266T	acC > acT	798	88	85	3	0.017	3.4%
Reported variants c
A9D	gcC > gcA	26	63	55	0	ND	ND
R231C	Cgt > Tgt	691	90	90	0	ND	ND
L345V	Ctc > Gtc	1,033	82	82	0	ND	ND

Abbreviation: ND, not detected.

^aOnly heterozygous variants were observed in this cohort.

^bVariants that were observed at least once in our cohort.

^cVariants reported previously in dbSNP, EVA, or iDOG databases but not observed in this study.

3.2. Sequence identity of canine P2X4

The canine P2X4 amino acid sequence was compared to six cloned P2X4 orthologues (Table 2). Canine P2X4 had the highest sequence identity (90%) to human P2X4 (Table 2) with the highest regions of sequence identity occurring around and within the transmembrane domains (Figure 1a). A single M348V substitution exists within the second transmembrane domain of canine P2X4, the only difference between the transmembrane domains of canine and human P2X4 receptors (Figure 1a). A stretch of 45 amino acids from P176 to A221, which contain a number of residues that form the ATP‐binding sites of P2X4 (Chataigneau, Lemoine, & Grutter, 2013), is also conserved between the mammalian species (Figure 1a). P2X4 sequences are relatively well conserved between canine and four other mammalian P2X4 orthologues (>85%), but this sequence identity drops below 70% (Xenopus) and 60% (zebrafish) when compared to taxa outside of the mammalian lineage (Table 2, Figure 1b).

TABLE 2.

Sequence identity between protein sequences of cloned P2X4 orthologues

	Sequence identity (%)
	Canine a	Human b	Bovine c	Mouse d	Rat e	Xenopus f	Zebrafish g
Canine	100	90	88.7	86.1	85.6	69.5	56.5
Human		100	87.4	86.9	87.4	69.7	58.3
Bovine			100	85.6	85.8	68.2	56.5
Mouse				100	95.6	67.7	57.8
Rat					100	68.2	59.1
Xenopus						100	56.6
Zebrafish							100

Note. Sequences were obtained from the NCBI protein database with the exception of the full‐length canine P2X4 sequence which was unavailable from NCBI and was retrieved from ENSEMBL. Sequences were aligned with Clustal O (v1.2.4), and sequence identity was determined using the per cent identity matrix tool.

^aENSCAFT00000013312.3.

^bNP_002551.2.

^cNP_001029221.1.

^dNP_035156.2.

^eNP_113782.1.

^fNP_001082067.1.

^gNP_705939.1.

FIGURE 1.

Multiple sequence alignment of known mammalian P2X4 receptors. (a) P2X4 sequences were aligned using Clustal O v1.2.4 (https://www.ebi.ac.uk/Tools/msa/clustalo/; last accessed November 2019). Asterisks (*) underneath the alignment indicate residues that are identical between all five orthologues, and blank spaces indicate where differences are observed. Sections highlighted grey indicate where canine and human residues are identical but differ compared to other non‐human orthologues, and sections highlighted blue indicate where canine and human residues differ. Predicted transmembrane domains based on the Expasy TMpred prediction program (https://www.expasy.org/; last accessed November 2019) are highlighted yellow. The amino acid residue involved in P2X4 orthologue sensitivity to the P2X4 receptor antagonist BX430 (identified by Ase et al., 2019) is highlighted green. Residues where two of the above occur are indicated by dual‐highlighted split boxes. Amino acids involved in ATP binding are underlined. Dog (dP2X4; P2RX4‐201 ENSCAFT00000013312.3), human (hP2X4; NP_002551.2), bovine (bP2X4; NP_001029221.1), mouse (mP2X4; NP_035156.2), and rat (rP2X4; NP_113782.1) P2X4 protein sequence entries were obtained from ENSEMBL or NCBI (last accessed November 2019). (b) Phylogenetic tree calculated from Clustal O multiple sequence alignment of above P2X4 sequences, as well as zebrafish P2X4 (NP_705939.1) and Xenopus P2X4 (NP_001082067.1). Tree was constructed using the average distance method in Jalview 2.11.0

3.3. Expression and localisation of cloned canine P2X4 receptors

Human and rodent P2X4 receptors localise primarily to lysosomal compartments (Huang et al., 2014; Stokes & Surprenant, 2009). To determine the subcellular localisation of canine P2X4 receptors, 1321N1 cells were transfected with canine or human P2X4‐EmGFP constructs and analysed by confocal microscopy. 1321N1 cells transfected with canine or human P2X4‐EmGFP revealed a distinct vesicular localisation and to a lesser extent cell‐surface expression, both of which were similar for canine and human P2X4 receptors, visualised either by the EmGFP‐tag (Figures 2a and S1) or an anti‐P2X4 antibody (Figure S1). In contrast, 1321N1 cells transfected with the EmGFP empty vector did not display a distinct vesicular localisation (cytoplasmic/nuclear distribution) nor were immunolabeled with the anti‐P2X4 antibody at levels above the background fluorescence. Furthermore, both canine and human P2X4 receptors appeared to colocalise with lysosomes (Pearson's coefficient = 0.735 and 0.610, respectively) revealed by anti‐LAMP1 antibody labelling (Figure 2a). Western blot analysis of transfected 1321N1 cells confirmed the presence of a ~80‐kDa band corresponding to the glycosylated P2X4 α‐subunit (~55 kDa) plus EmGFP‐tag (27 kDa) in transfected, but not wild type or EmGFP‐transfected 1321N1 cells (Figure 2b), validating the use of this antibody for future studies of canine and human P2X4 receptors. Similar results were observed for HEK293 cells transfected with canine or human P2X4‐EmGFP constructs (Figure S2A,B).

FIGURE 2.

Expression of canine and human P2X4 receptors in 1321N1 cells. (a) 1321N1 cells, transfected with canine or human P2X4‐EmGFP, or EmGFP vector alone, were fixed, permeabilised, and stained with anti‐LAMP1 antibody to label lysosomes. Cells were imaged by confocal microscopy, and colocalisation was analysed using ImageJ. Inset squares were digitally enlarged three times the original to examine P2X4 localisation in vesicular compartments. Scale bar = 20 μm. (b) Proteins of whole cell lysates of nontransfected 1321N1 cells (Nil) and 1321N1 transfected with canine (cP2X4) or human (hP2X4) P2X4‐EmGFP or EmGFP vector alone (EmGFP) were examined by western blotting using an anti‐P2X4 or anti‐actin antibody prior to imaging. (a, b) Data are representative of three independent experiments

3.4. Pharmacological characterisation of canine P2X4 receptor agonists

To pharmacologically characterise canine P2X4 receptors, the Ca²⁺ permeability properties of P2X4 channels were used to measure increases in intracellular Ca²⁺ bound to the ratiometric, fluorescent dye, Fura‐2. Incubation with 10‐μM ATP or 300‐μM BzATP induced rapid Ca²⁺ responses which peaked within 10–30 s of application, followed by a slowly declining, sustained period, which was not observed upon addition of ECS vehicle alone (Figure 3a,b). Furthermore, these nucleotides evoked peak Ca²⁺ responses only in 1321N1 cells transfected with canine or human P2X4‐EmGFP, but not with the EmGFP expression vector alone (Figure 3c).

FIGURE 3.

Ca²⁺ response agonist profile of canine and human P2X4 receptors in 1321N1 cells. (a–g) 1321N1 cells, transfected with canine or human P2X4‐EmGFP or EmGFP (control) plasmid DNA, were loaded with Fura‐2 and incubated in the absence or presence of each nucleotide in ECS (as indicated). (a, b) Representative ΔCa²⁺ traces for each nucleotide at respective maximal concentration. Solid and dashed lines indicate Ca²⁺ traces in presence or absence of nucleotide, respectively. (c) Peak ΔCa²⁺ response to maximal concentrations of ATP (10 μM) and BzATP (300 μM). (d–g) Nucleotide‐induced ΔCa²⁺ responses are presented as per cent of maximum response to 10‐μM ATP, with data fit to the Hill equation to produce concentration–response curves. (f, g) ADP or ATP were preincubated with hexokinase (4.5 U·ml⁻¹) for 1 hr at 37°C prior to addition. Red open and closed circles represent 10‐μM ATP in the presence or absence of hexokinase, respectively. (a–h) Data shown are mean ± SEM from five independent experiments (or eight for ATP concentration–response curve). *P < .05, significantly different from nucleotide in absence of hexokinase; two‐way ANOVA with Bonferroni post hoc test following a D'Agostino–Pearson test for normality

ATP was the most potent agonist of canine P2X4 receptors (Table 3), evoking a concentration‐dependent Ca²⁺ response with maximal activity near 10 μM (Figure 3d). ATP produced a similar pharmacological profile in 1321N1 cells expressing human P2X4 receptors (Figure 3e). BzATP also induced concentration‐dependent Ca²⁺ responses in 1321N1 cells expressing canine and human P2X4 receptors, although with reduced potency towards the canine receptors (Figure 3d,e). Peak Ca²⁺ responses achieved with 300‐μM BzATP at canine and human P2X4 receptors were 41 ± 7% and 92 ± 14% of 10‐μM ATP, respectively. Unlike ATP, there was a significant difference in EC₅₀ for BzATP between canine and human receptors (Table 3). Unexpectedly, ADP induced concentration‐dependent Ca²⁺ responses in cells expressing canine and human P2X4 receptors (Figure 3f,g); however, previous studies have reported that commercial stocks of ADP can contain small amounts of ATP (Mahaut‐Smith, Ennion, Rolf, & Evans, 2000). Therefore, ADP (and ATP as a positive control) was preincubated with hexokinase to remove contaminating ATP which resulted in significantly reduced ADP‐induced Ca²⁺ responses in canine and human P2X4‐transfected 1321N1 cells (Figure 3f,g). These experiments support previous reports of ATP contamination in ADP stocks and suggest that this contaminant ATP may account for the unexpected Ca²⁺ response to ADP.

TABLE 3.

Pharmacological activity of compounds mediating changes in intracellular Ca²⁺ in 1321N1 cells expressing canine or human P2X4 receptors

	Canine P2X4		Human P2X4
	pEC50 or pIC50	Hill coefficient	pEC50 or pIC50	Hill coefficient
Agonists (pEC50) a
ATP	6.59 ± 0.13	1.89	6.72 ± 0.06	1.59
BzATP	3.44 ± 0.13* , † (41%)	1.72	5.04 ± 0.26 † (92%)	1.68
ADP b	NR (<10%)	—	NR (<10%)	—
Modulators (pEC50) c
IVM	6.98 ± 0.05 † (111%)	1.60	6.91 ± 0.13 † (137%)	1.85
ATP	6.52 ± 0.12	2.09	6.69 ± 0.07	1.66
Antagonists (pIC50) d
BX 430	5.11 ± 0.06* (43%)	1.65	5.71 ± 0.08 (29%)	1.29
5‐BDBD	5.24 ± 0.24 (53%)	0.89	5.28 ± 0.36 (54%)	0.99
Duloxetine	4.82 ± 0.20 (13%)	1.00	4.77 ± 0.18 (13%)	1.00
Paroxetine	4.88 ± 0.13* (28%)	0.76	4.11 ± 0.20 (37%)	0.84
TNP‐ATP	5.09 ± 0.27 (18%)	1.00	5.36 ± 0.24 (8%)	1.00

Note. Data are mean ± SEM from five independent experiments.

Abbreviation: NR, negligible response (pEC₅₀ not calculated).

^aValues in parentheses indicate the per cent of max response compared to 10‐μM ATP.

^bADP in the presence of hexokinase to remove contaminating ATP.

^cpEC₅₀ of ATP in the absence or presence of 3‐μM ivermectin ; values in parentheses indicate the per cent of response compared to 10‐μM ATP.

^dpIC₅₀ of antagonists calculated using the approximate pEC₈₀ of ATP (0.75 μM); values in parentheses indicate the per cent of response in the presence of 100‐μM antagonist compared to max ATP in absence of antagonist.

^* P < .05, significantly different from the pEC₅₀ or pIC₅₀ of respective compound at human P2X4 receptors;

^† P < .05, significantly different from ATP alone for corresponding P2X4 receptor (non‐parametric Mann–Whitney U test for agonists and ivermectin and parametric Student's t test for antagonists).

To extend these findings, P2X4‐mediated Ca²⁺ response agonist profiles were also characterised in HEK293 cells, a commonly used cell line for studying heterologously expressed P2X receptors (Jiang & Roger, 2020). Canine and human P2X4‐transfected HEK293 cells displayed similar Ca²⁺ response agonist profiles (Figure 4a–d) to those observed for transfected 1321N1 cells (ATP pEC₅₀ canine 6.09 ± 0.04, human 6.47 ± 0.03; BzATP pEC₅₀ canine <3, human 4.14 ± 0.06). Furthermore, electrophysiological characterisation of canine and human P2X4‐mediated responses by automated patch clamp recordings in HEK293 cells was consistent with our Ca²⁺ response data. Robust inward currents were activated with a charge density of 21 ± 7 and 34 ± 16 pC·pF⁻¹ at 100‐μM ATP for canine and human P2X4 receptors, respectively (Figure 4e,f). This resulted in similar ATP response profiles in HEK293 cells transfected with canine (pEC₅₀ of 5.02 ± 0.11) and human P2X4 receptors (pEC₅₀ of 5.17 ± 0.10; Figure 4e–g). No inward currents were elicited by 300‐μM ATP in patch clamped mock EmGFP‐transfected HEK293 cells (Figure S3A), yet ATP‐induced Ca²⁺ responses were observed in EmGFP‐transfected HEK293 cells (Figure S3B) consistent with the activity of endogenous metabotropic https://www.guidetopharmacology.org/GRAC/ObjectDisplayForward?objectId=324 receptors reported previously in these cells (Schachter, Sromek, Nicholas, & Harden, 1997). Further investigation of HEK293 cells transfected with EmGFP vector alone in the absence or presence of 2‐mM EGTA (to chelate extracellular Ca²⁺) or 10‐μM AR‐C118925 (a P2Y2‐selective inhibitor) showed that the observed ATP‐induced Ca²⁺ response was partially reduced (53 ± 6.4%) in the presence of EGTA and almost completely reduced (90.6 ± 5.4%) upon AR‐C118925 treatment (Figure S3C). Therefore, further characterisation of Ca²⁺ responses was largely limited to P2X4‐expressing 1321N1 cells.

FIGURE 4.

Agonist profile of canine and human P2X4 receptors in HEK293 cells. (a–g) HEK293 cells, transfected with canine or human P2X4‐EmGFP plasmid DNA. (a–d) Cells were loaded with Fura‐2 and incubated in the absence or presence of each nucleotide (as indicated). (a, b) Representative ΔCa²⁺ traces for each nucleotide at respective maximal concentration. Solid and dashed lines indicate Ca²⁺ traces in presence or absence of nucleotide in ECS, respectively. (c, d) Nucleotide‐induced ΔCa²⁺ responses are presented as per cent of maximum response to 10‐μM ATP. Data shown are mean ± SEM from five independent experiments. (e, f) Representative automated whole‐cell patch clamp current traces of canine or human P2X4 in absence or presence of increasing concentrations of ATP (as indicated by the bars above the current traces) in standard external solution. (g) ATP‐induced inward currents are presented as charge density (pC·pF⁻¹) normalised to current at 1‐mM ATP, with data fit to the Hill equation to produce concentration–response curves. Data shown are mean ± SEM from eight (canine P2X4) or six (human P2X4) independent experiments

3.5. Ivermectin potentiates ATP‐induced Ca²⁺ flux through canine P2X4 receptors

Ivermectin is an allosteric modulator of human and rodent P2X4 receptors (Khakh, Proctor, Dunwiddie, Labarca, & Lester, 1999; Priel & Silberberg, 2004), but its effect on canine P2X4 receptors is unknown. In order to determine if ivermectin is also a positive allosteric modulator of these receptors, cells expressing canine or human P2X4 receptors were incubated in the presence or absence of 3‐μM ivermectin (Priel & Silberberg, 2004), prior to activation with increasing concentrations of ATP. Ivermectin significantly potentiated peak Ca²⁺ responses observed at 0.1‐ and 0.3‐μM ATP in 1321N1 cells expressing canine (Figure 5a,b) or human (Figure 5c,d) P2X4 receptors, resulting in 2.9‐ and 1.7‐fold decreases in the ATP EC₅₀ for each orthologue, respectively (Table 3). Consistent with these results, ATP‐induced inward currents recorded in HEK293 cells expressing canine P2X4 receptors were potentiated by 1‐μM ivermectin as demonstrated by a 2.0‐ to 2.5‐fold increase in charge density following increased peak inward currents and marked decreases in desensitisation kinetics (Figure S4A,B). The current enhancement observed during our automated patch clamp characterisation of canine P2X4‐mediated currents supports previous whole‐cell patch clamp recordings of human P2X4 receptors obtained in the presence of ivermectin (Priel & Silberberg, 2004).

FIGURE 5.

Modulation of ATP‐induced Ca²⁺ responses by ivermectin (IVM) on canine and human P2X4 receptors in 1321N1 cells. (a–d) 1321N1 cells, transfected with canine or human P2X4‐EmGFP, were loaded with Fura‐2 and preincubated in the absence (vehicle, 0.1% DMSO in ECS) or presence of 3‐μM ivermectin for 20 min. Cells were incubated in the absence or presence of increasing concentrations of ATP. (a, c) Representative ΔCa²⁺ traces for 0.3‐μM ATP, where solid and dashed lines indicate Ca²⁺ traces in absence or presence of ivermectin, respectively. (b, d) ATP‐induced ΔCa²⁺ responses are presented as per cent of maximum response to 10‐μM ATP in the absence of ivermectin, with data fit to the Hill equation to produce concentration–response curves. (a–d) Data shown are mean ± SEM from five independent experiments. *P < .05, significantly different from respective ATP concentration in absence of ivermectin; two‐way ANOVA with Bonferroni post hoc test following a D'Agostino–Pearson test for normality

3.6. Characterisation of canine P2X4 receptor antagonists

A number of human and rodent P2X4 receptor antagonists have been identified, but no data have been published to date regarding their effect on canine P2X4 receptors. To address this, Ca²⁺ responses in 1321N1 cells transfected with canine P2X4 receptors were assessed using two P2X4‐selective antagonists (BX430 and 5‐BDBD), as well as three nonselective antagonists of human or rodent P2X4 receptors (duloxetine, paroxetine, and TNP‐ATP; Stokes et al., 2017). At 0.75‐μM ATP (corresponding to the approximate EC₈₀), all five antagonists demonstrated concentration‐dependent inhibition of both peak and sustained ATP‐induced Ca²⁺ responses of canine and human P2X4 receptors (Figure 6a–j). Moreover, with the exception of BX430 and paroxetine, each compound displayed similar efficacy against canine and human receptors (Figure 6k–o, Table 3). The P2X4‐selective antagonist BX430 was less active against canine P2X4 receptors at concentrations greater than 3 μM resulting in a significant reduction in potency, compared with that for human P2X4 receptors (Figure 6k, Table 3). In contrast, the non‐selective inhibitor paroxetine displayed significantly greater potency against canine P2X4 when compared to human P2X4 receptors (Figure 6n, Table 3). However, unlike BX430, higher concentrations of paroxetine appear to have comparable inhibitory effects at canine and human P2X4 receptors.

FIGURE 6.

Ca²⁺ response antagonist profiles of canine and human P2X4 receptors in 1321N1 cells. (a–o) 1321N1 cells, transfected with canine or human P2X4‐EmGFP, were loaded with Fura‐2 and incubated in the absence (ECS alone or 1% DMSO in ECS) or presence of increasing concentrations of BX430 (1% DMSO), 5‐BDBD (1% DMSO), duloxetine (ECS), paroxetine (1% DMSO), or TNP‐ATP (ECS) for 20 min and then exposed to 0.75‐μM ATP (approximate to the EC₈₀ of ATP). Representative ΔCa²⁺ traces in absence or presence of antagonists at (a–e) canine and (f–j) human P2X4 receptors. (k–o) ATP‐induced ΔCa²⁺ responses are presented as per cent of maximum response to 10‐μM ATP in the absence of antagonists, with data fit to the Hill equation to produce concentration–response curves. Data points indicate the mean ± SEM from five independent experiments

Given the similarity in efficacy of the P2X4‐selective antagonist 5‐BDBD between canine and human P2X4 receptors (Figure 6l), and its increasing use in studies of P2X4 receptor activity, the mechanism of inhibition of 5‐BDBD was further assessed against maximal concentrations of P2X4 receptor agonists. 1321N1 cells transfected with canine or human P2X4 receptors were preincubated in the presence or absence of 30‐μM 5‐BDBD and then activated with 10‐μM ATP or 300‐μM BzATP. However, despite previously observing inhibition of ATP‐induced Ca²⁺ responses with 0.75‐μM ATP (~EC₈₀, Figure 6l), 5‐BDBD failed to reduce the Ca²⁺ responses of canine or human P2X4 receptors, evoked by saturating concentrations of 10‐μM ATP (Figure 7a). In contrast, 5‐BDBD caused a significant reduction in Ca²⁺ responses evoked by 300‐μM BzATP in cells expressing canine or human P2X4 receptors (81 ± 8% and 48 ± 7% reduction, respectively; Figure 7b). Taken together, the data presented in Figures 6k and 7a are consistent with 5‐BDBD being a more potent inhibitor of P2X4 receptors at sub‐saturating concentrations of ATP, consistent with a competitive mode of inhibition. Accordingly, 300‐μM BzATP‐induced responses of P2X4 receptors were effectively blocked by 5‐BDBD, suggesting that BzATP at this concentration does not saturate P2X4 receptors from either species.

FIGURE 7.

Inhibitory effect of 5‐BDBD at maximal P2X4 receptor agonist concentrations. (a, b) 1321N1 cells, transfected with canine or human P2X4‐EmGFP, were loaded with Fura‐2, preincubated in the absence (0.3% DMSO in ECS) or presence of 30‐μM 5‐BDBD for 20 min, and then incubated with maximal or near maximal concentrations of (a) ATP or (b) BzATP, respectively. Data were plotted as a percentage of the maximal response to 10‐μM ATP in absence of antagonist. Data presented are the means ± SEM from six independent experiments. *P < .05, significantly different from nucleotide alone; parametric Student's t test

4. DISCUSSION

The current study described for the first time, the pharmacology of canine P2X4 receptors and indirectly supports the concept that dogs express functional P2X4 receptors. Canine P2X4 receptors displayed similar in vitro pharmacology to that of human P2X4 receptors, with two notable differences: canine P2X4 receptors were less sensitive to activation by BzATP and to inhibition by BX430. Our work revealed that canine P2X4 receptors, like the human receptors, largely reside within intracellular compartments that are immunoreactive to LAMP1, consistent with lysosomal localisation. Concurrently, we also show that the protein coding region of the P2RX4 gene is highly conserved amongst dogs.

It was determined here that ATP and BzATP are agonists and partial agonists of canine P2X4 receptors, respectively. Under our experimental setting, the EC₅₀ for ATP was similar for both canine and human P2X4 receptors, consistent with previously published EC₅₀ values for human, mouse, and rat P2X4 receptors (Abdelrahman et al., 2017; Garcia‐Guzman, Soto, Gomez‐Hernandez, Lund, & Stuhmer, 1997; Jones et al., 2000; Layhadi, Turner, Crossman, & Fountain, 2018; Soto et al., 1996). Given the high sequence identity between canine and human P2X4 (90%), it was not unexpected to identify similar pharmacological profiles towards ATP. This is supported by studies demonstrating conservation of crucial ATP‐binding residues (underlined in Figure 1) across P2X receptor subtypes (Chataigneau et al., 2013; Jiang, Rassendren, Surprenant, & North, 2000; Roberts et al., 2008) and amongst human and rodent P2X4 receptors (Abdelrahman et al., 2017). The BzATP EC₅₀ reported here for canine P2X4 receptors is consistent with that reported for rat P2X4 (EC₅₀ > 100 μM), but not human or mouse P2X4 receptors (Abdelrahman et al., 2017). The data reported here also suggest that BzATP is a partial agonist of human P2X4 receptors, in agreement with previous studies (Abdelrahman et al., 2017; Bianchi et al., 1999; He, Gonzalez‐Iglesias, & Stojilkovic, 2003). The reasons for observed differences in BzATP sensitivity between species are yet to be fully understood but may be attributed to variations in BzATP‐sensitive residues between species leading to less favourable binding conformations (Browne, Jiang, & North, 2010; Pasqualetto, Brancale, & Young, 2018; Tvrdonova, Rokic, Stojilkovic, & Zemkova, 2014).

The current study demonstrated that ADP is not an agonist of canine P2X4 receptors as reported for human P2X4 receptors in which 100‐μM ADP failed to elicit significant responses (Garcia‐Guzman et al., 1997). However, subsequent studies report ADP acting as a partial agonist at P2X4 receptors (Bianchi et al., 1999; Carpenter et al., 1999; He et al., 2003). Such discrepancies may be due to the presence of ATP as a contaminant in commercially available ADP stocks (Mahaut‐Smith et al., 2000). Thus, consistent with previous studies (Micklewright, Layhadi, & Fountain, 2018), ADP was preincubated with hexokinase to remove any contaminating ATP, which revealed negligible ADP‐induced Ca²⁺ responses in 1321N1 cells expressing recombinant canine or human P2X4 receptors. Thus, as reported by Garcia‐Guzman et al. (1997) for the human receptors, ADP does not activate canine P2X4 receptors. Moreover, caution should be taken when attributing ADP effects on ATP‐responsive P2 receptors when using commercially obtained stocks.

The current study found ivermectin to be a positive modulator of canine P2X4 receptors. This commonly used anti‐parasitic drug resulted in significant increases in the efficacy of ATP at canine and human P2X4 receptors. This increase is consistent with previous reports that ivermectin positively modulates human P2X4 receptors (Priel & Silberberg, 2004). Moreover, the positive modulation of canine P2X4 receptors by ivermectin is supported by conservation of critical residues for ivermectin recognition and binding such as Trp46, Trp47, Val60, and Val357 (Jelinkova et al., 2006; Popova et al., 2013). The evidence presented here regarding the sensitivity of canine P2X4 receptors to ivermectin, coupled with the frequent use of ivermectin as a preventative treatment for heartworm in dogs, provides a strong platform for dogs as natural models of human disease. Recently, ivermectin has been proposed as a potential P2X4‐active drug for the treatment of demyelinating diseases like multiple sclerosis (Zabala et al., 2018); however, such diseases are seldom reported in domestic dogs (Vandevelde & Zurbriggen, 2005). Observations of canine health following lifelong treatment with ivermectin could provide valuable insight as to the effect of targeting P2X4 receptors for treatment or prevention of associated diseases such as multiple sclerosis in humans.

The current study has shown that three nonselective compounds, duloxetine, paroxetine, and TNP‐ATP, can inhibit canine P2X4 receptors. The selective 5‐HT reuptake inhibitor, paroxetine, and the 5‐HT‐noradrenaline reuptake inhibitor, duloxetine, are reported to non‐selectively inhibit human and rodent P2X4 receptors (Nagata et al., 2009; Yamashita et al., 2016), as well as to alleviate allodynia in models of chronic pain (Iyengar, Webster, Hemrick‐Luecke, Xu, & Simmons, 2004; Zarei, Sabetkasaei, & Moini Zanjani, 2014). Consistent with these previous studies, both compounds demonstrated effective inhibition of canine and human P2X4 receptors in our models, albeit at relatively high concentrations (>100 μM). Moreover, the broad‐spectrum P2X receptor antagonist TNP‐ATP blocked canine and human P2X4 receptors with similar potency, which agrees with previous work describing TNP‐ATP as an antagonist of human and rodent receptors (Abdelrahman et al., 2017; Nagata et al., 2009). These compounds lack selectivity for P2X4 receptors, and as such, their use for studying canine P2X4 receptors may be limited. However, given that duloxetine and paroxetine are approved human drugs, they may provide potential therapeutic benefits in dogs, in which P2X4 receptors may be involved.

This study also demonstrated that two selective inhibitors of P2X4 receptors, BX430 and 5‐BDBD, can inhibit the canine receptors. The data presented here identified the phenylurea BX430 as a moderately potent antagonist of canine P2X4 receptors, although with a 4‐fold higher IC₅₀ than for the human receptors. This is consistent with species differences reported with BX430 where this compound has displayed potent inhibition of human and bovine P2X4, moderate inhibition of zebrafish and Xenopus P2X4, and no inhibition of rodent P2X4 receptors (Ase, Honson, Zaghdane, Pfeifer, & Seguela, 2015; Ase, Therrien, & Seguela, 2019). Ase et al. (2019) have attributed this difference to a single P2X4 residue (Ile312 human numbering, highlighted green in Figure 1a) which forms a docking site with Asp88 and Tyr300 and plays a crucial role in determining sensitivity to BX430. The conservation of this docking site between P2X4 orthologues such as canine, human, and zebrafish and the apparent differences in sensitivity to BX430 may indicate a role for additional residues in mediating the inhibitory effects of BX430.

In contrast, the benzodiazepine derivative 5‐BDBD inhibited canine and human P2X4 receptors with similar potency but with limited effectiveness compared to other inhibitors. Consistent with these data, 5‐BDBD has been reported as a moderately potent inhibitor of human and rodent recombinant P2X4 receptors, with limited effectiveness at submicromolar concentrations (Abdelrahman et al., 2017; Ase et al., 2015; Balazs et al., 2013; Coddou, Sandoval, Hevia, & Stojilkovic, 2019). The data presented here are also in agreement with recent studies demonstrating inhibition of endogenous P2X4 receptor activity in human monocyte‐derived macrophages (Layhadi & Fountain, 2017) and human CD4⁺ T cells (Ledderose et al., 2018). Furthermore, the current study reported 5‐BDBD to be ineffective at inhibiting Ca²⁺ responses induced by saturating concentrations of ATP. This is consistent with reports describing 5‐BDBD as a competitive inhibitor of P2X4 receptors, assessed by radioligand binding assays (Balazs et al., 2013), suggesting that this compound may have an allosteric inhibitory mechanism (Abdelrahman et al., 2017). Nonetheless, the limited potency and efficacy of BX430 and 5‐BDBD highlight the need to identify more potent, selective P2X4 receptor antagonists for treating disease in humans and dogs.

The current study also demonstrated that canine P2X4, like human and rodent P2X4 receptors, is predominantly localised to intracellular compartments of cells, with limited expression at the plasma membrane. Consistent with what has previously been shown for human and rodent P2X4 (Huang et al., 2014; Stokes & Surprenant, 2009), canine P2X4 receptors colocalised with LAMP1‐labelled lysosomes. These data are further supported by observations that di‐leucine (L₂₂I₂₃) and tyrosine‐based (Y₃₇₂xxV and Y₃₇₈xxGL) endo‐lysosomal targeting motifs (Qureshi, Paramasivam, Yu, & Murrell‐Lagnado, 2007) are conserved in canine P2X4 receptors. Moreover, expression of functional canine P2X4 receptors on lysosomal compartments supports growing evidence for a physiological role of these receptors in lysosomal membrane trafficking and fusion (Murrell‐Lagnado & Frick, 2019). In addition, despite transfection with equal amounts of plasmid DNA, the Ca²⁺ response and current magnitude for human P2X4 receptors were consistently higher than those for the canine receptors . This difference is likely to reflect differences in transcription and translation rates of canine and human P2X4 receptors, or the expression of a canine protein in a human cell line. Notably, uncharacterised effects on ion channel physiology and/or pharmacology arising from attachment of EmGFP to the C‐terminal of the P2X4 protein cannot be discarded, despite representing a technique commonly utilised in functional characterisation (Sadovnick et al., 2017).

The current study determined that the coding sequence of P2RX4 is highly conserved amongst 101 dogs. The lack of genetic variation in canine P2RX4 contrasts that of canine P2RX7 (Sophocleous et al., 2019; Spildrejorde et al., 2014) which encodes four characterised missense variants. The difference between canine P2RX4 and P2RX7 parallels the frequencies of missense variants between human P2RX4 (Stokes et al., 2011) and P2RX7 (Stokes et al., 2010). Since the publication of the canine genome (Lindblad‐Toh et al., 2005), few missense P2RX4 variants have been reported in canine genome databases (dbSNP, EVA, and iDOG); however, none so far have been validated by publication or characterised in functional studies. Nonetheless, the reduced polymorphic variation in canine P2RX4 compared to human P2RX4 is consistent with current observations of genome‐wide loss in genetic variation due to domestication and selective breeding (Ostrander, Wayne, Freedman, & Davis, 2017). Currently, there are around 400 recognised domestic pedigrees worldwide (Ostrander et al., 2017). With such a diverse range of pedigrees worldwide, there is potential for regional or pedigree bias within the present study, with Labrador, Staffordshire, and Maltese associated pedigrees collectively forming over one third of the total cohort. Just over one quarter of the pedigrees in this cohort are represented by a single sample and, as such, the identification of P2RX4 variants may have been precluded, particularly if they exhibit a very low minor allele frequency or are more commonly associated with particular pedigrees.

The unique establishment of dogs as close human companions and the high level of sequence identity between the two species has resulted in the recognition of dogs as natural models of human ageing and disease (Sandor & Kubinyi, 2019). This, in part, can be attributed to breeding programs and implementation of veterinary medicine, resulting in aging canine populations, and with this, diseases that are prevalent in aging human populations such as chronic pain, inflammation, and neurodegenerative diseases. The P2X4 receptors have known roles in controlling inflammation, chronic pain, remyelination, and pulse pressure in humans and rodents (Bernier et al., 2018; Stokes et al., 2011; Su et al., 2019; Zabala et al., 2018). The current functional characterisation of canine P2X4 receptors suggests potential roles in canine physiology and disease. However, expression of P2X4 receptors in canine tissues is limited to P2X4 transcripts reported in brain and gastrointestinal tissue from dogs (Lee et al., 2005) and P2X4 protein detected in canine arteries (Delorey et al., 2012). Thus, future studies would benefit from utilising known patterns of P2X4 receptor expression in human and rodent tissues such as blood vessels, brain, gut, lungs, and immune cells (Burnstock & Knight, 2004), to aid in identifying canine tissues that express P2X4 receptors.

In conclusion, the P2X4 receptor is an important signalling molecule with potential for therapeutic targeting for the treatment of major health issues including chronic pain, nerve damage, and cardiovascular disease. The data presented here will aid the development of more potent and selective P2X4 receptor‐targeting compounds and will assist with preclinical validation of potential therapeutics for P2X4 receptor‐related diseases in both human and veterinary medicine.

AUTHOR CONTRIBUTIONS

R.A.S. designed and performed the majority of experiments, analysed all the data, and wrote the manuscript. R.K.F.‐U. designed, conducted, and analysed automated patch clamp experiments. R.S. conceived the study. R.S. and L.S. obtained funding for the study. T.B. generated the expression constructs. V.S., S.K., and L.B. performed additional gene sequencing. S.J.C., B.L.C., and A.S. provided canine blood samples and technical advice. S.K., L.B., and R.B. performed additional cell experiments. M.D. and L.S. provided technical advice. R.S. and L.O. supervised the project, designed experiments, reviewed the data, and co‐wrote the manuscript. All authors edited the manuscript.

CONFLICT OF INTEREST

The authors declare no conflict of interest.

DECLARATION OF TRANSPARENCY AND SCIENTIFIC RIGOUR

This Declaration acknowledges that this paper adheres to the principles for transparent reporting and scientific rigour of preclinical research as stated in the BJP guidelines for https://bpspubs.onlinelibrary.wiley.com/doi/full/10.1111/bph.14207, https://bpspubs.onlinelibrary.wiley.com/doi/full/10.1111/bph.14208, and https://bpspubs.onlinelibrary.wiley.com/doi/full/10.1111/bph.14206, and as recommended by funding agencies, publishers, and other organisations engaged with supporting research.

Supporting information

Table S1. Primer pairs used for amplification and sequencing of P2RX4 from canine genomic DNA, and sequencing of canine and human plasmid DNA

Table S2. Distribution of P2RX4 gene variants in a random sample of 101 dogs

Table S3. Synonymous P2RX4 variants reported on the iDOG online database.

Figure S1. Expression of canine and human P2X4 in 1321N1 cells. 1321N1 cells, transfected with canine or human P2X4‐EmGFP, or EmGFP vector alone, were fixed, permeabilised and stained with anti‐P2X4 antibody to co‐label P2X4‐EmGFP fusion protein. Cells were imaged by confocal microscopy. Inset squares were digitally enlarged three times the original to examine P2X4 localisation in vesicular compartments. Scale bar = 20 μm. Images are representative of five independent experiments.

Figure S2. Expression of canine and human P2X4 in HEK293 cells. (A) HEK293 cells, transfected with canine or human P2X4‐EmGFP, or EmGFP vector alone, were fixed, permeabilised and stained with anti‐LAMP1 antibody to label lysosomes. Cells were imaged by confocal microscopy. Inset squares were digitally enlarged three times the original to examine P2X4 localisation in vesicular compartments. Scale bar = 20 μm. (B) Proteins of whole cell lysates of non‐transfected HEK293 cells (Nil) and HEK293 transfected with canine (cP2X4) or human (hP2X4) P2X4‐EmGFP or EmGFP vector alone (EmGFP) were examined by western blotting using an anti‐P2X4 or anti‐actin antibody prior to imaging. Data are representative of five independent experiments.

Figure S3. Pharmacological characterisation of endogenous P2 receptor function in EmGFP‐transfected HEK293 cells. (A) Representative automated whole‐cell patch clamp current trace of EmGFP‐transfected HEK293 in presence of 300 μM ATP (as indicated by the bar above the current trace) in standard external solution. (B, C) HEK293 cells, transfected with EmGFP plasmid DNA alone, were loaded with Fura‐2 and incubated in the absence or presence of each nucleotide (as indicated). (B) Representative ΔCa²⁺ trace for each nucleotide at respective maximal concentration. Solid and dashed lines indicate Ca²⁺ traces in presence or absence of nucleotide in ECS, respectively. (B) To determine P2Y‐mediated Ca²⁺ flux, cells were incubated in the presence or absence of EGTA (2 mM) and the P2Y2‐selective inhibitor AR‐C118925 (10 μM) prior to activation with 10 μM ATP, presented as percent of max response to 10 μM ATP. Data shown are mean ± SEM from (A, B) five or (C) three independent experiments.

Figure S4. Modulation of ATP‐induced inward currents by IVM on canine P2X4 in HEK293 cells. (A, B) HEK293 cells, transfected with canine P2X4‐EmGFP, were resuspended in standard external solution and whole‐cell inward currents were measured by automated patch clamp. Cells were bathed in (A) standard external solution or (B) 1 μM IVM for 10 minutes and then recordings were taken in the absence or presence ATP (± IVM) as indicated by the bars above the current traces. (B) Data are representative traces from five independent experiments. Representative trace shown in (A) are the full 5‐second scale traces presented in Figure 4E for comparison of current amplitude and desensitisation in absence of IVM.

Sophocleous RA, Berg T, Finol‐Urdaneta RK, et al. Pharmacological and genetic characterisation of the canine P2X4 receptor. Br J Pharmacol. 2020;177:2812–2829. 10.1111/bph.15009