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J Biol Chem. Author manuscript; available in PMC 2006 April 12.
Published in final edited form as:
PMCID: PMC1435685
EMSID: UKMS5278

DISRUPTION OF LIPID RAFTS INHIBITS P2X1 RECEPTOR MEDIATED CURRENTS AND ARTERIAL VASOCONSTRICTION.

P2X1 receptors for ATP are ligand gated cation channels expressed on a range of smooth muscle preparations and blood platelets. The receptors appear to be clustered close to sympathetic nerve varicosities and mediate the underlying membrane potential changes and constriction following nerve stimulation in a range of arteries and resistance arterioles. In this study we have used discontinuous sucrose density gradients, Western blot analysis and cholesterol measurements to show that recombinant and smooth muscle (rat tail artery, vas deferens and bladder) P2X1 receptors are present in cholesterol rich lipid rafts and co-localize with the lipid raft markers flotillin-1 and -2. Lipid rafts are specialised lipid membrane microdomains involved in signalling and trafficking. To determine whether lipid raft association was essential for P2X1 receptor channel function we used the cholesterol depleting agent methyl-β-cyclodextrin (10 mM for 1 hour). This lead to a redistribution of the P2X1 receptor throughout the sucrose gradient and reduced P2X1 receptor mediated (α,β-methylene ATP 10 μM) currents in HEK293 cells by >90% and contractions of the rat tail artery by ~ 50%. However contractions evoked by potassium chloride (60 mM) were unaffected by methyl-β-cyclodextrin and the inactive analogue α-cyclodextrin had no effect on P2X1 receptor mediated currents or contractions. P2X1 receptors are subject to ongoing regulation by receptors and kinases and the present results suggest that lipid rafts are an essential component in the maintenance of these localised signalling domains and play an important role in P2X1 receptor mediated control of arteries.

Introduction

ATP released from nerves, following tissue damage or shear stress acts at P2 receptors to regulate cardiovascular function (1). P2 receptors are divided into ligand gated P2X receptor channels and G-protein coupled P2Y receptors. P2X receptor mediated vasoconstriction has been described in a range of arteries in the periphery (1) and in the brain (2). Sympathetic nerves co-store and co-release ATP and noradrenaline and the relative contribution of purinergic and noradrenergic mechanisms to vasoconstriction is dependent on the size of the vessel and the parameters of stimulation (3,4). The P2X receptor mediated component predominates in small diameter arteries (4,5) and in submucosal resistance arterioles P2X receptor activation is solely responsible for sympathetic neurogenic vasoconstriction with noradrenaline acting through pre-synaptic mechanisms to regulate transmitter release (6). In addition P2X receptor channels in smooth muscle are permeant to calcium (~10% of current flowing through the channel under physiological conditions (7,8)) and a substantial component of the calcium required for contraction enters by this route (4,9). P2X receptors therefore provide a mechanism for sympathetic nerve mediated regulation of vascular tone that is resistant to alpha adrenoceptor and calcium channel antagonists.

Seven P2X receptor subunits have been identified (P2X1–7) and these can assemble as homo- and heterotrimeric receptors with a range of properties (10). The characteristic features of artery smooth muscle P2X receptors; (i) sensitivity to the ATP analogues α,β-methylene and L-β,γ-methylene ATP and (ii) responses that desensitise during agonist application, are consistent with the expression of P2X1 receptor subunits. In arterial smooth muscle the P2X1 receptor is the dominant isoform and P2X receptor mediated responses are abolished in arteries from P2X1 receptor deficient mice (11,12). Studies with these P2X1 receptor deficient mice have established a role for these receptors in sympathetic nerve mediated vasoconstriction (11) and autoregulation of blood flow in the kidney (12,13). The level of P2X1 receptors can be regulated by congestive heart failure (14), cardiomyopathy (15) and shear stress (16). In addition it has been shown recently that P2X receptors are involved in sensitising responses following heart failure (17). P2X1 receptors are also expressed on blood cells, including platelets and P2X1 receptors contribute to platelet activation (18) and aggregation (1921) and that P2X1 receptor deficiency is protective against thromboembolism (22). Thus evidence is building that P2X1 receptors can play important roles in the cardiovascular system and regulation of blood flow.

In arteries P2X1 receptors do not appear to be randomly distributed throughout the plasma membrane as P2X1 receptor immunoreactivity appears in clusters (23). This clustering of receptors is supported by electrophysioliogical studies on dissociated artery muscle excised membrane patches; some patches had multiple P2X receptor channels whilst channel activity was absent in others (24). Recent studies also indicate that the P2X1 receptors can be regulated by phosphorylation of interacting proteins (25) suggesting that the P2X1 receptor exists in an organised signalling domain. One possible explanation for the clustering of P2X1 receptors could be the inclusion in membrane lipid rafts (26). Lipid rafts are rich in cholesterol and glycosphingolipids that results in liquid ordered microdomains within the liquid-disordered glycerophospholipid membrane bilayer (27,28). Recent evidence suggests that there is heterogeneity in lipid rafts and that a range of different domains can be separated based on differences in detergent solubility (for a review see (27)). A wide range of proteins, including many signalling molecules have been shown to be preferentially associated with rafts (29) including a range of ion channels (for a review see (30)). In this study we have shown that P2X1 receptors are concentrated on lipid rafts and that disruption of rafts reduces P2X1 receptor signalling in arteries.

Materials and methods

Reagents

Adenosine 5’-[α,β-methylene]triphosphate (α,β-meATP), cholesterol, filipin III, KCl, α-cyclodextrin (α-CD), β-cyclodextrin (β-CD), methyl-β-cyclodextrin (Mβ-CD), were obtained from Sigma (Sigma-Aldrich, Poole, UK).

Cell culture and transient transfection

Native human embryonic kidney 293 (HEK293) cells and HEK293 cells subcloned after transfection with the human wild-type P2X1 receptor (P2X1cl-1 cells), were maintained in culture as previously described (25). For some studies native HEK293 cells were transiently transfected with plasmid cDNA encoding either the rat wild-type P2X1 receptor or human wild-type P2X1 receptor tagged with EGFP at its carboxy terminus (P2X1-EGFP cells) using LipofectAMINE™ 2000 Reagent (Invitrogen)/Opti-MEM (Invitrogen, Paisley, UK). Control and β-CD treated cells (10 mM for 1h at 37ºC) were imaged live using a 60X oil immersion lens mounted on a Fluoview FV300 confocal microscope (excitation wavelength of 488 nm for EGFP and filters set to capture emission at wavelengths greater than 510 nm) (Olympus, Tokyo, Japan). Fluorescence was captured using Olympus Fluoviewver 4.2 software.

Rat tissue collection

Male Wistar rats (250–350 g) were killed by stunning and cervical dislocation. For membrane fractionation, tail artery, vas deferens and bladder were excised and processed immediately or frozen in liquid nitrogen for later use. For contraction experiments, tails arteries were kept at 4ºC in physiological saline solution (150 mM NaCl, 2.5 mM KCl, 10 mM HEPES, 2.5mM CaCl2, 1 mM MgCl2, pH to 7.3 with NaOH) for a maximum of 3 hours before use.

Membrane fractionation

Cells and rat tissues were fractionated using a detergent-free method adapted from (31,32). Two 80 cm2 tissue flasks of HEK293 P2X1cl-1 cells were washed 3 times with PBS and scraped into 2 ml of 500 mM sodium carbonate, pH 11 and left on ice for 20 min. Either seven tail arteries, 3 vas deferens or 3 bladders from rat were crushed in liquid nitrogen using a pestle and a mortar. When reduced to powder, the tissues were then homogenized into 2 ml of 500 mM Na2CO3, pH 11 using a tissue grinder and left on ice for 20 min. The lysed cells and rat tissues were then subjected to three 20-s bursts of sonication. Homogenates were brought to 45% sucrose by addition of an equal volume of 90% sucrose in MBS (25 mM MES and 150 mM NaCl, pH 6.5) and loaded in an ultracentrifuge tube. A discontinuous sucrose gradient was layered on top of the sample by placing 4 ml of 35 % sucrose prepared in MBS with 250 mM Na2CO3 then 4 ml of 5% sucrose (also in MBS/Na2CO3). The gradient was centrifuged at 39 000 rpm on a TH-641 rotor in a Sorvall OTD65B ultracentrifuge (Kendro Laboratory Products Plc, Bishop’s Stortford, UK) for 16h at 4ºC. In some studies HEK293 P2X1c1-1cells were lysed in 2 ml MBS containing either 0.1 % Triton-X-100 or 1 % Triton-X-100 and were homogenized using a tissue grinder and left for 20 min on ice. The homogenates were brought to 45 % sucrose by addition of an equal volume of 90 % sucrose in MBS and loaded in an ultracentrifuge tube. A discontinuous sucrose gradient (35 and 5 % sucrose in MBS, lacking Triton X-100) was formed above the homogenate and centrifuged as described above for the detergent-free condition. After centrifugation, 11 fractions of 1 ml were collected from the top to the bottom of each tube. Cholesterol measurements were assessed with the Amplex Red Cholesterol assay kit (Molecular Probes Europe BV, Leiden, The Netherlands). The cell volume or tissue quantities used for the sample preparations described above where each representative of one individual experiment. Each experimental conditions where repeated 3 times.

Western blots

The protein level of fraction 4 was used as a loading reference. To do so, the same volumes of samples (different fractions) were loaded for each gel after normalization for protein quantity in fraction 4 (0.04 μg for tail artery, vas deferens or bladder and 0.1 μg for HEK293 P2X1cl-1 cells). After separation of the samples on 10–12% SDS-PAGE gels and transfer onto nitrocellulose, the membrane was processed for staining with the primary antibody (anti-P2X1 receptor antibody (1:2000) (Almoner, Jerusalem, Israel) or anti-Caveolin1 (1:1000), anti-Caveolin-2 (1:250), anti-Flotillin-1 (1:250), anti-Flotillin-2 (1:5000) (BD Biosciences Europe, Erembodegem, Belgium). Protein bands were visualised using an ECL+plus kit and Hyperfilm MP (Amersham Bioscexperiment.

Surface cell expression and immunoprecipitations

Cell surface proteins were biotinylated as previously described (33) and washed with PBS. Cells from two 80cm2 tissue flasks were processed for membrane fractionation as described above and blotted with Immunopure streptavidin/horseradish peroxidase conjugated (0.5 μg/mL) (Pierce, Perbio Science, Tattenhall, UK). Cells contained in a 35 mm Ø Petri dish were processed for immunoprecipitation (33). The samples were resuspended in 15 μl of gel sample buffer before running on SDS-PAGE gels. In parallel an aliquot of total lysate (0.5 μg of total protein) was blotted with anti-p44/42 MAP kinase antibody (1:1000) (New England Biolabs, Hitchin, UK)). Each experimental condition was repeated 3 times.

Patch-clamp recordings

Conventional whole-cell and amphotericin permeabilised patch-clamp experiments were performed at a holding potential of 60 mV at room temperature (21ºC) as described previously (11,34). The agonist (α,β-meATP, 10 μM) was rapidly applied via a U-tube. When looking at the effect of cyclodextrins on the P2X1 receptor responses to α,β-meATP, the cells were incubated with the drugs at a concentration of 10 mM for 1h at 37ºC, for cholesterol repletion cells were subsequently incubated for 1h at 37ºC with 660 μg/ml cholesterol. Filipin (10 μM) was incubated with the cells for 30 min at 37ºC.

Contraction studies

Rat tail artery rings were mounted in a Mulvany myograph and perfused with physiological saline solution at 34ºC and vasoconstrictions to applied drugs measured as described previously (35). Reproducible responses to 10 μM α,β-meATP could be recorded when the agonist was added at 30 min intervals. The superfusate was stopped when the arteries were incubated in presence of cyclodextrin drugs (10 mM for 1h at 34ºC). In the control condition, the superfusate was also stopped for 1h. Before to test subsequent contraction responses to α,β-meATP and KCl (60 mM), the superfusate was reinitiated and the arteries were washed with physiological saline solution.

Data analysis

Data are expressed as mean±s.e.mean throughout and n=number of cells, number of arteries. Differences between means were determined by Student’s t-test and a P value of < 0.05 was considered statistically significant.

Results

P2X1 receptors are associated with lipid rafts

Membrane proteins can show a variable distribution relating to the lipid composition of the membrane, with a number of proteins being preferentially located in cholesterol rich lipid rafts. Stable expression of P2X1 receptors in HEK293 cells provides an ideal model system to investigate the role lipid rafts in receptor function. Cell lysates were extracted under detergent free conditions and ultracentrifuged on a discontinuous sucrose gradient (31). P2X1 receptor protein was predominantly found co-localised with the lipid raft markers flotillin-1 and flotillin-2 in the fractions 3 and 4 (36)(Figure 1a) and was absent from non-transfected cells (Figure 1d). Fractions 3 and 4 contain more than 75% of the total cellular cholesterol (and only ~ 10% of the total protein, Figure 1b) confirming the presence of the P2X1 receptor protein in the cellular enriched-cholesterol fractions. As the P2X1 receptor is a transmembrane protein one possibility was that the fractionation concentrates the plasma membrane in fractions 3 and 4 and this could account for the predominant localisation of the P2X1 receptors in these fractions and not lipid raft association. To test this we used membrane impermeant sulfo-NHS-LC-biotin to label surface proteins and determine their distribution on the gradient. Biotinylated proteins were distributed throughout the gradient demonstrating that the fractionation does not just concentrate membrane proteins. Moreover, the biotinylation process was specific for surface proteins as no trace of the cytosolic p44/42 MAP kinase was observed at the cell surface although present in the total lysate (Figure 1e). Taken together these data demonstrate that P2X1 receptors are localised to enriched-cholesterol lipid rafts.

Figure 1
Presence of the P2X1 receptor in the lipid raft-enriched fractions prepared from HEK-293 cells

P2X1 receptors remain in the lipid raft fraction following activation and desensitisation

Activation of some receptors, for example the β2-adrenoceptor leads to movement out of the lipid rafts and a reduction in signalling (37,38). P2X1 receptors desensitise rapidly following agonist stimulation, and require several minutes for recovery, raising the possibility that the recovery process is dependent on movement between lipid rafts and the rest of bulk membrane. Activation of the P2X1 receptor with α,β-meATP (10 μM) for 10 min should fully desensitised the receptor, however it had no effect on the distribution of the receptor in the lipid raft fractions (Figure 1f) indicating that movement out of the lipid rafts is not associated with desensitisation of the P2X1 receptor channel.

Lipid raft disruption regulates P2X1 receptor properties

Depletion of cellular cholesterol leads to dissociation of the lipid rafts. The cholesterol depleting agent Mβ-CD (10 mM for 1h) reduced by 51 ± 28 % the total cellular cholesterol content and induced a redistribution of the P2X1 receptor along the gradient (Figure 2a). Following cholesterol depletion, the P2X1 receptor was now detected readily in fractions 5 to 11. Mβ-CD is membrane impermeable and does not deplete cholesterol from intracellular membranes, this may in part account for the P2X1 receptor detected in fractions 3 and 4 that could correspond to receptors on intracellular membranes being trafficked.

Figure 2
Cholesterol depletion moves the P2X1 receptor out from the lipid raft fraction and inhibited agonist evoked currents but had no effect on the surface localisation of the receptor expressed in HEK293 cells.

In patch clamp studies, the ATP analogue α,β-meATP (10 μM a maximal concentration) evokes transient inward currents through P2X1 receptors from HEK293 P2X1cl-1 cells (Figure 2b). Depletion of cellular cholesterol with either Mβ-CD or β-CD (10 mM for 1h) reduced the amplitude of agonist evoked responses by >90 % with no effect on either the rise time or rate of decay of the response (Figure 2b). The reduced amplitude of response is unlikely to result from a decrease in agonist potency at the receptor, as suggested for the effects of cholesterol depletion on cyclic nucleotide gated channels (39), as there was no effect on the time-course of the response and a supramaximal concentration of ATP (1 mM, 1,000 fold greater than the EC50 concentration) was also reduced by >90 % (data not shown). Mβ-CD or β-CD treatment also has no effect on the recovery of P2X1 receptors from the desensitised state as reproducible responses to α,β-meATP were evoked at 5 minute intervals using the amphotericin permeabilised patch technique (as shown previously (34)) from control cells and following Mβ-CD or β-CD treatment (data not shown). The inactive cyclodextrin stereoisomer α-CD (10 mM for 1h)(40) had no effect on agonist-evoked responses (Figure 2b). Following Μβ–CD treatment responses to α,β-meATP (10 μM) were partially restored (~ 4 fold increase in amplitude compared to Μβ–CD, p<0.05) by cholesterol repletion (660 μg/ml for l hr)−3222 ± 549, −212 ± 130 and −818 ± 240 p A for control, Μβ–CD and cholesterol repletion respectively, n=<8). The reduction in current amplitude in response to Mβ-CD is unlikely to result from a decrease in the number of P2X1 receptor channels at the cell surface as the localisation of EGFP-tagged P2X1 receptors at the cell membrane (peak currents through these receptors are also reduced by ~85% by Μβ–CD treatment; −5224 ± 1063 and −887 ± 200 pA for control and Μβ–CD respectively, n>6) was unaffected by β-CD treatment (observation of 17 and 14 cells respectively for control and β-CD treatment)(Figure 2d).

Lipid rafts may also be disrupted by filipin that acts to form multimeric globular complexes with membrane cholesterol (41,42). Filipin (10 μM for 30 mins) reduced by >90% peak currents following P2X1 receptor activation with no effect on the time-course of the response (−5809 ± 790 and 119 ± 70 pA for control and filipin treatment respectively, n+12,5). Taken together our results show that lipid raft disruption in HEK293 cells significantly depressed P2X1 channel function.

What type of lipid rafts are P2X1 receptors associated with?

Lipid rafts are thought to be heterogeneous with a range of different types of domain dependent on the protein and lipid content of the rafts (27). The only sub-type of rafts that can be identified morphologically are caveolae. These are small plasma membrane invaginations associated with the expression of the caveolins (27). In this study caveolins –1 and –2 were below the limit of detection in HEK293 cell lines (data not shown, however caveolins were detected in arterial smooth muscle cells – see later) suggesting that P2X1 receptors are not associated with caveolae in lipid rafts.

The composition of lipid rafts appears to be dependent on the method of isolation and different types of lipid raft domain have been postulated based on differences in detergent solubility of individual raft proteins. Lipid rafts prepared by non-detergent methods are enriched in glycerophospholipids while in contrast detergent extraction with Triton X-100 leads to a depletion of these glycerophospholipids and in addition non-detergent methods lead to a greater retention of inner-leaflet-membrane lipids (for review see (27)). Thus comparing results from different isolation procedures may give an insight into which sub-population of lipid rafts the P2X1 receptor associates with. We therefore compared rafts isolated by non-detergent methods with extraction with 0.1 and 1% Triton X-100 (Figure 3). At 0.1% Triton X-100 P2X1 receptors were associated with the buoyant fraction, however in addition the P2X1 receptor was also significantly detected in bottom fractions. When 1% Triton X-100 was used the P2X1 receptor was detectable at low levels in the buoyant fractions and predominantly in bottom fractions, this is consistent with a previous report on Triton X-100 (1%) extracted P2X1 receptors (43). The Triton X-100 concentration dependent localisation of the P2X1 receptor was mirrored by the fractions in which the lipid raft marker flotillin-2 was found. This suggests that the association of P2X1 receptors in lipid rafts, like for other proteins, for example the T cell receptor in Jurkat cells (44)(and for a more general review see (27)), are sensitive to Triton X-100 extraction. This raises the possibility that the association of P2X1 receptors with lipid rafts may be regulated by glycerophospholipids or inner-leaflet-membrane lipids as these are depleted by Triton X-100 treatment.

Figure 3
P2X1 receptor and flotillin-2 association with lipid rafts in HEK293 cells is sensitive to the concentration of Triton X-100 detergent

P2X1 receptors in rafts in smooth muscle cells

We have shown that recombinant P2X1 receptors associated with lipid rafts and disruption of these leads to an inhibition of P2X1 receptor mediated currents. In this series of studies we determined whether native P2X1 receptors are expressed in lipid rafts in smooth muscle preparations and whether disruption of the rafts can regulate the functional properties of native P2X1 receptors. Rat tail arteries were lysed under detergent free conditions and ultracentrifuged on a discontinuous sucrose gradient. P2X1 receptor protein was identified in the fractions 4 and 5 of the gradient just as the lipid raft markers, flotillin-1 and flotillin-2. P2X1 receptor protein also co-localised with caveolae (specialised subtype of lipid rafts) proteins caveolin-1 and caveolin-2 (Figure 4a). The enriched-cholesterol fractions represented by fractions 3,4 and 5, contains more than 70% of the total cellular cholesterol (with a peak ~30% for fraction 4) but less than 10% of the total cellular protein (Figure 4b). Taken together, these results confirm that P2X1 receptor protein is present in the cellular enriched-cholesterol fractions of rat tail arteries. P2X1 receptor protein and lipid raft marker caveolin-1 were similarly distributed along rat vas deferens and bladder gradients in fractions 4 and 5 (Figure 4c). It is interesting to note that there were slight differences in the distribution of lipid rafts between HEK293 cells and smooth muscle cells; P2X1 receptor and lipid raft associated proteins were present in fractions 3 and 4 of HEK293 cells while in fractions 4 and 5 of smooth muscle cells. This localisation reflected the distribution of cholesterol (for comparison see figure 1b and and4b).4b). Since HEK293 cells and smooth muscles are distinct cell types, the membrane fractionation divergences are likely to be due to cell membrane composition differences.

Figure 4
Presence of the P2X1 receptor in the lipid raft-enriched fractions prepared from rat smooth muscles (rat tail artery, vas deferens and bladder)

Raft disruption reduces P2X1 receptor mediated arterial constriction

The treatment of rat tail arteries with the cholesterol depleting agent Mβ-CD (10 mM for 1h) disrupts lipid rafts and resulted in a redistribution of the P2X1 receptor along the gradient (Figure 5a) similar to its effects observed previously on HEK293 P2X1cl-1 cells. Tail artery constriction amplitudes to α,β-meATP (10 μM for 10 min) were dramatically reduced by half after Mβ-CD treatment (10 mM for 1h), while the inactive analogue α-CD (10 mM for 1h) had no effect (110.5±5.8%, 52.5±4.1% and 115.4±6.9% of the initial control response, respectively for control, Mβ-CD and α-CD, n=4–7 from 3 and 4 different animals) (Figure 5b, c). In contrast, vasoconstrictions induced by 60 mM KCl were unaffected either by treatment with Mβ-CD or α-CD (97.3±4.2%, 104.2±10.9% and 96.1±2.5% of the initial control response, respectively for control, Mβ-CD and α-CD, n=4–8 from 3 and 5 different animals) demonstrating that the cyclodextrins do not interfere with the ability of the smooth muscle to contract. Taken together these data suggest that P2X1 receptors are associated with lipid rafts in arteries and that disruption of these rafts inhibits responses through the channel.

Figure 5
Lipid raft disruption and inhibition of P2X receptor mediated contractions of the rat tail artery.

Discussion

The localisation of the P2X1 receptor to the buoyant cholesterol rich fractions and the co-localisation with lipid raft markers (flotillins 1 and 2, and in addition for the smooth muscle preparations caveolins 1 and 2), combined with the reduction of functional P2X1 receptor mediated responses following interference with lipid raft cholesterol, provides strong evidence that P2X1 receptors are present in lipid rafts in HEK293 cells as well as smooth muscles (arteries, vas deferens and bladder).

Lipid rafts are characterised as cholesterol and sphingolipid rich regions of membrane. Density fractionation of cholesterol rich lipid rafts has shown that there is heterogeneity in the rafts (45) and demonstrates that there are likely to be different populations of microdomains that vary in lipid and protein composition. The only clearly identifiable subtype of lipid rafts are the caveolae (46), these membrane invaginations are characterised by the presence of caveolins. Many proteins are associated with caveolae through interaction with these caveolins; for example the ATP-sensitive potassium channel pore forming subunit Kir6.1 co-immunoprecipitates with caveolin in arterial smooth muscle (47). We were unable to co-immunoprecipitate the P2X1 receptor with anti-caveolin antibodies (unpublished observations) suggesting that caveolin is not directly associated with the localisation of the P2X1 receptor to lipid rafts. This is further supported by our work in HEK293 cells where levels of caveolins 1&2 were below the limit of detection (data not shown). A similar lack of caveolins has been reported in other HEK293 cell lines and used to suggest that β-adrenoceptor raft localisation is not caveolin dependent (37). However caveolins have been described in some HEK293 cell lines (39) and it seems likely that their expression may vary between different sub-clones of these cells. Myristoylation and palmitoylation of proteins may be involved in associations with cavaeolae (48,49) however the lack of motifs for such forms of protein modification suggests that this is unlikely for the P2X1 receptor. Additional support that caveolin 1 is unlikely to be involved in P2X1 receptor function comes from studies with knockout mice; P2X1 receptor mice show male infertility due to a reduced contractile responses of the vas deferens (50), and in this study we show that disruption of lipid rafts with β-CD reduced P2X1 receptor currents by >90%, however caveolin 1 knockout mice show normal levels of fertility (51). Finally as P2X1 receptors are associated with rapid nerve mediated depolarisation of smooth muscle it seems counterintuitive that the receptor would be localised to membrane invaginations (caveolae) as the ATP would have to diffuse into the caveolae before activating the channel. It therefore seems likely that the P2X1 receptor is associated with non-caveolar lipid rafts.

Pike (27) has recently suggested a model where lipid rafts can be divided into three classes based on their sensitivity to extraction by a variety of methods (i) Triton X-100 and CHAPS resistant (ii) Brij96, Brij98 and non-detergent isolation resistant and (iii) lubrol, tween resistant fractions. The findings that the P2X1 receptor is found in the buoyant lipid raft fraction when non-detergent methods are used and is sensitive to the concentration of Triton X-100 used suggests that the P2X1 receptor is associated with class (ii) lipid rafts, however the motif/region of the receptor associated with raft localisation, either directly or through an interacting protein remains to be determined.

Cholesterol depletion with Mβ-CD resulted in a re-distribution of P2X1 receptors throughout the membrane fractions consistent with a disruption of the lipid rafts. Similar movements have been described for a range of ion channels and raft associated proteins e.g. (28,39,52). In addition Mβ-CD treatment or the use of filipin to sequester cholesterol in the membrane and disrupt the lipid rafts (41,42) reduced P2X1 receptor mediated responses. Recombinant human P2X1 receptor currents recorded in HEK293 cells were reduced by >90% (also for recombinant rat P2X1 receptors, control 799 ± 137 pA and 31 ± 7 following Mβ-CD treatment, p<0.001, n=10, 5) and contractions of arteries were reduced by ~50% (this difference most likely results from the reduced access of Mβ-CD in whole artery studies compared to patch clamp studies on isolated single cells). It is unlikely that these effects of Mβ-CD result from non-specific effects or a block of the ability of the muscle to contract as responses evoked by depolarisation with 60 mM KCl were unaffected by Mβ-CD treatment (this study) and Mβ-CD has previously been shown to have no effect on L-type calcium currents expressed in HEK293 cells (53). Similarly P2X1 receptor currents or contractions were unaffected by α-CD, the stereoisomer that has no effect on membrane cholesterol (40). In addition cholesterol repletion following Mβ-CD treatment resulted in a significant restoration of P2X1 receptor mediated responses. Taken together these data strongly suggest that the reduction in P2X1 receptor responsiveness is due to depletion of cholesterol and disruption of the lipid rafts. Similar reductions in function following lipid raft disruption have been described for a number of other ion channels for example voltage dependent potassium channel Kv2.1 (28) and cyclic nucleotide gated channels (39).

Lipid raft association of the P2X1 receptor may be a method for bringing the receptor into an organised signalling domain. We have previously shown that P2X1 receptors are basally phosphorylated, P2X1 receptor currents are reduced following treatment with the tyrosine kinase inhibitor genistein, and can be potentiated by kinase activation or stimulation of Gαq coupled receptors e.g. mGlur1α (54). Mβ-CD treatment dissociates proteins from rafts and inactivates signalling cascades (26), for example the Src family of tyrosine kinases are present in rafts (30). Therefore disruption of the localised signalling pathways could result in disruption of ongoing regulatory mechanisms and account for the decrease in P2X1 responses following cholesterol depletion.

The association of P2X1 receptors in lipid rafts may not only be important in maintaining the responsiveness of the receptor but may also be a mechanism to achieve clustering of the receptor. Immunohistochemical and single channel recording studies have indicated that the P2X1 receptor is not randomly distributed in the membrane but forms clusters (23,24) and that in smooth muscle preparations these clusters are often seen close to sympathetic nerve varicosities; the site of transmitter release. Thus lipid rafts may be a way of concentrating the P2X1 receptor at signalling hot-spots. Calcium imaging studies on the rat tail artery showed the initial transient rise in calcium was due to purinergic receptor stimulation and this was quickly followed by the noradrenergic component that oscillates and propagates as a wave through the vessel (55). Subsequent, higher resolution studies from vas deferens (56), mesenteric arteries (9,57), and bladder (58) showed discrete, localised, P2X receptor mediated calcium rises in the smooth muscle cells following nerve stimulation. The sympathetic nerves that innervate arteries and the vas deferens release the co-transmitters ATP and noradrenaline. It is interesting that nerve evoked P2X receptor, but not α-adrenoceptor mediated rises in calcium are recorded in response to short trains of stimulation from the vas deferens (56) and mesenteric arteries (9). A similar finding was reported in contractile studies on submucosal arterioles where following sympathetic nerve stimulation the vasoconstriction is mediated solely by P2X receptors, noradrenaline is released from the nerves but acts pre-junctionally through α2-adrenoceptors to regulate transmitter release (6). These data suggest that although sympathetic nerves release both ATP and noradrenaline under certain conditions P2X receptors appear to be preferentially activated. One possible reason for this could be the differential membrane localisation of P2X and α-adrenoceptors. This seems likely as in this study we have shown that the P2X1 receptor is localised in lipid rafts and in previous work it has been shown that α1-adrenoceptors are not present in the lipid raft fractions and noradrenaline mediated contractions are not affected by cholesterol depletion with Mβ-CD (59). This would provide a mechanism where P2X1 receptors in lipid rafts could be concentrated close to the site of transmitter release whilst α-adrenoceptors would be excluded from these domains and thus be less responsive and require longer periods of nerve stimulation to be activated (57).

In summary the present study shows for the first time that P2X1 receptors are associated with lipid rafts and that disruption of these rafts compromises P2X1 receptor responsiveness probably through interference with organised signalling microdomains. This has significant implications for cardiovascular function where P2X1 receptors have been shown to have important signalling roles, for example in the control of resistance artery vasoconstriction following sympathetic nerve stimulation (6,11), pressure induced autoregulation in the kidney (12) and platelet regulation (1822).

Acknowledgments

We thank the Wellcome Trust for support, Drs C. Dart and L.J. Sampson for discussions and help with raft protocols and Dr B.D.Grubb for help with the confocal microscopy.

References

1. Ralevic V, Burnstock G. Pharmacol Rev. 1998;50:413–492. [PubMed]
2. Lewis CJ, Ennion SJ, Evans RJ. J Physiol. 2000;527:315–324. [PubMed]
3. Evans RJ, Cunnane TC. Naunyn Schmiedeberg’s Arch Pharmacol. 1992;345:424–430. [PubMed]
4. Gitterman DP, Evans RJ. Br J Pharmacol. 2001;132:1201–1208. [PMC free article] [PubMed]
5. Ramme D, Regenold JT, Starke K, Busse R, Illes P. Naunyn Schmiedeberg’s Arch Pharmacol. 1987;336:267–273. [PubMed]
6. Evans RJ, Surprenant A. Br J Pharmacol. 1992;106:242–249. [PMC free article] [PubMed]
7. Benham CD, Tsien RW. Nature. 1987;328:275–278. [PubMed]
8. Benham CD. J Physiol. 1989;419:689–701. [PubMed]
9. Lamont C, Wier WG. Circ Res. 2002;91:454–456. [PubMed]
10. North RA. Physiol Rev. 2002;82:1013–1067. [PubMed]
11. Vial C, Evans RJ. Mol Pharmacol. 2002;62:1438–1445. [PubMed]
12. Inscho EW, Cook AK, Imig JD, Vial C, Evans RJ. J Clin Invest. 2003;112:1895–1905. [PMC free article] [PubMed]
13. Inscho EW, Cook AK, Imig JD, Vial C, Evans RJ. Acta Physiol Scand. 2004;181:445–453. [PubMed]
14. Malmsjo M, Bergdahl A, Moller S, Zhao XH, Sun XY, Hedner T, Edvinsson L, Erlinge D. Cardiovasc Res. 1999;43:219–227. [PubMed]
15. Berry DA, Barden JA, Balcar VJ, Keogh A, dos Remedios CG. Electrophoresis. 1999;20:2059–2064. [PubMed]
16. Wang L, Andersson M, Karlsson L, Watson MA, Cousens DJ, Jern S, Erlinge D. Arterioscler Thromb Vasc Biol. 2003;23:1370–1376. [PubMed]
17. Li J, Sinoway AN, Gao Z, Maile MD, Pu M, Sinoway LI. Circulation. 2004;110:3049–3054. [PubMed]
18. Vial C, Rolf MG, Mahaut-Smith MP, Evans RJ. Br J Pharmacol. 2002;135:363–372. [PMC free article] [PubMed]
19. Toth-Zsamboki E, Oury C, Cornelissen H, De Vos R, Vermylen J, Hoylaerts MF. J Biol Chem. 2003;278:46661–46667. [PubMed]
20. Oury C, Sticker E, Cornelissen H, De Vos R, Vermylen J, Hoylaerts MF. J Biol Chem. 2004;279:26266–26273. [PubMed]
21. Oury C, Kuijpers MJ, Toth-Zsamboki E, Bonnefoy A, Danloy S, Vreys I, Feijge MA, De Vos R, Vermylen J, Heemskerk JW, Hoylaerts MF. Blood. 2003;101:3969–3976. [PubMed]
22. Hechler B, Lenain N, Marchese P, Vial C, Heim V, Freund M, Cazenave JP, Cattaneo M, Ruggeri ZM, Evans RJ, Gachet C. J Exp Med. 2003;198:661–667. [PMC free article] [PubMed]
23. Hansen MA, Dutton JL, Balcar VJ, Barden JA, Bennett MR. J Auton Nerv Sys. 1999;75:147–155. [PubMed]
24. Lewis CJ, Davies NW, Evans RJ. Drug Dev Res. 2001;52:164–169.
25. Vial, C., Tobin, A. B., and Evans, R. J. (2004) Biochem J
26. Simons K, Toomre D. Nat Rev Mol Cell Biol. 2000;1:31–39. [PubMed]
27. Pike LJ. Biochem J. 2004;378:281–292. [PubMed]
28. Xia F, Gao X, Kwan E, Lam PP, Chan L, Sy K, Sheu L, Wheeler MB, Gaisano HY, Tsushima RG. J Biol Chem. 2004;279:24685–24691. [PubMed]
29. Foster LJ, De Hoog CL, Mann M. Proc Natl Acad Sci U S A. 2003;100:5813–5818. [PubMed]
30. Martens JR, O’Connell K, Tamkun M. Trends Pharmacol Sci. 2004;25:16–21. [PubMed]
31. Ostrom RS, Liu X, Head BP, Gregorian C, Seasholtz TM, Insel PA. Mol Pharmacol. 2002;62:983–992. [PubMed]
32. Ostrom RS. Mol Pharmacol. 2002;61:473–476. [PubMed]
33. Ennion SJ, Evans RJ. FEBS Lett. 2001;489:154–158. [PubMed]
34. Lewis CJ, Evans RJ. Br J Pharmacol. 2000;131:1659–1666. [PMC free article] [PubMed]
35. Lewis CJ, Gitterman DP, Schluter H, Evans RJ. Br J Pharmacol. 2000;129:124–130. [PMC free article] [PubMed]
36. Bickel PE, Scherer PE, Schnitzer JE, Oh P, Lisanti MP, Lodish HF. J Biol Chem. 1997;272:13793–13802. [PubMed]
37. Rybin VO, Xu X, Lisanti MP, Steinberg SF. J Biol Chem. 2000;275:41447–41457. [PubMed]
38. Ostrom RS, Gregorian C, Drenan RM, Xiang Y, Regan JW, Insel PA. J Biol Chem. 2001;276:42063–42069. [PubMed]
39. Brady JD, Rich TC, Le X, Stafford K, Fowler CJ, Lynch L, Karpen JW, Brown RL, Martens JR. Mol Pharmacol. 2004;65:503–511. [PubMed]
40. Grassme H, Jekle A, Riehle A, Schwarz H, Berger J, Sandhoff K, Kolesnick R, Gulbins E. J Biol Chem. 2001;276:20589–20596. [PubMed]
41. Reungpatthanaphong P, Marbeuf-Gueye C, Le Moyec L, Salerno M, Garnier-Suillerot A. J Bioenerg Biomembr. 2004;36:533–543. [PubMed]
42. Bang B, Gniadecki R, Gajkowska B. Exp Dermatol. 2005;14:266–272. [PubMed]
43. Vacca F, Amadio S, Sancesario G, Bernardi G, Volonte C. J Neurosci Res. 2004;76 :653–661. [PubMed]
44. Janes PW, Ley SC, Magee AI. J Cell Biol. 1999;147:447–461. [PMC free article] [PubMed]
45. Hering H, Lin CC, Sheng M. J Neurosci. 2003;23:3262–3271. [PubMed]
46. Lai EC. J Cell Biol. 2003;162:365–370. [PMC free article] [PubMed]
47. Sampson LJ, Hayabuchi Y, Standen NB, Dart C. Circ Res. 2004;95:1012–1018. [PubMed]
48. Lee H, Woodman SE, Engelman JA, Volonte D, Galbiati F, Kaufman HL, Lublin DM, Lisanti MP. J Biol Chem. 2001;276:35150–35158. [PubMed]
49. Galbiati F, Volonte D, Meani D, Milligan G, Lublin DM, Lisanti MP, Parenti M. J Biol Chem. 1999;274:5843–5850. [PubMed]
50. Mulryan K, Gitterman DP, Lewis CJ, Vial C, Leckie BJ, Cobb AL, Brown JE, Conley EC, Buell G, Pritchard CA, Evans RJ. Nature. 2000;403:86–89. [PubMed]
51. Drab M, Verkade P, Elger M, Kasper M, Lohn M, Lauterbach B, Menne J, Lindschau C, Mende F, Luft FC, Schedl A, Haller H, Kurzchalia TV. Science. 2001;293:2449–2452. [PubMed]
52. Martens JR, Navarro-Polanco R, Coppock EA, Nishiyama A, Parshley L, Grobaski TD, Tamkun MM. J Biol Chem. 2000;275:7443–7446. [PubMed]
53. Lohn M, Furstenau M, Sagach V, Elger M, Schulze W, Luft FC, Haller H, Gollasch M. Circ Res. 2000;87:1034–1039. [PubMed]
54. Vial C, Tobin AB, Evans RJ. Biochem J. 2004;382:101–110. [PubMed]
55. Iino M, Kasai H, Yamazawa T. Embo J. 1994;13:5026–5031. [PubMed]
56. Brain KL, Jackson VM, Trout SJ, Cunnane TC. J Physiol. 2002;541:849–862. [PubMed]
57. Lamont C, Vainorius E, Wier WG. J Physiol. 2003;549:801–808. [PubMed]
58. Heppner TJ, Bonev AD, Nelson MT. J Physiol. 2005;564:201–212. [PubMed]
59. Dreja K, Voldstedlund M, Vinten J, Tranum-Jensen J, Hellstrand P, Sward K. Arterioscler Thromb Vasc Biol. 2002;22:1267–1272. [PubMed]