P2X receptors are ATP-gated cation channels expressed in nerve, muscle, bone, glands, and the immune system. The seven family members display variable Ca2+ permeabilities that are amongst the highest of all ligand-gated channels (Egan and Khakh, 2004). We previously reported that polar residues regulate the Ca2+ permeability of the P2X2 receptor (Migita et al., 2001). Here, we test the hypothesis that the formal charge of acidic amino acids underlies the higher fractional Ca2+ currents (Pf%) of the rat and human P2X1 and P2X4 subtypes. We used patch-clamp photometry to measure the Pf% of HEK-293 cells transiently expressing a range of wild-type and genetically altered receptors. Lowering the pH of the extracellular solution reduced the higher Pf% of the P2X1 receptor but had no effect on the lower Pf% of the P2X2 receptor, suggesting that ionized side chains regulate the Ca2+ flux of some family members. Removing the fixed negative charges found at the extracellular ends of the transmembrane domains also reduced the higher Pf% of P2X1 and P2X4 receptors, and introducing these charges at homologous positions increased the lower Pf% of the P2X2 receptor. Taken together, the data suggest that COO side chains provide an electrostatic force that interacts with Ca2+ in the mouth of the pore. Surprisingly, the glutamate residue that is partly responsible for the higher Pf% of the P2X1 and P2X4 receptors is conserved in the P2X3 receptor that has the lowest Pf% of all family members. We found that neutralizing an upstream His45 increased Pf% of the P2X3 channel, suggesting that this positive charge masks the facilitation of Ca2+ flux by the neighboring Glu46. The data support the hypothesis that formal charges near the extracellular ends of transmembrane domains contribute to the high Ca2+ permeability and flux of some P2X receptors.

Introduction

A change in calcium concentration is an essential intracellular signal that initiates fundamental physiological processes such as secretion and contraction. To succeed in this role, the intracellular free Ca2+ concentration ([Ca2+]i) is tightly controlled, and nature has devised an impressive array of proteins that regulate Ca2+ transport across cell membranes (Berridge et al., 2003). Neurotransmitter receptors play an essential role in this process. Metabotropic receptors increase [Ca2+]i by modulating voltage-gated Ca2+ channels or emptying internal Ca2+ stores (Ross et al., 2005). Ionotropic receptors, like the ATP-gated P2X receptor, take a more direct approach. P2X receptors are ligand-gated cation channels that increase [Ca2+]i by forming a cation-permeable pore (Benham and Tsien, 1987). They are expressed in a diverse range of tissues and are particularly abundant in the nervous system (Khakh, 2001; Illes and Ribeiro, 2004; Khakh and North, 2006). Seven full-length subtypes (P2X1–7) are expressed in mammals, and most are capable of forming both homo- and heteromeric receptors (Torres et al., 1999; North, 2002). The percentage of the ATP-gated current attributed to Ca2+(Pf%) varies from ∼3 to 15% depending on the subunit composition of the oligomeric receptor and the species of origin (Egan and Khakh, 2004). It is large enough to stimulate transmitter release from neurons (Pankratov et al., 2006), hormone release from endocrine glands (Troadec et al., 1998), and contraction of vascular smooth muscle (Ramme et al., 1987; Gitterman and Evans, 2001) and vas deferens (Brain et al., 2002).

Identifying the functional domains responsible for regulating Ca2+ flux is complicated by the lack of a clear picture of channel architecture. Each receptor is an oligomer of three subunits, with individual subunits comprised of intracellular amino and carboxy termini linked by two transmembrane domains and an extracellular loop (North, 2002). Mutagenesis studies indicate a role for the pore-lining second transmembrane segment (TM2) in the cation permeability (Migita et al., 2001) and Ca2+ current (Egan and Khakh, 2004) of at least one family member, the P2X2 receptor. The first transmembrane segment (TM1) also lines the pore, and mutagenesis of this domain is known to alter permeability and gating (Samways et al., 2006).

We previously reported that ∼6% of the total current through the P2X2 receptor is carried by Ca2+ (Egan and Khakh, 2004), and that the ability of this receptor to select amongst cations involves a Ca2+-sensing domain made of three polar amino acids of TM2 (Migita et al., 2001). This domain is absent in the two P2X receptors (P2X1 and P2X4) that display higher Pf%s of 11–15%, suggesting that distinct loci underlie the divergent Ca2+ currents of different family members. The aim of the present study was to identify the site(s) responsible for the elevated Ca2+ current of the P2X1 and P2X4 receptors.

Materials And Methods

Molecular Biology and Cell Culture

We used wild-type rat P2X3, human P2X4 (hP2X4), human P2X5 (hP2X5), and zebrafish P2X4.1 (zP2X4.1) receptors, and C-terminal epitope-tagged rat P2X1 and P2X2 receptors (P2X1-EYMPME, P2X2-DYKDDDDK) that were made and expressed using conventional techniques. Addition of epitope did not change the EC50 for ATP, the kinetics of the ATP-gated response (Torres et al., 1999), or, most importantly, the contribution of Ca2+ flux to total current through the channel pore of either parent receptor (Table I). Point mutations were introduced with the Quikchange II site-directed mutagenesis kit (Stratagene) and verified by automated DNA sequencing (Retrogen, Inc.). Chimerae were made as previously described (Haines et al., 2001a). All constructs were coexpressed with eGFP in human embryonic kidney-293 cells (HEK-293 or HEK-293T cells; American Type Culture Collection) using Effectene (QIAGEN) or Lipofectamine (Invitrogen) according to the manufacturers protocols. Transfected cells were maintained in Dulbecco's modified Eagle's medium (Invitrogen) supplemented with 10% FBS (Invitrogen or HyClone), 2 mM glutamine, 50 U/ml penicillin G, and 50 μg/ml streptomycin (Invitrogen), and incubated for 24–48 h at 37°C in a humidified, 5% CO2 atmosphere. They were subsequently replated at low density onto poly-l-lysine–coated glass coverslips (Gold Seal; Becton Dickinson) the night before the experiment.

TABLE I

Pf% for Wild Type, Tagged, and Mutant P2X1, P2X2, P2X3, and P2X4 Receptors


Protein
[ATP] (μM)
pA/pF
Pf%
n
Δ from P2X1?
Δ from P2X2?
Δ from hP2X4?
P2X1 WT P2X1 77 ± 17 12.1 ± 0.8 – Yes Yes 
 P2X1-EE 152 ± 34 11.6 ± 0.6a 11 – Yes Yes 
 (pooled P2X1120 ± 20 11.8 ± 0.5 19 – Yes Yes 
P2X2 WT P2X2 30 314 ± 22b 5.7 ± 0.4c 18 Yes – Yes 
 P2X2-FLAG 302 ± 41 6.7 ± 0.2d 55 Yes – Yes 
Chimerae P2X2-TM1X1 329 ± 66 9.3 ± 0.6 Yes Yes Yes 
 P2X2-TM2X1 103 ± 23 2.6 ± 0.3 Yes Yes Yes 
 P2X2-TM1/2X1 10 39 ± 16 8.9 ± 0.4 Yes Yes Yes 
pH on WT P2X1 (pH 5) 10 111 ± 29 5.8 ± 0.8 Yes No Yes 
 P2X2 (pH 5) 257 ± 30 5.8 ± 0.8 Yes No Yes 
P2X1 mutants P2X1-E52Q 130 ± 20 8.4 ± 0.5 13 Yes No Yes 
 P2X1-E52Q (pH 5) 30 225 ± 65 6.0 ± 0.7e Yes No Yes 
 P2X1-D327S 182 ± 35 10.5 ± 0.5 11 No Yes Yes 
 P2X1-DM 10 82 ± 20 6.1 ± 0.6f 11 Yes No Yes 
 P2X1-DM (pH 5) 30 39 ± 7 5.8 ± 1.6 Yes No Yes 
 P2X1-Y51H 90 ± 28 6.3 ± 0.8 Yes No Yes 
 P2X1-Y51V 85 ± 23 13.2 ± 1.3 No Yes No 
P2X2 mutants P2X2-Q52E 185 ± 57 11.2 ± 0.8 No Yes Yes 
 P2X2-S326D 140 ± 68 11.0 ± 0.9 No Yes Yes 
 P2X2-DM 10 107 ± 15 13.4 ± 0.9 12 No Yes No 
 P2X2-DM (pH 5) 132 ± 25 6.6 ± 1.3g Yes No Yes 
P2X3 P3X3 69 ± 21 4.8 ± 0.3 Yes Yes Yes 
 P2X3-H45Y 132 ± 39 6.7 ± 0.5h Yes No Yes 
P2X4 hP2X4 10 154 ± 53 16.0 ± 1.0 11 Yes Yes – 
 hP2X4-E51Q 10 101 ± 33 14.6 ± 1.1 No Yes No 
 hP2X4-D332S 10 199 ± 60 13.0 ± 0.6 No Yes No 
 hP2X4-DM 30 111 ± 22 8.6 ± 0.6i Yes No Yes 
 zP2X4.1 1000 11 ± 3 7.3 ± 1.3 Yes No Yes 

Protein
[ATP] (μM)
pA/pF
Pf%
n
Δ from P2X1?
Δ from P2X2?
Δ from hP2X4?
P2X1 WT P2X1 77 ± 17 12.1 ± 0.8 – Yes Yes 
 P2X1-EE 152 ± 34 11.6 ± 0.6a 11 – Yes Yes 
 (pooled P2X1120 ± 20 11.8 ± 0.5 19 – Yes Yes 
P2X2 WT P2X2 30 314 ± 22b 5.7 ± 0.4c 18 Yes – Yes 
 P2X2-FLAG 302 ± 41 6.7 ± 0.2d 55 Yes – Yes 
Chimerae P2X2-TM1X1 329 ± 66 9.3 ± 0.6 Yes Yes Yes 
 P2X2-TM2X1 103 ± 23 2.6 ± 0.3 Yes Yes Yes 
 P2X2-TM1/2X1 10 39 ± 16 8.9 ± 0.4 Yes Yes Yes 
pH on WT P2X1 (pH 5) 10 111 ± 29 5.8 ± 0.8 Yes No Yes 
 P2X2 (pH 5) 257 ± 30 5.8 ± 0.8 Yes No Yes 
P2X1 mutants P2X1-E52Q 130 ± 20 8.4 ± 0.5 13 Yes No Yes 
 P2X1-E52Q (pH 5) 30 225 ± 65 6.0 ± 0.7e Yes No Yes 
 P2X1-D327S 182 ± 35 10.5 ± 0.5 11 No Yes Yes 
 P2X1-DM 10 82 ± 20 6.1 ± 0.6f 11 Yes No Yes 
 P2X1-DM (pH 5) 30 39 ± 7 5.8 ± 1.6 Yes No Yes 
 P2X1-Y51H 90 ± 28 6.3 ± 0.8 Yes No Yes 
 P2X1-Y51V 85 ± 23 13.2 ± 1.3 No Yes No 
P2X2 mutants P2X2-Q52E 185 ± 57 11.2 ± 0.8 No Yes Yes 
 P2X2-S326D 140 ± 68 11.0 ± 0.9 No Yes Yes 
 P2X2-DM 10 107 ± 15 13.4 ± 0.9 12 No Yes No 
 P2X2-DM (pH 5) 132 ± 25 6.6 ± 1.3g Yes No Yes 
P2X3 P3X3 69 ± 21 4.8 ± 0.3 Yes Yes Yes 
 P2X3-H45Y 132 ± 39 6.7 ± 0.5h Yes No Yes 
P2X4 hP2X4 10 154 ± 53 16.0 ± 1.0 11 Yes Yes – 
 hP2X4-E51Q 10 101 ± 33 14.6 ± 1.1 No Yes No 
 hP2X4-D332S 10 199 ± 60 13.0 ± 0.6 No Yes No 
 hP2X4-DM 30 111 ± 22 8.6 ± 0.6i Yes No Yes 
 zP2X4.1 1000 11 ± 3 7.3 ± 1.3 Yes No Yes 

P2X1-EE and P2X2-FLAG are the epitope-tagged receptors (P2X1-EYMPME and P2X2-DYKDDDDK, respectively) that served as templates for mutagenesis. The concentrations of agonist ([ATP]) used in the study were chosen to give measurable responses while avoiding dye saturation, and the pA/pFs are the average current densities measured at these concentrations. Wild-type (WT) P2X1 and epitope-tagged P2X1-EE receptors did not differ in Pf%s, and the pooled data were used as control in subsequent statistical evaluations. Although the Pf% of the tagged P2X2-FLAG receptor measured here is the same as that of the WT P2X2 receptor measured in our previous work reported elsewhere (Egan and Khakh, 2004), we did not pool these two datasets because they were obtained in separate studies performed at distinct locations (Cambridge, UK versus St. Louis, MO). Thus, all comparisons in the present study reflect differences from the P2X2-FLAG receptor. In the three rightmost columns, we compare the Pf%s of mutants and chimerae to the parent receptors. Data that are significantly different from P2X1, P2X2, and hP2X4 receptors are marked “Yes” and data that show no difference are marked “No.” All data were obtained at extracellular pH 7.4 except where noted.

a

Not different from the WT P2X1 receptor.

b

Unpublished data (n = 10); Li, personal communication.

c

Data (n = 18) from Egan and Khakh (2004).

d

Not different from the WT P2X2 receptor.

e

Significantly different from P2X2-E52Q (pH 7.4).

f

Significantly different from WT P2X1, P2X1-E52Q, and P2X1-S327S.

g

Significantly different from P2X2-DM (pH 7.4).

h

Significantly different from the WT P2X3.

i

Significantly different from wild-type hP2X4, hP2X4-E51Q, and hP2X4-S332S.

Patch-Clamp Photometry

Our use of the dye-overload method (Schneggenburger et al., 1993) is described in detail in Egan and Khakh (2004). In brief, we measured fractional calcium current (Pf%) by simultaneously measuring total membrane current and fluorescence in cells loaded with a high concentration (2 mM) of the calcium-sensitive dye, K5fura-2 (for examples, see Fig. 2 A). Cells on coverslips were placed in a recording chamber mounted on the stage of a TE3000 epifluorescence microscope (Nikon Instruments). ATP-gated current was recorded from single cells held at −60 mV using borosilicate glass recording electrodes (1B150F, World Precision Instruments), an Axopatch 200B amplifier (Molecular Devices), ITC-16 data acquisition hardware (Instrutech), IGOR Pro software (Wavemetrics), and a G4 computer (Apple Computers). Electrodes had open-tip resistances of 1–3 MΩ and contained an intracellular solution of the following composition (in mM): 140 CsCl, 10 tetraethylammonium Cl, 10 HEPES, 2 K5fura-2 (Molecular Probes/Invitrogen), 4.8 CsOH, pH 7.35. Light emitted from the 100-W xenon lamp was passed through a 380-nm bandpass filter and directed through the microscope objective (HMC 40X ELWD Plan Fluor; Modulation Optics, Inc.) into the recording chamber by reflection from the surface of a 400-nm dichroic long pass mirror. Light emitted by fura-2 was gathered by the objective, passed through the dichroic mirror and a 510-nm bandpass filter, and measured by a Model 714 Photomultiplier Detection System (Photo Technology International). We controlled the day-to-day variation in the sensitivity of the microscope/PMT by normalizing the fura-2 signal to a “bead unit” (BU). One BU equaled the average fluorescence of seven Carboxy Bright Blue 4.6-μm microspheres (Polysciences) measured one at a time on the morning of that day's experiment (Schneggenburger et al., 1993; Frings et al., 2000). Subsequent measurements of ATP-evoked changes in the fura-2 fluorescence of living cells were recorded as multiples of the daily BU. Cells were loaded with fura-2 by passive diffusion through the tip of the recording electrode. The time course of diffusion was measured in a subset of cells by monitoring the increase in intracellular fluorescence that results from fura-2 entry (Pusch and Neher, 1988). The time constant of loading was 144 ± 16 s (n = 30), and a near steady-state concentration of intracellular fura-2 was reached within ∼9–10 min of patch disruption. These measurements suggest that 10 min is the minimum time needed to equilibrate the intracellular compartment with the contents of the pipette, and we waited at least this long before acquiring data. The extracellular bath solution was (in mM) 140 NaCl, 2 CaCl2, 1 MgCl2, 10 glucose, and 10 HEPES, titrated to pH 7.4 with 4 NaOH. HEPES was replaced by MEPS in experiments performed at pH 5. ATP was applied for 0.2–2.0 s once every 2–3 min using triple-barreled theta glass and a Perfusion Fast-Step System SF-77 (Warner Instruments). For each construct and experimental algorithm, we used a concentration of ATP that evoked an easily measured decrease in fluorescence of fura-2 excited at 380 nm (ΔF380) but avoided the problem of dye saturation (Table I). As previously reported, we saw no effect of short applications of submaximal concentrations of ATP on the fura-2 fluorescence of mock-transfected cells using the methods described above (Egan and Khakh, 2004).

The Pf% was calculated as follows:

\[Pf\%{=}\frac{Q_{Ca}}{Q_{T}}{\ast}100.\]

QT is total charge and equal to the integral of the leak-subtracted ATP-gated transmembrane current. QCa is the part of QT carried by Ca2+, and is equal to ΔF380 divided by the calibration factor Fmax. Fmax was determined in a separate set of experiments as previously described (Egan and Khakh, 2004) and equaled 0.0185 ± 0.002 BU/nC (n = 12).

Ca2+ and Cl Permeability Measurements

We measured the relative permeabilities of Ca2+ and Cl to Cs+ (PCa/PCs and PCl/PCs, respectively) using a reversal potential-based method and the Goldman equations (Hille, 2004). Membrane current was recorded using an AxoPatch 200B amplifier, indifferent electrodes suspended in 3 M KCl agar bridges in contact with the bath solution, and the broken-patch configuration of the whole-cell voltage-clamp technique. The solution in the recording pipette was (in mM) 150 CsCl, 10 EGTA, and 10 HEPES, brought to pH 7.4 with CsOH. Open tip electrode resistances measured 0.7–2.0 MΩ. Giga-ohm seals were established in a standard bath solution of (in mM) 150 NaCl, 1 CaCl2, 1 MgCl2, 10 glucose, and 10 HEPES, pH 7.4 with NaOH, and subsequently switched to test solutions of different ionic compositions.

To measure PCa/PCs, the bath solution was first changed to one that contained predominately CsCl (in mM: 150 CsCl, 0.1 CaCl2, 1 MgCl2, 10 glucose, 10 HEPES), followed by one that contained predominately Ca2+ (in mM: 112 CaCl2, 1 MgCl2, 10 glucose, 10 HEPES). We included 0.1 mM Ca2+ in the CsCl solution to retard pore dilation because this process causes a time-dependent change in cation permeability that could alter PCa/PCs (Khakh et al., 1999; Chaumont and Khakh, 2006); in theory, addition of 0.1 mM Ca2+ to a solution of 150 mM CsCl solution is expected to have a negligible effect on the reversal potential of ATP-gated current (Lewis, 1979). We changed the membrane voltage of cells bathed in each solution from −80 to 60 mV at a constant rate (1.4 V/s) before and during applications of ATP, and measured the membrane voltage corresponding to the zero-current level (i.e., the Erev) from the leak-subtracted currents. We calculated PCa/PCs as

\[\frac{P_{Ca}}{P_{Cs}}{=}\frac{(aC_{s}{\ast}[Cs]_{i}){\ast}\mathrm{exp}({\Delta}E_{rev}{\ast}F/RT){\ast}(1{+}\mathrm{exp}({\Delta}E_{rev}{\ast}F/RT))}{4{\ast}aC_{a}{\ast}[Ca]_{o}},\]

where aCs and aCa are the activity coefficients of Cs+(0.75) and Ca2+(0.25), respectively, ΔErev is Erev,CaErev,Cs, T is temperature (295.15°K, equal to 22°C), and F and R are universal constants.

We used bath solutions containing different concentrations of CsCl to measure PCl/PCs. The first solution had the same concentrations (154 mM) of Cs+ and Cl as the pipette solution. We applied a voltage ramp to obtain Erev of the ATP-gated current and calculated the junctional offset as the deviation of the measured Erev from the expected zero current level of 0 mV. This was then subtracted from the Erev obtained in a “low” CsCl bath solution that contained (in mM) 30 CsCl, 0.1 CaCl2, 1 MgCl2, 210 sucrose, 10 glucose, and 10 HEPES. PCl/PCs was calculated as

\[\frac{P_{Cl}}{P_{Cs}}{=}\frac{1{-}{\{}[X]_{i}/[X]_{o}{\}}{\ast}\mathrm{exp}({\Delta}E_{rev}{\ast}F/RT)}{\mathrm{exp}({\Delta}E_{rev}{\ast}F/RT){-}{\{}[X]_{i}/[X]_{o}{\}}},\]

where [X]i/[X]o is the ratio of the intracellular and extracellular ion activities (Mitchell et al., 1997). Activity coefficients were determined by interpolation of the tabulated data of Robinson and Stokes (1970) and equaled 0.82 and 0.72 for 30 and 150 mM CsCl, respectively.

Data Analysis

All data are presented as the mean ± SEM for the number of experiments stated in the text. Significant differences amongst groups were determined using InStat (GraphPad Software) by one-way ANOVA with Tukey's post hoc, or Student's t test where appropriate. The P values of individual datasets are quoted in the text; values ≤0.05 were considered significant. We used the pooled data of wild-type P2X1 and P2X1-EE receptors as controls for comparisons to mutated receptors because we measured no difference in the Pf%s of these two groups (Table I).

Results

The percentage of ATP current carried by Ca2+ varies significantly amongst members of the P2X receptor family. In the present study, we first focused on the P2X1 and P2X2 receptors because they transduce appreciably different Ca2+ fluxes. We measured Pf%s of 11.8 ± 0.5 and 6.7 ± 0.2% for the rat P2X1 and P2X2 receptors, respectively (Table I). These values are significantly different (P < 0.001) from each other and in good agreement with previously published results (Rogers and Dani, 1995; Egan and Khakh, 2004). With this baseline established, we examined the effect of genetic manipulation of channel structure on Pf% to gain insight into the molecular basis of the variability in Ca2+ flux throughout the P2X family.

To What Extent Do Individual Transmembrane Segments Influence Ca2+ Flux?

Empirical data suggest that both TM1 and TM2 line the ion-conducting pore of the P2X2 receptor (Samways et al., 2006). Although no such data exist for the P2X1 receptor, ATP activates cationic currents through chimeric P2X1/P2X2 proteins with swapped transmembrane domains (Werner et al., 1996; Haines et al., 2001a), and therefore it is reasonable to assume that the pores of both channels share a roughly similar design. If so, then the disparity in Ca2+ flux amongst receptors might reflect subtype-specific differences in the primary structures of their pore-lining domains. To determine the influence of individual segments, we investigated three chimeric channels in which one (P2X2-TM1x1 and P2X2-TM2x1) or both (P2X2-TM1/2x1) of the transmembrane segments of the P2X2 channel were replaced by equivalent stretches of the P2X1 channel (Fig. 1, A and B). All three chimerae formed functional channels when expressed in HEK-293 cells (Fig. 1 C).

Figure 1.

P2X2 receptor chimerae. (A) Schematic representations of the receptor sequence for three P2X2 chimerae. Native sections of the background P2X2 receptor are labeled “X2” and sections substituted with corresponding parts of the P2X1 receptor are labeled “X1.” The chimerae contained amino acids Gly30-Lys53 and/or Asp327-Leu354 of the P2X1 receptor. (B) Sequence alignments comparing the putative transmembrane segments of the P2X1 (TM1, Gly30-Lys53; TM2, Asp327-Leu354), P2X2 (TM1, Gly30-Lys53, TM2, Ser326-Leu353), P2X3 (TM1, Gly24-Lys47; TM2, Asn317-Leu344), hP2X4 (TM1, Gly29-Lys52; TM2, Asp331-Leu358), and zP2X4 (TM1, Gly32-Lys55; TM2, Asn340-Leu367) receptors. The acidic amino acids that are the focus of this study (Glu52-P2X1, Glu51-P2X4, Asp327-P2X1, and Asp331-P2X4) are marked with bold red letters. The conserved aspartate (corresponding to P2X1-Asp350) found in all family members is marked with bold black letters. The polar residues of TM2 that regulate Ca2+ current through the P2X2 receptor (Migita et al., 2001) are marked with bold blue letters. Note that a polar residue occupies only one of the three corresponding positions of the P2X1 (Ser337, underlined) and hP2X4 (Ser350, underlined) receptors. The neighboring amino acids (His45 and Glu46) of the P2X3 receptor are shown in bold green letters. (C) Currents recorded in the presence of ATP for the P2X1, P2X2, and the three chimeric receptors described in the text. ATP was applied for 2 s (open horizontal bars) at concentrations of 10 μM for the P2X2-TM1/2X1 chimera and 3 μM for the others.

Figure 1.

P2X2 receptor chimerae. (A) Schematic representations of the receptor sequence for three P2X2 chimerae. Native sections of the background P2X2 receptor are labeled “X2” and sections substituted with corresponding parts of the P2X1 receptor are labeled “X1.” The chimerae contained amino acids Gly30-Lys53 and/or Asp327-Leu354 of the P2X1 receptor. (B) Sequence alignments comparing the putative transmembrane segments of the P2X1 (TM1, Gly30-Lys53; TM2, Asp327-Leu354), P2X2 (TM1, Gly30-Lys53, TM2, Ser326-Leu353), P2X3 (TM1, Gly24-Lys47; TM2, Asn317-Leu344), hP2X4 (TM1, Gly29-Lys52; TM2, Asp331-Leu358), and zP2X4 (TM1, Gly32-Lys55; TM2, Asn340-Leu367) receptors. The acidic amino acids that are the focus of this study (Glu52-P2X1, Glu51-P2X4, Asp327-P2X1, and Asp331-P2X4) are marked with bold red letters. The conserved aspartate (corresponding to P2X1-Asp350) found in all family members is marked with bold black letters. The polar residues of TM2 that regulate Ca2+ current through the P2X2 receptor (Migita et al., 2001) are marked with bold blue letters. Note that a polar residue occupies only one of the three corresponding positions of the P2X1 (Ser337, underlined) and hP2X4 (Ser350, underlined) receptors. The neighboring amino acids (His45 and Glu46) of the P2X3 receptor are shown in bold green letters. (C) Currents recorded in the presence of ATP for the P2X1, P2X2, and the three chimeric receptors described in the text. ATP was applied for 2 s (open horizontal bars) at concentrations of 10 μM for the P2X2-TM1/2X1 chimera and 3 μM for the others.

We found that swapping transmembrane segments had quantifiable effects on Pf% (Fig. 2). The chimeric channel containing the first transmembrane segment of the P2X1 channel in the P2X2 background (P2X2-TM1X1) had a Pf% equal to 9.3 ± 0.6%, which was significantly greater (P < 0.001) than that of the wild-type P2X2 channel. Substitution of both transmembrane segments (P2X2-TM1/2X1) had no greater effect (Pf% of 8.9 ± 0.5%) than replacing TM1 alone. In stark contrast, the chimeric P2X2 channel containing the TM2 of P2X1, (P2X2-TM2x1) exhibited a Pf% of 2.6 ± 0.3% that was significantly lower (P < 0.001) than both wild-type channels (Table I). These results suggest either that: (1) both transmembrane domains contribute to the regulation of Ca2+ current, each in their own way; or (2) construction of the chimerae produced nonspecific changes in the pore-forming domains that affect Pf%. To explore these possibilities, we used the less invasive methods described below.

Figure 2.

Pf% of chimerae. (A) Each panel shows a cartoon of the construct used to measure Pf%. The vertical open rectangles indicate transmembrane domains, and the sources of the different segments (P2X1 or P2X2) are marked with numbers. Next to the cartoons are current and fluorescence traces measured from cells expressing the indicated construct. The black traces are the ATP-evoked whole-cell currents (in amperes), the red traces are integrated current (in coulombs), and the gray traces are the change in F380 (in bead units). The duration of the ATP application is indicated by the horizontal open rectangles. (B) Here, and in Figs. 3 and 4, the sampling rate of the fluorescence signals are reduced by decimation to 10 Hz, converted to QCa (where QCa=F380/(Fmax*BU)), normalized to peak QT, and then multiplied by 100%. The amplitude of the steady-state value of each trace is equal to Pf%. (C) The bar graph shows the mean and standard error of the Pf% values. Values significantly different from the P2X1 and P2X2 receptors are marked with “a” and “b,” respectively.

Figure 2.

Pf% of chimerae. (A) Each panel shows a cartoon of the construct used to measure Pf%. The vertical open rectangles indicate transmembrane domains, and the sources of the different segments (P2X1 or P2X2) are marked with numbers. Next to the cartoons are current and fluorescence traces measured from cells expressing the indicated construct. The black traces are the ATP-evoked whole-cell currents (in amperes), the red traces are integrated current (in coulombs), and the gray traces are the change in F380 (in bead units). The duration of the ATP application is indicated by the horizontal open rectangles. (B) Here, and in Figs. 3 and 4, the sampling rate of the fluorescence signals are reduced by decimation to 10 Hz, converted to QCa (where QCa=F380/(Fmax*BU)), normalized to peak QT, and then multiplied by 100%. The amplitude of the steady-state value of each trace is equal to Pf%. (C) The bar graph shows the mean and standard error of the Pf% values. Values significantly different from the P2X1 and P2X2 receptors are marked with “a” and “b,” respectively.

The Ca2+ Flux of the P2X1 Receptor Is pH Sensitive

The considerable Ca2+ fluxes of many ligand-gated channels result from an interaction with negatively charged acidic amino acids in the mouth of the pore (Keramidas et al., 2004; Jensen et al., 2005). We looked for an effect of ionized side chains by measuring the Pf% of P2X1 and P2X2 receptors at a lower pH where a greater proportion of the COO side chains would be protonated and hence neutral (Hille et al., 1975; Green and Andersen, 1991) (Fig. 3 A). The Pf% of the wild-type P2X1 receptor at pH 5.0 was 5.8 ± 0.8%, a significant (P < 0.001) reduction when compared with that measured at pH 7.4. In contrast, the Pf% of the wild-type P2X2 receptor was unaffected by changing pH (Fig. 3 D). These data suggest that the electrostatic attraction of Ca2+ by one or more acidic amino acids is responsible for the high Pf%s of the P2X1 receptor, and that titratable carboxylates are unlikely to play a role in Ca2+ flux through the P2X2 receptor.

Figure 3.

The role of acidic residues in regulating the Ca2+ flux of P2X1 and P2X2 receptors. Panel A shows representative traces recorded for P2X1 and P2X2-DM receptors at physiological (7.4) and acidic pH (5.0). Panel B shows the effects of removing the fixed charge of the P2X1 receptor. Panel C shows the effects of adding fixed charge to the P2X2 receptor. (D) The bar graph presents mean Pf% data for all of the experiments. Values significantly different from the P2X1 and P2X2 receptors are marked with “a” and “b,” respectively.

Figure 3.

The role of acidic residues in regulating the Ca2+ flux of P2X1 and P2X2 receptors. Panel A shows representative traces recorded for P2X1 and P2X2-DM receptors at physiological (7.4) and acidic pH (5.0). Panel B shows the effects of removing the fixed charge of the P2X1 receptor. Panel C shows the effects of adding fixed charge to the P2X2 receptor. (D) The bar graph presents mean Pf% data for all of the experiments. Values significantly different from the P2X1 and P2X2 receptors are marked with “a” and “b,” respectively.

Acidic Residues Regulate the Ca2+ Flux of the P2X1 Receptor

Next, we used site-directed mutagenesis to locate the acidic amino acid(s) responsible for the high Ca2+ flux of the P2X1 channel. The data obtained using chimeric receptors suggested that the search should first center on the transmembrane domains. We discounted the conserved aspartate of TM2 because it is present in all family members (bold black residues of Fig. 1 B), and instead looked for differences in the primary sequences of the transmembrane domains of channels that show higher (P2X1, P2X4) and lower (P2X2) Ca2+ fluxes. P2X1 and hP2X4 channels have the highest Pf%s of all members of the extended family (Egan and Khakh, 2004). Both channels have acidic residues at homologous positions in TM1 (Glu52-P2X1 and Glu51-hP2X4) and TM2 (Asp327-P2X1 and Asp331-hP2X4) that are missing in the P2X2 channel (bold red residues of Fig. 1 B). We engineered P2X1 mutants in which one or both of these were replaced by the amino acids that occupy the equivalent positions in the P2X2 receptor (Gln52 and Ser326). Neutralizing fixed charge in TM1 (P2X1-E52Q) significantly (P < 0.001) reduced Pf% to 8.4 ± 0.4% (Fig. 3 B and Table I). Removing the fixed charge in TM2 (P2X1-D327S) had no significant effect (Pf% equals 10.5 ± 0.5%). However, neutralizing both charges (P2X1-DM) caused a greater reduction (6.1 ± 0.6%) than neutralizing the charge of either TM1 (P < 0.01) or TM2 (P < 0.001) alone, and the sum of the effects of the single mutations (42% reduction in Pf%) was roughly equal to the effect of the double mutation alone (48% reduction in Pf%). Further, lowering the pH to 5.0 significantly (P < 0.01) reduced the Pf% of P2X1-E52Q to 6.0 ± 0.7% but had no effect on P2X1-DM (5.8 ± 1.6%), suggesting that the resident charge at position 327 that is present in P2X1-E52Q but missing in P2X1-DM does affect Pf%. Taken together, these data argue that both Glu52 and Asp327 interact with Ca2+ in the mouth of the pore, although the relative contribution of each amino acid is different.

We also performed the converse experiment of placing carboxylates at appropriate sites in the P2X2 receptor. We generated three mutant P2X2 receptors in which one or both of Gln52 and Ser326 were replaced by glutamate or aspartate, respectively (Fig. 3, C and D). The Pf% for the single mutants P2X2-Q52E and P2X2-S326D were 11.2 ± 0.8 and 11.0 ± 0.9%, respectively. Both values are significantly greater (P < 0.001) than the wild-type P2X2 receptor and approximately equal to the wild-type P2X1 channel. Placing both mutations in a single construct (P2X2-DM) did not increase Pf% more than either mutation alone. Lowering the pH to 5.0 significantly (P < 0.001) reduced the Pf% of P2X2-DM to 6.6 ± 1.3%, (Fig. 3 A), suggesting that the increase in Pf% results from an electrostatic effect of the added charge and not an unintended change in protein structure.

The Pf% of the P2X4 Receptor Decreases when Fixed Charge Is Removed

Next, we looked at the effects of mutagenesis on the hP2X4 receptor because it shows the highest Pf% of all P2X receptors (Egan and Khakh, 2004) and, like the rat P2X1 receptor, has fixed negative charge at the outer edges of its transmembrane segments. The Pf% of the wild-type hP2X4 receptor was 16.0 ± 1.0% (Fig. 4). We found that neutralizing both of the fixed charges of the transmembrane segments by mutagenesis gave a receptor (hP2X4-DM) with a significantly (P < 0.001) reduced Pf% of 8.6 ± 0.6%. In contrast, removing a single charge had no effect. Specifically, the hP2X4-E51Q and hP2X4-D332S mutants had Pf%s of 14.6 ± 1.1 and 13.0 ± 0.6%, respectively, that were no different than the wild-type receptor. In this respect, the single charge deletion mutants of the hP2X4 receptors resemble the single charge addition mutants of the P2X2 receptor; the presence of one charge per subunit is enough to maintain a high Ca2+ flux through the pore.

Figure 4.

The role of acidic residues in regulating the Ca2+ flux of human and zebrafish P2X4 receptors. Panel A shows representative traces recorded for single and double mutants of hP2X4 receptors. Panel B presents mean Pf% data for all of the experiments. Values significantly different from the P2X2, hP2X4, and hP2X4-DM receptors are marked with “a,” “b,” and “c,” respectively.

Figure 4.

The role of acidic residues in regulating the Ca2+ flux of human and zebrafish P2X4 receptors. Panel A shows representative traces recorded for single and double mutants of hP2X4 receptors. Panel B presents mean Pf% data for all of the experiments. Values significantly different from the P2X2, hP2X4, and hP2X4-DM receptors are marked with “a,” “b,” and “c,” respectively.

As a final test, we measured the Pf% of the wild-type zebrafish P2X4.1 receptor (zP2X4.1) (Kucenas et al., 2003). It lacks the two relevant formal charges found in hP2X4 receptors (Fig. 1 B) and therefore naturally replicates the charge reduction of P2X4-DM. In keeping with our hypothesis, the Pf% of the zP2X4.1 receptor at 7.3 ± 1.3% was significantly smaller than the hP2X4 receptor and not different from the rat P2X2 and hP2X4-DM receptors (Fig. 4 B).

Neutralizing Fixed Charge Alters PCa/PCs but Not PCl/PCs

A caveat to using patch-clamp photometry to draw conclusions about the Ca2+ flux of cation channels is that it does not account for the possibility that some mutations may render the channels anion permeable. With the exception of the human and chick P2X5 receptors, most wild-type P2X receptors are cation selective and therefore impermeable to anion (Egan et al., 2006). However, it is possible that some mutations increase Cl permeability, and under the conditions of our experiments (equimolar [Cl] on either side of the membrane and a holding potential of −60 mV), such an effect would lead to an apparent inward current caused by efflux of Cl, a larger QT, and a reduction in Pf% that does not reflect a change in Ca2+ flux per se. To differentiate a change in Ca2+ permeability from a change in Cl permeability, we acquired reversal potential data from ATP-gated currents obtained from cells bathed in a range of extracellular solutions (see Materials and methods).

Switching the bath solution from one that contained 150 mM CsCl to one that contained 112 mM CaCl2 caused a rightward shift in the reversal potential of the ATP-gated current of the wild-type P2X1 and hP2X4 receptors (Fig. 5 A) that translated to PCa/PCs values of 3.6 ± 0.2 and 4.6 ± 0.4, respectively (Table II). The shifts were smaller for P2X1-DM and hP2X4-DM receptors that lack the relevant formal charges, and the resulting PCa/PCss were significantly (P < 0.001) lower than the wild-type templates at 2.6 ± 0.1 and 2.8 ± 01, respectively. They were not significantly different from that of the P2X2 receptor that measured 2.9 ± 0.1 (Table II). Again, these data support our contention that formal charge influences the Ca2+ dynamics of P2X1 and P2X4 receptors.

Figure 5.

Voltage–current curves measured in different concentrations of extracellular Ca2+ or Cl. (A) The raw data traces show ATP-gated currents evoked in solutions that contained either Cs+ (blue traces) or Ca2+ (red traces) as the dominant extracellular cation. The change in Erev measured after switching from the Cs+ to Ca2+ solution was smaller for the double mutant receptors (right traces) than it was for their wild-type templates (left traces), suggesting that eliminating formal charge decreased PCa/PCs. (B) The raw data traces show ATP-gated currents evoked in extracellular solutions that contained either 150 mM (blue traces) or 30 mM CsCl (red traces). For the wild-type P2X1 and hP2X4 receptors (top and middle left graphs), the change in Erev measured upon switching to the low CsCl solution was ∼30–33 mV as expected for a channel that showed a low Cl permeability. The same was true for the double mutant receptors that lacked the relevant formal charges of acidic amino acids (right traces), showing that mutagenesis did not change PCl/PCs. The change in Erev was smaller in hP2X5 receptors that show a significant Cl permeability (bottom left graph).

Figure 5.

Voltage–current curves measured in different concentrations of extracellular Ca2+ or Cl. (A) The raw data traces show ATP-gated currents evoked in solutions that contained either Cs+ (blue traces) or Ca2+ (red traces) as the dominant extracellular cation. The change in Erev measured after switching from the Cs+ to Ca2+ solution was smaller for the double mutant receptors (right traces) than it was for their wild-type templates (left traces), suggesting that eliminating formal charge decreased PCa/PCs. (B) The raw data traces show ATP-gated currents evoked in extracellular solutions that contained either 150 mM (blue traces) or 30 mM CsCl (red traces). For the wild-type P2X1 and hP2X4 receptors (top and middle left graphs), the change in Erev measured upon switching to the low CsCl solution was ∼30–33 mV as expected for a channel that showed a low Cl permeability. The same was true for the double mutant receptors that lacked the relevant formal charges of acidic amino acids (right traces), showing that mutagenesis did not change PCl/PCs. The change in Erev was smaller in hP2X5 receptors that show a significant Cl permeability (bottom left graph).

TABLE II

Relative Ca2+ and Cl Permeabilities Measured from Shifts in Erev

Protein
ΔErev,Ca
PCa/PCs
n
ΔErev,Cl
PCl/PCs
n
P2X1 9.3 ± 0.9 3.6 ± 0.2 −30.3 ± 1.1 0.09 ± 0.02 12 
P2X1-DM 3.9 ± 0.6 2.6 ± 0.1 −32.0 ± 1.2 0.07 ± 0.01 12 
P2X2 6.1 ± 0.4 2.9 ± 0.1  ND  
hP2X4 12.6 ± 1.6 4.6 ± 0.4 −29.7 ± 1.4 0.09 ± 0.02 
hP2X4-DM 4.5 ± 1.0 2.8 ± 0.1 −32.9 ± 1.1 0.06 ± 0.02 
hP2X5  ND  −5.8 ± 2.5 0.66 ± 0.12 
Protein
ΔErev,Ca
PCa/PCs
n
ΔErev,Cl
PCl/PCs
n
P2X1 9.3 ± 0.9 3.6 ± 0.2 −30.3 ± 1.1 0.09 ± 0.02 12 
P2X1-DM 3.9 ± 0.6 2.6 ± 0.1 −32.0 ± 1.2 0.07 ± 0.01 12 
P2X2 6.1 ± 0.4 2.9 ± 0.1  ND  
hP2X4 12.6 ± 1.6 4.6 ± 0.4 −29.7 ± 1.4 0.09 ± 0.02 
hP2X4-DM 4.5 ± 1.0 2.8 ± 0.1 −32.9 ± 1.1 0.06 ± 0.02 
hP2X5  ND  −5.8 ± 2.5 0.66 ± 0.12 

The control solution contained predominantly 150 mM CsCl, and the test solutions contained either 112 CaCl2 or 30 CsCl (see text for details). ΔErev,Ca = Erev,112 CaErev,150 CsCl. ΔErev,Cs = Erev,30 CsClErev,150,CsCl.

In contrast, Cl permeability was not affected by the mutagenesis (Fig. 5 B). We first measured the PCl/PCs of the hP2X5 receptor because it shows a significant Cl permeability (Bo et al., 2003). As expected, we found that switching to the 30 mM CsCl bath solution caused a ∼6-mV shift in Erev; the calculated PCl/PCs was 0.66 ± 0.12 (Fig. 5 B). We then measured the Cl permeability of wild-type and double mutant P2X1 and P2X4 receptors. Shifting the extracellular solution from one that contained 150 mM CsCl to one that contained 30 mM CsCl caused an approximately −30 mV shift in the ATP-gated current in all four constructs, demonstrating that both the wild-type and mutant receptors are largely impermeable to anions (Table II). These data support the hypothesis that the changes in Pf% that we measure after mutagenesis reflect specific effects on Ca2+ flux.

P2X3 Has Fixed Charge but a Relatively Low Pf%

The glutamate of TM1 is conserved in only one other P2X subtype as Glu46 of the P2X3 receptor. However, the P2X3 receptor exhibits the lowest Ca2+ flux of all family members (Egan and Khakh, 2004). Unlike P2X1 and P2X4 receptors, the P2X3 receptor contains a positively charged amino acid, His45, immediately upstream to the conserved Glu46 residue (green residues of Fig. 1 B). His45 may provide a countercharge in the pore that serves to partially shield the neighboring Glu46 or simply repel Ca2+, thus attenuating the ability of the carboxylate to attract Ca2+. To test this hypothesis, we constructed a mutant P2X3 receptor in which His45 was replaced by the tyrosine found at the homologous position of the P2X1 receptor. This produced a small but significant (P < 0.05) increase in Pf% to 6.7 ± 0.5% compared with the wild-type P2X3 control of 4.8 ± 0.3% (Table I).

We also generated mutant P2X1 receptors in which Tyr51 was replaced by either a basic histidine (P2X1-Y51H) or a neutral valine (P2X1-Y51V). We found that the Pf% of P2X1-Y51H was significantly reduced (6.3 ± 0.8%) compared with the wild-type P2X1 receptor. This decrease was unlikely to be caused by a gross disruption of channel structure because substituting neutral valine (the corresponding residue of P2X2) for Tyr51 had no effect on Pf% (Table I). Taken together, the data suggest that the identity of the amino acid just upstream of the conserved charges of P2X1 and P2X3 receptors helps to determine the magnitude of the Ca2+ flux through the channel pore.

Discussion

We identified acidic residues at the outer ends of the transmembrane domains that contribute to the high fractional Ca2+ current of ATP-gated P2X1 and P2X4 receptors. We found that acidification decreased the relatively large Ca2+ current of the P2X1 receptor but had no effect on the smaller Ca2+ current of the P2X2 receptor, suggesting that only the former uses acidic amino acids to discriminate amongst permeant cations. (We also looked at the effect of a lower pH on the hP2X4 receptor. The concentration–response curve of the P2X4 receptor shifts to the right at lower pHs, necessitating the use of a very high (>1 mM ATP) concentration of ATP to evoke a measurable ligand-gated current. At these high concentrations, applications of ATP evoked biphasic changes in intracellular calcium that did not follow the time course of the integrated current, making measurements of Pf% untenable.) The magnitude of the reduction is surprising when one considers the predicted degree of protonation of the COO side chains. We expected to measure a modest reduction in Pf% upon lowering the pH to 5.0, a value close to the pKa of glutamate and aspartate in aqueous solution, and where half of the carboxyl groups are protonated and thus unable to attract Ca2+ (Falke et al., 1994). Instead, we found that acidification produced a Pf% equal to that of mutant receptors lacking both formal charges (i.e., P2X1-DM). These data suggest that the pKas of Glu52 and Asp327 of the P2X1 receptor may be significantly higher than predicted from measurements of pure aqueous solutions of amino acids, an idea in keeping with published reports that the local microenvironment of a protein greatly influences the degree of ionization of its constituent amino acids (Fersht, 1985; Klockner et al., 1996; Seifert et al., 1999; Petsko and Ringe, 2004). Of particular interest is the positive charge provided by a conserved lysine (e.g., Lys53 of the P2X1 receptor) that sits immediately downstream of Glu52. Placing positive charge close to an acidic amino acid is enough to shift the latter's pKa to 7.0 or higher (Falke et al., 1994). We have not pursued a study of this lysine because it is fully conserved and therefore unlikely to play a role in determining the dissimilar Ca2+ fluxes measured across the family. However, its role in the absolute magnitude of the Ca2+ flux should be considered in future experiments.

The idea that individual P2X receptors use distinct domains for cation selection is supported by the marked changes in Pf% observed in P2X1/P2X2 chimerae. Replacing the TM1 of the less Ca2+-permeable P2X2 receptor with that of the more permeable P2X1 receptor significantly enhanced Ca2+ flux, as did addition of a single formal negative charge to the extracellular end of TM1 (P2X2-Q52E). These data are important for three reasons: first, they add weight to the proposition that TM1 lines the outer segment of the ion channel (Haines et al., 2001b); second, they show for the first time that TM1 makes a measurable contribution to cation selection; and third, they suggest that the divergent Ca2+ currents of the P2X1 and P2X2 subtypes result in part from differences in the primary sequences of their TM1s.

The case of TM2 is more complex. We found that adding fixed negative charge to TM2 of the P2X2 receptor increased Pf% only when Thr336, Thr339, and Ser340 are unperturbed (compare P2X2-S326D to P2X2-TM2X1), a finding that supports previous reports that the polar residues of TM2 play an important role in Ca2+ flux in this receptor subtype (Egan and Khakh, 2004). Two of these are replaced by neutral amino acids in the P2X2-TM2X1 chimera, thus depriving the parent P2X2 receptor of polar side chains that may provide the countercharge needed to partially dehydrate Ca2+ at a constriction within the pore (Migita et al., 2001). If so, then P2X1 receptors either use other means to overcome the energy barrier provided by the constriction or lack it entirely. An alternative explanation is that the decrease in Pf% in the P2X2-TM2X1 chimera reflects nonspecific changes in the topology of the pore brought about by swapping one TM2 with another. This latter possibility is difficult to ignore, and additional experiments are needed to firmly establish a role for polar amino acids in ion permeation in some P2X receptors.

Many classes of ligand-gated cation channels use the formal charge of acidic amino acids positioned near the pore to facilitate Ca2+ transport into cells (Keramidas et al., 2004; Jensen et al., 2005). P2X receptors have three sites that could serve a similar function. One of these is fully conserved in TM2 (see bold black letters in Fig. 1 B) and therefore unlikely to explain the differences in Ca2+ flux measured here. Two nonconserved sites are present in P2X1 and P2X4 receptors but not in the P2X2 receptor, and we found that removing both charges significantly decreased the Pf% and PCa/PCs without affecting PCl/PCs. Thus, our data are consistent with a model that uses the oxygen atoms of COO side chains to form an electrostatic ring that attracts cations. The attraction is stronger for Ca2+ than for Na+, and therefore could explain the high Ca2+ permeability and flux of P2X1 and P2X4 receptors. The location of the ring in the permeation pathway is unknown because the inner and outer limits of the pore are poorly defined (Egan et al., 2006). We favor the hypothesis that it lies in the extracellular vestibule for the following two reasons. First, such a model is consistent with those that explain the high Ca2+ fluxes of other ligand-gated cation channels (Barry and Lynch, 2005). Second, models that incorporate the formal charges of aspartate and glutamate side chains into Ca2+ binding sites within the pore often result in channels that show Ca2+ block of monovalent current (Voets et al., 2004), and no such effect is reported for either the P2X1 or P2X4 receptors.

We suggest the simple hypothesis that the formal charges of both TM1 and TM2 influence the flux of Ca2+ through P2X1 and hP2X4 channels in a nonadditive fashion. We found that adding charge to either transmembrane domain significantly enhanced Ca2+ flux of the P2X2 receptor, whereas removing the charge of either Glu51 or Asp332 had only a small effect on the Pf% of the hP2X4 receptor. The results are therefore consistent, for together they suggest that (1) a single charge is sufficient to cause the elevated Ca2+ permeability noted in the P2X2-DM and wild-type hP2X4 receptors; and (2) there is very little summation in terms of increasing Pf% when both charges are present in the outer parts of TM1 and TM2.

For the P2X1 receptor, it appears that Glu52 of TM1 might play a larger role than Asp327 of TM2. We found that removing Glu52 significantly reduced the Pf% of the P2X1 receptor, whereas removing Asp327 had no effect, as might be expected if Asp327 is innocuous. Although these findings seem to invalidate the dual-charge hypothesis discussed above, two experiments suggest that Asp327 does influence Ca2+ flux through the channel. First, subtracting both charges (i.e., P2X1-DM) has a significantly greater effect on the Pf% than removing either Glu52 or Asp327 alone. Second, acidification decreases the Pf% of P2X1-E52Q but has no effect on P2X1-DM, and the only difference between these two mutants is the presence of Asp327. It is difficult to explain why eliminating the glutamate of TM1 significantly decreases the Pf% of P2X1 but not P2X4 receptors, except to say that structural differences in TM2 may make the Asp327 of the P2X1 receptor either less accessible to Ca2+ or less charged than either the Asp332 of the hP2X4 receptor or the Asp326 of the P2X2-S326D mutant. P2X1, P2X2, and P2X4 receptors have remarkably different phenotypes and display unique agonist and antagonist profiles, rates of desensitization, Ca2+ permeabilities, and trafficking (North, 2002). If function follows structure, then structural differences between these subtypes are to be expected.

Fig. 6 shows cartoon representations of P2X1, P2X2 and P2X4 receptors that incorporate two different models of Ca2+ transport. We assume that each channel is comprised of three subunits (Nicke et al., 1998; Aschrafi et al., 2004; Barrera et al., 2005; Mio et al., 2005), and that each subunit contributes two transmembrane segments to the pore (North, 2002). In P2X1 and P2X4 receptors, Ca2+ current is enhanced by a ring of charge made of glutamate and aspartate residues positioned in the outer mouth of the pore. These residues could function to accumulate extracellular Ca2+ in the permeation pathway by a simple electrostatic attraction, or form a binding site that selects Ca2+ over monovalent cations (Hille, 2004; Jensen et al., 2005). As mentioned above, the latter hypothesis seems doubtful because, unlike P2X2 receptors, Ca2+ does not block monovalent cation current through P2X1 receptors as expected for a binding site model (Evans et al., 1996). The extracellular ring of charge is missing in the P2X2 receptor (Fig. 6, C and D). Instead, polar residues in the middle of the pore provide the countercharge needed to dehydrate Ca2+ in a narrow part of the pore (Migita et al., 2001). In this model, the narrow constriction provides an energy barrier that impedes current flow through the channel, and the relative conductance of cations is determined by their ability to momentarily “bind” to the polar side chains as they shed water. The presence of an intrapore binding site in P2X2 receptors is supported by two findings. First, ATP-gated currents saturate with increasing concentrations of extracellular Na+ (Ding and Sachs, 1999b). Second, monovalent current is blocked by low millimolar concentrations of extracellular Ca2+ (Nakazawa and Hess, 1993; Ding and Sachs, 1999a).

Figure 6.

Schematic representations of the ion permeable pores of trimeric P2X1, P2X2, and P2X4 receptors. Residues in TM1 are marked with black letters, and residues in TM2 are marked with red letters. The leftmost panels show side (A and E) and top (B and F) views of P2X1 and P2X4 receptors. The mouth of the pore is surrounded by an outer ring of six acidic amino acids (two from each of three subunits). The outer ring is missing in the P2X2 receptor (C and D). Also shown are the three polar residues within TM2 of the P2X2 receptor that are thought to form a polar ring at a narrow section of the channel pore. Only one of these sites is occupied by a polar amino acid in the P2X1 and P2X4 receptors. TM1 also contains a conserved tyrosine (Tyr43) that may be properly positioned to contribute to the polar ring.

Figure 6.

Schematic representations of the ion permeable pores of trimeric P2X1, P2X2, and P2X4 receptors. Residues in TM1 are marked with black letters, and residues in TM2 are marked with red letters. The leftmost panels show side (A and E) and top (B and F) views of P2X1 and P2X4 receptors. The mouth of the pore is surrounded by an outer ring of six acidic amino acids (two from each of three subunits). The outer ring is missing in the P2X2 receptor (C and D). Also shown are the three polar residues within TM2 of the P2X2 receptor that are thought to form a polar ring at a narrow section of the channel pore. Only one of these sites is occupied by a polar amino acid in the P2X1 and P2X4 receptors. TM1 also contains a conserved tyrosine (Tyr43) that may be properly positioned to contribute to the polar ring.

One question then remains: how well do these two mechanisms describe the Ca2+ current of other family members? Fixed charge is conserved in the highly Ca2+-permeable P2X1 and P2X4 receptors, but is also present in the less permeable P2X3 and P2X7 receptors. We have not investigated P2X7 receptors so far, but experiments with P2X3 receptors suggest again that each subunit uses a slightly different method to select amongst cations. We suggest that the Pf% of the P2X3 receptor is relatively low because of the presence of a unique His45 just upstream of Glu46. The positive charge it supplies might serve to partially neutralize the negative charge of Glu46 and hinder the electrostatic interaction with Ca2+. However, it is worth noting that neutralizing His45 did not elevate the Pf% of the P2X3 receptor to a value equal to that of P2X1 receptor, suggesting that additional factors contribute to the lower Pf% measured for P2X3 receptor.

Acknowledgments

We gratefully acknowledge the help and guidance of Rosalie Uchanski, Mark Voigt, and Zhiyuan Li, the comments of Stephen Baylor, Baljit Singh Khakh, Keisuke Migita, and Florentina Soto, and the technical assistance of Angela Lipka and Steven Harris. The hP2X4, zP2X4.1, and hP2X5 plasmids were gifts of Drs. Soto (University of Washington, Seattle, WA), Voigt (St. Louis University, St. Louis, MO), and R. Alan North (Manchester University, Manchester, UK), respectively.

This work was supported by the National Institutes of Health.

Olaf S. Andersen served as editor.

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Abbreviations used in this paper: BU, bead unit; HEK, human embryonic kidney; Pf%, fractional Ca2+ current; TM, transmembrane.