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The Journal of Neuroscience, April 1, 1998, 18(7):2350-2359
A Domain Contributing to the Ion Channel of ATP-Gated
P2X2 Receptors Identified by the Substituted Cysteine
Accessibility Method
Terrance M.
Egan,
William R.
Haines, and
Mark M.
Voigt
Department of Pharmacological and Physiological Sciences, St. Louis
University Health Sciences Center, St. Louis, Missouri 63104
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ABSTRACT |
P2X receptors are a family of ATP-gated ion channels thought to
have intracellular N and C termini and two transmembrane segments separating a large extracellular domain. We examined the involvement of
the second putative transmembrane domain (TM2) of the P2X2 subunit in ion conduction, using the substituted cysteine accessibility method (SCAM). This method tests the ability of hydrophilic reagents such as Ag+ or the methanethiosulfonates to modify
covalently the sulfhydryl side chains exposed to aqueous environments.
ATP-gated current was measured in HEK293 cells transiently expressing
either wild-type or functional mutant P2X2 receptors
containing a cysteine substitution in or around TM2. Application of
Ag+ to gating channels had no sustained
effect on wild-type P2X2 (WT) but irreversibly altered
whole-cell currents in 15 mutants. By contrast, bath
application of (2-aminoethyl)methanethiosulfonate (MTSEA) to
closed channels inhibited 8 of the 15 residues affected by
Ag+ when the channel was gating. Inhibition of the
closed channel was prevented in seven of eight mutants when
membrane-permeant MTSEA was scavenged by 20 mM
intracellular cysteine, indicating that these seven mutants lie on the
intracellular side of the channel gate. Further, MTSEA inhibited
current through G342C in the absence of intracellular cysteine but
augmented the current when cysteine was present, suggesting that this
residue may be part of the gate. Taken together, the data help to the
identify a functional domain of the channel pore by mapping residues on either side of the channel gate.
Key words:
ATP receptor; P2X subtype; scanning cysteine mutagenesis; sulfhydryl-modifying reagents; ion channel; transmembrane domain
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INTRODUCTION |
The ability of extracellular ATP to
modulate cellular functions involves the activation of a large family
of purinergic (P2) receptors that fall into two discrete subclasses
(Burnstock, 1996 ). Members of the first class are the metabotropic P2
receptors (P2U, P2Y, and others) that belong to the heptahelical
G-protein-coupled receptor superfamily. Members of the second class are
ligand-gated ion channels, designated P2X receptors (Fredholm et al.,
1994). Functional studies of these ionotropic receptors identify them as cationic channels that possess appreciable Ca2+
permeability (Benham and Tsien, 1987; Bean, 1992; Evans et al., 1996).
Recent molecular cloning studies suggest that the P2X receptors are
architecturally distinct from other known ionotropic receptor proteins
such as nicotinic-cholinergic, glutamatergic, and GABAergic receptors
(Surprenant et al., 1995; Buell et al., 1996a; North, 1996). They are
thought to have two transmembrane domains (TM1 and TM2) separated by a
large extracellular region containing 10 positionally conserved
cysteine residues and variable numbers of consensus sites for N-linked
glycosylation, with both the N and C termini located inside the cell
(Brake et al., 1994; Valera et al., 1994). This topology is novel for
ligand-gated channels (Ortells and Lunt, 1995) but is similar to that
of some K+ channels (Aldrich, 1993; Rossier et al.,
1994; Suzuki et al., 1994; Krapivinsky et al., 1995; Ortells and Lunt,
1995), amiloride-sensitive sodium channel subunits (Canessa et al.,
1993; Jentsch, 1994), a proton-gated cation channel (Waldmann et al.,
1997), and putative mechanosensitive channel subunits of nematode (Hong
and Driscoll, 1994; Huang and Chalfie, 1994). Although experimental
evidence from our laboratories (Torres et al., 1998) and others (Buell et al., 1996b; Collo et al., 1996) supports this model, the functional domains of the protein that form the ion conducting pore are poorly understood. The goal of the work reported here was to map such a
domain. We chose to study TM2 because it resembles similar domains of
other ion channels in its potential to form an amphipathic  -helix
having a hydrophilic face that lines part of the water-filled ion-conducting pore (Brake and Julius, 1996).
Pore-lining domains of other channel proteins have been identified by
the substituted cysteine accessibility method (SCAM) (Jakes et al.,
1990; Akabas et al., 1992). This method involves replacing individual
amino acids with cysteine and then testing the ability of
sulfhydryl-reactive reagents to modify the side chains of the
substituted residues. It assumes that the modifying reagents react only
with sulfhydryls exposed to aqueous environments and that modifications
of hydrated residues within the water-filled pore of the ion channel
lead, in some cases, to a change in current. We measured the ability of
Ag+ and (2-aminoethyl)methanethiosulfonate (MTSEA)
to alter current through wild-type (WT) and 27 individual
cysteine-substituted mutants. Our results suggest that TM2 is long
enough to traverse the lipid membrane and forms part of the
ion-conducting pore of P2X2. Further, the data predict that
TM2 is unlikely to form either a stable -helix (Surprenant et al.,
1995) or  -sheet (Rassendren et al., 1997), as previously
suggested.
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MATERIALS AND METHODS |
Construction of cysteine-substituted mutants.
Site-directed mutagenesis of the P2X2 receptor was
performed by the overlap primer extension method (Ausubel et al.,
1995). PCR was performed with VENT polymerase (New England Biolabs,
Beverly, MA), and all mutations were verified by sequencing with the
de-aza-GTP Sequenase kit from Amersham (Arlington Heights, IL).
Cell culture and transfection. Human embryonic kidney
(HEK293) cells were plated 1 d before transfection at a density of
4 × 105 cells per 35 mm culture dish and
placed in a humidified atmosphere containing 5% CO2 at
37°C. A mixture of 1 µg of cDNA and 6 µl of Lipofectamine (Life
Technologies, Gaithersburg, MD) in 1 ml of Opti-MEM (Life Technologies)
was added to each plate for 2-5 hr, after which the medium was
replaced with Minimal Essential Medium with Earle's salts supplemented
with 10% fetal calf serum, glutamine, penicillin, and streptomycin.
The cells were returned to the 5% CO2 incubator for 24-72
hr, at which time they were used for electrophysiology assays.
Transfection efficiency, judged by the presence of ATP-gated currents,
ranged from ~80% for WT and many of the mutants to ~16% for
others. Only S340C and D349C failed to respond to ATP (see below).
Electrophysiology. A suspension of transiently transfected
cells was made by agitating the solution bathing the cells attached to
the bottom of a single culture dish with a fire-polished Pasteur pipette. An aliquot of this cell suspension was transferred to a
recording chamber mounted on the stage of an inverted microscope. Whole-cell current was recorded from single cells (average whole-cell capacitance, 55 ± 2 pF; mean ± SEM) with an AxoPatch 200A
amplifier (Axon Instruments, Foster City, CA) and low-resistance
electrodes. The average uncompensated series resistance was 4 ± 0.2 M , of which up to 80% was nullified by using the internal
circuitry of the amplifier. The typical holding voltage was  40 mV,
although holding voltages of 20 to 80 mV were used occasionally to
compensate for relatively large or small currents, respectively. We saw
no unusual voltage-dependent behavior of the ATP-gated currents or the
ability of sulfhydryl-reactive reagents to alter these currents in this
voltage range. In most experiments the pipettes were filled with the
following intracellular solution (in mM): 140 CsCl, 10 tetraethylammonium-Cl, 5 EGTA, and 10 HEPES, pH 7.3 with CsOH. The concentration of CsCl was reduced to 130 mM when
20 mM cysteine was included in the filling solution. The
composition of the extracellular solution was determined by the nature
of the sulfhydryl-specific modifying reagent. When
Ag+ was used, the extracellular solution was (in
mM): 150 NaNO3, 1 Ca(NO3)2, 1 Mg(NO3)2, 10 glucose, and 10 HEPES, pH 7.3 with NaOH. When MTSEA was used, the extracellular
solution was (in mM): 140 NaCl, 0.5 CaCl2, 1 MgCl2, 10 glucose, and
20 HEPES, pH 7.0 with NaOH. This latter solution was designed to slow
hydrolysis of MTSEA and to prevent changes in pH caused by hydrolytic
byproducts of MTSEA. Stabilizing pH is particularly important because
ATP-gated current through P2X2 is augmented by
acidification (King et al., 1996). Drugs were applied by manually
moving the electrode and attached cell into the line of flow of
solutions exiting one of an array of inlet tubes. Every other tube
contained a test solution separated by a tube containing the normal
bath solution. Short applications (1-5 sec) of ATP were accomplished
by moving cells in a linear manner from control solution through an
ATP-containing solution to another control solution. Successive
applications were separated by 2-5 min to minimize receptor
desensitization. Modifying reagents were applied either in the presence
or absence of ATP. The percentage of change, measured from the averages
of an equal number (three or more) of steady-state responses before and
after application of a modifying reagent, was calculated as [(IATP,after/IATP,
before 1) × 100]. Each experiment was repeated at least
three times, and the results are displayed as the mean ± SEM.
Data were analyzed by one-way ANOVA. Significance was determined from
the Tukey's protected multiple comparison test by using GB-STAT (Dynamic Microsystems). p  0.01 was considered
significant. Inhibitions and potentiations were analyzed separately
because the large potentiations (up to 500%; see Results) made even
the largest inhibitions (which can be no more than 100%) appear
insignificant. Solutions of MTSEA were made from solids immediate
before the start of application. A stock solution of 1 mM
Ag(NO3) was made fresh daily and kept in the dark in
the refrigerator; an aliquot of this stock was added to the
solution in one barrel of the inlet array at the time of each
experiment. MTSEA was purchased from Toronto Research Chemicals
(Downsview, Ontario, Canada). All other reagents were purchased from
Sigma (St. Louis, MO).
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RESULTS |
The P2X receptor family is composed of at least seven subunits
that, although conserved at the structural level, have unique pharmacological and electrophysiological profiles. Of these seven, the
P2X2 subunit was chosen for these studies for the following two reasons. First, transient transfections of HEK293 cells with cDNA
encoding this subunit result in large ATP-gated currents in most cells.
Second, P2X2 shows little desensitization during short drug
applications and therefore is amenable to the repetitive drug
application protocol used in this study.
Wild-type P2X2 and 27 mutants were characterized in
voltage-clamped HEK293 cells. Cysteine mutations (L327C-L347C,
D349C-L353C, and M356C) were made throughout the predicted length of
the putative TM2 (Fig. 1). Every mutant
except S340C and D349C generated robust inward currents in response to
ATP (1-100 µM) applied to cells voltage-clamped at
negative transmembrane potentials. This lack of functional expression
contrasts with recent findings by Rassendren et al. (1997), who
reported that ATP evoked small currents from these two mutants. We
cannot explain this difference. We did not attempt a detailed
comparison of WT and mutant receptors, however, currents through all of
the responsive mutants superficially resembled those through WT in
their current densities, kinetics, desensitization, resensitization,
and current-voltage relationships, with one exception; currents
through S345C decayed at a slower rate than WT on washout of ATP. In 40 cells transfected with either S340C or D349C, ATP (10-1000
µM) failed to evoke membrane currents in cells
voltage-clamped at membrane holding potentials between 100 and 60 mV.
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Figure 1.
Amino acid sequence around putative transmembrane
region TM2 of P2X2. The region thought to lie within the
membrane is underlined. Each residue mutated to cysteine
is marked with a C. The asterisk indicates the position of endogenous C348.
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The SCAM was used to identify potential pore-lining residues.
Functional homomeric and heteromultimeric WT and mutant receptors were
tested by using both ionic Ag+ and MTSEA as
modifiers. Ag+ is a small inorganic monovalent
cation that forms a strong covalent S-Ag bond with thiolates exposed
to aqueous environments (Dance, 1986). MTSEA forms a disulfide bond and
attaches
-SCH2CH2NH4+.
Our experiments were designed to answer three questions. First, which
residues are modified by Ag+ when the channel is
gating? We assume that Ag+ permeates the channel and
modifies accessible residues on both sides of the channel gate. Second,
which residues are modified by MTSEA when the channel is not
gating? We assume that MTSEA enters the cell by passive diffusion and
labels both intracellular and extracellular residues of the closed
channel state(s). Third, are the residues modified by MTSEA in the
extracellular and/or the intracellular environments? We assume that
MTSEA modifies only extracellular residues when a high concentration of
free intracellular cysteine is present (Holmgren et al., 1996).
Identification of mutated residues exposed to the aqueous
environment during gating
Coapplications of ATP (10 or 30 µM) and 500 nM Ag+ were used to identify amino acid
residues exposed during gating. The atomic radius of
Ag+ is close to K+ and
Na+, and Ag+ permeates other
types of cation nonselective channels [Lü and Miller (1995) and
references therein], so it seems likely that P2X2 is also
permeable. This hypothesis was difficult to test because long (more
than ~10 sec) applications of Ag+ increased leak
current, and this prevented the accurate measurement of current through
P2X2 (Fig.
2A). This increase was
seen in both transfected and nontransfected HEK293 cells and was not
related to expression of the constructs used in this study. Shorter
applications seldom evoked nonspecific currents but did not allow
enough time to measure relative ionic permeabilities accurately.
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Figure 2.
Nonspecific and reversible effects of
Ag+. Both examples are from HEK293 cells transiently
transfected with cDNA encoding WT. In this and all figures of raw data,
the approximate times of ATP application are shown as solid
lines. Holding voltage was 40 mV. A,
Ag+ increased leak current in a reversible manner
when it was applied for longer than ~10 sec. B,
Coapplication of Ag+ and ATP (open
bar) reversibly potentiated current through WT.
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Wild-type P2X2 contains 13 endogenous cysteines, 11 of
which are predicted to be located either in the large extracellular loop or in TM2 (Surprenant et al., 1995; Brake and Julius, 1996). Coapplication of ATP and 500 nM Ag+ to
WT reversibly potentiated ATP-gated currents (Fig.
2B). A similar potentiation by other transitional
metals was reported previously (Li et al., 1993, 1996). Of greater
importance was the finding that no irreversible modification of
ATP-gated current was seen, suggesting that the native cysteines,
including C348 in TM2, either were not accessible to
Ag+ or that, if modified, did not affect current
flow through the pore. The lack of an irreversible effect on
ligand-gated current through WT permits a clearer interpretation of
results obtained via the cysteine-substituted mutant receptors.
Ten of the mutants (P329C, T330C, I331C, I332C, A335C, I341C, F346C,
W350C, I351C, and M356C; for example, see Fig.
3A) similarly were unaffected
by single 5 sec applications of 500 nM
Ag+ and ATP. By contrast, ATP-gated currents through
12 mutants (L327C, I328C, N333C, L334C, T336C, A337C, L338C, T339C,
V343C, S345C, L352C, and L353C; for example, see Fig. 3B)
were reduced significantly by concurrent administration. In most cases
a reduction in peak amplitude was the only irreversible effect seen
when Ag+ inhibited current. However, in some cases
(T336C, A337C, V343C, and S345C) coapplication of ATP and
Ag+ resulted in both a reduction in peak current and
the production of a small sustained inward current (see Fig.
3B). The production of sustained inward current may reflect
a population of ATP-gated channels that have been locked open by an
action of Ag+. We favor this explanation because we
never saw irreversible induction of inward currents in WT or most
mutant receptors and because the sustained inward current was reversed
by applications of 5 mM DTT (see Fig. 3B).
Currents through three mutants (G342C, G344C, and L347C; for example,
see Fig. 3C) were augmented irreversibly. In all cases the
inhibitions or potentiations began immediately after the start of
coapplication and progressed during the course of a 5 sec application.
The time course of the irreversible modification was obscured by the
concurrent reversible potentiation, precluding reliable measurement of
reaction rates of Ag+ at the various sites. Neither
the sustained augmentations nor inhibitions reversed during long (up to
45 min) washouts of Ag+, suggesting that a covalent
modification at the site of the substituted cysteine was responsible
for the change in the size of the current through the pore. A summary
of the irreversible effects of Ag+ on gating WT and
mutant P2X2 receptors is presented in Figure 4. Longer applications of
Ag+ may have produced greater effects, and the
augmentations and inhibitions that we detail here therefore must be
considered to be the lower limits of these effects.
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Figure 3.
Irreversible effects of Ag+
applied to gating channels. ATP (30 µM) was applied once
every 3 min to cells expressing mutant receptors. Each
trace shows three successive applications of ATP, followed by coapplication of ATP and 500 nM
Ag+, and then three more ATP applications after the
washout of Ag+. Coapplication of
Ag+ and ATP (open bars) had no
irreversible effect (A), irreversibly inhibited
(B) or irreversibly potentiated
(C) ATP-gated responses. DTT (5 mM;
hatched bar) partially reversed both the sustained inward current and the inhibition of current through V343C by Ag+ (B). See Results for
details. Calibration: 1000 pA, 10 sec.
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Figure 4.
Average data for wild-type and mutant
P2X2 receptors. Average data for the number of experiments
are indicated. Results significantly different
(p < 0.01) from wild-type
(WT) are marked with solid bars.
Inhibitions and potentiations are shown in the top and
bottom graphs, respectively.
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A change in current amplitude can reflect modification of a
residue within the pore. Another possibility is that modification of a
residue located at a distance from the pore changes current by
affecting agonist affinity or efficacy. To exclude this possibility, we
measured the dose-response relationship before and after covalent modification of four mutants (I328C, G342C, V343C, and L353C) having
substitutions spaced throughout TM2. Irreversible modification by
Ag+ increased (G342C) or decreased (I328C, V343C,
and L353C) the maximum response to a high concentration of ATP but did
not alter the EC50 values significantly (Fig.
5), suggesting that the addition of
Ag+ did not change agonist effcacy or
affnity.
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Figure 5.
ATP dose-response curves before and after
Ag+. Dose-response curves were generated before and
after a single 5 sec coapplication of 30 µM ATP and 500 nM Ag+ to cells expressing the indicated
mutants.
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Identification of mutated residues exposed to the aqueous
environment when the channel is closed
Some residues that are accessible when the channel is gating may
not be accessible when the channel is closed. This is because the
channel moves among several conformations (open, closed, and desensitized) in the presence of agonist but exists only in the closed
state in the absence of agonist. The 5 sec applications of 500 nM Ag+ applied in the absence of ATP
were used to label extracellular residues accessible in the closed
state. This protocol did not result in significant
(p < 0.01) modification of current through any
of the residues tested (I328C, N333C, L334C, T336C, A337C, L338C,
T339C, G342C, V343C, S345C, L352C, and L353C) (data not shown),
indicating either that these residues are entirely inaccessible in the
closed state or that access is limited. Longer applications of
Ag+ may have produced more significant effects but
caused the nonspecific leak currents described above.
MTSEA (1 mM), another sulfhydryl-reactive reagent, in our
hands did not activate leak currents when it was applied for up to 5 min in the absence of ATP. An added advantage of MTSEA is that it
exists at neutral pH as both cationic and uncharged species. The cation
is hydrophilic and readily reacts with sulfhydryls exposed to water.
The uncharged species is lipophilic and easily crosses cell membranes
by passive diffusion (Holmgren et al., 1996). Once it is inside the
cell, equilibrium between the charged and uncharged species is
reestablished, resulting in a significant intracellular concentration
of reactive MTSEA+. This means that it can attack
the protein from both sides of the membrane, resulting in the
modification of residues on both sides of the channel gate even when
the channel is closed.
Baseline ligand-gated current was measured by applying ATP for 1 sec
every 5 min for a total of three to five applications. Then, MTSEA was
applied for 5 min in the absence of ATP, the cells were washed with
normal bath solution, and periodic ATP applications were restarted.
ATP-gated current through WT was unaffected by MTSEA applied in this
way. We then tested 22 cysteine-substituted mutants of TM2, including
all 15 mutants accessible to Ag+ when the channel
was gating. Of these, current through eight (I328C, L334C, L338C,
T339C, G342C, S345C, L352C, and L353C) were inhibited significantly
(Fig. 6), and the rest were unchanged. These inhibitions persisted in an irreversible manner for at least 30 min, in keeping with the predicted covalent modification of the
cysteinyl side chain. As expected, all of the mutants modified by MTSEA
when the channels were in the closed state also were modified by
Ag+ when the channels were gating (compare Figs. 4
and 7). The seven mutants (L327C, N333C,
T336C, A337C, V343C, G344C, and L347C) that were not modified by MTSEA
apparently do not face an accessible aqueous environment when the
channel is closed. This may be because these residues are buried when
the channel is closed and swing out into water when the channel opens,
or it may be because they occupy a position accessible to
Ag+ but relatively inaccessible to the larger
MTSEA.
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Figure 6.
MTSEA inhibits some, but not all,
cysteine-substituted P2X2 receptors. MTSEA (1 mM) was applied alone for 5 min after baseline responses to
10 µM ATP were established. ATP was applied once every 5 min. Current through unmodified mutants (left-hand
sweeps of each pair of traces) resembled that of WT except that
current through S345C took longer to return to baseline on the
withdrawal of ATP. ATP-gated currents were reduced after application of
MTSEA (right-hand sweeps of each pair). Calibration: 500 pA, 3 sec.
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Figure 7.
Average data for wild-type and mutant
P2X2 receptors. Average data for the number of experiments
are indicated. Results significantly different
(p < 0.01) from wild-type (WT) are marked
with solid bars.
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Sidedness of closed-state reactive residues
The simplest model of TM2 is one that places I328C and L353C near
opposite ends of a membrane-spanning -helix with L353C close to the
intracellular surface of the membrane. To test the orientation of TM2
in the membrane, we used the method of Holmgren et al. (1996) to
scavenge intracellular sulfhydryl-reactive compounds. This approach
uses high levels of free cysteine in the electrode-filling solution to
inactivate MTSEA that enters the cell either through the patch leak or
diffusion through the membrane. If the free intracellular cysteine
prevents the change in current seen in its absence, then it is
reasonable to assume that the mutated residue lies on the intracellular
side of the channel gate. Although we assume that cysteine acts as a
scavenger, another possibility is that it protects by covalently
modifying accessible intracellular residues, making them unsusceptible
to subsequent MTSEA applications. In either case, only intracellular
residues would be affected when the channel is closed, and our
interpretation of the results would remain the same. An additional
concern is that extracellular residues are modified covalently by
cysteine that flows out of the cell through the pore of the channel
during establishment of baseline responses to ATP. Again, residues
modified in such a manner would not react with subsequent applications
of MTSEA and thus would appear to be intracellular. If cysteine flows
out through the channel (and against the electrical gradient), then we
expect to see a change in the size and/or shape of the first few
ATP-gated currents as cysteine modifies accessible residues. No such
change was evident.
The ability of 5 min applications of 1 mM MTSEA to modify
current through I328C, L334C, L338C, T339C, G342C, S345C, L352C, and
L353C was reinvestigated with 20 mM cysteine in the pipette solution. Lower concentrations of cysteine gave variable results. The
only residue significantly inhibited by MTSEA in the presence of
intracellular cysteine was I328C (Fig.
8A), positioning this residue on the extracellular side of the gate. L334C, L338C, T339C, S345C, L352C, and L353C were not inhibited significantly, which suggests that they are intracellular. Of greatest interest was the
effect on G342C. MTSEA potentiated current through G342C in the
presence of cysteine (Fig. 8B) but inhibited current
when cysteine was absent (see Fig. 6). These findings suggest that G342C can be attacked from either side of the membrane. The effect of
intracellular cysteine on the ability of MTSEA to modify current through mutant receptors is summarized in Figure
9.
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Figure 8.
MTSEA inhibits I328C and potentiates G342C
in the presence of 20 mM intracellular cysteine. Three
consecutive traces before (left side) and after
(right side, under the open bar) a 5 min application of 1 mM MTSEA are shown for I328C
(A) and G342C (B). Note
that the potentiation of G342C was reversed by a 5 min application of
5 mM DTT.
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Figure 9.
Average data for MTSEA in the presence and absence
of 20 mM intracellular cysteine. Solid bars
are significant data from Figure 7 obtained without cysteine and are
shown again here for direct comparison. Hatched bars are
results obtained in the presence of cysteine. Only I328C and G342C were
significantly different (asterisks) from WT during
cysteine perfusion. Note that G342C is potentiated by MTSEA in the
presence of intracellular cysteine but is inhibited in its
absence.
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Heteromerization allows nonfunctional homomeric mutants to
be tested
Substitution of the -COOH of D349 or the -CH2OH of
S340 with the -SH of cysteine appears in our hands to result in the
loss of channel function. There are four possible explanations for this
result. First, these mutations might result in a low efficiency in
protein translation/translocation. Second, substitutions at these
residues change agonist affinity. This idea is not supported by our
finding that even very high concentrations of ATP (up to 1 mM) did not evoke current in any cells. Third, side chains
important for cation conduction have been deleted. Fourth, disulfide
bonds form between adjoining cysteinyl side chains of homomeric
complexes of S340C and D349C, and these disulfide bonds disrupt current through the mutants. The latter two problems might be overcome by
studying heteromultimeric complexes of WT and S340C or D349C in which
some, but not all, of the subunits contain substituted residues. These
complexes might retain channel activity and at the same time acquire
susceptibility to sulfhydryl-specific reagents. We tested this
hypothesis by measuring the ability of MTSEA to alter ATP-gated current
in cells cotransfected with a mixture of either S340C/WT or D349C/WT
cDNAs. Both mixtures produced robust cation currents that resembled
those of WT in response to 10 µM ATP. MTSEA alone or in
combination with ATP had no irreversible effect on cells transfected
with a mixture of WT and S340C. Because we saw no effect, we draw no
conclusion about the accessibility of S340C. By contrast, MTSEA did
produce a substantial and irreversible block of current through
D349C/WT when it was applied along with ATP (Fig.
10) but had no effect on subsequent ATP
applications if it was applied to closed channels. The data argue that
D349C is on the intracellular side of the channel gate, but it is less accessible than other intracellular residues (L334C, L338C, T339C, S345C, L352C, and L353C) to the MTSEA that enters the cell by passive
diffusion through the lipid membrane when the channel is closed.
Coapplication of ATP and MTSEA had no irreversible effect on WT alone
(data not shown).
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Figure 10.
Heteromultimers of D349C and WT are blocked by
MTSEA when the channel is gating. Data are from a cell transfected with
a 2:1 mixture of D349C/P2X2 cDNAs. ATP (10 µM) was
applied once every 5 min to establish a baseline response. A sustained
application of ATP (filled bar) was begun, during
which time some receptor desensitization was apparent, followed
immediately by a 20 sec coapplication of ATP and MTSEA (open
bar). MTSEA caused an immediate decrease in whole-cell current
that was easily distinguished from the slower decrease in current
caused by receptor desensitization. In control experiments the
desensitization of WT followed an exponential time course. Assuming
this is also true of D349C/WT, we have indicated the expected
progression of whole-cell current in the absence of MTSEA with a
fine-dotted line. Short ATP applications were restarted
5 min after application of MTSEA. The peak current amplitude never
returned to its pre-MTSEA level. Holding voltage was 50 mV.
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DISCUSSION |
We used scanning cysteine mutagenesis to identify and map a domain
of P2X2 involved in ion conduction. This method carries with it certain assumptions that influence how results are interpreted (Akabas et al., 1992). The guiding principles are that functional mutants do not have unduly perturbed secondary structure, that channels
are water-filled pores lined in part by transmembrane-spanning domains,
and that Ag+ and MTSEA+ react
much more readily with ionized thiolates exposed to water than with
nonionized thiols exposed to lipid. Thus, residues that project into
the lumen of the channel might be accessible, whereas residues buried
in the lipid membrane are not. A change in current indicates that the
residue in question is accessible and modified. The SCAM has been used
to refine models of topology for numerous proteins in which the pore
domain was identified previously by other site-directed mutagenic
approaches (Akabas et al., 1992; Lü and Miller, 1995; Cheung and
Akabas, 1996; Kuner et al., 1996; Sun et al., 1996), and under these
conditions it is reasonable to conclude that reactive residues line the
channel pore. It is harder to justify this conclusion when the general
location of the pore domain is unknown, because it is also possible
that modification of a nonpore-lining residue alters current, perhaps
by changing receptor efficacy. In the case of P2X2,
this possibility must be considered carefully, because the locations of
the pore and the ATP-binding site are mainly unknown.
The hypothesis that TM2 does, indeed, line the pore of P2X2
is supported by six lines of evidence. First, hydropathy-plot analysis
of P2X2 predicts two putative transmembrane spanning domains, and one or both of these must constitute part of the pore.
Second, cysteine substitution within TM2 results in mutants sensitive
to hydrophilic modifying reagents, and a part of TM2 must face water.
Third, agonist potency was unchanged in all four mutants (I328C, G342C,
G344C, and L353C) in which ATP dose-response curves were generated
before and after modification by Ag+, arguing
against the possibility that current is changed by modifying agonist
EC50. Fourth, some mutants (L334C, L338C, S345C, L352C, and
L353C) fail to react with MTSEA in the presence of intracellular cysteine; these sites must be inside the cell and therefore cannot contribute to an extracellular ATP-binding site. By contrast, I328C
reacts equally well in the presence and absence of cysteine perfusion
and therefore must be outside. Taken together, these data confirm that
TM2 spans the membrane. Fifth, G342C is attacked from both sides of the
membrane, suggesting that it is exposed to water and contacts both the
extracellular and intracellular environments. This can happen
only if G342C resides in a water-filled transmembrane pore. Sixth,
D349C is modified only when the channel opens, implying that it lies at
a restricted site on the intracellular side of a channel gate.
Our initial strategy was to investigate gating channels with
Ag+ as the modifying reagent. We observed two
reversible effects that were unrelated to the presence of an engineered
cysteine, and these nonspecific effects somewhat limited the scope of
the experiment. Specifically, the transient potentiation of the
ATP-gated current prevented accurate measurement of the reaction times
of accessible mutants, and induction of a background current restricted the exposure time to Ag+. However, these nonspecific
effects disappeared on washout of Ag+, and in 15 mutants (L327C, I328C, N333C, L334C, T336C, A337C, L338C, T339C, G342C,
V343C, G344C, S345C, L347C, L352C, and L353C) an irreversible change in
ATP-gated current remained. These data are remarkable for the sheer
number of reactive positions. Indeed, it is somewhat surprising that
the endogenous cysteine at position 348 was unreactive in WT. It is
possible that some of the effects that we assign to
cysteine-substituted receptors are attributable to altered positioning
of C348 in the mutants. However, our data suggest that this is
unlikely, because current through most of the mutant receptors (S345C
being the exception) resembles WT and because it is hard to understand
how modification of a single residue could account for the variable
effects seen with the sulfhydryl-reactive reagents. This issue could be
resolved by making cysteine substitutions of a mutant receptor in which
C348 is changed to another amino acid. The present results are also
remarkable for the lack a discernible pattern in the reactive sites.
For example, all (excluding S340C, which was nonfunctional) but three
positions in the span from N333C to L347C reacted strongly, including
two tracts of four contiguous sites (T336C-T339C, G342C-S345C). This
contiguity seems to rule out stable -helical and -sheet secondary
structures for TM2 during gating. It does not, however, eliminate the
possibility that the channel assumes either of these conformations at
some time, for example, when the channel is closed.
The closed channel was studied by applying the modifying reagent in the
absence of ATP, and only eight (I328C, L334C, L338C, T339C, G342C,
S345C, L352C, and L353C) of the 15 residues that reacted with
Ag+ during gating were sensitive to MTSEA. Although
it is tempting to postulate that Ag+ reacts with
more residues during gating because additional sites are exposed when
the channel opens, an equally plausible explanation is that the smaller
Ag+ experiences less steric hindrance than the
larger MTSEA+, thereby gaining access to more
residues. We attempted to address this problem by studying the effect
of MTSEA on gating channels, but we found that coapplication of ATP and
MTSEA caused nonspecific effects in some (but not all) mutants, which
made interpretation of the results unreliable. These effects were not
unexpected, because studies investigating other ligand-gated ion
channels also have reported nonspecific effects of the
methanethiosulfonates applied to open channels (Kuner et al., 1996; Sun
et al., 1996). We did not observe such effects when MTSEA was applied
in the absence of ATP. Regardless, a pattern of reactivity does seems to emerge from the closed channel data in that all of the reactive sites lie on one face of an -helical model of TM2 having 3.6 amino
acids per turn. If so, then the protein must undergo a major conformation change when the channel opens, because the pattern of
reactivity is unordered during gating. Another possibility is that the
apparent helical pattern of reactivity in the closed state is
deceptive. This appears to be the case as ascertained in experiments
that used intracellular cysteine.
We determined the transmembrane orientation of reactive residues of the
closed state. Mapping extracellular residues was
straightforward; they were detected by bath application of
MTSEA and intracellular perfusion of cysteine. Mapping the
intracellular vestibule of P2X2 was problematic. One
approach is to measure single-channel current from inside-out membrane
patches, using electrodes filled with an extracellular solution
containing ATP and an intracellular (bath) solution containing MTSEA.
However, the continued presence of ATP in the pipette leads to
significant and progressive receptor desensitization, making the
absolute measurement of the effect of the modifying reagent tenuous. As
an alternative approach, we took advantage of the fact that uncharged
MTSEA easily crosses cell membranes and modifies residues on the
opposite side from which it is applied and that intracellular perfusion
of cysteine can be used to scavenge the permeant MTSEA (Holmgren et
al., 1996). We assume that residues that fail to react with MTSEA in
the presence of cysteine are contiguous with the intracellular
compartment, whereas those that do react are extracellular. From these
experiments we learned that, first, I328C is on the extracellular side
of the channel gate because it reacts equally well with MTSEA in the
presence and absence of cysteine. Second, six mutants (L334C, L338C,
T339C, S345C, L352C, and L353C) are intracellular because MTSEA was
ineffective when cysteine was present. Third, G342C resides at or near
the gate because it is accessible from both sides of the membrane. It
is difficult to envision how either an -helix or a -sheet could
give this pattern of reactivity. A topology that fits the data is an
outwardly facing loop with G342C at its apex (Fig.
11). Loop structures recently have been proposed for other ionotropic receptors (Kuner et al., 1996; Sun et
al., 1996). The role of the loop in P2X2 is unknown, but it is tempting to speculate that it forms part of a channel gate that
moves when the channel opens. Indeed, G342C is one of three amino acids
in TM2 completely conserved in all members of the P2X2 receptor family
(I328C and D349C are the others), and it may play a key role in channel
function. Further, it is interesting to note that modification by
Ag+ of residues at and around G342C produced some of
the greatest changes in current when the channel was gating. For
example, the largest inhibitions were seen after modification of T339C
and V343C, whereas currents through G342C, G344C, and L347C were
strongly potentiated. G342C itself must be capable of positioning its
cysteinyl side chain in either an inwardly or outwardly facing
direction, because MTSEA causes qualitatively polar effects (e.g.,
inhibition or potentiation), depending on whether the side chain is
attacked from the inside or the outside. We do not know how MTSEA
causes these two effects, and indeed the mechanism does not influence how the results presented here are interpreted. What is important is
that G342C is accessible from both sides.
View larger version (21K):
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|
Figure 11.
A possible topology for TM2. A, A
linear representation of accessibility of TM2 of homomeric constructs
of P2X2 to Ag+ applied to gating
channels (shaded circles) and MTSEA applied to closed
channels (solid circles). All residues affected by MTSEA when closed also were affected by Ag+ when gating.
The accessibility of other residues (open circles) is
unclassified either because they did not respond to the sulfhydryl modifying reagents or because they did not respond to ATP (S340C and
D349C). Currents through heteromultimeric complexes of D349C and WT were blocked by MTSEA applied to gating channels, and this residue most likely is on the intracellular side of the channel gate.
B, The intracellular and extracellular pattern of
accessibility to MTSEA applied to closed channels in the presence or
absence of 20 mM intracellular cysteine. G342C was
accessible from both sides.
|
|
Our data suggest that residues in TM2 line the pore. A similar
conclusion was reported recently in a paper published while this
manuscript was in preparation (Rassendren et al., 1997), although our
results are not in complete agreement with theirs; these discrepancies
may arise from the conditions used to study the cysteine-mutant
receptors, including the presence or absence of
methanthiosulfonate-induced background current and differences in the
concentrations of cysteine in the recording pipettes. In addition, our
results show that TM2 crosses the membrane in a nonhelical manner. The
fact that we find a large number of residues that react with
Ag+ suggests that this domain makes a major
contribution to the structure of the pore. These findings do not rule
out the possibility that additional parts of the protein contribute or
that there is an as yet unknown additional subunit. The results
presented here demonstrate that P2X receptors should not be modeled
after other, better characterized ion channels and that the structural
motifs underlying function remain to be determined empirically.
| |
FOOTNOTES |
Received Oct. 22, 1997; revised Dec. 15, 1997; accepted Jan. 15, 1998.
This work was supported in part by National Institutes of Health Grants
NS35534 (M.M.V.) and HL56236 (T.M.E.). We thank Drs. A. Brake and D. Julius for the kind gift of the P2X2 receptor cDNA, Gonzalo
Torres for discussion and technical support, Dr. A. Karlin (Columbia
University, NY) for advice about using methanethiosulfonates, Dr. Alan
Stephenson for help with data analysis, and Laura Hobart for Saturday
transfections.
Correspondence should be addressed to Dr. Mark M. Voigt, Department of
Pharmacological and Physiological Sciences, St. Louis University Health
Sciences Center, 1402 South Grand Boulevard, St. Louis, MO 63104.
| |
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W. R. Haines, K. Migita, J. A. Cox, T. M. Egan, and M. M. Voigt
The First Transmembrane Domain of the P2X Receptor Subunit Participates in the Agonist-induced Gating of the Channel
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Top
Abstract
Introduction
Compound via MeSH
