| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
Biophys J, July 2002, p. 206-218, Vol. 83, No. 1
Division of Biology 156-29, California Institute of Technology, Pasadena, California 91125 USA
 |
ABSTRACT |
|---|
 TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES
|
|---|
The mammalian serotonin transporters rSERT or hSERT were expressed in oocytes and labeled with sulforhodamine-MTS. The endogenous Cys-109 residue contributes most of the signal, and the labeled transporter shows normal function. The SERT fluorescence decreases in the presence of 5-HT and also depends on the inorganic substrates of SERT. The fluorescence also increases with membrane depolarization. During voltage-jump experiments, fluorescence relaxations show little inactivation or history dependence. The fluorescence signal has a voltage dependence similar to that of the prepriming step of the previously described voltage-dependent transient current. However, the fluorescence relaxations are the fastest voltage-dependent events yet studied at SERT; their time constants of ~8-30 ms are severalfold faster than the prepriming or inactivation phases of the transient currents. These fluorescence signals are interpreted within the framework of the gate-lumen-gate model. The signals may monitor initial events at the outer gate.
| |
INTRODUCTION |
|---|
|
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES
|
|---|
Despite recent strides in understanding the
relation between structure and function at neurotransmitter
transporters, relatively little is known about the conformational
changes that accompany ion-coupled transport. Several tools are
available to extend this knowledge. For the mammalian serotonin
transporter SERT, previous studies have used ligand binding,
radiotracer flux, substituted cysteine accessibility, and
electrophysiology, often in conjunction with site-directed mutagenesis.
State-dependent fluorescence changes represent another possible source
of data. Such measurements have not yet been applied to SERT, but were
previously applied to the Na+-coupled glucose
transporter SGLT1 (Loo et al., 1998
) and to the GABA transporter GAT1
(Li et al., 2000), which have topological or sequence similarity to
SERT. Furthermore, previous fluorescence measurements at GAT1 revealed
a signal with time course, voltage dependence, and
Na+ dependence similar to that of the SERT
voltage-dependent transient current (Li et al., 2000), but GAT1 does
not display SERT-like transient currents. We therefore asked whether
fluorescence signals at SERT would resemble those at GAT1 and/or would
parallel the SERT voltage-dependent transient current.
In planning site-specific fluorescence experiments for SERT, we noted
that several previous studies have utilized methanesulfonate probes at
the native Cys-109 group, which probably lies in an extracellular loop
between TM1 and TM2 (Chen et al., 1997a; Ni et al., 2001). This
residue reports an interaction between Li+ and
SERT (Chen et al., 1997a), is near regions in TM1 thought to
participate in 5-HT binding (Adkins et al., 2001), and even appears to
report conformation changes in regions of the protein (especially TM7)
that are distant from Cys-109 in sequence space (Kamdar et al., 2001).
Cys-109 therefore appeared to be a good candidate for fluorescence
studies. Alkylation at Cys-109 with small residues such as MTSET
inactivates the transporter (Cao et al., 1998; Chen et al.,
1997a, 1998); but we hoped that alkylation with a bulkier,
zwitterionic fluorescent group would add charge at a lower density,
perhaps retaining function.
We report here that this hope is fulfilled and that it is possible to
study the fluorescence of a covalently bound sulforhodamine group as
the functional SERT molecule undergoes transitions that depend on
membrane potential and on substrate concentration. This paper begins to
analyze such signals. We first gathered data for fluorescence changes
due to bath-applied substrates and for relaxations due to voltage jumps
at rSERT and hSERT; we then reexamined current relaxations during
voltage jumps, and we then compared electrophysiological and
fluorescence data. Voltage-dependent transient currents at rSERT were
previously reported (Mager et al., 1994), but this paper reports the
first study on the hSERT transient current.
| |
MATERIALS AND METHODS |
|---|
|
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES
|
|---|
Reagents and solutions
Two sulfhydryl-reactive MTS reagents, the fluorophore
sulforhodamine methanethiosulfonate (sulforhodamine MTS, S699150) and the preblocking reagent MTSET, were purchased from Toronto Research Chemicals Inc. (www.trc-canada.com). The control recording solution, ND96, contained 96 mM NaCl, 2 mM KCl, 1 mM MgCl2,
and 5 mM HEPES, pH 7.4. The NMDG Cl solution contained 96 mM NMDG
instead of Na+ in the ND96 solution. The sodium
gluconate solution contained 96 mM gluconate instead of
Cl
in the ND96 solution. The phosphate-buffered
saline (PBS) solution was purchased from Irvine Scientific (Irvine,
CA). Other chemicals were purchased from Sigma (St. Louis, MO).
Expressing hSERT and rSERT in Xenopus oocytes
The high-efficiency expression system for human and rat
serotonin transporters in Xenopus oocytes (Mager et al.,
1994; Cao et al., 1998) was used. In brief, 30 ng cRNA of WT rat or
human SERT (rSERT, hSERT) or the respective C109A mutant (Mager et. al., 1994; Cao et al., 1998; Y. Tong and M. Li, unpublished results) was injected into stage VI oocytes, and cells were incubated at 18°C
for 7-10 days. The incubation solution contained ND96 plus 2% horse serum.
Labeling hSERT and rSERT expressed in oocyte membrane
Seven days after oocytes are injected, expression is optimal for
fluorescence labeling. As a result, only exceptionally viable batches
of oocytes were eligible for these experiments. All manipulations were
performed at 22°C. Oocytes expressing hSERT and rSERT were pretreated
with 10 µM MTSET in ND96 solution for 10 min and washed three times.
This step, similar to one performed to isolate fluorescence from
K+ channels (Mannuzzu et al., 1996) or GAT1,
substantially reduced nonspecific sulforhodamine labeling. Oocytes were
then exposed for 15 min to a solution containing 96 mM LiCl instead of
NaCl, 100 µM sulforhodamine-MTS, and 1% DMSO (used to solubilize the sulforhodamine-MTS). The fluorescence signal was greater when the
oocytes were labeled in a Li+ solution than in
Na+ (Ni et al., 2001). The labeled oocytes were
then washed and incubated in the ND96, ready for recording. The result
of labeling was examined by measuring the fluorescence intensity of the
oocyte membrane at the animal pole.
Electrophysiology and fluorescence measurement
We used the instruments described in a previous study (Li
et al., 2000). In brief, two-electrode voltage clamp procedures were
used (Quick and Lester, 1994). The fluorescence of the labeled oocytes
is measured with a photomultiplier tube attached to the side port of an
inverted fluorescence microscope (Olympus IX-70-FLA) with a stabilized
100 W mercury light source and an objective of 40×, NA1.3. The filter
cube is the high-Q TRITC 410012b set (excitation 545 nm, half-width 30 nm; emission 610 nm, half-width 75 nm) from Chroma Technology Corp.
(Brattleboro, VT). The oocyte sits on the microscope stage and is
visualized for electrophysiology by a separate stereomicroscope. The
exciting beam was attenuated by factors approaching 100. Bleaching of
the fluorophore amounted to ~0.5% during a typical trial of 100 episodes that lasted a total of 1 min. The emission signal from the
oocytes was appropriately amplified and filtered. A HumBug (Qwest
Scientific, N. Vancouver, BC) cleaned up the remaining 60 Hz. Signals
were acquired and averaged by an Axon Digidata interface and pCLAMP 8 (Axon Instruments, Union City, CA). Traces shown were digitally
filtered at 200 Hz except where indicated.
Data analysis
The kinetics of the fluorescence and transient current were fitted to single or two-exponential processes with nonlinear routines in ORIGIN and CLAMPFIT 8.
| |
RESULTS |
|---|
|
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES
|
|---|
Fluorescence measurements
The fluorescence signals arise at labeled Cys-109 and depend on 5-HT
We labeled oocytes expressing human or rat SERT (hSERT or rSERT, respectively) cRNA with the sulfhydryl-reactive fluorophore, sulforhodamine-MTS, according to the protocol described in Materials and Methods. Oocytes expressing rSERT exhibit ~53% increase in fluorescence intensity compared to uninjected oocytes (Table 1); a similar percentage increase was observed for hSERT (data not shown). The rSERT-C109A mutant is fully functional and displays transient currents similar to those of wild-type rSERT (Fig. 2 B below and Cao et al., 1998), yet
oocytes injected with rSERT-C109A cRNA exhibit only ~23% increase in
fluorescence intensity compared to uninjected oocytes (Table 1). From
this result we conclude that more than half the fluorescence of the
SERT-specific fluorescence arises from labeling of Cys-109.
|
, 1998; Li et al., 2000; Lin et al., 1996; Mager et al., 1994).
This indicates that sulforhodamine-labeled rSERT is functional by
electrophysiological criteria. The amplitude of the
transport-associated current is ~20 nA at a holding potential of 
60
mV. The application of 5-HT also reproducibly decreased steady-state
fluorescence (Fig. 1 B); in the experiment of Fig. 1, this
decrease amounted to ~8% within ~50 s at a holding potential of
60 mV. 5-HT-induced changes in current and fluorescence of oocytes
expressing SERT have a different time course: when 5-HT is perfused
into the recording chamber, the fluorescence changes more slowly than
the current. In five oocytes studied systematically 100-120 s after
application of 20 µM 5-HT, the fluorescence decrease was 13 ± 2% (mean ± SEM). In the Discussion below, we comment on these
differences in time course between the optical and electrophysiological signals. We verified that uninjected oocytes displayed no change in
fluorescence when exposed to 5-HT.
|
Kinetics of the fluorescence relaxations
We measured simultaneous current and fluorescence relaxations in oocytes injected with rSERT, rSERT-C109A, or hSERT cRNA and subjected to voltage jumps (Fig. 2). The electrophysiological responses have been presented in previous publications (Cao et al., 1997, 1998; Lin et al., 1996; Mager et al.,
1994). At this point, we note only that hyperpolarizing jumps lead to
transient inward currents (denoted by arrows in Fig. 2) in
sulforhodamine-labeled rSERT and hSERT-injected oocytes, again showing
that the labeled oocytes are electrophysiologically functional. In a
later section we describe additional electrophysiological measurements
designed for direct comparison with the fluorescence studies.
|
40 mV to a test
potential of +60 mV, the fluorescence increases to a new steady state
~0.2% higher. The amplitude of the fluorescence increase varied
among batches of oocytes, between 0.1% and 0.8%. The fluorescence
increase is observed neither with the functional mutant C109A (Fig. 2
B), nor with uninjected oocytes (Fig. 2 C), suggesting that the voltage-dependent fluorescence originates from the
fluorophore label on the Cys-109 residue of SERT. Thus, although the
fluorescence relaxations are rather small and required us to average
100 or more sweeps for accurate measurement, they can be unambiguously
measured and assigned to a site-specifically labeled
fluorophore. Incidentally, these fluorescence relaxations are similar
in amplitude to those measured for the tetramethylrhodamine-labeled GABA transporter GAT1 (Li et al., 2000), but the kinetics of the relaxations, and even the sign of the change in response to voltage jumps, differ for the two transporters, again suggesting that the
signals arise directly from the transporters rather than from endogenous or artifactual sources.
The two transporters, rSERT and hSERT, have similar relaxation
amplitudes when studied under similar conditions (Fig. 2, D and E). However, the kinetics of the fluorescence increase
differ between rSERT and hSERT. In qualitative terms, most of the hSERT relaxation is much faster than the rSERT relaxation. When the fluorescence relaxations are fitted to two-exponential processes, the
rSERT signal gives time constants of 50 ± 10 ms and 460 ± 30 ms, with amplitude contributions of 30% and 70%, respectively. For
hSERT, the two components are 8 ± 5 ms and 570 ± 40 ms,
with amplitude contributions of 80% and 20% (Table
2). Several factors may distort these
estimates of time constants. First, the slower component rSERT and
hSERT may not be accurate because the fluorescence did not reach
steady state under our experimental protocol (longer pulses caused
damage to the cells when repeated several hundred times). The time
constants given may be underestimates. Second, the faster time constant
(8 ms) for hSERT may have been distorted by the settling time of our
voltage-clamp circuit. Therefore, this time constant may be an
overestimate.
|
Voltage dependence of the rSERT fluorescence relaxations
In addition to changing the steady-state fluorescence of SERT (Fig. 1), 5-HT also changes the voltage dependence of fluorescence. In the experiment of Fig. 3, fluorescence relaxations of rSERT were measured when the membrane potential was stepped from a holding potential of40 mV to test potentials between +60 mV and 140 mV at
40-mV decrements. The fluorescence versus voltage (F-V) plots are shown in the bottom panels. In the absence of 5-HT, the
fluorescence saturated at the most negative potentials studied, 140
to 120 mV (Fig. 3 A). This saturation was removed when 10 µM 5-HT was added to the recording chamber (Fig. 3 B). The
result is best illustrated by normalizing the fluorescence value at +60 mV to 1 and that at 140 mV to 0 (Fig. 3 D). If the
F-V plot in the absence of 5-HT resembles the beginning of a
rather shallow Boltzmann relation, the more linear F-V in
the presence of 5-HT resembles the midregion of a Boltzmann relation.
|
40 mV to +60 mV, then to 140 mV,
and back to 40 mV. 5-HT (10 µM) introduced a faster component in
the rise of the fluorescence relaxation produced by a depolarizing jump
and a slower component in the falling phase produced by a hyperpolarizing jump.
Ionic dependence of the rSERT relaxations
We also noted the effects of the ionic SERT substrates, Na+ and Cl, on
fluorescence. When Na+ was replaced by NMDG in
experiments like those of Fig. 1, rSERT fluorescence increased by
5 ± 3% over a period of several min (mean ± SEM,
n = 5). There was a slow phase of this increase; but it
was too small for systematic measurement. Replacement of Cl by gluconate decreased fluorescence by
5 ± 4% (mean ± SEM, n = 4); there was no
slow phase. Replacement of Na+ by
Li+ did not change fluorescence detectably
(<2%).
We also noted the effects of ionic substitutions on voltage-jump
relaxations at rSERT (Fig. 4). When
Na+ was replaced by NMDG, the normalized
F-V relation was steepest at extreme negative potentials
(Fig. 4, B and E). If the F-V plot in
the presence of Na+ resembles the beginning of a
rather shallow Boltzmann relation, the more linear F-V in
the presence of NMDG resembles the positive end of a Boltzmann
relation. The F-V relations were not changed detectably when
Na+ was replaced by Li+
(Fig. 4, C and F) or when
Cl was replaced by gluconate (Fig. 4,
D and E).
|
40 mV to +60 mV, then to 140 mV, and back to 40 mV.
The most significant effect of replacing Na+ by
NMDG is to slow the falling phase of the fluorescence relaxation (Fig.
4 H). Replacing Na+ by
Li+ does not have a significant effect on the
kinetics of the fluorescence relaxation (Fig. 4 I). The
effect of replacing Cl by gluconate resembles
that of replacing Na+ by NMDG, but to a lesser
extent (Fig. 4 J).
Substrate dependence of the hSERT relaxations resembles that of rSERT
Despite the fact that hSERT exhibits faster kinetics in fluorescence relaxations compared to rSERT, the effects of varying substrates on fluorescence relaxations are similar in hSERT and rSERT (Figs. 5 and 6). 5-HT decreased steady-state fluorescence of hSERT (data not shown) and rendered the F-V curve more linear (compare Fig. 5 with Fig. 3). Na+ replacement by NMDG or Li+, and Cl replacement
by gluconate, affect the F-V relation of hSERT like the
analogous replacements at rSERT (compare Fig. 6 with Fig. 4). The
effects of manipulating SERT substrates on the kinetics of the
fluorescence relaxations were also similar in hSERT and rSERT: 5-HT
introduced a slower component in the falling phase of the fluorescence
relaxation. Replacement of Na+ by NMDG or
Li+ and replacing Cl with
gluconate introduced a slower component in the falling phase of the
fluorescence relaxation.
|
|
Electrophysiological measurements
We now present a reexamination of current relaxations during
voltage jumps at rSERT and hSERT, followed by a comparison with the
fluorescence relaxations. In the course of this study we also gathered
the first data about hyperpolarization-activated transient currents in
hSERT-expressing oocytes (Fig. 7).
As previously reported for rSERT (Mager et al., 1994), a jump from a
positive potential to a large negative potential in the absence of 5-HT
produces a distinctive inward transient current. The transient current is eliminated by 5-HT (Mager et al., 1994), and we used this fact to
isolate the transient current in the traces of Fig. 7 A. The transient currents of hSERT and rSERT have similar dependence on both
prepulse and test pulse potentials. However, hSERT seems to display a
faster decay in the transient current. Two-exponential fits to the
transient currents at 140 mV give time constants of 45 ± 10 and
325 ± 19 ms for rSERT, and 63 ± 8 and 218 ± 15 ms for
hSERT (Fig. 7, B and D; see also Table 2).
Because the decay phase of the transient current is dominated (~70%)
by the slower component, the hSERT transient currents display a
half-decay time (~76 ms at 140 mV) roughly half as large as
the rSERT transient currents. The slow component of time constant 325 ms in rSERT may be due to HEPES and other amines binding to SERT,
because this component is decreased when HEPES is removed from the
recording solution (Table 2).
|
We detected fluorescence relaxations associated with both
hyperpolarizing and depolarizing steps. We therefore designed and conducted additional experiments to probe electrophysiological events
during depolarizing pulses. Little or no current actually flows, but
previous data show that a sojourn at positive potentials increases the
amplitude of a subsequently measured hyperpolarizing transient current
(Mager et al., 1994). This phenomenon may be described as an
inactivation at negative potentials or as a prepriming at positive
potentials. We use the latter term: a depolarizing prepulse preprimes
the SERT to a conducting state upon membrane hyperpolarization.
The voltage dependence of the prepriming step in rSERT was
previously studied (Mager et al., 1994). In the present experiments we
studied the time course of the prepriming process, using prepulses of
variable duration (Fig. 8). The
single-exponential time constant for the prepriming process at +60 mV
is ~86 ms for rSERT (Fig. 8). The same two-pulse protocol was also
used to measure the time course of the prepriming process in hSERT
(data not shown). In normal solutions the analysis is complicated by
the presence of a use-dependent blockade of the transient current in
hSERT; this blockade was eliminated by removing HEPES from the
recording solution. We found that the time constants of the prepriming
process in hSERT is near that of rSERT, i.e., ~95 ms at +60 mV
membrane potential (Table 2).
|
Comparing fluorescence and current measurements
The transient current depends on prepulse voltage because of two
presumably linked phenomena: inactivation at negative potentials and
prepriming at positive potentials. The detailed dependences are
presented in Fig. 9 A in a
format like that of the activation-inactivation plots for ion channels.
However, we find that there is little history-dependence or
inactivation of the fluorescence signals (Figs. 3-6). Fig. 9
B compares F-V plots for prepulses to either +60
mV, which fully preprimes the transient current, versus prepulses to
40 mV, which inactivates >80% of the transient current. There is
little difference between the F-V relations for these two
conditions. Fig. 9 B also shows that the F-V
relation is like the "prepriming" relation between prepulse
potential and transient current.
|
Table 2 gathers the information we have obtained about kinetic processes of hSERT and rSERT. Both fluorescence changes and transient currents are summarized. Evidently the situation is much simpler in the absence of HEPES, and these data will be emphasized in the Discussion. The time constants measured by fluorescence signals are all less than those measured by the transient current; e.g., for the prepriming step, the time constants measured by fluorescence are 8 ms for hSERT and 30 ms for rSERT; those measured by transient current are ~95 ms for both hSERT and rSERT. For the conducting step, the time constants measured by fluorescence are <20 ms and those measured by transient current are >50 ms (Table 2).
| |
DISCUSSION |
|---|
|
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES
|
|---|
We have detected a component of sulforhodamine fluorescence that occurs when SERT (either rSERT or hSERT) is expressed in oocytes. Specificity is assured by the fact that the C190A transporter shows normal function, but less SERT-dependent fluorescence. The SERT fluorescence also depends on transmembrane potential and on the organic and inorganic substrates of SERT. The fluorescence relaxations are the fastest voltage-dependent events that we have studied at SERT; they are faster than either the prepriming or the inactivation step of the transient current, but they have a voltage dependence similar to that of the prepriming step. Because the fluorescence relaxations amount to <1% of the background fluorescence even after 7 days of expression, experiments on these relaxations are tedious and limited only to exceptionally viable batches of oocytes. Nonetheless, we can draw some initial conclusions.
Working model
Fig. 10 presents a working model
for interpretation of the fluorescence and current measurements within
the framework of the gate-lumen-gate scheme (Cao et al., 1998). The
gate-lumen-gate model basically assumes that the transporter contains a
channel-like lumen with gates at each end. Coupled transport involves
"alternating access," or cycling of the gates so that the
transporter changes the compartmentalization of the substrates.
Conducting states, however, involve simultaneous opening of each gate.
The voltage-dependent transient current is the conducting state with
the largest known currents. We assume that prepriming current is
limited by events at the outer gate, i.e., the inner gate would remain
continually open during this process. We make this assumption partially
because externally applied drugs, such as HEPES, cocaine, and some
local anesthetics, can block the transient current and/or modify its waveform. Some of these pharmacological data have been published (Mager
et al., 1994); other data will be reported separately. Unfortunately,
we have no information about the mechanism(s) of inactivation of the
transient current at negative potentials. Our fluorescence signals have
no component that matches this inactivation.
|
We also believe that the outer gate is labeled by MTS-sulforhodamine
(SR), because the Cys-109 residue is externally accessible (see
Introduction). On this assumption we have assigned fluorescence changes
to transitions
either binding or conformational changeat the outer
gate; these fluorescence-detected transitions are represented by blue
arrows, which point to increased fluorescence. The largest decrease in
fluorescence (~13%) occurs when 5-HT binds at the outer gate. The
slow fluorescence changes in the presence of external 5-HT (Fig. 1)
could arise because this binding site is also somewhat accessible to
intracellular 5-HT, which accumulates during prolonged 5-HT
application. Because we have not studied whether the slow fluorescence
changes occur in the C109A mutant, it is formally possible that the
slow fluorescence changes occur at sulforhodamine-labeled residues
other than Cys-109; such non-Cys-109 labeling appears to account for
nearly half the MTSR signal (Table 1).
We suggest that the fluorescence also increases when the outer gate is primed to open, especially in the absence of 5-HT. That is, the transporter would enter the set of states named 2§. This suggestion arises because depolarization is associated both with increased fluorescence and with prepriming of the transient currents. The major components of the fluorescence relaxations are the fastest voltage-dependent events that we have studied at SERT; they are faster than either the prepriming or the activation step of the transient current, but they have a voltage dependence similar to the prepriming step. A hyperpolarizing step activates the transient current at a rate that probably exceeds the resolution of our voltage-clamp circuit. We believe this activation is probably an ohmic phenomenon like an instantaneous I-V relation; that is, the hyperpolarizing step generates current through states that are conducting but blocked, or just about to open. In short, we suggest that the fluorescence monitors early, local events near Cys-109 in the prepriming of the outer gate. Analogies to gating currents at ion channels suggest themselves. The state named 2§ is presumably a set of interconverting states, including primed, conducting, and inactivated states. Because voltage influences the equilibria among these states, the slower components of the fluorescence relaxations would arise from these changes.
The specific change in the environment of sulforhodamine that produces
the signals is unknown. The two major candidates are changes in
polarity and changes in quenching. The latter mechanism appears to
account for voltage-dependent changes in tetramethylrhodamine fluorescence at K+ channels (Cha and Bezanilla,
1997, 1998). Thus, environmental changes around the chromophore would
result from conformational changes induced by voltage changes or by
5-HT binding. The 5-HT-evoked fluorescence decrease is the largest
fluorescence signal yet observed with ion-coupled transporters. Some
(Adkins et al., 2001; Barker et al., 1999) but not all (Barker et al.,
1994; Buck and Amara, 1994; Chen et al., 1997b; Giros et al.,
1994; Kitayama et al., 1992; Lee et al., 2000; Smicun et al., 1999)
data suggest that 5-HT, dopamine, or norepinephrine bind at TM1 in
their respective transporters. However, 5-HT itself does not absorb
detectably at wavelengths >320 nm, and it seems quite unlikely that
binding to SERT would shift the spectrum by >200 nm, to cause direct
quenching of sulforhodamine. However, 5-HT might favor the binding of
an endogenous quenching fluorophore. Na+ or
Li+ would be required for such binding; NMDG
would remove this quenching by disfavoring the binding.
Although the precision of the present data may not merit an explicit scheme like Fig. 10, we include the scheme in this discussion because it has served well in our previous studies. In general, however, we believe that the data can usefully be evaluated against a large class of alternating-access models, in which ion-coupled transport is accomplished by conformational changes that control the compartmentalization of substrates.
Voltage and ionic dependence of the fluorescence relaxations
The voltage dependence of the fluorescence bears comment, but only
in a quantitative sense. We assume that the fluorescence behaves as a
sigmoid function of voltage and is characterized by a rather shallow
Boltzmann. If so, in the absence of 5-HT the midpoint of this relation
is more positive than the voltage range we have investigated (140 to
+60 mV). In the presence of 5-HT, the fluorescence becomes linear with
voltage over the entire range of our measurements, suggesting that the
midpoint is within this range. When Na+ is
replaced by NMDG, the relation begins to saturate at more positive
potentials, as though the midpoint of the Boltzmann relation is in a
region more negative than the range we have investigated. Assuming that
the midpoint of the Boltzmann relation is also the midpoint of a
conformational equilibrium sensed by the fluorescence, this midpoint is
shifted in the negative direction by 5-HT and even more in this
direction by NMDG. Recent data have anticipated the observation that
Li+ and NMDG, two Na+
substitutes, have distinguishable effects on the environment sensed by
Cys-109 (Ni et al., 2001).
Contrast with GAT1 relaxations
The florescence relations that we have measured at SERT-expressing
oocytes differ in several ways from those we have previously recorded
with covalently labeled tetramethylrhodamine at GAT1, even though 1)
the same apparatus was used (Li et al., 2000), and 2) GAT1 was labeled
at Cys-74, which aligns with SERT-Cys-109. The sign of the voltage
dependence differs: the fluorescence increases and decreases with
depolarization, respectively, at GAT1 and SERT. The kinetics of the
major phases of the relaxations are roughly 10-fold slower at GAT1 than
at SERT. The organic substrate GABA has little effect on the GAT1
fluorescence relaxations, but 5-HT has clear effects on both
steady-state and transient SERT fluorescence. Thus we can clearly
reject our previous suggestion (Li et al., 2000) that the state
revealed by the GAT1 fluorescence signals represents an occult
transient current. The speed and voltage dependence of the SERT
fluorescence relaxations actually resemble those of SGLT1 labeled in
TM11 (Loo et al., 1998) more than those of GAT1 labeled in TM1 (Li et
al., 2000).
This contrast between the fluorescence properties of labeled GAT1 and
SERT does, however, serve to emphasize the several differences between
the transport mechanisms of GAT1 and SERT (Lester et al., 1996).
Partially because of this contrast between the GAT1 and SERT
fluorescence signals, we have evaluated the SERT fluorescence signals
in terms of a physiological state that occurs in SERT, but not in GAT1,
rather than in terms of a mechanism or state thought to be common
between the two transporters. This unique state is the
voltage-dependent transient current.
GAT1 and SERT share the property that they are inactivated (at
GAT1-Cys-74 and SERT-Cys-109) by MTSET (Yu et al., 1998) but not by
rhodamine derivatives (Li et al., 2000). A reasonable explanation would
be that the bulkier, rhodamine group is added charge at a lower density
and is therefore not directly occluding the permeation pathway. In
addition, sulforhodamine is zwitterionic, so that no net charge is
added to the protein.
| |
CONCLUSIONS |
|---|
|
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES
|
|---|
These hypotheses about the significance of the SERT fluorescence signals are reasonable interpretations of our data, but alternative explanations are not yet excluded. Regardless of the detailed interpretations, we believe that our observations do establish the principle that substrate- and voltage-dependent movements of the SERT proteins (both rSERT and hSERT) do occur and that the voltage-dependent movements are faster than previously described events. We believe that more detailed schemes should be evaluated after experiments with additional labels and additional labeled residues.
| |
ACKNOWLEDGMENTS |
|---|
We thank Yanhe Tong for making the C109A mutant of hSERT.
This research was supported by National Institutes of Health Grant DA-019121 and by a National Research Service Award to Ming Li.
| |
FOOTNOTES |
|---|
Address reprint requests to Henry A. Lester, Division of Biology 156-29, California Institute of Technology, Pasadena, CA 91125. Tel.: 626-395-4946; Fax: 626-564-8709; E-mail: lester{at}caltech.edu.
Submitted September 24, 2001, and accepted for publication February 13, 2002.
| |
REFERENCES |
|---|
|
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES
|
|---|
-carbomethoxy-3-(4-fluorophenyl)tropane binding of dopamine transporter.
Mol. Pharmacol.
57:883-889
-aminobutyric acid transporter GAT1 revealed by simultaneous voltage clamp and fluorescence.
J. Gen. Physiol.
115:491-508
Biophys J, July 2002, p. 206-218, Vol. 83, No. 1
© 2002 by the Biophysical Society 0006-3495/02/07/206/13 $2.00
This article has been cited by other articles:
 |
|
 |
|
  D. G. Gagnon, C. Frindel, and J.-Y. Lapointe Voltage-Clamp Fluorometry in the Local Environment of the C255-C511 Disulfide Bridge of the Na+/Glucose Cotransporter Biophys. J., April 1, 2007; 92(7): 2403 - 2411. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 D. D.F. Loo, B. A. Hirayama, M. H. Karakossian, A.-K. Meinild, and E. M. Wright Conformational Dynamics of hSGLT1 during Na+/Glucose Cotransport J. Gen. Physiol., December 1, 2006; 128(6): 701 - 720. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 L. V. Virkki, H. Murer, and I. C. Forster Mapping Conformational Changes of a Type IIb Na+/Pi Cotransporter by Voltage Clamp Fluorometry J. Biol. Chem., September 29, 2006; 281(39): 28837 - 28849. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
L. V. Virkki, H. Murer, and I. C. Forster Voltage Clamp Fluorometric Measurements on a Type II Na+-coupled Pi Cotransporter: Shedding Light on Substrate Binding Order J. Gen. Physiol., April 24, 2006; 127(5): 539 - 555. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J. Zheng Patch Fluorometry: Shedding New Light on Ion Channels Physiology, February 1, 2006; 21(1): 6 - 12. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
D. D.F. Loo, B. A. Hirayama, A. Cha, F. Bezanilla, and E. M. Wright Perturbation Analysis of the Voltage-sensitive Conformational Changes of the Na+/Glucose Cotransporter J. Gen. Physiol., December 28, 2004; 125(1): 13 - 36. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. Blunck, D. M. Starace, A. M. Correa, and F. Bezanilla Detecting Rearrangements of Shaker and NaChBac in Real-Time with Fluorescence Spectroscopy in Patch-Clamped Mammalian Cells Biophys. J., June 1, 2004; 86(6): 3966 - 3980. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 H. P. Larsson, A. V. Tzingounis, H. P. Koch, and M. P. Kavanaugh Fluorometric measurements of conformational changes in glutamate transporters PNAS, March 16, 2004; 101(11): 3951 - 3956. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE |
F is expressed as a percentage of resting
fluorescence.
3 cells. (E) The
F-V plot in normal Na+ solution
(squares) and NMDG solution (circles).
(F) F-V plot in normal Na+
solution (squares) and Li+ solution
(circles). (G) F-V plot in
normal Cl
