Correspondence to: Henry A. Lester, Division of Biology 156-29, California Institute of Technology, 1201 East California Boulevard, Pasadena, CA 91125. Fax:626-564-8709 E-mail:lester{at}caltech.edu.
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Abstract |
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The rat -aminobutyric acid transporter GAT1 expressed in Xenopus oocytes was labeled at Cys74, and at one or more other sites, by tetramethylrhodamine-5-maleimide, without significantly altering GAT1 function. Voltage-jump relaxation analysis showed that fluorescence increased slightly and monotonically with hyperpolarization; the fluorescence at -140 mV was ~0.8% greater than at +60 mV. The time course of the fluorescence relaxations was mostly described by a single exponential with voltage-dependent but history-independent time constants ranging from ~20 ms at +60 mV to ~150 ms at -140 mV. The fluorescence did not saturate at the most negative potentials tested, and the midpoint of the fluorescencevoltage relation was at least 50 mV more negative than the midpoint of the chargevoltage relation previously identified with Na+ binding to GAT1. The presence of
-aminobutyric acid did not noticeably affect the fluorescence waveforms. The fluorescence signal depended on Na+ concentration with a Hill coefficient approaching 2. Increasing Cl- concentration modestly increased and accelerated the fluorescence relaxations for hyperpolarizing jumps. The fluorescence change was blocked by the GAT1 inhibitor, NO-711. For the W68L mutant of GAT1, the fluorescence relaxations occurred only during jumps to high positive potentials, in agreement with previous suggestions that this mutant is trapped in one conformational state except at these potentials. These observations suggest that the fluorescence signals monitor a novel state of GAT1, intermediate between the E*out and Eout states of Hilgemann, D.W., and C.-C. Lu (1999. J. Gen. Physiol. 114:459476). Therefore, the study provides verification that conformational changes occur during GAT1 function.
Key Words: voltage clamp, Xenopus oocyte, tetramethylrhodamine, conformational change
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INTRODUCTION |
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Specific Na+-coupled transporters are present on neuronal and glial membranes for all known lowmolecular weight neurotransmitters or, in the case of acetylcholine, for their catabolites. These transporters are thought to be responsible for removal of the neurotransmitter from the vicinity of receptors and are therefore important for termination of synaptic transmission. To influence the time course of synaptic transmission, this removal would be expected to occur on a time scale of milliseconds for ligand-gated channels, and on a time scale of hundreds of milliseconds for G-proteincoupled receptors. This postulated role for neurotransmitter transporters in neurotransmission calls for direct measurements of the time course of neurotransmitter transporter action.
Besides their postulated physiological function, neurotransmitter transporters are pharmacologically important: they appear to be the sites of action for important abused (cocaine) and therapeutic (antidepressants, psychostimulants, antiepileptics) drugs (
Therefore it is an important goal to study the physical mechanism of neurotransmitter transporters at the molecular level. The driving force for transporting neurotransmitter across the cell membrane comes from the electrochemical gradient of the cotransported ions, primarily Na+. Na+, Cl-, and sometimes K+ are cosubstrates with a stoichiometry of 1 or 2 mol/mol of neurotransmitter (-aminobutyric acid (GABA) transporter GAT1 (
In this research, we used combined electrophysiological and optical techniques to study the molecular mechanism of neurotransmitter transporter function. Electrophysiology can be used for transporter studies because ion binding and translocation steps, which are partial reactions in the transport cycle, produce electrical signals. For GAT1, electrophysiology is employed to assess rates, reaction steps, and turnover numbers (
For the experiments, we built an apparatus for simultaneous electrophysiological and optical recording from Xenopus oocytes. We detected and analyzed conformational transitions at GAT1 with a time resolution of milliseconds. These results may contribute to explaining the molecular mechanism of transporter function and drug action, which are important for understanding the physiological and pathological role of neurotransmitter transporters in native cells.
Our studies were enhanced by the use of two previously characterized GAT1 mutants. (a) The C74A mutant functions rather similarly to wild type (WT),1 yet is insensitive to the sulfhydryl reagent MTS-ethyltrimethylammonium (MTSET) (
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MATERIALS AND METHODS |
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Reagents and Solutions
The fluorescent dye tetramethylrhodamine-5-maleimide was purchased from Molecular Probes, Inc. The blocking reagent 3-maleimidopropionic acid was purchased from Aldrich Chemical Co. MTSET was purchased from Toronto Research Chemicals. The GAT1 inhibitor NO-711 was purchased from Research Biochemicals, Inc. Other reagents were purchased from Sigma Chemical Co. The recording solution, ND96, contained 96 mM NaCl, 2 mM KCl, 1 mM MgCl2, and 5 mM HEPES, pH 7.4. The incubation solution contained ND96 plus 2% horse serum. The NMDG substitution for Na+ contained 96 mM NMDG instead of Na+ in the ND96 solution. The gluconate substitution of Cl- contained 96 mM gluconate instead of Cl- in the ND96 solution.
Oocyte Expression
The high-efficiency expression system for GABA and serotonin transporters in Xenopus oocytes (
Fluorescence Labeling and MTSET Reaction
At the end of the sixth day of incubation, the oocytes were reacted with 10 mM 3-maleimidopropionic acid in ND96 for 1 h at 18°C, to block the endogenous reactive sulfhydryl groups in the oocyte membrane. After this blocking reaction, the oocytes were washed and brought to 18°C incubation for 1 d. On the eighth day after cRNA injection, the oocytes were incubated in ND96 solution containing 5 mM tetramethylrhodamine-5-maleimide (TMRM) for 1 h on ice as described (
The result of TMRM labeling was examined by measuring the fluorescence intensity of the oocyte surface at the animal pole with the apparatus described below, and by fluorescence confocal imaging of the oocyte surface using a MRC-600 confocal microscope (MRC-600; Bio-Rad Laboratories) with a 10x, NA 0.5 objective.
Apparatus
The recording setup, as shown in Fig 1, consists of a microscope, a photomultiplier tube (PMT) attached to the side port of the microscope, and conventional two-electrode voltage clamp instruments. The inverted fluorescence microscope (IX-70-FLA; Olympus Corp.) is fitted with a stabilized 100-W Hg light source and an oil-immersion objective of 40x, NA 1.3. The dichroic mirror is the high Q TRITC set from Chroma Technology Corp. The PMT (R928P; Hamamatsu Phototonics) is in a housing originally built by Photon Technology, Inc. The oocyte is placed on the microscope stage and is visualized for electrophysiology by a separate stereomicroscope. The exciting beam was attenuated by factors approaching 300 by neutral density filters; therefore, an incandescent lamp would probably suffice. A digitally controlled (i.e., finger-operated) shutter blocked the beam, except during actual data trials, to minimize bleaching. The emission signal from the oocytes was appropriately amplified and filtered at 200 Hz by an eight-pole low-pass filter (902-LPF; Frequency Devices, Inc.). Each trace was acquired and averaged over 30 sweeps by an Axon Digidata interface and pCLAMP 7 (Axon Instruments). A HumBug (Quest Scientific) removed the remaining 60 Hz. Two-electrode voltage clamp procedures were used as described (
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Off-line Data Analysis
Steady state fluorescence values were measured as the average over the final 200 ms at the test potential. For kinetic analyses such as those shown in Fig 6 Fig 7 Fig 8 Fig 9 (below), signals were subjected to further averaging across cells, digital filtering (50 Hz eight-pole Bessel), baseline alignment, and linear detrending where appropriate. Waveforms were fit to single or double exponentials with routines in ORIGIN 5 and CLAMPFIT 8. In preliminary analyses, we verified that (a) the baseline alignment did not result in systematic voltage-dependent shifts, and (b) the linear detrending did not result in systematic elimination of slow exponential components with time constants of ~500 ms or less. We cannot rule out the possibility that relaxations with larger time constants would be detected by test potentials longer than those used here.
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Because the fluorescence relaxations were small and noisy, accurate analysis of these relaxations is a major topic of this paper. Therefore, several details of the analysis are evaluated in the RESULTS. Fig 5 and Fig 6 show the progression from raw to analyzed traces for two important data sets. Fig 7 and Fig 8 present tests for history dependence. Fig 9 and Fig 10 compare two methods for analyzing the voltage dependence of the fluorescence relaxations.
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GABA Uptake Assay
[3H]GABA uptake experiments were performed as follows. Oocytes expressing GAT1 were incubated in ND96 solution containing various concentrations of GABA and trace amounts of [3H]GABA for 20 min, and then washed with ND96 solution five times. Each individual oocyte was then dissolved in 1 ml 10% SDS and the radioactivity was measured using a scintillation counter (LS 5000 TD/TA; Beckman Instruments Inc.).
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RESULTS |
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TMRM Treatment of GAT1 Does Not Affect GABA Uptake
Fig 2 presents results of [3H]GABA uptake experiments on oocytes expressing WT-GAT1 and exposed either to TMRM or MTSET. TMRM labeling did not significantly affect GABA uptake: for eight cells measured at 128 mM extracellular GABA, the uptake was 0.32 ± 0.02 and 0.33 ± 0.02 pmol/oocyte per 20 min in unlabeled and TMRM-labeled oocytes, respectively. When the data in Fig 2 were fit by hyperbolic (or Michaelis-Menten) doseresponse relations, the unlabeled and TMRM-labeled oocytes yielded virtually identical values: Km = 83 µM and Vmax = 0.54 pmol/oocyte per 20 min. These Km values are several times higher than the values previously observed for GAT1 expressed in oocytes (
GAT1 and C74A-GAT1 Are Labeled by TMRM
Fluorescent labeling of GAT1 by TMRM was verified by confocal microscopy (Fig 3) and was quantified by PMT measurements of the fluorescence of the oocyte surface at the animal pole, where the autofluorescence was partially absorbed by the pigment granules. We also studied C74A-GAT1, a mutant that is less susceptible to sulfhydryl reagents (
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TMRM Labeled WT-GAT1 and C74A-GAT1 Have Very Similar Electrophysiological Properties
Fig 2 showed that reaction with TMRM had little effect on GABA uptake by GAT1. We also found only small reductions in GABA-induced currents in electrophysiological studies. In measurements of transport-associated currents (200 µM GABA, -60 mV, measured at the end of a 40-s application), exposure to TMRM decreased currents by 17 ± 18% for WT-GAT1 and by 17 ± 5% for C74A-GAT1 (mean ± SEM, n = 7 and 2, respectively). Fig 4 presents additional data on electrophysiological properties of TMRM-labeled WT-GAT1 and C74A-GAT1. Fig 4 A shows that the time course and amplitude of GABA-induced current are quite similar. Average WT-GAT1 and C74A GABA-induced currents (100 µM, -60 mV, 40-s application) in the experiments of Fig 4 A were 120 ± 27 and 140 ± 42 nA, respectively (mean ± SEM, n = 8).
We also compared charge movements during voltage-jump relaxations in the absence of GABA for uninjected oocytes, WT-GAT1-injected oocytes, and C74A-injected oocytes. After each jump, the oocytes expressing WT-GAT1 and C74A display transient currents, which relax to new steady state currents over a time course of hundreds of milliseconds. The transient currents have previously been analyzed in detail ( = 25 mV. V1/2 is the voltage at which charge movements are half completed, Qmax corresponds to the complete movement of charges between the membrane and the medium, z is the charge of the particle moving,
is the fraction of the membrane field through which the charge moves, q is the elementary charge, and k and T have their usual meanings. The values for V1/2, Qmax, and kT/qz
are similar to previously reported values for WT-GAT1 (-27 mV, 80 nC, and 28 mV, respectively;
Interactions between TMRM and MTSET in Fluorescent Labeling
For the experiments presented in Table 2, fluorescence was measured on oocytes reacted with TMRM and/or MTSET in all possible sequences. As expected from the data of Table 1, the fluorescence intensity of TMRM was less in C74A-GAT1 than in WT-GAT1-expressing oocytes. Furthermore, the fluorescence intensity of TMRM-reacted WT-GAT1 oocytes was reduced by almost 50% when TMRM treatment was followed by MTSET reaction. For C74A-GAT1 oocytes, the percent reduction in the TMRM fluorescence intensity by MTSET was greater, 76%; however, the absolute value of the decrease in fluorescence signal was roughly equal (34 V) in the two cases. Furthermore, the fluorescence intensity for TMRM-labeled WT-GAT1 also exceeded that for TMRM-labeled C74A-GAT1 by 34 V. These results indicate (a) that the fluorescence signal in WT-GAT1 is caused by TMRM reaction with Cys74 and also with another residue, and (b) that TMRM fluorescence at this second residue is reduced by subsequent reaction with MTSET, via an unknown mechanism. Because the C74A transporter activity is not blocked by MTSET, it is clear that reaction of MTSET with cysteines other than Cys74 does not inhibit transporter function. Table 2 also presents results of the reverse series of exposures. After MTSET pretreatment, exposure to TMRM results in an increment of ~4 V in fluorescence signals at both WT-GAT1 and C7A-GAT1. In the most straightforward interpretation of this result, MTSET alkylation prevents TMRM from labeling Cys74, but allows labeling at non-Cys74 sites.
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We attempted to extend these observations in a set of electrophysiological measurements on oocytes before and after sequential TMRM and MTSET exposure. However, these dual exposures and repeated impalements rendered the oocytes so unhealthy that the results were inconclusive. Our experiments yield no decisive information about the fraction of Cys74 residues that are labeled by TMRM; we assume, but cannot prove, that this fraction is near unity. Nevertheless, the data do suggest that roughly half the steady state fluorescence signal arises from TMRM bound to Cys74. Furthermore, C74A-GAT1 provides a useful contrast to WT-GAT1 because C74A-GAT1 functions like WT-GAT1, but is not labeled by TMRM at position Cys74. However, both MTSET and TMRM appear to react with one or more additional sites on GAT1. These observations provide the background for time-resolved measurements on TMRM-labeled GAT1 fluorescence during voltage jumps, reported in detail below.
Voltage Jumps Induce Fluorescence Changes in TMRM-labeled rGAT1
Fig 5 presents a survey of simultaneous electrophysiological and optical signals recorded from oocytes in the apparatus described in Fig 1. We used the voltage-jump protocol introduced by
In the C74A mutant, there was a detectable but smaller fluorescence change, <0.4% over the range from +60 to -160 mV (Fig 5 C). Again, depolarization decreased and hyperpolarization increased the fluorescence.
Fig 6 shows the voltage and time dependence of the fluorescence signals in more detail. In Fig 6A and Fig B, the fluorescence relaxations are fit to single exponentials after suitable averaging and filtering (see MATERIALS AND METHODS). For C74A-GAT1, the amplitude of the fluorescence change was approximately linear with voltage (Fig 6 C). For example, the change over the 100-mV ranges from +60 to -40 mV and from -40 to -140 mV was 0.22 ± 0.01% and 0.20 ± 0.01%, respectively (mean ± SEM, n = 5 oocytes). In contrast, the fluorescence signals from WT-GAT1 were nonlinear with membrane potential: jumps to potentials more negative than -60 mV resulted in larger increases in fluorescence intensity than jumps to potentials more positive than -60 mV. When expressed as a percentage of background fluorescence, WT-GAT1 fluorescence relaxations at voltages more negative than -60 mV were 1.5- to 3-fold greater than C74A fluorescence relaxations; but because the absolute fluorescence intensity was 1.66-fold higher for oocytes expressing WT-GAT1 than for the C74A transporter, the absolute values of relaxations in WT-GAT1 oocytes were 2.5- and 4.5-fold larger than in C74A-GAT1 oocytes for jumps to -60 and -140 mV.
The time constant of the fluorescence relaxations showed much greater voltage dependence for WT-GAT1 than for the C74A mutant (Fig 6 D). The average time constant of five C74A cells was between 75 and 103 ms for all voltages from +60 to -140 mV (Fig 6 D), whereas the average time constant of five WT-GAT1 cells increased monotonically from 22 ± 1 ms at +60 mV to 151 ± 6 ms at -140 mV, a 6.5-fold change. An exponential fit of the voltage dependence of the WT-GAT1 time constants gave an e-fold change per 120 mV, but the value for C74A-GAT1 was at least 600 mV. Thus, although differences in fluorescence amplitude between WT-GAT1 and C74A-GAT1 are apparent only at membrane voltages from -60 to -140 mV (Fig 6 C), the relaxation kinetics differ between WT-GAT1 and the C74A mutant over almost the entire voltage range accessible to experiment. In other words, the WT-GAT1 relaxation waveforms are not a simple sum of a Cys74-independent component plus an additional Cys74-specific component.
Tests for History Independence
In relaxation analysis, it is a fundamental concept that transition probabilities depend on the present value of parameters such as membrane potential, drug concentration, and temperature, but not on the history of these parameters. The apparently simple behavior of the relaxations allowed a test of this concept for the fluorescence signals (Fig 7 and Fig 8). The time constant of the fluorescence relaxation for WT-GAT1 did not depend strongly on the prepulse potential for steps to a constant test potential (-140 mV in the experiment of Fig 7). The single-exponential time constant remained in the range from 122 to 155 ms as the prepulse voltage ranged from -100 to +60 mV (Fig 7 C). The amplitude of the exponential component did of course depend on the prepulse potential, and decreased from 0.38% at a prepulse potential of +60 mV to 0.12% at a prepulse potential of -100 mV (Fig 7 B). There was an additional component, with a time constant <10 ms, for the largest jumps (from voltages more positive than zero). This was a consistent finding among each of 40 cells tested. This rapid component has not been examined systematically in our experiments.
In the complementary experiment, we studied the kinetics of the fluorescence decrease due to a voltage jump from a constant hyperpolarizing voltage (-140 mV) to varying depolarized levels (between -60 and +60 mV). The results are shown in Fig 8. Unlike the previous experiment, the time constant of the fluorescence decrease was dramatically affected by the membrane potential of the test pulse: the time constant increased from 30 ± 1 ms at +60 mV to 100 ± 3 ms at -60 mV, a threefold increase (Fig 8 C). At each voltage, these time constants are similar to the values measured for jumps from a prepulse potential of -40 mV (Fig 6). Fluorescence relaxations, therefore, appear to conform to the concept that the relaxation rates depend on the present value of the membrane potential rather than on its history.
Comparison of Fluorescence and Charge Movement
As noted above, voltage jumps evoke transient capacitive currents at WT-GAT1. The voltage dependence and kinetics of these charge movements have been analyzed in detail previously (
Fig 9 compares the voltage dependence and kinetics of the capacitive charge movements with those of the fluorescence relaxations. Data from five cells were averaged, and the amplitudes of the fluorescence change and charge movement were plotted as a function of membrane voltage. As seen in the figure, the plot of fluorescence lies to left of the plot of charge movement by at least 50 mV (Fig 9 A). Furthermore, the fluorescence shows no sign of saturation with hyperpolarization, so that the midpoint of the fluorescencevoltage relation cannot be determined. Although we know that the midpoint of the chargevoltage relation is approximately -26 mV (Fig 4), we know only that the midpoint of the fluorescencevoltage relation is more negative than approximately -75 mV. The actual difference between charge and fluorescence is thus at least 50 mV on the voltage axis.
Despite the similar range of time constants for the charge movements and the fluorescence signals, they differ significantly in value at almost every voltage tested between +60 and -140 mV (Fig 9 B). Furthermore, the time constants characterizing the fluorescence change and charge movement have distinct dependences on membrane potential (Fig 9 B). While the time constant of the fluorescence change increases monotonically with hyperpolarizing voltages (with perhaps a hint of saturation at the highest negative potentials), the time constant of charge movement shows a maximum at -40 mV (Fig 9 B; see also
An Alternative Subtraction Procedure Confirms the Distinct Voltage Dependences of Charge Movement and Fluorescence
Because the fluorescence signals are small, we sought additional tests of the conclusion that the fluorescence signal occurs at membrane potentials more hyperpolarized than the charge movement. The amplitude of the fluorescence relaxations for C74A, expressed as F/F, equals that of WT-GAT1 for most of the voltage range (Fig 6). As discussed above, WT-GAT1 fluorescence waveforms cannot be expressed simply as a sum of Cys74-dependent and -independent terms. Nevertheless, we subtracted the fluorescence signal of C74A from that of WT-GAT1 (both expressed as
F/F) to approximate a site-specific fluorescence signal from residue Cys74. The resulting waveforms are shown in Fig 10 A. Using this analysis, fluorescence relaxations due to TMRM labeling at C74 appear to occur only at hyperpolarized voltages (more negative than -40 mV). The fluorescence increased with hyperpolarizing membrane potentials; there were no signs of saturation at the most negative membrane potential tested, -140 mV (Fig 10 B). The fluorescence relaxation fit well to a single exponential process with time constant ranging from 75 to 150 ms at potentials from -40 to -140 mV (Fig 10 C). These observations on voltage dependence and kinetics agree well with the characteristics of the unsubtracted traces (Fig 6 Fig 7 Fig 8 Fig 9). Thus, the details of the signal analyses do not strongly affect the conclusion that distinct voltage dependences characterize the fluorescence change and the charge movements.
Effect of Substrates
The effect of GAT1 substrates, GABA, Na+ and Cl- on the electrophysiological properties of GAT1 was studied previously in our laboratory (
The effects of changing these substrates on the fluorescence signal of GAT1 are shown in Fig 11. The top panels show exemplar fluorescence traces for the voltage-jump relaxations from -40 to -140 mV; The bottom panels present the effects of GAT1 substrates on the fluorescence amplitude averaged across several complete voltage-jump experiments, like those of Fig 5, at test potentials in the range +40 to -140 mV. GABA produced virtually no change in the amplitude of the fluorescence relaxations (although the simultaneous voltage-clamp measurements revealed that the charge movements were shunted and became GABA-induced currents as expected).
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Cl- substitution with gluconate modestly decreased the amplitude of fluorescence relaxations, by 1050%, at voltages more negative than -60 mV (Fig 11, B1 and B2). If the effect of Cl- replacement by gluconate is treated as a shift in the fluorescencevoltage relation, the shift amounts to ~15 mV in the negative direction. This is considerably less than the ~44-mV negative shift reported for the same ionic replacement in charge movement experiments (
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On the other hand, Na+ substitution with NMDG blocked most of the fluorescence signal. Of >20 cells studied, for the jump from -40 to -140 mV, the amplitude of the fluorescence change was decreased by ninefold in the absence of Na+, and there were similar reductions at all voltages more negative than -60 mV. The remaining relaxation had a time constant of 32 ± 4 ms (n = 4). Table 3 also shows that the time course of the fluorescence decrease for the voltage jump from -140 to -40 mV was not significantly affected by Cl- but was affected by Na+. In the presence of both Na+ and Cl-, the fluorescence decrease is a single exponential process with time constant of 59 ± 1 ms (n = 4). In the absence of Cl- (substitution by gluconate) and presence of Na+, the time constant of the fluorescence decrease was 56 ± 4 ms (n = 4). The time course of the fluorescence decrease was dramatically affected by the absence of Na+: the time constant decreased to 9 ± 1 ms (n = 4).
Na+ Concentration Dependence of the Fluorescence Relaxations
The Na+ dependence of the fluorescence was further studied by varying the Na+ concentration in the extracellular medium. The results are shown in Fig 12. To study the Na+-dependent fluorescence signal, the residual small fluorescence that is Na+ independent was subtracted from each trace. Upon voltage jump from -40 to -140 mV, the fluorescence increased in a single-exponential process (Fig 12 A). Both the amplitude and the time constant of the fluorescence change depended on the Na+ concentration. Because there was no apparent saturation with Na+ concentration, the plot of fluorescence amplitudeNa+ concentration was fit to a power law (as though it were the foot of a Hill function). The exponent was 1.8 (Fig 12 B), which is consistent with the observation from previous studies that two Na+ interact with GAT1 (-140 mV, vs. [Na+] was fit to a straight line, resulting in a slope of 64 M-1 s-1 (Fig 12 C). Although complete removal of Na+ did dramatically accelerate the fluorescence decrease for the jump from -140 to -40 mV (Table 3), there was no systematic effect of Na+ concentration in the range from 48 to 96 mM; the rate constant, 1/
-40 mV, varied between 13 and 18 s-1 (Fig 12 D).
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Effect of a Transport Inhibitor
The GAT1 inhibitor, NO-711 (Fig 13 A), was studied for its effect on the fluorescence and charge movement of GAT1. NO-711 blocked the charge movements of GAT1 during voltage jumps, as found in earlier studies (
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The W68L Mutation Shows Fluorescence Relaxations Only at High Positive Potentials
We also labeled oocytes expressing the GAT1-W68L mutation (
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That W68L-GAT1 produces detectable fluorescence relaxations under jumps to high positive potentials fits well with the idea that the fluorescence relaxations monitor transitions of active transporters. In previous studies, W68L-GAT1 showed GABA transport and transport-associated currents <10% those of WT-GAT1 (
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DISCUSSION |
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These experiments introduce simultaneous measurements on electrophysiology and fluorescence for the study of a neurotransmitter transporter. The data show that specific fluorescence signals are obtained when the GABA transporter GAT1 is subjected to jumps in membrane potential. Proof that the signals arise directly from functional GAT1 comes from several facts. (a) The signals depend absolutely on GAT1 expression. (b) The point mutant C74A produces distinct signals. (c) The poorly functional W68L transporter displays WT levels of steady state fluorescent labeling, but only small relaxations and only under the same restricted conditions that also produce charge movements. The results for W68L-GAT1 would be significant even if there were no fluorescence signals at all; the argument does not depend on the small signals for jumps to +40 and +60 mV, but does depend on the normal level of fluorescence labeling combined with the absence of most fluorescence relaxations. (d) The GABA uptake inhibitor NO-711 blocks the fluorescence relaxations. These observations eliminate a number of admittedly unlikely artifacts arising from (a) nonspecific effects of membrane protein expression or (b) accessory proteins that might be brought to the membrane by GAT1.
The data also show that part of the fluorescence signal arises at Cys74, a residue thought to lie in the first extracellular loop (
There are non-Cys74 signals as well; these appear to depend linearly on membrane potential. However, the fact that we measure single but different time constants in the presence and absence of Cys74 suggests that the signals are not a simple sum of Cys74 and non-Cys74 components, vitiating a clear view of possible interactions among fluorescent labels. Interpretation of the non-Cys74 signals is also complicated by four additional observations about effects of sulfhydryl reagents. (a) In the present experiments, TMRM labeling could occur at non-Cys74 sites despite previous reaction with MTSET, but could also be reversed by subsequent exposure to MTSET (Table 2). (b) Reactivity at Cys residues other than Cys74 may depend on the functional state of the transporter (
The fluorescence signals are small, limiting our ability to resolve their detailed amplitude and time course under varying conditions. The largest change in fluorescence that we observed, corresponding to voltage jumps between +60 and -140 mV, amounted to 0.8%. Our apparatus works well, as judged by the robust signals (31%) obtained with a similarly labeled K+ channel (
In addition to the challenges posed by temporal analysis of the relaxations, other uncertainties are raised in evaluating the steady state fluorescence intensities. Previous studies show that changing the concentration of inhibitors, of Na+, or of Cl- at constant voltage produces charge movements associated with ion binding to GAT1 (
Despite these limitations, the experiments clearly show that the fluorescence signals differ in important ways from the previously known signals, primarily electrophysiological, associated with GAT1 function. The dependence on membrane potential (Fig 9 and Fig 10), GABA (Fig 11), Cl- concentration (Fig 11), and Na+ concentration (Fig 12) all differ markedly from the characteristics of the transport-associated currents and transient charge movements described in previous studies (
Nature of the Voltage-induced Fluorescence Signal
The effects of jumping the membrane voltage on GAT1 could have one or more of three physical bases. (a) Membrane potential changes the electrochemical gradient of the charged GAT1 substrates, Na+ and Cl-. Under most of our conditions, a hyperpolarizing voltage jump increases the electrochemical driving force for Na+, thus possibly changing the balance of transporters among intermediate states in the transport cycle. (b) A jump in the membrane electrical field could change in the energy barrier for some conformational changes during GAT1 function. These conformational changes could involve entire domains of the protein, or individual side chains. (c) In a related mechanism, the covalently attached fluorophore itself, which has a substantial dipole moment, could move in the field. Changes in fluorescence of reporter groups are thought to arise from changes either in the polarity of the immediate environment and/or in quenching by discrete neighboring moieties. Any of the mechanisms described above could involve such changes. In addition, one should consider (d) altered quenching by ions bound nearby. We also cannot rule out changes in quenching due to interactions between neighboring fluorophores, although modern studies have shown no evidence that transporters similar to GAT1 exist as multimers (
The fact that the fluorescence signal from WT-GAT1 is a nonlinear function of voltage argues against hypotheses c and d, which would be expected to produce effects that are linear, at least to first order, with the membrane field. A related observation, that W68L-GAT1 displays fluorescence relaxations only at the high positive potentials that also permit conformational changes (Fig 14) (
Nature of the Fluorescent State
A conformational change produces the fluorescence signal. Conformational changes form the basis of kinetic-state models for GAT1 function studied by
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We were initially surprised that GABA does not strongly affect the relaxations. However, we emphasize that most transporters are thought to reside in the states that include E*out, E*out-fluo, and Eout, both in the presence and absence of GABA (
Our study thus supports kinetic models in which multiple conformational changes occur during neurotransmitter transporter function, and our data suggest that one such conformational change comprises part of the transition previously thought to limit the rate of transport (
Relationship to Other Work
We believe that we have detected a conformational change in GAT1 near Cys74. The importance of this region in GAT1 function was also revealed by other studies. Amino acid residues near Cys74 are proposed to be the Na+ or Cl- binding site: this region is most conserved in the Na+, Cl--dependent neurotransmitter transporter family, and Arg 69 and Trp 68 in this region, the putative role of which is to interact with ions, are absolutely required for GAT-1 function (
That the fluorescence does not directly monitor the GABA-transporter interaction agrees with previous suggestions that the extracellular loops between TM7 and TM8 and between TM11 and TM12 are involved in GABA binding (
The GAT1 Fluorescence Signal Recalls a Phenomenon at Serotonin Transporters
The serotonin transporter SERT has ~40% sequence similarity to GAT1, and therefore the two transporters presumably share overall structural details. We feel justified in comparing some functional phenomena as well between SERT and GAT1. At SERT, jumps to hyperpolarizing voltages induce the transient activation of a channel-like conducting pathway (
If a state like E*out-fluo does produce the transient resistive current in SERT, then E*out-fluo contains a channel-like pathway in SERT but not in GAT1. Perhaps this partially occluded state, which we have already drawn as extending nearly through the molecule (Fig 15), is actually open to both sides in SERT. Indeed, several studies suggest that a channel-like pathway also exists in GAT1 (
Outlook
Over the past decade, electrophysiological experiments have revealed the existence of several previously unsuspected states at neurotransmitter transporters. We have shown how to conduct fluorescence measurements on a neurotransmitter transporter. We conclude that our measurements monitor a novel conformational state of GAT1, but this state has properties intermediate between those of two known states, and it may resemble a previously described state of SERT. We expect that it will be possible to measure fluorescent signals associated with labeling of other GAT1 residues. We hope that some of these future signals will be larger and therefore more amenable to quantitation.
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Footnotes |
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2 On the other hand, the linear voltage dependence of the fluorescence signal from the C74A mutant would be explained by contributions from any or all of these four mechanisms. However, W68L-GAT1 both undergoes conformational changes and displays fluorescence relaxations, only at positive potentials, suggesting that even the C74A signals arise from specific conformational changes. We have little data to suggest the nature of the conformational changes monitored by the C74A signals and will not consider them further.
1 Abbreviations used in this paper: GABA, -aminobutyric acid; MTSET, methanethiosulfonate-ethyltrimethylammonium; PMT, photomultiplier tube; TMRM, tetramethylrhodamine-5-maleimide; WT, wild type.
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Acknowledgements |
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We thank Mike Walsh for excellent technical assistance and Ehud Isacoff, Micah Siegel, and Yong-Xin Li for advice and reagents.
This work is supported by grants from the National Institutes of Health (NS-11756, DA-09121).
Submitted: 7 December 1999
Revised: 11 February 2000
Accepted: 22 February 2000
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