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Originally published In Press as doi:10.1074/jbc.M408038200 on July 20, 2004

J. Biol. Chem., Vol. 279, Issue 38, 39505-39512, September 17, 2004
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A Trimeric Quaternary Structure Is Conserved in Bacterial and Human Glutamate Transporters*

Sandra Gendreau{ddagger}§, Stephan Voswinkel§, Delany Torres-Salazar¶, Niklas Lang{ddagger}, Hannelore Heidtmann¶, Silvia Detro-Dassen{ddagger}, Günther Schmalzing{ddagger}||, Patricia Hidalgo¶||**, and Christoph Fahlke¶||**{ddagger}{ddagger}

From the Departments of {ddagger}Molecular Pharmacology and Physiology, Rheinisch-Westfälische Technische Hochschule Aachen, 52057 Aachen, Germany and the **Centro de Estudios Científicos, 509000 Valdivia, Chile

Received for publication, July 16, 2004


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Neuronal and glial glutamate transporters play a central role in the termination of synaptic transmission and in extracellular glutamate homeostasis in the mammalian central nervous system. They are known to be multimers; however, the number of subunits forming a functional transporter is controversial. We studied the subunit stoichiometry of two distantly related glutamate transporters, the human glial glutamate transporter hEAAT2 and a bacterial glutamate transporter from Escherichia coli, ecgltP. Using blue native polyacrylamide gel electrophoresis, analysis of concatenated transporters, and chemical cross-linking, we demonstrated that human and prokaryotic glutamate transporters expressed in Xenopus laevis oocytes or in mammalian cells are assembled as trimers composed of three identical subunits. In an inducible mammalian cell line expressing hEAAT2 the glutamate uptake currents correlate to the amount of trimeric transporters. Overexpression and purification of ecgltP in E. coli resulted in a homogenous population of trimeric transporters that were functional after reconstitution in lipid vesicles. Our results indicate that an evolutionarily conserved trimeric quaternary structure represents the sole native and functional state of glutamate transporters.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Glutamate is the major excitatory neurotransmitter in the mammalian central nervous system. After the release from glutamatergic nerve terminals, glial and neuronal glutamate transporters remove glutamate from the synaptic cleft to ensure low resting glutamate levels and to prevent neuronal damage by excessive glutamate receptor activation. Five mammalian glutamate transporters, EAAT1–5,1 have been cloned (1–5) and shown to belong to a large family of membrane transport proteins, the sodium dicarboxylate symporter family (6).

EAAT glutamate transporters sustain two fundamentally distinct transport mechanisms. They function as co-transporters of glutamate, sodium, potassium, and protons ions ("coupled transport") (7, 8) and as anion channels ("uncoupled transport") (4, 9–12). The molecular basis for these diverse transport functions is not understood. Eskandari et al. (13) suggested that the coupled and the uncoupled transport functions are mediated by distinct oligomeric states of the same protein subunit; i.e. a multimeric assembly conducts anions, while a single subunit suffices for coupled glutamate transport. EAAT transporters are known to be multimers (14), and a pentameric assembly has been proposed based on results obtained with freeze-fracture electron microscopy (13). However, freeze-fracture electron microscopy is only suitable to determine the subunit stoichiometry of membrane proteins with known transmembrane topology (15), a property that has not yet been established for glutamate transporters (16).

Here we determined the subunit stoichiometry of two distantly related glutamate transporters, the human glial glutamate transporter hEAAT2 (2) and the bacterial glutamate transporter ecgltP from Escherichia coli (17). The results from a variety of experimental approaches indicate that the two transporters assemble as homotrimers demonstrating an evolutionarily conserved trimeric quaternary structure of glutamate transporters.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Expression of His6-tagged Polypeptides in Xenopus Oocytes and in Mammalian Cells—A pTLN2-hEAAT2 plasmid (18) was modified by adding a cDNA fragment encoding six histidine residues either NH2-(HisNTEAAT2) or COOH-terminal (HisCTEAAT2) to the hEAAT2 coding region by PCR. A cDNA fragment encoding a His-tagged ecgltP was amplified by PCR from genomic E. coli DNA and subcloned into a pGEMHE vector using BamHI and HindIII restriction sites. To generate the ecgltP-ecgltP concatameric construct (pGEMHE-ecgltP-ecgltP), an NH2-terminal His-tagged coding region of ecgltP was linked to a non-tagged ecgltP with a cDNA sequence encoding a 20-amino acid linker sequence (SPLHPGLYPYDVPDYAISAV) in a single open reading frame. Mutations were inserted using the QuikChangeTM site-directed mutagenesis kit (Stratagene) and confirmed by sequencing. Transcription of cRNAs and handling of oocytes was performed as described previously (19).

To generate an inducible stable cell line, a cDNA fragment encoding an NH2-terminal His-tagged hEAAT2 was subcloned into the pcDNA5/FRT/To vector (Invitrogen). Flp-In T-Rex 293 cells (Invitrogen) were co-transfected with pcDNA5/FRT/To-HisNTEAAT2 and pOG44 (Invitrogen) using the calcium phosphate method, and oligoclonal cell lines were obtained by selection for the antibiotic hygromycin (Invitrogen). After 28 days, hygromycin-resistant clones were picked and tested for uptake of radioactive L-[3H]glutamate after 24 h of incubation with 1 µg/ml tetracycline. Six cell lines were positive, and one was used for the experiments reported here.

Electrophysiological Examination of Injected Xenopus Oocytes and Stably Transfected Mammalian Cells—EAAT-associated currents in oocytes were recorded by two-electrode voltage clamp using a CA1 amplifier (Dagan, Minneapolis, MN). Oocytes were held at –30 mV, and currents elicited by 200-ms voltage steps between –130 and +40 mV were filtered at 2 kHz and digitized with a sampling rate of 10 kHz using a Digidata AD/DA converter (Axon Instruments, Union City, CA). The standard external solution contained 96 mM NaCl, 4 mM KCl, 0.3 mM CaCl2, 1 mM MgCl2, 5 mM HEPES, pH 7.4, and the glutamate-containing solution was supplied with 0.5 mM L-glutamate. Anion currents were determined after exchanging the external solution to 96 mM NaSCN, 4 mM KCl, 0.3 mM CaCl2, 1 mM MgCl2, 5 mM HEPES, pH 7.4, in the absence and presence of 0.5 mM external glutamate without any current subtraction procedure (12). Anion currents were normalized by dividing current amplitudes by the Glu/Na+/H+/K+ uptake current amplitude measured at –140 mV. Glutamate uptake currents in mammalian cells were measured through standard whole-cell patch clamp recordings using an Axopatch 200B (Axon Instruments) amplifier as described previously (12). Currents were filtered at 5 kHz and digitized with a sampling rate of 50 kHz using a Digidata (Axon Instruments). Cells were clamped to 0 mV for at least 2 s between test sweeps. The intracellular solution contained 115 mM KCl, 2 mM MgCl2, 5 mM EGTA, 10 mM HEPES, pH 7.4, and the extracellular solution contained 140 mM NaCl, 4 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 5 mM HEPES, pH 7.4. To elicit glutamate transport-associated currents, cells were moved into a stream of an external solution supplemented with 0.5 mM L-glutamate. The glutamate uptake current was determined as the difference between the current amplitude in the presence and in the absence of glutamate measured at a test step to –175 mV. Data were analyzed using pClamp (Axon Instruments) and SigmaPlot (Jandel Scientific, San Rafael, CA) programs.

Purification of [35S]Methionine-labeled Protein Form Xenopus Oocytes and Mammalian Cells—cRNA-injected and non-injected control oocytes were incubated for the indicated time with RevidueTM L-[35S]methionine (>37 TBq/mmol, Amersham Biosciences) at ~25 MBq/ml (~0.1 MBq/oocyte) in frog Ringer's solution at 19 °C for metabolic labeling. Either immediately after the pulse or after an additional chase period, the radiolabeled oocytes were extracted with digitonin (1.0%) in 0.1 M sodium phosphate buffer, pH 7.4. His-tagged proteins were isolated by Ni2+-NTA-agarose (Qiagen, Köln, Germany) chromatography as detailed previously (19) with the modification that iodoacetamide was routinely included at 10 mM and 1 mM in the lysis and washing buffers (20).

Flp-In T-Rex HEK293 cells stably expressing the His-EAAT2 transporter were cultured at a range of tetracycline concentrations (0–1 µg/ml) for 24 h at 37 °C, then starved for 30 min in methionine- and serum-free minimum Dulbecco's modified Eagle's medium, and subsequently metabolically labeled with RevidueTM L-[35S]methionine at ~4 MBq/ml in methionine- and serum-free minimum Dulbecco's modified Eagle's medium for 2 h at 37 °C. His-EAAT2 was natively purified from digitonin extracts of these cells by Ni2+-NTA chromatography as described above.

Chemical Cross-linking—His-tagged proteins bound to Ni2+-NTA beads (packed volume, ~15 µl) were washed twice with imidazole-free sodium phosphate buffer (pH 8.0) supplemented with 0.2% digitonin. The Ni2+-NTA beads were resuspended in 50 µl of 0.2 M triethanolamine/HCl (pH 8.5), 0.5% digitonin. The cross-linking reaction was initiated by adding dimethyl adipimidate (DMA·2HCl, Pierce) or dimethyl suberimidate (DMS·2HCl, Pierce) from a freshly prepared stock solution in distilled water or Me2SO, respectively. After 60 min at the indicated temperature, the cross-linking reaction was terminated by washing the Ni2+-NTA-agarose beads once with imidazole-free sodium phosphate buffer, 0.2% digitonin. Bound protein was released from the beads with the non-denaturing elution buffer.

SDS-PAGE and Blue Native (BN)-PAGE Analysis—[35S]Methionine-labeled proteins were denatured for 10 min at 56 °C with SDS sample buffer containing 20 mM dithiothreitol and electrophoresed in parallel with 14C-labeled molecular mass markers (Rainbow, Amersham Biosciences) on linear SDS-polyacrylamide gels. To investigate the glycosylation status of the proteins, samples were treated for 1–2 h with either endoglycosidase H (Endo H) or PNGase F (New England Biolabs, Beverly, MA) in the presence of reducing SDS sample buffer and 1% (w/v) Nonidet P-40 to counteract SDS inactivation of PNGase F. BN-PAGE was performed as described by Nicke et al. (19) immediately after protein purification. Molecular mass markers (Combithek II, Roche Applied Science) were run in two different lanes of the gel and subsequently visualized by Coomassie and silver staining. Gels containing purified ecgltP were stained with silver. Radioactive proteins were visualized by autoradiography using BioMax MS films (Eastman Kodak Co.) at –70 °C. Both SDS- and BN-polyacrylamide gels were fixed and dried. For quantification, the dried gels were exposed to a Phosphor-Imager screen and scanned using a Storm 820 PhosphorImager (Amersham Biosciences). Individual bands were quantified with the ImageQuant software.

Expression, Purification, and Reconstitution of ecgltP—The pASK-ecgltP construct was generated by subcloning the cDNA encoding ecgltP into a pASK-IBA5 (IBA, Göttingen, Germany) vector to add an NH2-terminal strep tag. Transformed BL-21 (DE-3) E. coli bacteria were induced with 200 µg/l anhydrotetracycline at 37 °C for 2 h, harvested by centrifugation, and stored at –80 °C until use. Membranes were collected by centrifugation at 100,000 x g for 60 min at 4 °C in a Beckman 45 Ti rotor. The protein was extracted by solubilization in 15 mM dodecylmaltoside (DDM) for 2 h at 4 °C followed by centrifugation at 100,000 x g for 60 min at 4 °C. ecgltP was purified in one step by strep-tactinTM (IBA) affinity chromatography according to the manufacturer's instruction manual. Western blots were carried out using a strep-tag AP detection kitTM (IBA) following the manufacturer's instructions.

Strep-ecgltP was reconstituted into liposomes using standard methods (21, 22). Proteoliposomes were resuspended in buffer A (20 mM Mes, 100 mM potassium acetate, 5 mM MgSO4, pH 6) and incubated for 2 h on ice. The uptake was initiated by diluting 25 µl of the proteoliposomes in 650 µl of buffer B (120 mM Mes, 100 mM NaOH, 5 mM MgSO4, 2 µM L-[3H]glutamate, pH 6) at room temperature (22). Control experiments were performed with vesicles without ecgltP or by diluting the proteoliposomes in 650 µl of buffer A supplemented with 2 µM L-[3H]glutamate. 100-µl probes were taken after various time periods and then poured into a 10-fold excess of ice-cold 0.1 M LiCl solution, followed by immediate filtration over cellulose nitrate filters. After washing with 0.1 M LiCl, filters were assayed for radioactivity.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
His-tagged hEAAT2 Transporters Exhibit Unaltered Functional Properties—We added an NH2- or a COOH-terminal hexahistidine tag to hEAAT2 and ecgltP to purify the transporters by a single Ni2+-metal affinity chromatography step. To test whether these sequence alterations affect transport functions, WT and His-tagged transporters were expressed in Xenopus oocytes, and currents were examined with a two-electrode voltage clamp. We performed experiments in two distinct external anion compositions to separate the current components of the stoichiometrically coupled transport of glutamate Na+, H+, and K+ from the pore-mediated anion conductance. In Cl–-based external solution, the conductance of the hEAAT2-associated anion channel is very small (9), and the Glu/Na+/H+/K+ current amplitudes can therefore be approximated to the difference between currents in the presence and absence of external glutamate substrate. The so-calculated current amplitudes are similar in magnitude and voltage dependence for WT and His-tagged hEAAT2 (Fig. 1A). In a SCN–-based external solution, hEAAT2 exhibited anion currents that largely exceeded the Glu/Na+/H+/K+ current component allowing to directly measure anion currents (12). For all tested hEAAT2 constructs, a comparable constitutive anion current was observed in the absence of glutamate (Fig. 1B) that was about 2-fold increased by external glutamate (12). We conclude that the addition of the His tag does not alter the coupled and the uncoupled current amplitudes of hEAAT2. Neither glutamate-induced inward currents nor glutamate-induced increases of anion currents were observed in oocytes injected with WT or His-tagged ecgltP cRNA.



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  		FIG. 1.
Basic functional and biochemical characterization of glutamate transporters expressed in Xenopus oocytes. A, voltage dependence of the absolute Glu/Na+/H+/K+ uptake current component for WT EAAT2, HisNT EAAT2, and HisCTEAAT2. B, voltage dependence of the normalized anion current amplitude in the absence (filled symbols) and in the presence (open symbols) of 0.5 mM L-glutamate. Results are given as the mean ± S.E., n > 7. C, EAAT2 and ecgltP transporters purified by Ni2+-NTA chromatography after a 4-h [35S]methionine pulse from Xenopus oocytes were resolved by reducing SDS-PAGE (10% acrylamide) in parallel with 14C-labeled molecular mass markers. D, plot of molecular masses of the hEAAT2 and ecgltP polypeptides determined by SDS-PAGE at various polyacrylamide concentrations versus the polyacrylamide concentration. E, SDS-PAGE analysis of WT EAAT2 and N206Q and N216Q EAAT2 purified from Xenopus oocytes after a 4-h [35S]methionine pulse. F, SDS-PAGE analysis after a 36-h chase interval subsequent to the 4-h [35S]methionine pulse reveals the additional presence of an Endo H-resistant EAAT2 polypeptide (¶), which can be reduced to the protein core by PNGase F. *, Endo H-sensitive EAAT2 polypeptide.

 
SDS-PAGE Analysis and Glycosylation of Glutamate Transporters Heterologously Expressed in Xenopus Oocytes—His-tagged hEAAT2 and ecgltP transporters expressed in Xenopus oocytes were metabolically labeled with [35S]methionine, extracted with 1% (w/v) digitonin, and purified by metal affinity chromatography. Both proteins expressed at high levels in Xenopus oocytes and were metabolically stable during a sustained chase. When denatured by SDS and resolved by reducing SDS-PAGE, the hEAAT2 and the ecgltP polypeptides both migrated at 10–20% lower masses than calculated from the amino acid sequences (48 kDa for ecgltP and 63 kDa for hEAAT2) (Fig. 1C). However, a Ferguson analysis determining the apparent molecular mass in SDS-polyacrylamide gels for several acrylamide concentrations (23) yielded molecular masses of 46 kDa for the His-tagged ecgltP and 69 kDa for His-hEAAT2 when extrapolated to high acrylamide concentration (Fig. 1D) demonstrating that the observed differences between the apparent and the calculated masses are caused by anomalous migration.

Membrane proteins are often N-glycosylated when expressed in eukaryotic cells. As oligosaccharide side chains are sequentially processed from a high mannose form in the endoplasmic reticulum (ER) to the complex-glycosylated form in the Golgi apparatus, the presence of complex oligosaccharides can be used to monitor the efficiency of the exit of the protein from the ER. The hEAAT2 sequence encompasses two glycosylation sequences, 206NATS and 216NETV, which are both located on the predicted large ectodomain (residues 143–239) between transmembrane regions TM3 and TM4 (16). Complete deglycosylation of newly synthesized hEAAT2 polypeptide in Xenopus oocytes during a 4-h pulse period resulted in a 3-kDa decrease of the molecular mass (Fig. 1E, lanes 1–2), corresponding to the mass of one single N-glycan. Moreover, glutamine substitution of only one of the two asparagine residues resulted in polypeptides migrating at the same position as the WT hEAAT2 (Fig. 1E, lanes 3 and 5) suggesting that one of the two glycosylation sites remains unused in WT EAAT2 presumably because of the small distance of only 10 amino acids. No mass shift by PNGase F was observed when the asparagines residues of both N-glycosylation sequons were replaced by glutamine (Fig. 1E, lanes 7 and 8).

Posttranslational processes during a 36-h chase interval led to the occurrence of a prominent broad band (75–95 kDa) well above that of the core-glycosylated hEAAT2 polypeptide (~60 kDa) (Fig. 1E). This band could be reduced to the hEAAT2 apoprotein by incubation with PNGase F (~57 kDa in Fig. 1E, lane 3), but not with Endo H. Resistance to Endo H treatment distinguishes high mannose from complex oligosaccharides and attributes the higher molecular weight band to the mature complex-glycosylated hEAAT2 polypeptide. Quantification by phosphorimaging analysis demonstrated that 66% of the total hEAAT2 protein leaves the ER within the 36-h chase interval. We conclude that the majority of hEAAT2 subunits are in a mature state located in post-ER compartments including the plasma membrane after the chase interval. The prokaryotic ecgltP polypeptide does not exhibit glycosylation sites precluding such an analysis for this particular transporter.

hEAAT2 Transporters Migrate as Trimers in Blue Native Polyacrylamide Gels—BN-PAGE analysis (24, 25) permits gel electrophoresis under non-denaturing conditions and thus the determination of the oligomeric structure of proteins (19, 26, 27). hEAAT2 transporters expressed in Xenopus oocytes migrated predominantly as a single band of ~200 kDa (Fig. 2A) in BN-polyacrylamide gels when compared with the defined membrane protein complexes generated by partial denaturing of the homopentameric {alpha}1 GlyR (26) or the homotrimeric P2X1 receptor (19). These molecular masses are well above those of the respective monomers suggesting that hEAAT2 transporters exist exclusively as an homogenous population of multimers in Xenopus oocytes.



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  		FIG. 2.
Oligomeric state of EAAT2 transporters in Xenopus oocytes and mammalian cells determined by BN-PAGE. A, autoradiography of a BN-polyacrylamide gel containing EAAT2 purified from Xenopus oocytes. Samples were treated as indicated to induce dissociation into lower order intermediates including monomers. B, quantitative profiles of the gel lanes shown in A were obtained by PhosphorImager analysis and are marked by the same lane numbers. The origin of the abscissa corresponds to the top of the polyacrylamide gel. Numbers specify the oligomeric state of the corresponding protein peak. The five bands that became visible upon partial denaturing of the GlyR are consistent with its pentameric state. C, autoradiography of a BN-polyacrylamide gel containing EAAT2 transporter isolated from Flp-In T-Rex HEK293 cells. Various EAAT2 protein levels were adjusted by inducing the cells with the indicated concentrations of tetracycline (Tet). D, variation of normalized trimeric EAAT2 protein levels (bars, means from two experiments) and of mean glutamate uptake current amplitudes (symbols, means ± S.E. from 3–8 cells) with the tetracycline concentration 24 h before the experiment. The solid line represents a fit of the tetracycline concentration dependence of the amount of the EAAT2 trimer with a Michaelis-Menten function.

 
The electrophoretic mobility of proteins is biased by dye binding and protein shape to an unclear extent, and therefore the exact number of monomers incorporated/protein complex cannot be readily deduced from mass estimates alone. A reliable approach to determine the number of polypeptide chains incorporated in one transporter complex is to dissociate protein complexes into lower order intermediates by weakening non-covalent subunit interactions by heat or low concentrations of SDS (19, 26). For hEAAT2, a 1-h incubation at 56 °C both in the presence (Fig. 2A, lane 4) and absence (lane 5) of Coomassie dye generated a ladder-like pattern of three protein bands. Incubation in the additional presence of increasing concentrations of SDS (Fig. 2A, lanes 6–10) led to a gradual disappearance of the ~200-kDa protein and an enhanced appearance of two additional proteins with masses of ~130 and ~65 kDa. At ≥0.05% SDS, the ~65-kDa band became the predominant one (Fig. 2A, lanes 9 and 10). Fig. 2B shows PhosphorImager analysis of the gel shown in Fig. 2A. All dissociating conditions caused the appearance of a total of three bands with masses corresponding to the assembly of three, two, and one unit with the monomer becoming the dominant species at increasing SDS concentrations (Fig. 2A). The complex-glycosylated hEAAT2 transporter also exhibits a trimeric structure as revealed by BN-PAGE (data not shown). The virtual absence of aggregated proteins and other multimerization states (Fig. 2) indicates that trimerization of hEAAT2 monomers occurs efficiently in oocytes during or shortly after synthesis of the individual subunits. A productive assembly process is further illustrated by the efficient exit of the hEAAT2 transporters from the ER inferred from the acquisition of complex-type carbohydrates (see above).

Oligomerization of EAAT transporters might be affected by the cellular environment or by unphysiologically high expression levels. To address these possibilities, we generated an inducible mammalian cell line that stably expresses NH2-terminal His-tagged hEAAT2 (Fig. 2, C and D). This cell line allowed us to study oligomerization of hEAAT2 at different protein expression levels by simply varying the tetracycline concentration added to the culture medium 24 h prior to the experiment. hEAAT2 transporters expressed in mammalian cells were metabolically labeled with [35S]methionine, extracted with 1% (w/v) digitonin, and purified by metal affinity chromatography. Incubation of the cells with inducing tetracycline concentration between 0 and 1000 ng/ml resulted in pronounced changes in the amount of the purified hEAAT2 protein and glutamate uptake currents. BN-PAGE analysis demonstrates that the hEAAT2 protein exists exclusively in a trimeric state over a broad range of expression levels in HEK293 cells and that monomers are entirely absent (Fig. 2C). Fig. 2D shows a plot of the intensity of the trimeric hEAAT2 band (bars) and the mean glutamate uptake currents (symbols) versus the tetracycline concentration. The solid line represents a fit of these data with a Michaelis-Menten relationship (Fig. 2D). Glutamate transport changes with the same dependence on the tetracycline concentration as the amount of purified trimeric hEAAT2 protein. Uptake and quantity of hEAAT2 trimers are highly correlated indicating that glutamate uptake is entirely mediated by hEAAT2 trimers. We conclude that a trimeric architecture of the hEAAT2 transporter is neither a result of overexpression nor of expression in non-mammalian host cells but represents the sole native and functional state of the hEAAT2 transporter.

ecgltP Transporters Migrate as Trimers in Blue Native Polyacrylamide Gels—The ecgltP protein migrated predominantly at ~150 kDa in BN-PAGE (Fig. 3A, lane 3). In addition, a slower migrating distinct band, presumably an ecgltP hexamer, and an amorphous mass of proteins, most likely ecgltP aggregates, were visible. Oligomerization of ecgltP thus appears to be less complete than that of hEAAT2. However, quantification of the various ecgltP forms by PhosphorImager analysis indicated that the 150-kDa band is the most prominent one (Fig. 3B). A 1-h incubation at 56 °C both in the absence (not shown) as well as in the presence of Coomassie dye (Fig. 3A, lane 4) and denaturing with increasing concentrations of SDS (lanes 5–7) led to the dissociation of the 150-kDa band into the dimeric and monomeric ecgltP species.



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  		FIG. 3.
Oligomeric state of ecgltP transporters in Xenopus oocytes determined by BN-PAGE. A, autoradiography of a BN-polyacrylamide gel containing ecgltP. Samples were treated as indicated to induce dissociation into lower order intermediates including monomers. B, quantitative profiles of the gel lanes shown in A were obtained by PhosphorImager analysis and are marked by the same lane numbers. The origin of the abscissa corresponds to the top of the polyacrylamide gel. Numbers specify the oligomeric state of the corresponding protein peak. C, reducing SDS-PAGE analysis of isolated ecgltP-ecgltP concatamers expressed in Xenopus oocytes. During the chase interval two polypeptides appeared (lane 3) with masses corresponding to the ecgltP monomer that are barely detectable after the pulse (lane 2). Co, non-injected control oocytes. D, oligomeric state of the concatenated ecgltP dimer determined by BN-PAGE. The oligomeric state attained by expression of monomeric ecgltP is shown for comparison. Dissociation into lower order intermediates was induced by partial denaturing with SDS as indicated. The monomeric by-products formed during the chase interval (*) assemble into non-covalently linked trimeric ecgltP proteins (#). E, schematic showing the assembly of full-length concatenated ecgltP dimers and monomeric by-products.

 
Our results demonstrate that both hEAAT2 and ecgltP glutamate transporters are assembled as trimers from a minimal unit that migrates close to the expected molecular mass of the monomer in BN-PAGE. To rule out the possibility that the lowest molecular band corresponds to an unusually stable dimer and correspondingly the intermediate and higher molecular mass bands to tetramers and hexamers, we engineered a concatenated cDNA construct for one of the transporters (ecgltP-ecgltP) by linking two ecgltP coding regions in a single open reading frame. By reducing SDS-PAGE the ecgltP-ecgltP polypeptide was resolved as a 74-kDa polypeptide (Fig. 3C, lane 2), i.e. twice the mass of the apparent molecular mass of 37 kDa for the ecgltP monomer (lane 4). In BN-PAGE two major bands were observed (Fig. 3D, lane 1), and dissociating treatment with SDS led to the appearance of a third major band of ecgltP-ecgltP (lane 2) that was not further dissociable and that migrated at approximately the same position as the non-covalently associated ecgltP dimer, (ecgltP)2 (lane 10). These results show that the intermediate molecular weight band dissociated from the ecgltP protein indeed corresponds to the dimeric form and, accordingly, the lower and higher molecular bands to the monomeric and trimeric forms, respectively.

Under non-denaturing conditions (Fig. 3D, lane 1) the two bands corresponding to (ecgltP-ecgltP)2 and (ecgltP-ecgltP)3 are prominent indicating that both conformations are stable and occur with comparable probability. This observation further corroborates a trimeric ecgltP structure that predicts that two oligomeric populations assemble from dimeric concatamers (Fig. 3E), i.e. an assembly of two concatamers, one of them providing two subunits ((ecgltP-ecgltP)2), or the association of three dimeric concatamers, each of them contributing one subunit ((ecgltP-ecgltP)3) to the trimer interface.

An additional faint band (indicated by (ecgltP)3 in lane 1 of Fig. 3D) migrating at exactly the same position as the ecgltP trimer assembled from three ecgltP monomers, (ecgltP)3 (lane 9), became more abundant after an additional chase period (Fig. 3D, lane 5). The occurrence of this oligomeric complex is most likely because of a proteolytic digestion of ecgltP concatamers into monomers (28) giving rise to a trimer from proteolysis-derived monomers (Fig. 3E). Upon treatment with SDS, this protein dissociated into a polypeptide migrating at the same position as the ecgltP monomer (Fig. 3D, lanes 6–8). This is confirmed in SDS-PAGE analysis where two by-products besides the full-length concatamer are observed after a chase period (Fig. 3C, lane 3), one migrating virtually at the same position as the ecgltP monomer and a second one with a 1–2-kDa larger mass probably corresponding to the ecgltP monomer plus the 18-residue linker. The finding that two distinct concatameric constructs, the one used in this study and the one of Nicke et al. (28), are both subject to proteolytic digestion in Xenopus oocytes demonstrates the limitations of using tandem constructs to study multimeric proteins with defined composition in this expression system.

Cross-linking of hEAAT2 or ecgltP Generates Covalently Bound Dimers and Trimers—We next used protein cross-linking to study intermolecular interactions within glutamate transporter subunits. Two homobifunctional imidoester reagents, DMA and DMS, were tested for their ability to covalently link transporter molecules extracted from Xenopus oocytes. At 22 °C as well as 37 °C, DMA cross-linked ecgltP to dimers (Fig. 4A, lanes 2–4) and to trimers at higher reagent concentrations (lanes 3–5). DMS differs from DMA by a slightly longer spacer arm (11 Å versus 8.6 Å) and was more efficient in cross-linking ecgltP to dimers and trimers at both 22 °C (Fig. 4B, lanes 2–4) and 37 °C (lanes 5–7). DMS also cross-linked hEAAT2 transporter subunits to dimers and to trimers preferentially at higher concentrations (Fig. 4C). Adducts larger than trimers were neither observed with ecgltP nor with hEAAT2 transporters, corroborating the trimeric quaternary structure determined by BN-PAGE analysis.



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  		FIG. 4.
Cross-linking of digitonin-solubilized purified glutamate transporters. ecgltP transporters (A and B) and hEAAT2 transporters (C) were purified from [35S]methionine-labeled oocytes and incubated with cross-linkers as indicated while still bound to Ni2+-NTA beads. After elution with non-denaturing buffer, samples were supplemented with SDS sample buffer and 20 mM dithiothreitol and resolved by SDS-PAGE (4–10% acrylamide gradient gel) followed by autoradiography. Numbered arrows indicate positions of monomers, dimers, and trimers.

 
Purification and Characterization of ecgltP—ecgltP tagged with a strep tag (strep-ecgltP) was expressed in E. coli and purified after detergent extraction by one affinity chromatography step using a strep-tactinTM superflow column. Reducing SDS-PAGE analysis of the purified protein demonstrated a single band with an apparent molecular mass of 37 kDa close to the metabolically labeled ecgltP expressed in Xenopus oocytes (Fig. 5A). The identity of the ecgltP was verified by Western blot (Fig. 5B) and amino acid analysis (data not shown). The homogeneity of the purified protein was evaluated by fast protein liquid chromatography-attached size exclusion chromatography using a Superdex 200 column preequilibrated with buffer containing 1 mM DDM (Fig. 5C). The strep-ecgltP eluted predominantly as a single symmetrical peak indicating that the majority of the purified protein (>95%) exists in one oligomeric conformation. The small peak eluting after the main fraction most likely corresponds to ecgltP monomers. Varying the DDM detergent concentration from 1 to 10 mM (corresponding to ~1–10x the critical micellar concentration) had no effect on the oligomeric state as judged by size exclusion chromatography, indicating that conformation of the purified ecgltP is not because of artificial association promoted by a low detergent concentration.



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  		FIG. 5.
Biochemical and functional characterization of recombinant ecgltP produced in E. coli. A, reducing SDS-PAGE of strep-tagged ecgltP purified from E. coli. Lane 1, protein markers; lane 2, ecgltP eluted from a strep-tactinTM column. MW, molecular weight. B, Western blot of strep-tagged ecgltP detected with strep-tactinTM alkaline phosphatase conjugate. C, elution profile monitored at 280 nm from size exclusion chromatography using a Superdex 200 column (24-ml bed volume) equilibrated with 50 mM phosphate buffer, 300 mM NaCl, 1 mM EDTA, 1 mM DDM, pH 7.0. The arrows show the void volume (Vo) and the elution volume of albumin (Alb.) (67 kDa). D, BN-PAGE (4–16% acrylamide) of purified strep-tagged ecgltP. The protein bands were visualized by silver staining. The numbers indicate the molecular weights of standards in kDa. E, time course of the accumulation of L-[3H]glutamate in vesicles reconstituted with 10 µg of ecgltP/mg of lipid ({blacksquare}, means ± S.E., n = 4) or without protein (, n = 1).

 
We employed BN-PAGE analysis to determine the quaternary structure of the purified ecgltP. Strep-ecgltP migrates in BN-PAGE (Fig. 5D) mostly as a prominent band at the same position observed for the oocyte-expressed ecgltP (lanes 1 and 2). Treatment with heat or SDS causes a dissociation into lower order assemblies (Fig. 5D, lanes 3–5) resembling the one into dimers and monomers of the oocyte-expressed transporter. Thus, the ecgltP purified from E. coli exhibits the same oligomeric structure as ecgltP and hEAAT2 expressed in Xenopus oocytes and mammalian cells.

The purified strep-ecgltP protein is functionally active when reconstituted into liposomes. Proteoliposomes containing strep-ecgltP were loaded with a potassium acetate solution, and the addition of L-[3H]glutamate to a sodium-containing external solution enabled glutamate uptake into the vesicles. Fig. 5E shows the time course of radioactive glutamate accumulation by proteoliposomes containing purified ecgltP transporters. No glutamate uptake was observed when the intra- and extravesicular solution had the same salt content or in control vesicles without protein.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Multimerization of EAAT transporters was first reported by Haugeto et al. (14). Using SDS-PAGE analysis and cross-linking of native rat brain or recombinant EAAT2 and EAAT3 transporters expressed in HeLa cells, dimeric and trimeric assemblies were observed. These results demonstrated that EAAT transporters are multimers in native cells; however, the authors did not deduce the subunit stoichiometry from these experiments, as a larger number of subunits/single transporter could not be excluded. Later, Eskandari et al. (13) determined a cross-sectional area/unitary EAAT3 transporter that corresponds to a number of 35 transmembrane helices by freezefracture electron microscopy. Assuming seven transmembrane helices/subunit, the authors assigned five subunits/transporter. In contrast to this pentameric assembly, a trimeric stoichiometry was reported recently for two prokaryotic glutamate transporters based on cross-linking and in-line laser light scattering, refractive index, and ultraviolet absorption measurements (29) raising the possibility that eukaryotic and prokaryotic transporters might exhibit distinct subunit stoichiometries.

We here demonstrate that this is not the case. BN-PAGE analysis and chemical cross-linking of transporters heterologously expressed in Xenopus oocytes revealed a conserved trimeric stoichiometry for prokaryotic and eukaryotic transporters. Expression of hEAAT2 in an inducible mammalian cell line demonstrated that trimerization is independent of the expression level and occurs in mammalian as well as in amphibian cells. No other oligomerization state was observed at all tested expression levels. Moreover, glutamate transport and hEAAT2 trimer quantity are highly correlated indicating that the trimer is the functional unit of glutamate transport under physiological conditions.

The experiments with single and concatenated ecgltP polypeptide units (Figs. 2 and 3) excluded a higher multimerization state than three. Under non-denaturing conditions, we did not observe monomers and dimers besides the trimeric state, and this result refutes the pentameric assembly suggested by Eskandari et al. (13). At present, we cannot explain the distinct outcome of biochemical and microscopic approaches in determining the glutamate transporter stoichiometry. A possible reason for the observed differences is the currently unresolved transmembrane topology for EAAT transporters (16). A larger number of transmembrane helices/single glutamate transporter subunit would decrease the number of subunits necessary for the observed 35 transmembrane helices and might thus resolve the inconsistencies between our results and those of Eskandari et al. (13). The results of our study and of earlier studies are best explained by a trimeric subunit stoichiometry general to prokaryotic and eukaryotic glutamate transporters.

The conserved trimeric structure of glutamate transporters is in clear contrast to the situation observed in another family of neurotransmitter transporters, the SLC6 transporter family including {gamma}-aminobutyric acid, dopamine, serotonin, norepinephrine, solutes, and amino acid transporters (30). A prokaryotic {gamma}-aminobutyric acid transporter (31) and a mammalian glycine transporter (27) were shown to form monomers. There is evidence for the formation of tetrameric dopamine transporters (32), and serotonin transporters were shown to be dimers or tetramers (33–35). Similar to EAAT transporters, SLC6 transporters exhibit not only a stoichiometric co-transport (36, 37) but also current components that appear to be conducted by permeation pathways similar to that of ion channels (38–42). The channel-like activity was reported to become more apparent with increasing expression levels of rat and human serotonin transporters heterologously expressed in Xenopus oocytes (43). Ramsey and DeFelice (43) interpreted this result as evidence for an endogenous regulatory protein that is necessary for the carrier transport mode. At low heterologous expression levels, virtually all serotonin transporters will contain this regulatory protein and thus function as carriers, while the endogenous protein is not sufficiently available at high expression levels resulting in homomultimeric serotonin transporters that function as channels. In support of this hypothesis, syntaxin A was identified recently as the first interacting protein that promotes stoichiometrically coupled serotonin transport (44). An alternative explanation might be the existence of multiple homo-oligomerization states with different functions. At low expression levels, serotonin transporters might be formed by a smaller number of subunits than at high levels thus explaining the observed dependence of transporter function and protein expression (43). This hypothesis would also account for the multiple experimentally determined subunit stoichiometries within the SLC6 family.

Glutamate transporters exhibit only a single oligomeric state formed by the assembly of three identical subunits. Our reconstitution experiments (Fig. 5) demonstrate that homotrimeric ecgltP is able to sustain coupled transport. The situation is not as clear for the pore-mediated uncoupled transport; however, several experimental results support the notion that accessory subunits are not involved in anion conduction by glutamate transporters. Coupled and uncoupled current components of heterologously expressed EAAT transporters differ little between distinct expression systems (9, 45), and anion pores associated with distinct EAAT isoforms exhibit distinct pore properties (12). In SDS- or BN-polyacrylamide gels, only a single protein could be observed after expression and purification of hEAAT2 and ecgltP in Xenopus oocytes as well as in mammalian cells. The trimeric state of glutamate transporters is attained immediately after biosynthesis and appears to be the only significant oligomeric state in homologous and heterologous systems. These results taken together indicate that one quaternary structure of glutamate transporters supports two distinct transport processes. At present, we cannot distinguish whether the three subunits contribute to the formation of a central anion conduction pathway or a central carrier domain mediating coupled transport or whether each subunit can mediate both transport modes.


   FOOTNOTES
 
* This work was supported by Deutsche Forschungsgemeinschaft Grants FOR450/1 (TP1 and TP4 to P. H. and Ch. F., respectively) and SCHM 536/6-1 (to G. S.) and by a START grant of the medical faculty of the Rheinisch-Westfälische Technische Hochschule (to P. H.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

§ Both authors contributed equally to this work. Back

|| These authors contributed equally to this work. Back

{ddagger}{ddagger} To whom correspondence should be addressed: Dept. of Physiology, RWTH Aachen, Pauwelsstr. 30, 52057 Aachen, Germany. Tel.: 49-241-80-888-10; Fax: 49-241-80-824-34; E-mail: chfahlke{at}physiology.rwth-aachen.de.

1 The abbreviations used are: EAAT, excitatory amino acid transporter; Ni2+-NTA, nickel-nitrilotriacetic acid; DMA, dimethyl adipimidate; DMS, dimethyl suberimidate; BN, blue native; PNGase F, glycopeptide N-glycosidase F; Endo H, endoglycosidase H; DDM, dodecylmaltoside; Mes, 4-morpholineethanesulfonic acid; WT, wild type; ER, endoplasmic reticulum; SLC, solute carrier. Back


   ACKNOWLEDGMENTS
 
We thank Dr. M. Hediger for providing the pTLN2-hEAAT2 expression construct; Dr. Benjamin Kaupp for the pGEMHE vector, insightful discussions, and critical reading of the manuscript; Drs. Simon Hebeisen, Nico Melzer, and Maike Warnstedt for help in generating the inducible EAAT2 cell line; and Barbara Poser for excellent technical assistance.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Storck, T., Schulte, S., Hofmann, K., and Stoffel, W. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 10955–10959[Abstract/Free Full Text]
  2. Pines, G., Danbolt, N. C., Bjoras, M., Zhang, Y., Bendahan, A., Eide, L., Koepsell, H., Storm-Mathisen, J., Seeberg, E., and Kanner, B. I. (1992) Nature 360, 464–467[CrossRef][Medline] [Order article via Infotrieve]
  3. Kanai, Y., and Hediger, M. A. (1992) Nature 360, 467–471[CrossRef][Medline] [Order article via Infotrieve]
  4. Fairman, W. A., Vandenberg, R. J., Arriza, J. L., Kavanaugh, M. P., and Amara, S. G. (1995) Nature 375, 599–603[CrossRef][Medline] [Order article via Infotrieve]
  5. Arriza, J. L., Eliasof, S., Kavanaugh, M. P., and Amara, S. G. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 4155–4160[Abstract/Free Full Text]
  6. Slotboom, D. J., Konings, W. N., and Lolkema, J. S. (1999) Microbiol. Mol. Biol. Rev. 63, 293–307[Abstract/Free Full Text]
  7. Levy, L. M., Warr, O., and Attwell, D. (1998) J. Neurosci. 18, 9620–9628[Abstract/Free Full Text]
  8. Zerangue, N., and Kavanaugh, M. P. (1996) Nature 383, 634–637[CrossRef][Medline] [Order article via Infotrieve]
  9. Wadiche, J. I., Amara, S. G., and Kavanaugh, M. P. (1995) Neuron 15, 721–728[CrossRef][Medline] [Order article via Infotrieve]
  10. Larsson, H. P., Picaud, S. A., Werblin, F. S., and Lecar, H. (1996) Biophys. J. 70, 733–742[Abstract/Free Full Text]
  11. Billups, B., Rossi, D., and Attwell, D. (1996) J. Neurosci. 16, 6722–6731[Abstract/Free Full Text]
  12. Melzer, N., Biela, A., and Fahlke, Ch. (2003) J. Biol. Chem. 278, 50112–50119[Abstract/Free Full Text]
  13. Eskandari, S., Kreman, M., Kavanaugh, M. P., Wright, E. M., and Zampighi, G. A. (2000) Proc. Natl. Acad. Sci. U. S. A. 97, 8641–8646[Abstract/Free Full Text]
  14. Haugeto, O., Ullensvang, K., Levy, L. M., Chaudhry, F. A., Honore, T., Nielsen, M., Lehre, K. P., and Danbolt, N. C. (1996) J. Biol. Chem. 271, 27715–27722[Abstract/Free Full Text]
  15. Eskandari, S., Wright, E. M., Kreman, M., Starace, D. M., and Zampighi, G. A. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 11235–11240[Abstract/Free Full Text]
  16. Danbolt, N. C. (2001) Prog. Neurobiol. 65, 1–105[CrossRef][Medline] [Order article via Infotrieve]
  17. Tolner, B., Poolman, B., Wallace, B., and Konings, W. N. (1992) J. Bacteriol. 174, 2391–2393[Abstract/Free Full Text]
  18. Trotti, D., Rolfs, A., Danbolt, N. C., Brown, R. H., Jr., and Hediger, M. A. (1999) Nat. Neurosci. 2, 427–433[CrossRef][Medline] [Order article via Infotrieve]
  19. Nicke, A., Baumert, H. G., Rettinger, J., Eichele, A., Lambrecht, G., Mutschler, E., and Schmalzing, G. (1998) EMBO J. 17, 3016–3028[CrossRef][Medline] [Order article via Infotrieve]
  20. Sadtler, S., Laube, B., Lashub, A., Nicke, A., Betz, H., and Schmalzing, G. (2003) J. Biol. Chem. 278, 16782–16790[Abstract/Free Full Text]
  21. Knol, J., Veenhoff, L., Liang, W. J., Henderson, P. J. F., Leblanc, G., and Poolman, B. (1996) J. Biol. Chem. 271, 15358–15366[Abstract/Free Full Text]
  22. Gaillard, I., Slotboom, D. J., Knol, J., Lolkema, J. S., and Poolman, B. (1996) Biochemistry 35, 6150–6156[CrossRef][Medline] [Order article via Infotrieve]
  23. Ferguson, K. A. (1964) Metabolism 13, 985–1002[Medline] [Order article via Infotrieve]
  24. Schägger, H., and von Jagow, G. (1991) Anal. Biochem. 199, 223–231[CrossRef][Medline] [Order article via Infotrieve]
  25. Schägger, H., Cramer, W. A., and von Jagow, G. (1994) Anal. Biochem. 217, 220–230[CrossRef][Medline] [Order article via Infotrieve]
  26. Griffon, N., Büttner, C., Nicke, A., Kuhse, J., Schmalzing, G., and Betz, H. (1999) EMBO J. 18, 4711–4721[CrossRef][Medline] [Order article via Infotrieve]
  27. Horiuchi, M., Nicke, A., Gomeza, J., Aschrafi, A., Schmalzing, G., and Betz, H. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 1448–1453[Abstract/Free Full Text]
  28. Nicke, A., Rettinger, J., and Schmalzing, G. (2003) Mol. Pharmacol. 63, 243–252[Abstract/Free Full Text]
  29. Yernool, D., Boudker, O., Folta-Stogniew, E., and Gouaux, E. (2003) Biochemistry 42, 12981–12988[CrossRef][Medline] [Order article via Infotrieve]
  30. Blakely, R. D., De Felice, L. J., and Hartzell, H. C. (1994) J. Exp. Biol. 196, 263–281[Abstract/Free Full Text]
  31. Li, X. D., Villa, A., Gownley, C., Kim, M. J., Song, J., Auer, M., and Wang, D. N. (2001) FEBS Lett. 494, 165–169[CrossRef][Medline] [Order article via Infotrieve]
  32. Hastrup, H., Sen, N., and Javitch, J. A. (2003) J. Biol. Chem. 278, 45045–45048[Abstract/Free Full Text]
  33. Schmid, J. A., Scholze, P., Kudlacek, O., Freissmuth, M., Singer, E. A., and Sitte, H. H. (2001) J. Biol. Chem. 276, 3805–3810[Abstract/Free Full Text]
  34. Jess, U., Betz, H., and Schloss, P. (1996) FEBS Lett. 394, 44–46[CrossRef][Medline] [Order article via Infotrieve]
  35. Kilic, F., and Rudnick, G. (2000) Proc. Natl. Acad. Sci. U. S. A. 97, 3106–3111[Abstract/Free Full Text]
  36. Hilgemann, D. W., and Lu, C. C. (1999) J. Gen. Physiol. 114, 459–476[Abstract/Free Full Text]
  37. Galli, A., Jayanthi, L. D., Ramsey, I. S., Miller, J. W., Fremeau, R. T., Jr., and DeFelice, L. J. (1999) J. Neurosci. 19, 6290–6297[Abstract/Free Full Text]
  38. Mager, S., Min, C., Henry, D. J., Chavkin, C., Hoffman, B. J., Davidson, N., and Lester, H. A. (1994) Neuron 12, 845–859[CrossRef][Medline] [Order article via Infotrieve]
  39. Cammack, J. N., and Schwartz, E. A. (1996) Proc. Natl. Acad. Sci. U. S. A. 93, 723–727[Abstract/Free Full Text]
  40. Galli, A., Petersen, C. I., deBlaquiere, M., Blakely, R. D., and DeFelice, L. J. (1997) J. Neurosci. 17, 3401–3411[Abstract/Free Full Text]
  41. Lin, F., Lester, H. A., and Mager, S. (1998) Biophys. J. 71, 3126–3135
  42. Ingram, S. L., Prasad, B. M., and Amara, S. G. (2002) Nat. Neurosci. 10, 971–978
  43. Ramsey, I. S., and DeFelice, L. J. (2002) J. Biol. Chem. 277, 14475–14482[Abstract/Free Full Text]
  44. Quick, M. W. (2003) Neuron 40, 537–549[CrossRef][Medline] [Order article via Infotrieve]
  45. Otis, T. S., and Kavanaugh, M. P. (2000) J. Neurosci. 20, 2749–2757[Abstract/Free Full Text]

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