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Department of Molecular Physiology and Biophysics, Center for Molecular Neuroscience, Vanderbilt University, Nashville, Tennessee (K.M.K., B.J.L., Y.W., A.G.); Molecular Neuropharmacology Group, Department of Pharmacology, the Panum Institute, University of Copenhagen, Copenhagen, Denmark (C.J.L.; U.G.); and Departments of Psychiatry and Pharmacology, Center for Molecular Recognition, College of Physicians and Surgeons, Columbia University, New York, New York (J.A.J.)
Received March 1, 2006; Revision received May 5, 2006.
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Abstract |
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; Giros and Caron, 1993). The DA transporter (DAT) coordinates the spatial and temporal regulation of dopaminergic neurotransmission by mediating the reuptake of DA into presynaptic neurons. Although diffusion and enzymatic degradation contribute to reducing the synaptic concentration of DA, knockout studies have established DAT as the primary mechanism controlling extracellular DA levels (Giros et al., 1996; Jones et al., 1998). The regulatory mechanisms governing DAT are critical to DA signaling/homeostasis. Altered dopaminergic signaling has been implicated in multiple brain disorders, including Parkinson's disease, schizophrenia, attention deficit hyperactivity disorder, and addiction (Jucaite, 2002).
DAT belongs to the Na+/Cl--dependent transporter gene family, which also consists of plasmalemmal carriers for the other monoamines, norepinephrine, epinephrine, and serotonin. Modulation of transporter surface expression may constitute a dynamic component of DA clearance (Melikian and Buckley, 1999; Loder and Melikian, 2003; Sorkina et al., 2005). Indeed, the clearance efficiency of DAT is determined not only by the transport rate of an individual transporter but also by the number of functional transporters on the cell surface. DAT trafficking has been shown to be regulated by receptor signaling, as well as by direct activation of protein kinase C (Zhang and Reith, 1996; Huff et al., 1997; Vaughan et al., 1997; Zhang et al., 1997; Zhu et al., 1997; Pristupa et al., 1998; Mayfield and Zahniser, 2001; Carvelli et al., 2002; Granas et al., 2003; Loder and Melikian, 2003; Moron et al., 2003; Sorkina et al., 2005), and by interaction with cytosolic proteins (Torres et al., 2001; Carneiro et al., 2002; Bjerggaard et al., 2004; Jiang et al., 2004; Lee et al., 2004). In addition, substrates (Saunders et al., 2000; Chi and Reith, 2003; Sorkina et al., 2003; Kahlig et al., 2004) and inhibitors (Daws et al., 2002; Little et al., 2002) also regulate DAT function by a trafficking mechanism.
The psychostimulant AMPH is a substrate for DAT. Thus, AMPH elevates extraneuronal DA concentrations through a variety of mechanisms, including uptake competition and stimulation of DAT-mediated DA efflux (Sulzer et al., 2005). In addition, we and others have demonstrated that AMPH causes DAT redistribution away from the plasma membrane (Saunders et al., 2000; Chi and Reith, 2003; Sorkina et al., 2003; Kahlig et al., 2004). We have also shown that DAT leaves the membrane as an active protein in response to AMPH (Kahlig et al., 2004). Therefore, AMPH decreases DA transport capacity by reducing the number of transporter proteins on the plasma membrane and not through inactivation of surface DAT (Kahlig et al., 2004). It is unclear, however, whether the activity of the transporter is necessary for this response to AMPH. Although AMPH can enter the cell to some extent by lipophilic diffusion, the presence of a catecholamine transporter dramatically increases the rate and extent of AMPH accumulation (Bonisch, 1984; Sitte et al., 1998). Moreover, because the inhibitor cocaine blocks AMPH-induced DAT trafficking (Daws et al., 2002), it is likely that AMPH interacts with the transporter to cause its cell surface redistribution. It is important to understand, however, whether this interaction and subsequent changes in DAT activity are sufficient or if AMPH must cross the membrane in a DAT-mediated active transport process to decrease DAT plasma membrane expression.
In this study, to investigate the significance of DAT-mediated AMPH transport, we examined the ability of AMPH to induce transporter trafficking in an uptake-deficient DAT (Y335A-hDAT) that is targeted normally to the plasma membrane and binds AMPH (Loland et al., 2002). Extracellular AMPH did not cause Y335A-hDAT trafficking, but Zn2+, which stimulates Y335A-hDAT-mediated substrate transport (Loland et al., 2002), restored AMPH-induced DAT trafficking. Moreover, bypassing the transport cycle and directly delivering AMPH inside the cell with a whole-cell patch pipette produced Y335A-hDAT trafficking. Taken together, these data suggest that intracellular AMPH is required for AMPH-induced DAT trafficking.
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Materials and Methods |
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). The constructs were verified by restriction enzyme mapping and DNA sequencing, followed by stable transfection into human embryonic kidney 293 cells with Lipofectamine (Invitrogen, Carlsbad, CA). Stably transfected pools were selected with geneticin as described previously (Javitch et al., 1997). Cells were grown in Dulbecco's modified Eagle's medium supplemented with 10% fetal bovine serum at 37°C and 5% CO2. Reagents were purchased from Sigma (St. Louis, MO) unless otherwise noted. Confocal Microscopy and Electrophysiology. Stably transfected cells were plated at a density of 105 per 35-mm culture dish. Before imaging, attached cells were washed twice with bath solution containing 130 mM NaCl, 10 mM HEPES, 1.5 mM CaCl2, 0.5 mM MgSO4, 1.3 mM KH2PO4, and 34 mM dextrose adjusted to pH 7.35. In experiments using Zn2+, 10 µM Zn2+ was added to the external solution with the other ions. Cells were visualized with an inverted microscope (Eclipse TE300; Nikon, Tokyo, Japan) running the confocal imaging software, EZ 2000 (Coord Automatisering, Utrecht, The Netherlands). Images were acquired using 488-nm excitation with a 510-nm long pass filter. To minimize fluorophore bleaching and the cytotoxic effects of confocal microscopy, images were acquired at intervals of at least 1 min. AMPH and DA were bath-perfused at the indicated concentrations.
To deliver AMPH intracellularly to the uptake-deficient Y335A-hDAT, we used a whole-cell patch pipette in the current-clamp mode. To reduce the effect of any extracellular AMPH that leaked from within the cell, cells were bath-perfused during the experiment. A programmable puller (model P-2000; Sutter Instruments, Novato, CA) was used to fabricate quartz recording pipettes with a resistance of 7 M
. The recording pipette was filled with an internal solution containing the 120 mM KCl, 2 mM MgCl2·6H2O, 10 mM HEPES, and 30 mM dextrose adjusted to pH 7.35. Ten micromolar AMPH or 10 µM DA was added to the internal patch solution, as indicated. Membrane potentials were recorded in current-clamp mode using an Axopatch 200B (Molecular Devices, Sunnyvale, CA) with a low-pass Bessel filter set at 5000 Hz. The basal confocal image was acquired immediately after establishing the whole-cell configuration. Images were acquired as described above.
Because membrane proteins such as GLUT4 can be localized just under the plasma membrane (Inoue et al., 2003) in a manner that cannot be resolved from plasma membrane expression using optical methods, we measured intracellular fluorescence to quantify DAT cell surface redistribution as described previously (Kahlig et al., 2004). The quantification of intracellular fluorescence was performed by normalizing each respective time point by the total fluorescence in the whole field image to account for photobleaching (Lippincott-Schwartz et al., 1999; Piston et al., 1999). In addition, we also accounted for cytosolic volume changes (Lippincott-Schwartz et al., 1999; Piston et al., 1999). ImageJ software (http://rsb.info.nih.gov/ij/) was used for this analysis. Normalized data were analyzed with Prism software (ver. 3.02; GraphPad Software, Inc.) and reported as mean ± S.E. unless otherwise indicated.
Cell Surface Biotinylation. Cell surface biotinylation experiments were performed as described previously (Apparsundaram et al., 1998) with slight modification. Stably transfected cells were seeded in six-well plates (106 cells/well) approximately 48 h before the experiment. The cells were washed twice with 37°C bath solution (with or without 10 µM Zn2+), and cells were treated as indicated at 37°C for 1 h. The cells were washed twice with ice-cold PBS containing 0.1 mM CaCl2 and 1 mM MgCl2 (PBS-Ca-Mg) and treated with sulfo-NHS-biotin (1.5 mg/ml; Pierce Chemical Co., Rockford, IL) in PBS-Ca-Mg on ice for 1 h. The reaction was quenched by washing twice with 4°C PBS-Ca-Mg containing 100 mM glycine (PBS-Ca-Mg-glycine) followed by an incubation with PBS-Ca-Mg-glycine for 30 min on ice. Cells were then washed twice with 4°C PBS-Ca-Mg before lysis with 1 ml of radioimmunoprecipitation assay buffer [20 mM Tris, 20 mM EGTA, 1 mM dithiothreitol, 1 mM benzamidine, 1% Triton X-100; supplemented with protease inhibitors (100 µM phenylmethylsulfonyl fluoride, 5 µg/ml leupeptin, and 5 µg/ml pepstatin)] for 30 min on ice with constant shaking. Lysates were centrifuged at 14,000g for 30 min at 4°C. The supernatants were isolated and biotinylated proteins were separated by incubation with ImmunoPure Immobilized Streptavidin beads (Pierce Chemical Co, Rockford, IL) for 1 h at room temperature with constant mixing. Beads were washed three times with radioimmunoprecipitation assay buffer, and biotinylated proteins were eluted with Laemmli loading buffer for 30 min at room temperature. Total cell lysates and biotinylated proteins (cell surface) were separated by SDS-polyacrylamide electrophoresis (7.5%) and transferred into polyvinylidene difluoride membranes (Millipore). Polyvinylidene difluoride membranes were incubated for 1.5 h in blocking buffer (5% dry milk, 0.1% Tween 20 in Tris-buffered saline) and immunoblotted with a rat monoclonal antibody directed against the N terminus of hDAT (1:2000 in blocking buffer; Chemicon Inc., Temecula, CA). Immunoreactive bands were visualized using horseradish peroxidase-conjugated goat anti-rat antibody (1:5000 in blocking buffer; Santa Cruz Biotechnology, Santa Cruz, CA) with ECL Plus on Hypersensitive ECL film (GE Healthcare, Little Chalfont, Buckinghamshire, UK). Band densities were calculated using Scion Image software (Scion Corporation, Frederick, MD) and normalized to the appropriate total extract to control for protein loading. Data were analyzed with Prism software and reported as mean ± S.E.
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Results |
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; Torres et al., 2003). Furthermore, addition of an N-terminal GFP tag to DAT does not alter transporter function or regulation, as assessed by uptake, membrane targeting, PKC-stimulated trafficking, AMPH-induced trafficking, inhibitor binding, and substrate-induced currents (Daniels and Amara, 1999; Sorkina et al., 2003; Kahlig et al., 2004; Garcia et al., 2005). Using confocal imaging, we selected transfected cells with easily visualized plasma membrane fluorescence for analysis. Consistent with previous reports (Loland et al., 2003; Sorkina et al., 2003; Kahlig et al., 2004; Garcia et al., 2005), under basal conditions, both WT-hDAT and Y335A-hDAT were primarily expressed at the plasma membrane (Fig. 1, A and C, left). As expected, 10 µM AMPH caused WT-hDAT to traffic away from the membrane and accumulate intracellularly (Fig. 1A, right) (Saunders et al., 2000; Sorkina et al., 2003; Kahlig et al., 2004). In contrast, AMPH failed to cause the cellular redistribution of Y335A-hDAT (Fig. 1C). To quantify the increase of intracellular DAT, we measured changes in intracellular fluorescence and plotted them as a function of time after addition of AMPH (Fig. 1, B and D). Treatment with vehicle alone did not alter the intracellular distribution of WT-hDAT or Y335A-hDAT (
, Fig. 1, B and D; intracellular fluorescence was 96 ± 2% and 97 ± 3% of the basal values after 60 min, respectively). In WT-hDAT, AMPH produced a significant increase in intracellular fluorescence at all times between 20 and 60 min (*, p < 0.05, one-way ANOVA). After 60-min exposure to AMPH, the intracellular fluorescence of WT-hDAT cells increased to 214 ± 33% of basal levels (Fig. 1B, 
). In contrast, in Y335A-hDAT, intracellular fluorescence was unchanged by AMPH exposure (98 ± 5% of basal after 60 min AMPH exposure) (Fig. 1D, ).
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Zn2+ has been shown to modulate DAT activity through binding to an endogenous metal binding site (Norregaard et al., 1998; Loland et al., 1999). In WT-hDAT, 10 µM Zn2+ decreased the Vmax for DA uptake by approximately 70% (Loland et al., 2002). Therefore, if active transport of AMPH, and not simply lipophilic diffusion, is required for DAT trafficking, one might expect inhibition of transport by Zn2+ to reduce the ability of AMPH to induce trafficking. Indeed, in WT-hDAT cells, 10 µM Zn2+ prevented 10 µM AMPH from significantly increasing intracellular fluorescence (Fig. 2, A and B). The intracellular pool of WT-hDAT increased to 129 ± 11% of basal at 60 min (Fig. 2B, ), an increase that was not significant and was much smaller than the 214% increase obtained in the absence of Zn2+ (Fig. 1B).
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). Here, coapplication of 10 µM AMPH and 10 µM Zn2+ led to an increase in the Y335A-hDAT intracellular pool, which became significant by 20 min (Fig. 2, C and D) and reached 165 ± 11% of basal levels after 60 min (Fig. 2, C and D, ). Treatment with 10 µM Zn2+ alone did not alter intracellular levels of WT-hDAT or Y335A-hDAT (Fig. 2, B and D; after 60 min, intracellular fluorescence was 86 ± 4 and 93 ± 3% of the basal value, respectively). In parallel, we used cell surface biotinylation studies to confirm that the changes in intracellular transporter pools measured by confocal imaging were indicative of alterations in DAT cell surface expression. Treatment with 10 µM AMPH for 60 min induced a 46 ± 5% reduction in WT-hDAT cell surface expression compared with control (Fig. 3, A and B). In contrast, coincubation of 10 µM AMPH with 10 µM Zn2+ prevented this effect (WT-hDAT density was 119 ± 11% relative to control) (Fig. 3, A and B). Consistent with the confocal imaging data (Fig. 2), AMPH treatment did not reduce cell surface expression of Y335A-hDAT (Fig. 3, C and D). However, in contrast to its effects on WT-hDAT, 10 µM Zn2+ restored the ability of 10 µM AMPH to cause Y335A-hDAT trafficking (Fig. 3, C and D); coincubation with Zn2+ and AMPH for 60 min resulted in a 27 ± 6% loss of cell surface Y335A-hDAT (Fig. 3, C and D). Treatment with 10 µM Zn2+ alone did not alter cell surface expression of WT-hDAT or Y335A-hDAT (Fig. 3, B and D; band densities were 103 ± 16 and 112 ± 8%, respectively).
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Discussion |
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; Kahlig et al., 2004). A number of signal transduction pathways have been implicated in this modulation. For example, PKC, which has been shown to be activated by AMPH (Giambalvo, 1992a,b), reduces the cell surface expression of DAT, resulting in reduced uptake capacity (Pristupa et al., 1998; Granas et al., 2003; Loder and Melikian, 2003; Sorkina et al., 2005). Moreover, we have shown that insulin signaling maintains DAT cell surface expression and that the serine/threonine kinase Akt/protein kinase B plays a key role in the hormonal modulation of AMPH-induced DAT trafficking (Garcia et al., 2005). The molecular mechanism by which AMPH induces DAT trafficking and whether the transport of AMPH across the plasma membrane is essential for this effect are not known. Although cocaine, a DAT reuptake blocker, inhibits AMPH-induced trafficking (Daws et al., 2002; Little et al., 2002), this might occur by preventing the interaction of AMPH with the transporter. Thus, AMPH-induced trafficking may not require intracellular accumulation of AMPH. Rather, the interaction of AMPH with DAT may alter DAT function, such as an ion conductance, which would in turn alter intracellular signaling. Because cocaine blocks interaction of extracellular AMPH with the transporter as well as its transport, we cannot differentiate between these two mechanisms.
Instead, we have taken advantage of a transporter mutant, Y335A-hDAT, in which transport activity is reduced to less than 1% of WT-hDAT levels but that binds AMPH normally (Loland et al., 2002). In contrast to WT-hDAT, this uptake-deficient mutant does not undergo trafficking in response to AMPH (Figs. 1 and 3), demonstrating that the lipophilic diffusion of AMPH into the cells is not sufficient to cause transporter trafficking.
DAT contains three endogenous residues (His-193, His-375, and Glu-396) that coordinate the binding of Zn2+ (Norregaard et al., 1998; Loland et al., 1999). Zn2+ binding to this site produces a potent, noncompetitive inhibition of WT-hDAT uptake (Norregaard et al., 1998; Loland et al., 2002; Meinild et al., 2004). In contrast to this inhibition of WT-hDAT, Zn2+ dramatically stimulates uptake activity in Y335A-hDAT (Loland et al., 2002). Therefore, we used the differential effects of Zn2+ on the activity of WT-hDAT and Y335A-hDAT to further explore whether transport of AMPH is implicated in its stimulation of DAT trafficking.
Zn2+ inhibited AMPH-induced trafficking of WT-hDAT (Figs. 2 and 3), consistent with its inhibition of uptake. We demonstrated, by means of confocal imaging and cell surface biotinylation, that Zn2+ restored the ability of AMPH to induce trafficking of Y335A-hDAT (Figs. 2 and 3). This is consistent with Zn2+ stimulation of uptake in this mutant and the possibility that AMPH must be accumulated intracellularly. It is noteworthy that Zn2+ alone did not affect WT-hDAT or Y335A-hDAT trafficking (Figs. 2 and 3), indicating that uncoupled ion conductances elicited by Zn2+ (Meinild et al., 2004) do not cause transporter trafficking.
Taken together, these results suggest that Zn2+ regulation of AMPH-induced Y335A-hDAT trafficking is due to modulation of transporter activity. However, whether this is due to intracellular accumulation of AMPH or to alterations in intracellular signaling as a result of DAT function cannot be determined by these data. To discriminate between these possibilities, we used the uptake-impaired Y335A-hDAT mutant and delivered AMPH directly into the cell via the whole-cell patch pipette, thereby bypassing the need for DAT-mediated AMPH transport (Fig. 4A). With pipette delivery, the cytosolic concentration of AMPH would be expected to equilibrate with the patch pipette solution relatively rapidly. This concentration of internal AMPH clearly is more effective than the concentration reached by diffusion alone, as direct delivery of intracellular AMPH via the patch pipette induced trafficking of Y335A-hDAT (Fig. 4C). This suggests that the accumulation of intracellular AMPH is sufficient to promote trafficking and that DAT activity-related signaling alone is not sufficient. Moreover, the membrane voltage, as determined during current clamp, was not significantly altered by intracellular AMPH (data not shown), suggesting the absence of alterations in signaling pathways as a consequence of ionic conductance stimulation in this paradigm.
Intracellular AMPH may induce trafficking via three nonexclusive mechanisms. First, it is possible that the intracellular concentration of AMPH reached with active transport (or by pipette perfusion), but not by lipophilic diffusion, directly modulates signaling pathways. For example, we have shown that AMPH inhibits serine/threonine kinase Akt/protein kinase B (Garcia et al., 2005), and this might simply require intracellular AMPH and not DAT activity per se. Moreover, AMPH has been shown to activate extracellular signal-regulated kinase 1/2 (Choe et al., 2002) and PKC (Giambalvo, 1992a,b) and might directly modulate the activity of intracellular signal transduction cascades to regulate DAT membrane expression.
Second, the interaction of intracellular AMPH with DAT itself may promote a conformational state of the transporter, such as reverse transport, that supports its trafficking. Although the inability of intracellular DA to cause DAT trafficking (Fig. 4D) argues against such a mechanism, we cannot exclude the possibility that intracellular AMPH promotes a different DAT conformation than does intracellular DA. The lack of an effect of intracellular DA is somewhat puzzling, given that extracellular DA can also induce DAT trafficking (Saunders et al., 2000; Chi and Reith, 2003), although it does so at higher concentrations (Saunders et al., 2000) and is cell-type dependent (Daniels and Amara, 1999). Our data suggest that the mechanisms of AMPH- and DA-induced trafficking might differ.
Third, our data may also be explained by the weak base action of AMPH, which may reduce the acidic gradients necessary to maintain rapid endosomal recycling back to the plasma membrane (Sulzer and Rayport, 1990; Johnson et al., 1993), resulting in the accumulation of intracellular transporter pools in the face of constitutive recycling (Loder and Melikian, 2003; Melikian, 2004).
It is unlikely that direct phosphorylation of DAT is involved in AMPH-induced trafficking. Although methamphetamine has been shown to induce phosphorylation of the DAT N terminus, this phosphorylation is not necessary for DAT trafficking (Granas et al., 2003; Cervinski et al., 2005) but is essential for AMPH-induced DA efflux (Khoshbouei et al., 2004).
In summary, AMPH uses DAT to gain entry into the cell, and our data suggest that intracellular AMPH stimulates DAT trafficking. The physiological relevance of our findings is supported by animal studies demonstrating that administration of AMPH at doses reported to cause hyperactivity and stereotyped behaviors leads to extracellular AMPH concentrations equal to or higher than 10 µM (Clausing et al., 1995), the concentration we used in our studies. Furthermore, seminal studies by Fleckenstein and colleagues (Kokoshka et al., 1998; Fleckenstein et al., 1999) demonstrated that a single dose of amphetamines leads to a long-lasting decrement in DA uptake in the face of preserved DAT protein expression, a finding that may be explained by AMPH-induced cell surface redistribution of DAT. Identifying the intracellular target of AMPH may enable new therapies for AMPH abuse by enabling the modulation of DAT cell surface expression and activity.
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Acknowledgements |
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Footnotes |
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K.M.K., B.J.L., J.A.J., and A.G. contributed equally to this article.
ABBREVIATIONS: DA, dopamine; DAT, dopamine transporter; AMPH, amphetamine; hDAT, human dopamine transporter; WT, wild type; GFP, green fluorescent protein; PBS, phosphate-buffered saline; PBS-Ca-Mg, PBS containing 0.1 mM CaCl2 and 1 mM MgCl2; ANOVA, analysis of variance; PKC, protein kinase C.
Address correspondence to: Aurelio Galli, Department of Molecular Physiology and Biophysics, Center for Molecular Neuroscience, Vanderbilt University, 465 21st Ave. South, Nashville, TN 37232-8548. E-mail: aurelio.galli{at}vanderbilt.edu
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