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TOXICOLOGY
Department of Neurology (G.A.R., A.O.M., J.Y., G.H., T.X., A.H.M.) and Psychiatry (U.D.M.), The Johns Hopkins Medical Institutions, Baltimore, Maryland
Received for publication
May 26, 2005
Accepted
July 12, 2005.
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Abstract |
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; Biederman and Faraone, 2004; Wilens et al., 2004). For years, psychomotor stimulant drugs have been the mainstay of ADHD treatment (Greenhill et al., 2002; Fone and Nutt, 2005), and in the last decade, their use has increased substantially (Olfson et al., 2003; Robison et al., 2004). Of the various stimulant drugs used in the treatment of ADHD, amphetamine is among the most often prescribed (Greenhill et al., 2002; Fone and Nutt, 2005), both in children and adults (Wilens et al., 2004; Dodson, 2005).
As the use of amphetamine in the treatment of ADHD has increased, a large body of preclinical data has accrued indicating that amphetamine has the potential to damage brain dopamine-containing neurons in experimental animals. In particular, animals treated with amphetamine develop lasting reductions in striatal dopamine, its major metabolite dihydroxyphenylacetic acid (DOPAC), its rate-limiting enzyme tyrosine hydroxylase, its membrane transporter (DAT), and its vesicular transporter (VMAT2) (Gibb et al., 1994; McCann and Ricaurte, 2004). Anatomic studies indicate that lasting dopaminergic deficits after amphetamine are due to damage of dopaminergic nerve endings in the striatum, with sparing of dopaminergic nerve cell bodies in the substantia nigra.
Despite these preclinical data and growing awareness of potential long-term adverse effects of stimulant ADHD medications (Volkow and Insel, 2003; Fone and Nutt, 2005), there has been little expressed concern over possible dopaminergic neurotoxicity in humans receiving amphetamine for the treatment of ADHD. In large measure, this appears to be due to the fact that, as noted by various authors (Vitiello, 2001a,b; Greenhill et al., 2002; Fone and Nutt, 2005), doses, routes, and regimens of administration used in amphetamine neurotoxicity studies in animals differ significantly from those used in the treatment of ADHD.
The purpose of the present study was to determine whether amphetamine treatment, similar to that used clinically in the therapy of adult ADHD, produces long-term effects on brain dopaminergic neurons in adult nonhuman primates. Initial studies used baboons as experimental subjects because their size (20–30 kg) allows for administration of amphetamine at an absolute dose similar to that used in ADHD. To further simulate the clinical use of amphetamine, we trained baboons to self-administer amphetamine by mouth. We tested a 3:1 mixture of dextro- and levo-amphetamine, because one of the more common formulations used in the treatment of ADHD consists of a combination of 75% dextro-amphetamine and 25% levo-amphetamine. In a final set of studies, we used squirrel monkeys to test the species generality of our findings.
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Materials and Methods |
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Drugs and Chemicals. S(+)- and R(–)-Amphetamine were obtained through the National Institute on Drug Abuse drug supply program (Bethesda, MD), and their chemical authenticity was confirmed by means of gas chromatography/mass spectroscopy (GC/MS). Doses were expressed as the base weight. For GC/MS determinations, racemic amphetamine was obtained from Lipomed (Cambridge, MA), racemic-D8-amphetamine was purchased from Cerilliant (Round Rock, TX), and pentafluoropropionic acid (PFPA) was purchased from Fluka (St. Louis, MO). Clean Screen with Clean Thru tips solid-phase extraction columns (6 ml) were obtained from United Chemical Technologies (Bristol, PA). Other drugs and chemicals were obtained from the following sources: dopamine hydrochloride, DOPAC, 5-hydroxytryptamine creatinine sulfate complex, 5-hydroxyindoleacetic acid dicyclohexylammonium salt (5-HIAA), sodium octyl sulfate, EDTA, and quinine hemisulfate salt were obtained from Sigma-Aldrich (St. Louis, MO); [3H]WIN 35,428 was purchased from PerkinElmer Life and Analytical Sciences (Boston, MA); and [3H]DTBZ was obtained from American Radiolabeled Chemicals (St. Louis, MO).
Drug Administration. The oral route of administration was used. For baboons, this was accomplished by first training the animals to self-administer the drug vehicle orally. The training procedure (Kaminski et al., 2003) involved administration of Tang orange fruit drink (Kraft Foods, Rye Brook, NY) containing quinine, in increasing concentrations, immediately followed by administration of unadulterated orange drink (40 ml). Training lasted until the baboons were reliably drinking the most concentrated quinine solution (approximately 43 mg of quinine/l orange drink). Subsequently, amphetamine treatment was initiated by dissolving the appropriate amounts of dextro-amphetamine and levo-amphetamine (as sulfate salts) in distilled water, then adding the amphetamine isomer mixture to the orange drink. A 3:1 mixture of dextro [S(+)]- and levo [R(–)]-amphetamine, respectively, was used for all animal treatments. Dose selection was based upon published reports (Spencer et al., 2001; Greenhill et al., 2002; Wilens et al., 2004). In our initial baboon study, the amphetamine mixture was given twice daily (at 9:30 AM and 3:30 PM) for approximately 4 weeks, using doses shown in Table 1.
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In a second baboon study using different animals, we measured plasma amphetamine concentrations at the end of each of the 4 weeks of treatment, and similar to our first study, baboons were trained to self-administer the amphetamine mixture orally twice daily (at 9:00 AM and 3:00 PM) for 4 weeks, using doses shown below. As before, a 3:1 mixture of dextro [S(+)]- and levo [R(–)]-amphetamine, respectively, was used (Table 2).
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In a subsequent study involving squirrel monkeys, oral administration of the 3:1 mixture of dextro [S(+)]- and levo [R(–)]-amphetamine was accomplished by orogastric gavage while the animals were gently restrained in a Plexiglas chair. Squirrel monkeys received the amphetamine isomer mixture twice daily (9:30 AM and 3:30 PM) for 4 weeks. In these studies, rather than increasing the dose of the amphetamine isomer mixture automatically at the end of each week (as in the baboon study above), dose increments were made contingent upon plasma drug concentrations. In particular, when the plasma amphetamine concentration reached a level comparable with that observed clinically (between 100 and 150 ng/ml; see Discussion), dosage increments were stopped, and the animals were maintained on that dose of the 3:1 mixture of dextro [S(+)]- and levo [R(–)]-amphetamine for the remainder of the 4-week treatment period, as below (Table 3).
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Determination of Plasma Amphetamine Concentrations. Samples were analyzed by solid-phase extraction and gas chromatography-mass spectrometry using a procedure from United Chemical Technologies (http://www.unitedchem.com/s/pdfs/13-Applications%20Manual.pdf) adapted for plasma. Briefly, 0.5 ml of specimen was added to 25 µl of a 10 µg/ml aqueous solution of internal standard (d8-AMP), followed by 125 µl of 1.6 M periodic acid solution. Samples were allowed to stand at room temperature for 10 min after vortexing. Thereafter, 150 µl of 1.6 M KOH and 1 ml 0.1 M monobasic phosphate buffer (pH 6.0) were added, again followed by vortexing. The pH was adjusted to 5.0 to 7.0, if necessary. The samples were then centrifuged at 1800 rpm for 3 min. The supernatant was collected in 5-ml, 75 x 12-mm plastic test tubes and decanted onto CSDAU Clean Screen Extraction columns (United Chemical Technologies) preconditioned with 1.5 ml of methanol followed by 1.0 ml of 0.1 M monobasic phosphate buffer (pH 6.0). Supernatant samples were added to the columns, and low vacuum was applied. Columns were washed sequentially with 0.5-ml aliquots of 1.0 M acetic acid and 1.5 ml of methanol. The columns were dried thoroughly at high vacuum for 5 min. The analytes were eluted with 1.5 ml of a mixture of methylene chloride, 2-propanol, and ammonium hydroxide (80:20:2 by volume). The eluates were collected in conical glass centrifuge tubes. To the collected eluates, 15 µl of acidified methanol was added. Samples were evaporated under a continuous nitrogen stream until completely dry. After samples were evaporated to dryness, 50 µl of the derivatizing agent PFPA was added, and centrifuge tubes were capped, vortex-mixed, and placed on a heat block for 30 min at 50°C. After derivatization, the samples were dried under a gentle stream of nitrogen. Samples were then reconstituted with 50 µl of ethyl acetate and transferred to autosampler vials. Standards were prepared as follows: a 10 µg/ml amphetamine standard solution was prepared by diluting 0.1 ml of a 1.0 mg/ml ampoule to 10 ml of methanol. The contents of 100 µg/ml ampoule of d8-amphetamine internal standard solution were diluted to 10 ml with methanol to prepare a 10 µg/ml d8-amphetamine internal standard solution. Final concentrations of standards used in the analysis of plasma samples were 5, 20, 100, and 500 ng/ml. Analysis was performed using a Agilent Technologies model 6890N gas chromatograph equipped with a 30 m x 0.25-mm (i.d.) HP-5ms (5% phenyl)-methylpolysiloxane capillary column (0.25-µm film thickness) interfaced with an Agilent Technologies model 5973 inert electron impact mass spectrometer (Agilent Technologies, Palo Alto, CA). Helium was used as the carrier gas (flow rate, 1.3 ml/min) for 1 µl of splitless sample injections. The initial oven temperature was 90°C with a 1-min hold followed by ramps at 20°C/min to 180°C with a 2-min final hold. Total run time was 7.5 min. Specimens were analyzed in selected ion monitoring mode for the following ions (where q is the quantitative ion): d8-AMP (pentafluoropropionic derivative), m/z 193 (q), 126, and 96; AMP (pentafluoropropionic derivative), m/z 190 (q), 118, and 91. Amphetamine peak areas were integrated using the Agilent ChemStation Software (rev. D.01.00), and the ratio of the area of the calibrator and its internal standard were used for calculations.
Brain Dissection. Two weeks after drug treatment, animals were sacrificed under deep sodium pentobarbital anesthesia (60 mg/kg i.p.), and the brain was removed from the skull in a cold room (4°C). Regional dissection of the brain was performed using the Emmers and Akert (1963) atlas as a guide for the squirrel monkey and the Riche et al. (1988) atlas for the baboon. Brain regions of interest were isolated from coronally cut sections (approximately 4–5 mm thick) by means of free dissection. Tissue from each brain region was wrapped in aluminum foil and then stored in liquid nitrogen until assay.
Determination of Regional Brain Monoamine Concentrations. Regional brain concentrations of dopamine and 5-hydroxytryptamine were determined by high-performance liquid chromatography with electrochemical detection, as described previously (Yuan et al., 2002).
Quantitative DAT Autoradiography. Frozen brains were sectioned using a Microm HM505E cryostat at –20°C. Half-hemisphere coronal sections (20 µm) were thaw-mounted onto gelatin-coated microscope slides, and the DAT was labeled with 50 pM [125I]RTI-121, using the method of Staley et al. (1995). Autoradiographs were digitized with a Dage CCD 72 camera and MCID Elite 6.0 image analysis system (GE Healthcare, Little Chalfont, Buckinghamshire, UK).
DAT Binding. [3H]WIN 35,428-labeled DATs were measured using the method of Madras et al. (1989), with minor modification. Briefly, frozen striatal tissue was weighed, homogenized for 15 s in 20 volumes (w/v) of a 0.32 M sucrose phosphate buffer (pH = 7.4) at 0–4°C, and centrifuged in a Sorvall RC2B centrifuge (Sorvall, Newton, CT) at approximately 45,000g for 15 min at 0–4°C. The supernatant was discarded, and the pellet was resuspended in 20 volumes of sucrose phosphate buffer and then centrifuged once again at approximately 45,000g for 15 min at 0–4°C. The resulting pellet was suspended in buffer for a final tissue concentration of 10 mg/ml wet weight. [3H]WIN 35,428 was used at a predetermined saturating concentration of 30 nM. Cocaine, at a final concentration of 30 µM, was used to displace specific [3H]WIN 35,428 binding, to estimate nonspecific binding. Tubes were incubated in sextuplicate for 60 min at 0–4°C. The incubation was terminated by rapid filtration, using a 48-well cell harvester (Brandell, Gaithersburg, MD) and Whatman GFB filters soaked with 0.05% polyethylenimine. Filters were washed three times using ice-cold sucrose phosphate buffer. Radioactivity was measured with a Packard-1500 Tricarb Liquid Scintillation Analyzer (PerkinElmer Life and Analytical Sciences). Specific [3H]WIN 35,428 binding was calculated by subtracting the average value of the six tubes containing excess cocaine from the average of the six tubes without cocaine. Specific [3H]WIN 35,428 binding was expressed in dpm/mg original wet weight tissue.
VMAT Binding. [3H]DTBZ was used to label type 2 vesicular monoamine transporter (VMAT) sites. [3H]DTBZ binding was measured using the method of Vander Borght et al. (1996), with minor modifications. Briefly, tissue samples were homogenized for 15 s in 20 volumes (w/v) of sodium phosphate buffer, (25 mM, pH 7.7) and then centrifuged in a Sorvall RC2B centrifuge at approximately 45,000g for 15 min at 0–4°C. The resulting pellet was resuspended in 20 volumes (w/v) of sodium phosphate buffer, then homogenized again for 15 s, and recentrifuged at approximately 45,000g for 15 min at 0–4°C. The supernatant was discarded, and the resulting pellet was resuspended in buffer at a final concentration of 10 mg of original wet weight tissue per milliliter. Membrane preparations were incubated with a predetermined saturating concentration of [3H]DTBZ (15 nM) in 25 mM sodium phosphate buffer, pH 7.7, for 90 min at 30°C in a shaking water bath. Each sample was run in sextuplicate, such that three tubes were used to define total binding, and three tubes were used to determine nonspecific binding. Nonspecific binding was determined in the presence of 1 µM tetrabenazine and represented approximately 8 to 10% of total binding. The incubation was terminated by rapid filtration as above. Filters were washed three times with 10 ml of sodium phosphate buffer, and residual radioactivity was measured using a Packard-1500 Tricarb Liquid Scintillation Analyzer. Specific binding, calculated by subtracting nonspecific binding from total binding, was expressed as dpm/mg original wet weight tissue.
Western Blot Analysis. A rat anti-DAT monoclonal antibody (Chemicon International, Temecula, CA) was used as primary antibody (1:1000), and goat anti-rat Ig horseradish peroxidase-linked antibody (GE Healthcare) was used as secondary antibody (1:1000). An enhanced chemiluminescence Western blotting detection system (Amersham Biosciences) was used to visualize the signals produced. Actin polyclonal antibody (Santa Cruz Biotechnology, Inc., Santa Cruz, CA) was used to correct the loading protein amount, and the NIH image program (Image J) was used to semiquantitatively analyze the band intensity (http://rsb.info.nih.gov/ij/), as described previously (Xie et al., 2004). Tissue used for these studies was from the same animals used for the other neurochemical studies reported here.
Statistics. Data were analyzed by ANOVA and, where appropriate, by independent samples, two-tailed Student's t test. Results were considered significant when p < 0.05. Data analysis was performed using the Statistical Program for the Social Sciences (SPSS for Windows, Release 10.5; SPSS Inc., Chicago, IL).
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To ensure that the dose and dosing parameters used in the above study accurately approximated those used clinically, we trained another group of baboons (n = 3) to self-administer escalating oral doses of the 3:1 mixture of dextro [S(+)]- and levo [R(–)]-amphetamine (see Materials and Methods) and measured plasma concentrations of amphetamine at the end of each week of treatment. As the dose of the amphetamine isomer mixture was increased, plasma levels of amphetamine rose (Fig. 3A). The mean (±S.E.M.) plasma amphetamine concentration at the end of the 4-week treatment period was 168 ± 25 ng/ml. Similar to the baboons in our first study, this group of baboons had significant reductions in brain dopaminergic neuronal markers when examined 2 weeks after cessation of amphetamine treatment (Fig. 3, B–D).
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To determine the species generality of our findings, we carried out an additional study using squirrel monkeys. However, in this study, once the plasma concentration of amphetamine reached a level comparable with that reported clinically (100 to 150 ng/ml; see Discussion), dose increments were stopped. As in baboon studies, plasma levels of amphetamine rose when the dose of the 3:1 mixture of dextro [S(+)]- and levo [R(–)]-amphetamine was increased from 0.25 to 0.5 mg (Fig. 4A). Interestingly, despite maintaining the amphetamine dose constant at 0.5 mg for the last 2 weeks of treatment, plasma amphetamine concentrations decreased modestly, but significantly, to a mean (±S.E.M.) plasma concentration of 125 ± 14 ng/ml at the end of the 4-week treatment period (Fig. 4A).
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When these squirrel monkeys were examined approximately 2 weeks after the final dose of the 3:1 mixture of dextro [S(+)]- and levo [R(–)]-amphetamine, they had reductions in striatal dopaminergic markers, although the reduction in [3H]WIN 35,428-labeled DAT did not achieve statistical significance (Fig. 4, C–E).
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Doses of amphetamine that are used clinically range from 5 to 60 mg (Greenhill et al., 2002; Wilens et al., 2004), and except for slow-release, longer-acting formulations that have recently become available (Greenhill et al., 2003; McGough et al., 2003), are typically prescribed for twice-daily use. Plasma concentrations of amphetamine that develop after chronic amphetamine treatment have rarely been reported. Indeed, to our knowledge, there are only two reports that provide such information. The first, an early study by Borcherding et al. (1989), indicates that hyperactive children given escalating oral, twice-daily (9:00 AM and 1:00 PM) doses of dextroamphetamine over a 3-week period develop plasma amphetamine concentrations of approximately 120 to 140 ng/ml. The second, a more recent study by McGough et al. (2003), shows that patients with ADHD given a single daily (morning) dose of an extended release formulation of mixed amphetamine salts for a 6-week period develop dose-related plasma amphetamine concentrations, with a total (d-plus l-isomers) plasma amphetamine concentration of approximately 120 ng/ml after the highest dose examined (30 mg). Thus, plasma concentrations of amphetamine achieved clinically in patients receiving chronic amphetamine for the treatment of ADHD appear to be on the order of those shown here to produce dopaminergic neurotoxic effects in adult baboons and squirrel monkeys. Importantly, the moderate magnitude of the reduction of dopaminergic axonal markers in these nonhuman primates suggests that plasma levels achieved approach the minimal plasma concentration "threshold" required to produce dopaminergic neurotoxic changes.
In contrast to the paucity of clinical reports on plasma amphetamine concentrations after chronic treatment, there are numerous reports on the pharmacokinetics of single, oral doses of amphetamine in humans (Brown et al., 1979; Angrist et al., 1987; McGough et al., 2003). Collectively, these studies indicate that, as the dose of amphetamine is increased, there is a corresponding increase in plasma amphetamine concentration. Furthermore, when frequency of dosing is increased from once to twice daily, plasma concentrations of amphetamine increase accordingly (Greenhill et al., 2003), most likely due to the fact that the elimination half-life of amphetamine in humans is on the order of 6 to 9 h (Cho and Kumagai, 1994).
Although the present preclinical observations may have clinical implications, it would be premature to extrapolate them to humans receiving amphetamine treatment for ADHD for several reasons. First, the dopaminergic neurotoxicity may only occur in the context of doses of amphetamine that result in plasma concentrations comparable with those found in these experiments; lower dosage regimens that engender lower plasma amphetamine concentrations may not be associated with toxic effects on central dopaminergic neurons. Second, the mechanisms of amphetamine-induced dopaminergic neurotoxicity are not known, and theoretically, could be operant in nonhuman primates (and rodents) but not in humans. Third, aspects of amphetamine metabolism in nonhuman primates may differ from those in humans, and such differences could potentially result in neurotoxicity in nonhuman primates but not in humans. Fourth, the relative sensitivity of brain dopaminergic neurons to amphetamine toxicity in nonhuman primates and humans is unknown. Fifth, it is possible that the effects observed in normal primates with amphetamine may not be observed in ADHD patients because such patients presumably have abnormal neurotransmitter function, and such abnormalities may influence the expression of amphetamine neurotoxicity. Finally, it is important to note that amphetamine neurotoxicity data from the present studies were obtained in adult nonhuman primates; as such, although they may have implications for adults receiving amphetamine for the treatment of ADHD, their implications for children are less clear, because studies assessing the influence of age on the ontogeny of amphetamine neurotoxicity suggest younger animals are less susceptible to the neurotoxic effects of amphetamine (Cappon et al., 1997; Miller et al., 2000). Future studies in young adolescent primates are needed.
It is reasonable to wonder why, if clinically relevant doses of amphetamine produce toxic effects on brain dopaminergic neurons in nonhuman primates, evidence of dopaminergic neurotoxicity has not been revealed in clinical or neuroimaging studies of patients with ADHD. With regard to clinical studies, it is important to recognize that parkinsonism may not become manifest until central dopaminergic function is reduced by approximately 80 to 90% (Koller et al., 1991), a degree of dopamine reduction that is approximately 2-fold greater than that observed in the current study. Abnormalities in cognitive function, another potential functional consequence of dopaminergic loss (Robbins, 2003), may be difficult to distinguish from the underlying symptoms of ADHD for which amphetamine is being prescribed. Indeed, given the fact that abnormalities in dopaminergic neural function are believed, in part, to underlie symptoms of ADHD (Fone and Nutt, 2005), any indication of abnormal dopaminergic function in amphetamine-treated patients might be attributed to underlying disease, rather than amphetamine neurotoxicity. With respect to neuroimaging studies, it is important to recognize that most subjects with ADHD who have been included in PET/SPECT studies of the DAT have been medication-naive individuals (Krause et al., 2000, 2003; Jucaite et al., 2005) and that in those isolated instances in which ADHD patients treated with stimulants have been included, those treated with amphetamine (rather than methylphenidate) were either excluded from the analysis (Krause et al., 2003) or were not analyzed separately (Dougherty et al., 1999; Krause et al., 2002). Notably, results of these imaging studies have not been entirely consistent, with both increases (Dougherty et al., 1999; Krause et al., 2000) and no change (van Dyck et al., 2002) in DAT density reported in the striatum of ADHD patients. Thus, for a variety of reasons, the absence of previous clear clinical or PET/SPECT data showing evidence of DAT changes consistent with dopaminergic neurotoxicity in patients with ADHD previously treated with amphetamine should not be construed as evidence that it does not occur.
In summary, the present results indicate that amphetamine treatment similar to that used clinically for the management of adult ADHD produces brain dopaminergic neurotoxicity in adult nonhuman primates and engenders plasma concentrations of amphetamine that are on the order of those reported in some ADHD patients. Although it would be premature to extrapolate the present findings to ADHD patients treated with amphetamine, they provide an impetus to conduct controlled studies aimed at determining the potential for amphetamine to produce dopaminergic neurotoxicity in ADHD cohorts (adult, adolescent, and childhood forms). The present results also suggest that dopaminergic neurotoxic effects of amphetamine could potentially occur in other disorders that require chronic amphetamine treatment (e.g., narcolepsy) and that it might be prudent to monitor plasma amphetamine concentrations in patients considered at risk (e.g., those receiving higher amphetamine doses for extended periods). Finally, the present findings may also have implications for the drug abuse field, because plasma amphetamine concentrations in some abusers substantially exceed those here shown to produce dopaminergic neurotoxicity in nonhuman primates (Nakashima et al., 2003; Peters et al., 2003).
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Acknowledgements |
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Footnotes |
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ABBREVIATIONS: ADHD, attention-deficit/hyperactivity disorder; DOPAC, dihydroxyphenylacetic acid; DAT, dopamine transporter; VMAT2, vesicular monoamine transporter 2; GC/MS, gas chromatography/mass spectroscopy; PFPA, pentafluoropropionic acid; 5-HT, 5-hydroxytryptamine; 5-HIAA, 5-hydroxyindoleacetic acid; RTI-121, 3
-(4-[125I]iodophenyl)tropane-2-carboxylic acid isopropyl ester; WIN 35,428, 3-[4-flurorophenyl]-tropane-2-carboxylic acid methyl ester tartrate; DTBZ, dihydrotetrabenazine; ANOVA, analysis of variance.
Address correspondence to: Dr. George A. Ricaurte, Department of Neurology, Johns Hopkins Medical Institutions, 5501 Hopkins Bayview Circle, Rm. 5B.71E, Baltimore, MD 21224. E-mail: ricaurte{at}jhmi.edu
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