U.S. flag

An official website of the United States government

NCBI Bookshelf. A service of the National Library of Medicine, National Institutes of Health.

Michael AC, Borland LM, editors. Electrochemical Methods for Neuroscience. Boca Raton (FL): CRC Press/Taylor & Francis; 2007.

Cover of Electrochemical Methods for Neuroscience

Electrochemical Methods for Neuroscience.

Show details

Chapter 3Presynaptic Regulation of Extracellular Dopamine as Studied by Continuous Amperometry in Anesthetized Animals

, , and .

Electrically Evoked Dopamine Release Monitored by Continuous Amperometry


The Quest for Speed

In the peripheral and central nervous systems, neurotransmission is achieved by the release of neurotransmitters from nerve terminals into the extracellular fluid bathing postsynaptic receptors. Therefore, locally monitoring neurotransmitters in the extracellular fluid is an important technical goal. Because neurotransmission in the brain is a rapid process, with the transfer of chemical messages between neurons occurring in milliseconds, monitoring techniques that provide high temporal resolution are especially significant. Dopamine (DA) is an important neurotransmitter in several structures of the central nervous system such as the striatum, which contains the highest density of dopaminergic terminals. Since the 1970s, two main approaches to monitoring DA in the extracellular fluid have been developed: in vivo microdialysis and various in vivo electrochemical techniques. Electrochemical techniques can be used to monitor DA because the DA molecule is easily oxidized at the surface of carbon electrodes. Among the electrochemical techniques, continuous amperometry provides the fastest available temporal response with a low selectivity for DA over other oxidizable molecules. This chapter explains how to take advantage of the high temporal resolution of continuous amperometry while simultaneously avoiding the potential pitfalls associated with its low selectivity.

Principle of Continuous Amperometry

Continuous amperometry involves the application of a constant potential difference between a working electrode and a reference electrode. This potential is fixed at a level sufficient to oxidize the molecule of interest. In the case of DA, the carbon electrode is held at +0.4 V versus an Ag/AgCl reference electrode. Molecules that come into contact with the electrode surface are oxidized, with each DA molecule transferring two electrons to the electrode, giving rise to a measurable oxidation current. Molecules are delivered to the electrode surface by diffusion, which occurs at a rate proportional to the concentration in the solution surrounding the electrode. Hence, the DA oxidation current is also proportional to the DA concentration in the solution. However, any substance present in the solution that oxidizes at or below the potential applied to the electrode may contribute to the total oxidation current. For this reason, the total oxidation current might include contributions from other oxidizable substances, so continuous amperometry is not selective and must be used accordingly.

Methods: Carbon Fiber Electrodes and Potentiostats

With all electrochemical techniques, the aim is to oxidize molecules in the solution; therefore, the material selected for the working electrode must be conductive and highly resistant to being oxidized itself. This latter requirement especially applies to continuous amperometry because the working electrode is continuously held at an oxidative potential; most other electrochemical techniques involve alternating between oxidative and reductive potentials. Carbon, which is both conductive and more resistant to oxidation than metals, is the best choice of electrode material for electrochemical techniques used in the oxidation range. Among the various forms of carbon, carbon fibers are the most suitable because they are made of pyrolitic graphite, a material akin to pure graphite crystal. Gonon and coworkers designed the first carbon fiber microelectrode in 1978 [1].

The flow of oxidation current between the working and reference electrode might alter the applied potential by ohmic drop or electrode polarization. If not avoided, this would lead to irreproducible results. Some workers use a three-electrode potentiostat design in which the current through the reference electrode is maintained at zero by means of an auxiliary electrode. However, the active surface area of a carbon fiber microelectrode is very small indeed, so the current actually recorded during continuous amperometry is likewise very small, i.e., 1 nA or less. Thus, the ohmic drops are too small to be of any concern, and polarization effects are eliminated by using a reference electrode with sufficient surface area to assure that the current density at the reference electrode is small. Both potentiostats designed for continuous amperometry (AMU 130 by Radiometer Analytical, Lyon, France and Micro-C by WPI, Sarasota, FL) are twoelectrode potentiostats.

Validity And Field Of Application

Monitoring Dopamine in the Presence of Other Oxidizable Compounds

The use of continuous amperometry is completely valid when the compound of interest represents the main oxidizable compound released in the vicinity of the carbon fiber electrode. This condition exists during the monitoring of adrenaline or noradrenaline release by single chromaffin cells [2] and of DA release by single PC12 cells [3]. Likewise, in vitro preparations of sympathetic nerves release only noradrenaline, which can be unequivocally monitored by continuous amperometry [4]. The technique can also be used for monitoring electrically evoked DA release in striatal slices because, in this preparation, oxidizable molecules in the extracellular space of the slice remain below the detection limit of continuous amperometry unless DA release is evoked either by electrical stimulation or by amphetamine [5].

In contrast, the in vivo monitoring of DA release, either in anesthetized or in freely moving animals, is a much more challenging task because the extracellular DA level remains below 100 nM, even after amphetamine stimulation, whereas the main metabolite of DA (i.e., DOPAC) is present in the extracellular space at a concentration of 20 μM, and ascorbic acid is present at a concentration of 200 μM [6,7]. Both DOPAC and ascorbic acid are easily oxidizable substances that contribute to the oxidation current when the carbon fiber electrode is continuously held at +0.4 V versus Ag/AgCl. Several voltammetric techniques have been developed to monitor released DA without interference from DOPAC or ascorbic acid [8–12]. These studies were performed in anesthetized rats and showed that brief electrical stimulations of dopaminergic neurons, at a distance from the striatum, evoked a detectable increase in extracellular DA level but did not alter the extracellular levels of DOPAC and ascorbic acid. Brief electrical stimulations, lasting for 40 s or less, evoked an immediate increase in the exocytotic release of DA, which did not overlap with a modest and delayed increase in extracellular DOPAC [12,13] because the half-life of DOPAC is about 10 min [6] and because DOPAC is not sequestered in synaptic vesicles of dopaminergic neurons [14].

In anesthetized animals, electrical stimulation of dopaminergic neurons does not enhance the extracellular ascorbic acid level because dopaminergic terminals do not release ascorbic acid. Indeed, in the striatum of rats rendered hemi-parkinsonian by a unilateral 6-hydroxydopamine lesion, the striatal extracellular ascorbic acid level is not altered in the lesioned striatum compared to the intact one [7]. Moreover, the contribution of ascorbic acid to the secretion spikes, corresponding to exocytotic release from single chromaffin cells, has been proved to be negligible compared to the contribution of catecholamines [15]; the same is likely true regarding exocytotic release from dopaminergic fibers in vivo [14]. Nevertheless, the extracellular ascorbic acid level is not always stable. For example, it is enhanced by amphetamine or dopaminergic agonists and depressed by D2 dopaminergic antagonists and anesthetics. These pharmacological effects are explained by the link between ascorbic acid release and the activity of the glutamate transporters [16]. These variations in the extracellular ascorbic acid level affect the basal oxidation current recorded with continuous amperometry; however, they do not contribute to the transient oxidation signal evoked by brief electrical stimulations because pharmacologically induced ascorbic acid variations are much slower [7].

Amperometric Monitoring of Dopamine Release Evoked by Brief Electrical Stimulation

From the considerations developed above, it follows that, in anesthetized animals, DA is the only oxidizable compound released to the striatal extracellular fluid by brief electrical stimulation of dopaminergic neurons. Therefore, although continuous amperometry cannot separate DA from other oxidizable compounds, it is suitable for monitoring evoked DA overflow in this particular condition. Indeed, the brief changes in oxidation current, evoked by electrical stimulation and monitored with continuous amperometry in the striatum of anesthetized rats, exhibited all the expected characteristics of an impulse flow-dependent DA release process: (1) The evoked responses closely fit the anatomy of the dopaminergic neurons. Indeed, electrical stimulation of sites just above the medial forebrain bundle (MFB) did not evoke any response [17]. Moreover, the amplitude of the evoked responses was closely related to the density of dopaminergic terminals [18]. (2) When the carbon fiber electrode was held at 0 V instead of +0.4 V, i.e., at a potential that does not cause DA oxidization, no change in the oxidation current was recorded except transient electrical artifacts associated with the stimulation pulses [17,19]. (3) A blockade of impulse flow-dependent DA release by 6-hydroxydopamine injection in the MFB completely suppressed the evoked responses [20]. Moreover, partial degeneration of striatal dopaminergic terminals induced a corresponding decrease in the amplitude of the evoked responses [21]. (4) The evoked responses were abolished by the local application of tetrodotoxin and strongly diminished by cadmium, a blocker of calcium channels [17]. (5) The evoked responses were closely time-correlated with the onset and duration of the stimulation [17,22]. (6) Inhibition of DOPAC synthesis by pargyline did not decrease the evoked responses [17]. (7) Inhibition of DA uptake strongly slowed down the decay phase of the evoked responses [17,23]. Moreover, in the striatum of mice lacking the DA transporter, this decay phase was 100 times slower than in normal mice [18]. (8) Potentiation of the evoked DA release by D2 antagonists enhanced the amplitude of the evoked responses [17,22], but this effect was not observed in mice lacking D2 receptors [24].

Field of Application

Continuous amperometry can be used only in those restricted experimental conditions where only one oxidizable compound is responsible for the variations in the oxidation current recorded by the carbon fiber electrode. In contrast, continuous amperometry cannot be used to monitor DA release in freely moving animals in behavioral situations or to monitor variations of the basal extracellular DA level induced by pharmacological treatments such as amphetamine or dopaminergic agonists. Continuous amperometry can be used in anesthetized animals to monitor DA release evoked by brief electrical stimulations. It is sensitive enough to record the DA overflow evoked by single pulse stimulations in brain regions densely innervated by dopaminergic terminals [17,18,20,25]. In brain regions exhibiting a lower density of dopaminergic innervation, such as the basolateral nucleus of the amygdala, high-frequency stimulus trains (24 pulses, 60 Hz) must be used to evoke detectable DA overflow [26]. Finally, continuous amperometry can also be used in brain regions densely innervated by noradrenergic terminals to monitor the noradrenaline overflow evoked by brief electrical stimulations [27].

Comparison With Other Approaches

Comparison with In Vivo Microdialysis

Microdialysis has proved to be suitable for monitoring the basal extracellular DA level in freely moving animals. However, the time resolution of microdialysis is in the range of one to several minutes even if faster sampling rates are used. Indeed, this low time resolution is inherent in the fact that variations in the DA content of collected samples reflect variations in the extracellular DA level in the intact tissue with a time distortion caused by: (1) DA diffusion from intact release sites to the microdialysis probe through tissues lesioned by the implantation of the probe; (2) DA diffusion through the dialysis membrane; and (3) DA dilution in the perfusion medium filling the probe and the tubing connecting the probe to the fraction collector [28]. Therefore, pronounced DA overflow evoked by brief electrical stimulation cannot be monitored with microdialysis unless animals are treated with an uptake inhibitor. Indeed, uptake inhibition promotes DA diffusion from intact release sites to the probe, as elegantly shown by studies combining microdialysis with fast scan cyclic voltammetry (FSCV) [29,30].

Comparison with Fast Scan Cyclic Voltammetry

Compared to continuous amperometry, FSCV is much more selective for DA over other oxidizable compounds. Therefore, FSCV can be used to monitor DA transients in freely moving rats in behavioral situations (see Chapter 2 by Robinson and Wightman). FSCV has also been used extensively in anesthetized animals to monitor the DA overflow evoked by electrical stimulation. Observations obtained with FSCV and with continuous amperometry are generally in excellent agreement [26,31,32]. Minor discrepancies between some of the earliest studies have since been resolved. For example, the uptake blocker GBR 12909 was found to potentiate DA release rather than affecting DA clearance according to an early FSCV study [33]. In contrast, continuous amperometry clearly showed that GBR 12909 slows down DA clearance [23]; this observation was confirmed by a subsequent FSCV study from Wightman’s group [34].

The main advantage of continuous amperometry over FSCV is that the former exhibits a faster temporal response [26,31,32]. First, the sampling rate usually used with FSCV is 100 ms; the amperometric response to a rectangular pulse of catecholamine is more rectangular than the FSCV response because of the temporal lag due to adsorption and desorption of DA to and from the electrode [32]. With continuous amperometry, the time resolution is that of the data acquisition system and might be also limited by the filter of the current amplifier. A sampling interval of 20 μs is used to monitor amperometric events corresponding to the exocytotic release of DA [14]. When the electrically evoked DA overflow is recorded with continuous amperometry, the sampling interval is usually 1 ms [25]. This is sufficient because the kinetics of evoked DA overflow is constrained by DA reuptake. Maximal DA uptake kinetics occur in the in the dorsal striatum, where the apparent DA half-life is in the range of 60–80 ms [18,20,23]. Models of DA diffusion from release sites to carbon fiber electrodes suggest that the real half-life of DA in the intact striatum is in the range of 25–40 ms [25]. Therefore, the decay phase of the evoked DA overflow reflects DA elimination by reuptake with a much better accuracy when it is monitored with continuous amperometry than with FSCV.

Comparison with Other Electrochemical Techniques

Apart from FSCV and continuous amperometry, two other electrochemical techniques have been used to monitor electrically evoked DA overflow: differential normal pulse voltammetry (DNPV) and differential pulse amperometry (DPA). Both techniques were mainly used in combination with electrochemically treated carbon fiber electrodes. With DNPV it is possible to record, every 30 s, two distinct oxidation peaks corresponding to ascorbic acid and catechols. In untreated rats, the main compound contributing to the catechol peak is DOPAC [6]. When rats are treated with pargyline to inhibit DOPAC synthesis, the remaining catechol peak is entirely due to extracellular DA [10,11]. With this approach, it was possible to show that electrical stimulation of the MFB evoked the release of DA but did not alter the extracellular ascorbic acid level [11]. Therefore, this approach provided definitive information to characterize the in vivo monitoring of the evoked DA overflow. Currently, the interest of DNPV is low because of its slow time resolution.

DPA represents an improvement of DNPV with similar selectivity and sensitivity and a sampling rate of 500 ms. When DPA is combined with electrochemically treated carbon fiber electrodes, it is sensitive enough to detect the evoked DA overflow in the prefrontal cortex [37]. However, the real-time resolution is limited by the response time of electrochemically treated carbon fiber electrodes (10–30 s, depending on the treatment) [12,13,38]. Comparison between DPA and continuous amperometry is more extensively discussed in a previous review [39].

DPA has also been used with untreated carbon fiber electrodes [40]. With this approach, the time resolution is given by the sampling rate of DPA, which is 500 ms. The selectivity of DPA for DA over ascorbic acid [40] is similar to that of continuous amperometry (Dugast, Suaud-- Chagny, and Gonon 1994). The only advantage of DPA is that the carbon fiber electrode is held at a negative potential (−240 mV) between measurement pulses (+200 mV versus Ag/AgCl). Therefore, with DPA, DA molecules, which are oxidized by the carbon fiber to the corresponding quinone, can be reduced back by the electrode to DA. In contrast, because with continuous amperometry the electrode is maintained at an oxidizing potential, the electrode may contribute, by oxidative degradation, to eliminate the released DA. Comparison between continuous amperometry and DPA, combined with the same untreated carbon fiber electrode in the same experimental conditions, made it possible to resolve this issue. In this experiment, the rate of elimination of the released DA was monitored in the striatum of mice lacking the DA transporter. With both techniques, the measured DA half-life was exactly the same [18]. This comparison demonstrates that, even in the absence of DA uptake, oxidation of DA by the electrode in the continuous amperometric mode does not contribute to DA elimination because, in the extracellular fluid, ascorbic acid is present at a much higher concentration than DA. Thus, the quinones formed by oxidation of DA at electrodes are immediately reduced back to DA by ascorbic acid, which is a potent reducing agent [18,26]. From these considerations, it follows that in vitro tests for measuring the sensitivity to DA in the continuous amperometric mode must be performed with solutions containing high levels (≥100 μM) of ascorbic acid [17,26,32].

Characteristics of Dopaminergic Transmission

Dopaminergic Transmission Is Extrasynaptic But Local

Whether dopaminergic transmission is confined to the synaptic cleft or occurs through stimulation of postsynaptic receptors far away from release sites has been a matter of debate for decades. However, experimental evidence in favor of extrasynaptic transmission is growing [36]. First, most dopaminergic receptors are not located inside dopaminergic synapses [41–44]. Second, dopaminergic transporters are also extrasynaptic [45]. Third, most of the released DA diffuses out of the synaptic cleft and can be detected with carbon fiber electrodes [20,35]. Diffusion of DA in the extrasynaptic extracellular space is limited by DA uptake. Therefore, in the striatum, where uptake is very active, half of the released DA is cleared at a diffusion distance of 7–12 μM [20,35,36]. Thus, dopaminergic transmission is extrasynaptic but also highly localized. This conclusion points out the value of carbon fiber electrodes over microdialysis probes. The size of the former (5–8 μM in diameter) is in the same range as the average diffusion distance of DA, whereas that of the microdialysis probes (200 μM in diameter) is much larger.

Release And Elimination Of Dopamine From The Extracellular Space

Release and Diffusion of Dopamine in the Extracellular Fluid to the Electrode

The exocytotic release of DA from dopaminergic terminals is extremely fast: it occurs within less than 1 ms, as reported from the amperometric monitoring of quantal release in cultured dopaminergic neurons [14]. Unfortunately, this fast quantal release cannot be recorded in vivo or in striatal slices because it is not possible to position a carbon fiber electrode in direct contact with one or several intact release sites. The electrode penetration creates a dead zone of lesioned tissue between the intact release sites and the active surface of the electrode. The thickness of this dead zone has been estimated to be between 6 and 9 μM [5,26,36]. These estimates seem reasonable given the fact that the diameter of the carbon fibers range from 5 to 8 μM.

Elimination of Released Dopamine by Reuptake

Many pharmacological studies have shown that reuptake by the DA transporter plays a major role in the clearance of released DA although other pathways of elimination, such as extracellular degradation, have been considered. Recent studies using mice lacking the DA transporter show that reuptake is the only mechanism responsible for the clearance of released DA. In the striatum and nucleus accumbens of these mice, the half-life of released DA is slowed down by two orders of magnitude [18,46]. The half-life is not affected by inhibitors of catechol-Omethyl transferase or noradrenaline or serotonin transporters and is only slightly slowed by inhibition of monoamine oxidase [18,46]. Likewise, in the basal nucleus of the amygdala, DA uptake seems to be the only mechanism responsible for the elimination of released DA [47]. In the prefrontal cortex it appears likely that the noradrenaline transporter also contributes to the clearance of DA [48]. DA reuptake has been modeled with Michaellis–Menten kinetics. It is a slow phenomenon compared to exocytotic release. Indeed, the half-life of DA has been estimated to range from 25 to 40 ms (see above).

Monitoring the Evoked Dopamine Overflow with Carbon Fiber Electrodes

The extracellular DA concentration results from a dynamic equilibrium between release and reuptake. When DA overflow is evoked by a brief electrical stimulation, the rising phase of the oxidation signal reflects release minus reuptake and the decay phase only reuptake. Because release is much faster than reuptake, the rising phase should be much sharper than the decay phase. However, diffusion between intact dopaminergic terminals and the electrode surface through the dead zone equally slows down both phases. This explains why the transient oxidation signal evoked by brief electrical stimulation appears only slightly asymmetrical when recorded in the striatum (Figure 3.1). The asymmetry increases in the nucleus accumbens and olfactory tubercle because reuptake is less efficient in these limbic areas than in the striatum [23] and is even more pronounced in the basal nucleus of the amygdala where reuptake is much slower [26]. Nevertheless, when compared under the same experimental conditions with the same electrode, transient oxidation signals appear more asymmetrical when recorded with continuous amperometry than with FSCV [5]. Therefore, continuous amperometry is more accurate than FSCV for the independent estimate of release and reuptake parameters from evoked oxidation responses.

FIGURE 3.1. Effects of train length and of cocaine on evoked DA overflow.


Effects of train length and of cocaine on evoked DA overflow. The transient changes in oxidation current were monitored with a carbon fiber electrode held at +0.4 V by continuous amperometry in the striatum of a C57 mouse as described [18]. MFB electrical (more...)

Technical Recommendations

Experimental Protocols

Carbon Fiber Electrodes

Two shapes of carbon fiber electrodes have been developed: disk electrodes, in which the active surface is the section of one carbon fiber, and cylindrical carbon fiber electrodes, in which the active surface is that of a single carbon fiber protruding for 25–250 μM out of an insulating glass pipette. The first advantage of cylindrical electrodes over disk electrodes is that their active area is much larger. Because the oxidation current is proportional to the active area, this shape facilitates the detection of low DA concentrations. The second advantage derives from the fact that the cylindrical surface represents the main active area and parallels the penetration axis. During implantation into brain tissues, the tip of the electrode is covered with tissue fragments, but most of the cylindrical surface remains uncovered (see Figure 1 in Gonon [49]). Early on, FSCV was performed with disk-shaped carbon fiber electrodes, but cylindrical electrodes are increasingly used in combination with FSCV in recent in vivo studies (see Chapter 2). Continuous amperometry has always been combined with cylindrical electrodes [17,26].

Several procedures have been developed to produce cylindrical carbon fiber electrodes. The main difficulty is that one must use a resin to ensure a good seal between the carbon fiber and the glass pipette, but the resin must not cover the active electrode surface. We developed a procedure in which a polyester resin is pushed into the glass pipette from its back, unpulled end [50]. The resin reaches the end of the pulled tip of the pipette by capillary action and, normally, does not cover the protruding part of the carbon fiber. However, this procedure is quite delicate. With other procedures, an epoxy resin fills the pipette tip by capillary action from the pulled end of the pipette, and the electrode is immediately washed in acetone to remove residual epoxy from the active part of the carbon fiber [26].

Several types of carbon fibers have been used to produce electrodes. Because carbon fibers are usually produced to strengthen composite products, producers treat them to improve the adhesion of resin to carbon. This treatment is called sizing and alters the electrochemical properties of carbon fibers. Therefore, electrodes must be produced with unsized carbon fibers. In our team, we compared three types of unsized carbon fibers for their efficacy in combination with continuous amperometry: (1) AGT 10000, produced by SEROFIM, France, with a diameter of 8 μM (these fibers are no longer produced); (2) T300J, produced by SOFICAR, France, under the TORAYCA license, with a diameter of 7 μM; and (3) unsized carbon fiber supplied by Goodfellow Corporation, Devon, PA (reference product C 005722), with a diameter of 7 μM. The third type of fibers was found to be the most efficient and was used in our most recent study [51].

Implantation of Carbon Fiber Electrodes In Vivo

The implantation procedure must minimize the covering of the electrode with tissue fragments. The surgical approach must be performed under a microscope to avoid damaging the cortical surface. In rats, the cortical surface is protected by the pia mater, a sticky and transparent layer adhering to the cortical surface, and by the dura mater, a very strong membrane separated from the pia mater by cerebrospinal fluid. The dura mater must be surgically removed to give carbon fiber electrodes access to brain tissue. Direct implantation of carbon fiber electrodes through the pia mater is mechanically possible but results in fouling the active tip of the electrode with pia matter. To minimize fouling, we puncture the pia mater with a previously used carbon fiber electrode that has been stored for a few months in the open air. The resulting hole is marked with a thin, black ring of dust at the cortical surface. Then, a new carbon fiber electrode is implanted through this ring. In mice, the dura mater is thinner and adheres to the cortical surface. Therefore, it cannot be surgically removed. New carbon fiber electrodes are implanted through a hole formed in the dura mater as described above.

After an initial in vivo experiment, carbon fiber electrodes may be reused after washing off any tissue fragments with a water solution containing 5% triton X100. However, the electrode performance tends to decrease with successive in vivo experiments. The maximal number of experiments performed in our lab with the same electrode is three in rats and five in mice.

Reference Electrode

As in almost all electrochemical studies applied to neuroscience, our reference electrode is of the Ag/AgCl type. The active part is a silver wire 0.5 mm in diameter and 10 mm long (reference product 7890, supplied by A-M Systems, INC, Everet, WA). It is electrochemically coated with silver chloride by anodizing it in hydrochloric acid (1 N) at +0.65 V for 1 min. During in vivo recording, the active portion of the reference electrode is put between two sponges (2 mm thick) moistened by physiological saline solution (NaCl 9 g/L) and spread out on the skull. Reference electrodes must be renewed every five to ten in vivo experiments.

In Vitro Calibration of Carbon Fiber Electrodes

After in vivo experiments, the sensitivity of the carbon fiber electrodes to DA is tested in a flow injection system, originally developed for FSCV [52] and used for continuous amperometry [17,32]. For reasons given above (section entitled Comparison with Fast Scan Cyclic Voltammetry), the perfusion fluid must contain at least 100 μM of ascorbic acid. Because the comparison of in vitro calibrations performed before and after in vivo implantation shows a substantial decrease in sensitivity induced by implantation, reliable in vitro calibration must be performed after in vivo implantation. However, the use of in vitro calibration to estimate the absolute amplitude of an evoked overflow in terms of DA concentration must be considered with caution. Indeed, electrochemical detection is affected by several parameters that cannot be easily controlled under in vivo conditions: diffusion coefficient, pH, and the presence of compounds that are not electroactive by themselves but which might interfere with DA detection.

Analysis Of The Data

Direct Estimate of Dopamine Release and of Dopamine Uptake

With high frequency stimulation (100 Hz), the maximal amplitude of the oxidation responses is linearly related to the number of pulses between one and four pulses, either in untreated animals (Figure 3.1; see also [18]), or when DA uptake is inhibited by nomifensine [19]. Indeed, during these brief stimulations, DA uptake is negligible over DA release. Therefore, the maximal amplitude of the responses evoked by these brief stimulations is a good index of the amplitude of the DA release per pulse. For example, this index was used to show that DA release was reduced by 65% in the striatum of mice treated with MPTP [21] and that STOP null mice exhibit a higher DA release than control mice [53].

As illustrated in Figure 3.1, the half-life, measured from the decay phase of the evoked responses, is a good index of the efficiency of DA uptake. Indeed, half-life is increased by up to one order of magnitude after pharmacological inhibition [23,51] and by two orders of magnitude in mice lacking the DA transporter [18]. However, uptake inhibition also enhances the maximal amplitude of the evoked responses, even if the stimulation parameters (high frequency and number of pulses ≤4) are chosen to minimize uptake influence (Figure 3.1) [51]. Indeed, uptake inhibition favors diffusion of DA from release sites to the electrode surface [26]. Therefore, variations in the maximal amplitude of the responses evoked by brief stimulations reflect changes in the DA release per pulse provided that they are not associated with alterations of DA uptake. For example, Brun et al. [53] observed that the response evoked by single pulse stimulation was larger in STOP null mice compared to wild type mice. They attributed this increase to enhanced DA release because they showed that, in a wide range of comparable overflow amplitudes, t1/2 was similar in both genotypes.

DA reuptake is often modeled with Michaellis–Menten kinetics, in which two parameters are defined: the affinity parameter (KM) and the maximal velocity (Vmax). The half-life (t1/2 in Figure 3.1) depends on both KM and Vmax. Estimation of KM from the evoked DA overflow requires a mathematical model (see below). However, a rough estimate of Vmax can be directly obtained from the evoked DA overflow. Indeed, Vmax roughly corresponds to the maximal slope of the decay phase after stimuli of sufficient intensity (e.g., 20–50 pulses at 50–100 Hz) to cause DA to reach levels sufficient to saturate the DAT [53].

Mathematical Models

As discussed above, the evoked DA overflow results from DA release minus DA uptake. However, Wightman et al. already reported in 1988 that models based only on these two processes poorly fit FSCV recordings. Better fit was obtained by taking into account the diffusion of DA from intact release sites to the electrode [54]. Three models have been recently developed to simulate recordings obtained with continuous amperometry [5,26,55]. All of them take into account the diffusion of DA from intact release sites to the electrode. The model proposed by Venton et al. [26] represents an improvement over the former ones because it takes into account the fact that ascorbic acid reduces DA molecules oxidized by the electrode. The fit of this model with experimental recordings performed under highly diverse experimental conditions is indeed impressive.


Continuous amperometry has been used in brain slices to monitor the electrically evoked DA overflow and compared to FSCV in the same experimental conditions. In slices, continuous amperometry provides the same pieces of information as FSCV but with a better time resolution [5]. The use of continuous amperometry, both in striatal slices and in the striatum of anesthetized rodents, might help to resolve a puzzling question: the characteristics of the electrically evoked DA release strongly differ when studied in vitro or in vivo. Up to now, attempts to resolve this issue have been unsuccessful [56].

Continuous amperometry, combined with untreated cylindrical carbon fiber electrodes, is a valid technique to monitor the DA overflow evoked by brief electrical stimulations in anesthetized rats and mice. In these restricted experimental conditions, continuous amperometry is superior to FSCV because it provides the same pieces of information but with better time resolution [26]. Moreover, continuous amperometry is an easier technique, with minimal requirement in terms of electrochemical equipment. Direct measurement of the maximal amplitude of the evoked signals, and of the time for 50% decay from the maximum, provides fairly independent measurements of the two mechanisms that control extracellular DA levels: DA release per pulse and DA uptake. These direct estimates are sufficient in most experimental conditions for studying the parameters that regulate release and uptake. In more demanding experimental conditions, which associate prominent changes in both release and uptake, mathematical models can be used for a more accurate and independent measurement of release and uptake [26]. Thus, FSCV, and, more recently, continuous amperometry, have greatly contributed to the knowledge of the presynaptic mechanisms regulating dopaminergic transmission (also see the review by Schmitz et al.) [57] In the future, the use of continuous amperometry to monitor electrically evoked DA release, either in slices or in anesthetized animals, will undoubtedly help to tackle unresolved issues such as shortterm plasticity of dopaminergic transmission [57].


Gonon F, et al. Mesure électrochimique continue de la libération de dopamine réalisée in vivo dans le néostriatum du rat. CR Acad Sci Paris . 1978;286:1203. [PubMed: 96981]
Wightman RM, et al. Temporally resolved catecholamine spikes correspond to single vesicle release from individual chromaffin cells. Proc Natl Acad Sci. 1991;88:10754. [PMC free article: PMC53009] [PubMed: 1961743]
Pothos EN, Desmond M, Sulzer D. L-3,4-dihydroxyphenylalanine increases the quantal size of exocytotic dopamine release in vitro. J Neurochem. 1996;66:629. [PubMed: 8592133]
Gonon F, Msghina M, Stjärne L. Kinetics of noradrenaline released by sympathetic nerves. Neuroscience. 1993;56:535. [PubMed: 8255421]
Schmitz Y, et al. Amphetamine distorts stimulation-dependent dopamine overflow: effects on D2 autoreceptors, transporters, and synaptic vesicle stores. J Neurosci. 2001;21:5916. [PMC free article: PMC6763160] [PubMed: 11487614]
Gonon F, et al. In vivo electrochemical detection of catechols in the neostriatum of anaesthetized rats: dopamine or DOPAC? Nature. 1980;286:902. [PubMed: 7412872]
Gonon F, et al. Voltammetry in the striatum of chronic freely moving rats: detection of catechols and ascorbic acid. Brain Res. 1981;223:69. [PubMed: 7284811]
Ewing AG, Bigelow JC, Wightman RM. Direct in vivo monitoring of dopamine release from two striatal compartments in the rat. Science. 1983;221:169. [PubMed: 6857277]
Kuhr WG, et al. Monitoring the stimulated release of dopamine with in vivo voltammetry. I: characterization of the response observed in the caudate nucleus of the rat. J Neurochem. 1984;43:560. [PubMed: 6736965]
Gonon F, Navarre F, Buda M. In vivo monitoring of dopamine release in the rat brain with differential normal pulse voltammetry. Anal Chem. 1984;56:573. [PubMed: 6711824]
Gonon F, Buda M. Regulation of dopamine release by impulse flow and by autoreceptors as studied by in vivo voltammetry in the rat striatum. Neuroscience. 1985;14:765. [PubMed: 2986044]
Gonon F. Nonlinear relationship between impulse flow and dopamine release by rat midbrain dopaminergic neurons as studied by in vivo electrochemistry. Neuroscience. 1988;24:19. [PubMed: 3368048]
Suaud-Chagny MF, Buda M, Gonon FG. Pharmacology of evoked dopamine release studied in the olfactory tubercle by in vivo electrochemistry. Eur J Pharmacol. 1989;164:273. [PubMed: 2788097]
Pothos EN, Davila V, Sulzer D. Presynaptic recording of quanta from midbrain dopamine neurons and modulation of the quantal size. J Neurosci. 1998;18:4106. [PMC free article: PMC6792796] [PubMed: 9592091]
Leszczyszyn DJ, et al. Secretion of catecholamines from individual adrenal medullary chromaffin cells. J Neurochem. 1991;5:6. [PubMed: 2027003]
Rebec GV, Pierce RC. A vitamin as neuromodulator: ascorbate release into the extracellular fluid of the brain regulates dopaminergic and glutamatergic transmission. Prog Neurobiol. 1994;43:537. [PubMed: 7816935]
Dugast C, Suaud-Chagny MF, Gonon F. Continuous in vivo monitoring of evoked dopamine release in the rat nucleus accumbens by amperometry. Neuroscience. 1994;62:647. [PubMed: 7870296]
Benoit-Marand M, Jaber M, Gonon F. Release and elimination of dopamine in vivo in mice lacking the dopamine transporter: functional consequences. Eur J Neurosci. 2000;12:2985. [PubMed: 10971639]
Chergui K, Suaud-Chagny MF, Gonon F. Nonlinear relationship between impulse flow, dopamine release and dopamine elimination in the rat brain in vivo. Neuroscience. 1994;62:641. [PubMed: 7870295]
Gonon F. Prolonged and extrasynaptic excitatory action of dopamine mediated by D1 receptors in the rat striatum in vivo. J Neurosci. 1997;17:5972. [PMC free article: PMC6573191] [PubMed: 9221793]
Bezard E, et al. Adaptative changes in nigrostriatal pathway in response to increased 1-methyl-4- phenyl-1,2,3,6-tetrahydropyridine-induced neurodegeneration in the mouse. Eur J Neurosci. 2000;12:2892. [PubMed: 10971632]
Benoit-Marand M, Borrelli E, Gonon F. Inhibition of dopamine release via presynaptic D2 receptors: time course and functional characteristics in vivo. J Neurosci. 2001;21:9134. [PMC free article: PMC6763925] [PubMed: 11717346]
Suaud-Chagny MF, et al. Uptake of dopamine released by impulse flow in the rat mesolimbic and striatal systems in vivo. J Neurochem. 1995;65:2603. [PubMed: 7595557]
Rougé-Pont F, et al. Changes in extraces lular dopamine induced by morphine and cocaine: crucial control by D2 receptors. J Neurosci. 2002;22:3293. [PMC free article: PMC6757537] [PubMed: 11943831]
Dugast C, et al. On the involvement of a tonic dopamine D2-autoinhibition in the regulation of pulseto- pulse-evoked dopamine release in the rat striatum in vivo. Naunyn Schmiedebergs Arch Pharmacol. 1997;355:716. [PubMed: 9205955]
Venton BJ, et al. Real-time decoding of dopamine concentration changes in the caudate-putamen during tonic and phasic firing. J Neurochem. 2003;87:1284. [PubMed: 14622108]
Dugast C, Cespuglio R, Suaud-Chagny MF. In vivo monitoring of evoked noradrenaline release in the rat anteroventral thalamic nucleus by continuous amperometry. J Neurochem. 2002;82:529. [PubMed: 12153477]
Westerink BH. Analysis of biogenic amines in microdialysates of the brain. J Chromatogr B Biomed Sci Appl. 2000;747:21. [PubMed: 11103897]
Lu Y, Peters JL, Michael AC. Direct comparison of the response of voltammetry and microdialysis to electrically evoked release of striatal dopamine. J Neurochem. 1998;70:584. [PubMed: 9453552]
Yang H, Peters JL, Michael AC. Coupled effects of mass transfer and uptake kinetics on in vivo microdialysis of dopamine. J Neurochem. 1998;71:684. [PubMed: 9681459]
Michael DJ, Wightman RM. Electrochemical monitoring of biogenic amine neurotransmission in real time. J Pharm Biomed Anal. 1999;19:33. [PubMed: 10698566]
Venton BJ, Troyer KP, Wightman RM. Response times of carbon fiber microelectrodes to dynamic changes in catecholamine concentration. Anal Chem. 2002;74:539. [PubMed: 11838672]
May LJ, Kuhr WG, Wightman RM. Differentiation of dopamine overflow and uptake processes in the extracellular fluid of the rat caudate nucleus with fast-scan in vivo voltammetry. J Neurochem. 1988;51:1060. [PubMed: 2971098]
Budygin EA, et al. Effect of tolcapone, a catechol-O-methyltransferase inhibitor, on striatal dopaminergic transmission during blockade of dopamine uptake. Eur J Pharmacol . 1999;370:125. [PubMed: 10323260]
Garris PA, et al. Efflux of dopamine from the synaptic cleft in the nucleus accumbens of the rat brain. J Neurosci. 1994;14:6084. [PMC free article: PMC6577011] [PubMed: 7931564]
Gonon F, et al. Geometry and kinetics of dopaminergic transmission in the rat striatum and in mice lacking the dopamine transporter. Prog Brain Res. 2000;25:291. [PubMed: 11098665]
Claustre Y, et al. SSR181507, a dopamine D(2) receptor antagonist and 5-HT(1A) receptor agonist I: neurochemical and electrophysiological profile. Neuropsychopharmacology. 2003;28:2064. [PubMed: 12902994]
Marcenac F, Gonon F. Fast in vivo monitoring of dopamine release in the rat brain with differential pulse amperometry. Anal Chem. 1985;57:1778. [PubMed: 4037340]
Suaud-Chagny MF. In vivo monitoring of dopamine overflow in the central nervous system by amperometric techniques combined with carbon fibre electrodes. Methods. 2004;33:322. [PubMed: 15183181]
Suaud-Chagny MF, et al. Fast in vivo monitoring of electrically evoked dopamine release by differential pulse amperometry with untreated carbon fibre electrodes. J Neurosci Methods. 1992;45:183. [PubMed: 1363483]
Levey AI, et al. Localization of D1 and D2 dopamine receptors in brain with subtype-specific antibodies. Proc Natl Acad Sci. 1993;90:8861. [PMC free article: PMC47460] [PubMed: 8415621]
Sesack SR, Aoki C, Pickel VM. Ultrastructural localization of D-2 receptor-like immunoreactivity in midbrain dopamine neurons and their striatal targets. J Neurosci. 1994;14:88. [PMC free article: PMC6576858] [PubMed: 7904306]
Yung KK, et al. Immunocytochemical localization of D1 and D2 dopamine receptors in the basal ganglia of the rat: light and electron microscopy. Neuroscience. 1995;65:709. [PubMed: 7609871]
Caillé I, Dumartin B, Bloch B. Ultrastructural localization of D1 dopamine receptor immunoreactivity in the rat striatonigral neurons and its relation with dopaminergic innervation. Brain Res. 1996;730:17. [PubMed: 8883884]
Nirenberg MJ, et al. The dopamine transporter: comparative ultrastructure of dopaminergic axons in limbic and motor compartments of the nucleus accumbens. J Neurosci. 1997;17:6899. [PMC free article: PMC6573281] [PubMed: 9278525]
Giros B, et al. Hyperlocomotion and indifference to cocaine and amphetamine in mice lacking the dopamine transporter. Nature. 1996;379:606. [PubMed: 8628395]
Garris PA, Wightman RM. Distinct pharmacological regulation of evoked dopamine efflux in the amygdala and striatum of the rat in vivo. Synapse. 1995;20:269. [PubMed: 7570359]
Valentini V, Frau R, Di Chiara G. Noradrenaline transporter blockers raise extracellular dopamine in medial prefrontal but not parietal and occipital cortex: differences with mianserin and clozapine. J Neurochem. 2004;88:917. [PubMed: 14756813]
Gonon F. Monitoring dopamine and noradrenaline release in central and peripheral nervous systems with treated and untreated carbon-fiber electrodes. In: Boulton A, Baker G, Adams RN, editors. Voltammetric Methods in Brain Systems. Humana Press; Clifton, NJ: 1995. (chap. 5)
Gonon F, Buda M, Pujol JF. Treated carbon fiber electrodes for measuring catechols and ascorbic acid. In: Marsden CA, editor. Measurement of Neurotransmitter Release In Vivo. Wiley, Chichester; U.K.: 1984. chapter 7.
Samaha AN, et al. The rate of cocaine administration alters gene regulation and behavioral plasticity: implications for addiction. J Neurosci. 2004;24:6362. [PMC free article: PMC6729536] [PubMed: 15254092]
Kristensen EW, Wilson RL, Wightman RM. Dispersion in flow injection analysis measured with microvoltammetric electrodes. Anal Chem. 1986;58:986.
Brun P, et al. Dopaminergic transmission in STOP null mice. J Neurochem. 2005;94:63. [PubMed: 15953350]
Wightman RM, et al. Real-time characterization of dopamine overflow and uptake in the rat striatum. Neuroscience. 1988;25:513. [PubMed: 3399057]
Schonfuss D, et al. Modelling constant potential amperometry for investigations of dopaminergic neurotransmission kinetics in vivo. J Neurosci Methods. 2001;112:163. [PubMed: 11716951]
Benoit-Marand M, Schmitz Y, Gonon F, Zhang H, Sulzer D. Monitoring dopamine release under in vivo and in vitro conditions. Monitoring Molecules in Neorosciences; Proceedings of the 9th International Conference on In Vivo Methods; National University of Ireland, Dublin. 2001. p. 227.
Schmitz Y, et al. Presynaptic regulation of dopaminergic neurotransmission. J Neurochem. 2003;87:273. [PubMed: 14511105]
Copyright © 2007, Taylor & Francis Group, LLC.
Bookshelf ID: NBK2576PMID: 21204390


  • PubReader
  • Print View
  • Cite this Page

Other titles in this collection

Related information

  • PMC
    PubMed Central citations
  • PubMed
    Links to PubMed

Similar articles in PubMed

See reviews...See all...

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...