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

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Chapter 11Hydrogen Peroxide as a Diffusible Messenger: Evidence from Voltammetric Studies of Dopamine Release in Brain Slices

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Introduction

Reactive oxygen species (ROS) are often considered to be toxic byproducts of cell metabolism. Increased ROS production and oxidative stress contribute to cell death after acute brain injury, as well as in slowly progressing neurodegenerative disorders, including Parkinson’s disease [1,2]. However, this view of ROS is rapidly evolving in light of increasing evidence that ROS also act as cellular messengers that can modulate processes from short-term ion-channel activation to gene transcription. Hydrogen peroxide (H2O2) is a particularly intriguing candidate as a signaling molecule because it is neutral and membrane-permeable [3], and therefore can diffuse freely from a site of generation, like the established diffusible messengers, nitric oxide and carbon monoxide. Moreover, H2O2 has no unpaired electrons and is therefore not a free radical, in contrast to superoxide (·O2) and the hydroxyl radical (OH), and so does not readily cause oxidative damage [4].

We have recently discovered that endogenously generated H2O2 regulates the release of dopamine (DA). DA is an important neurotransmitter and neuromodulator that participates in a wide range of brain functions, including learning and cognition [5], control of movement [6,7], and mediation of desire and reward [8]. Consequently, dysfunction of dopaminergic transmission has been implicated in several significant brain disorders, including the psychoses of schizophrenia, the movement deficits of Parkinson’s disease, and addiction to substances like alcohol and cocaine. Studies of DA release regulation using voltammetric methods with carbon-fiber microelectrodes have made significant contributions to the present understanding of DA regulation and the role of DA in normal and pathological conditions. A number of key mechanistic insights have been obtained using these methods in brain slices in vitro, which readily permits voltammetric detection of DA release in discrete brain regions, without complicating factors inherent to in vivo studies, including animal behavior, anesthesia, and indirect effects from distant structures via long pathways.

In this chapter, we will discuss various methodological considerations for the use of in vitro voltammetric recording in brain slices, which will indicate the types of questions that these methods are best suited to address. In addition, we will review evidence that H2O2 is a diffusible messenger that mediates glutamate-dependent inhibition of DA release in dorsolateral striatum, based on data obtained using fast-scan cyclic voltammetry (FCV) with carbon-fiber microelectrodes in brain slices [9,10]. These studies also indicate the mechanism of release suppression, which is the activation of ATP-sensitive potassium (KATP) channels. Companion electrophysiological studies show that H2O2 also regulates the spontaneous activity of DA neurons in the substantia nigra pars compacta (SNc) via KATP channels [11], consistent with the apparently abundant expression of these channels throughout the nigrostriatal pathway [12–14]. Together, these studies illustrate how voltammetric methods in brain slices, both alone and in combination with companion techniques, can be used to address important questions in neuroscience.

Overview

Physiology of DA Release: Why It Can Be Detected in the Extracellular Space

Dopaminergic cell body regions within the central nervous system (CNS) are subdivided into cell groups A8–A17 based on the classification system of Dahlström and Fuxe [15]. The principal DA neuronal pathways arise from midbrain nuclei designated as the A9 and A10 cell groups, which are the SNc and ventral tegmental area (VTA). These project ipsilaterally via the medial forebrain bundle (MFB) to various forebrain structures. Dopaminergic neurons in the SNc send axon projections via the nigrostriatal pathway to the dorsolateral striatum (caudate putamen, CPu), which is critically involved in the control of movement [6,7]. DA neurons in the VTA project via the mesolimbic pathway to the ventromedial striatum (nucleus accumbens, NAc), which is involved in motivation and reward [8]. Additionally, DA neurons of the VTA project via the mesocortical pathway to the amygdala and prefrontal cortex, which have been implicated in the control of emotion and the pathophysiology of schizophrenia, respectively.

Synthesis of DA occurs in the cytosol of DA cell bodies and axons from the amino acid tyrosine; the first step mediated by tyrosine hydroxylase (TH) is rate limiting. Cytosolic DA is taken up into intracellular storage vesicles via the vesicular monoamine transporter 2 (VMAT2). Release of DA from axon terminals, like that of the primary excitatory transmitter glutamate and the primary inhibitory transmitter GABA (γ-aminobutyric acid), occurs via two main mechanisms: impulse-dependent (vesicular) and carrier-mediated release. With impulse-dependent DA release, Na+-dependent action potentials trigger Ca2+ entry via the opening of voltage-gated Ca2+-channels [16]. In DA axons, this catalyzes a process in which VMAT2-expressing vesicles that contain DA fuse with the presynaptic membrane to release their contents into a synapse or the extracellular environment via exocytosis (Figure 11.1). The amplitude and time course of an increase in extracellular DA concentration ([DA]o), after stimulation of a population of DA cells or axons, are determined by the competing processes of release and uptake. Uptake of DA is mediated by the dopamine transporter (DAT), which is found only on the plasma membrane of DA cells and axons [17–20] (Figure 11.1). However, conditions that cause the cytoplasmic concentration of DA to exceed that in the extracellular environment can lead to carrier-mediated DA release via reversal of the DAT, which is a Ca2+-independent process. Under normal physiological conditions, impulse-dependent exocytosis is the primary mechanism of axonal DA release.

FIGURE 11.1. Relationship between distance and DATs on [DA]o transients following quantal release in striatum.

FIGURE 11.1

Relationship between distance and DATs on [DA]o transients following quantal release in striatum. Schematic representations of a releasing DA synapse and a neighboring synapse on dendritic spines of striatal medium spiny neurons showing extrasynaptic (more...)

In contrast to fast, synaptically acting neurotransmitters like glutamate, DA acts on a slower time scale as a neuromodulator that sets a tone for local excitability. Consequently, the actions and regulation of DA differ in range and time course from those of “conventional” transmitters [20]. Released DA activates two broad classes of DA receptors, classified as D1-like and D2- like [21,22]. Both classes of DA receptors are G-protein coupled, with response times exceeding 100 ms [21]. By contrast, glutamate primarily mediates fast synaptic transmission via ionotropic receptors that generate excitatory postsynaptic potentials (EPSPs) lasting only 1–2 ms [23,24]. Moreover, the locations of DA and glutamate receptors differ. Both classes of DA receptors are located predominantly outside a synapse (extrasynaptic) [17,18], whereas glutamate receptors are primarily found within a synapse (subsynaptic) [23,24].

The extrasynaptic location of DA receptors implies that DA must escape a synapse and diffuse to its receptors to mediate local effects [20,25,26]. Consistent with this functional requirement, the DAT is also found extrasynaptically on DA axons [27]. Immediately after release, a DA molecule diffusing in three-dimensional extracellular space will be removed only when it encounters a DA axon (Figure 11.1). By contrast, glutamate synapses are literally surrounded by glutamate transporters, which are located not only on pre- and postsynaptic sites within a synapse, but also on glial membranes that ensheath glutamate synapses [28,29]. This three-dimensional barrier strongly limits glutamate “spillover” from one synapse to another.

Overall, DA neurotransmission is designed to occur primarily in the extracellular space. This makes DA an ideal transmitter substance to monitor with sensors that greatly exceed the <10 nm diameter of a synaptic cleft, e.g., 8–30 μ carbon-fiber microelectrodes. Although there continues to be interest in further electrode miniaturization to enable DA detection within a synapse, this is not really “where the action is.” Rather, changes in [DA]o monitored with voltammetric methods provide a direct indication of DA transmission, since the extent of activation of extrasynaptic DA receptors will be governed by the amplitude and duration of a given increase in [DA]o.

A Brief History of In Vivo and In Vitro Voltammetry

The use of voltammetric methods to study biological substances in vivo was first described in the mid-1950s, when Leland Clark introduced a membrane covered Pt electrode to monitor oxygen levels in blood and other tissues using cathodic reduction of oxygen [30]. Other electrode materials, including glassy carbon [31], were introduced later to examine the anodic oxidation of exogenous marker substances, including ascorbic acid and hydrogen. Subsequently, Ralph Adams introduced carbon paste as an electrode material with better characteristics for oxidation reactions than either Pt or glassy carbon [32].

The first report of anodic voltammetry to study endogenous neuroactive substances in vivo was by Adams and his colleagues, who used miniaturized carbon paste electrodes to examine neurotransmission in the rat CPu [33]. Although the initial goal of those studies was to monitor DA, the primary substance detected in the absence of stimulation was ascorbate. Interference from ascorbate held back progress in voltammetric studies of DA release in vivo for several years while researchers explored a variety of methods to overcome this problem. Several other applications of voltammetry for in vivo monitoring were introduced during this time, including detection of activity-dependent increases in DA metabolites in cerebrospinal fluid [34]. Eventually, two successful approaches were developed to eliminate interference from ascorbate. First, Gonon et al. [35,36] introduced a method for anodic pretreatment of carbon fibers that modifies an electrode surface to permit detection of DA and ascorbate oxidation at distinct potentials. At unmodified carbon surfaces, there is normally an overpotential for irreversible ascorbate oxidation that raises the required potential to roughly that required for DA oxidation (typically 0.3–0.5 V versus Ag/AgCl with slow scan methods or amperometry), despite the lower thermodynamic oxidation potential for ascorbate versus DA. Anodic pretreatment decreases the ascorbate overpotential by ~200 mV so that ascorbate and DA can be monitored simultaneously, but independently. The second approach for eliminating the ascorbate interference was to coat electrodes with Nafion®, a perfluorosulfonated ionomer that excludes anionic ascorbate, as well as the DA metabolite DOPAC (3,4-dihydroxyphenylacetic acid), while extracting cationic DA [37,38]. Coating with Nafion® not only improves the selectivity of voltammetric electrodes for DA, but also protects the electrode surface during tissue recording, which helps maintain sensitivity for DA detection.

Three additional developments in the late 1970s and early 1980s revolutionized the use of voltammetric methods to examine DA release and have led to widespread acceptance of these methods in the neuroscience community today. The first of these was the introduction of carbon-fiber microelectrodes for in vivo voltammetry by two groups working independently [35,36,39,40]. The carbon fibers used were typically 7–10 μm in diameter, which produced much smaller microelectrodes than the 150–300-μm carbon paste-based electrodes first introduced for in vivo recording (e.g., [33,37]). Carbon-fiber microelectrodes approached physiological dimensions and caused minimal damage when implanted in living tissue; indeed recent studies indicate that there is little cellular damage at distances >7 μm from the center of a carbon-fiber microelectrode track [41]. The second development was the discovery that a pure DA release signal in the striatum could be elicited by electrical stimulation of the dopaminergic MFB pathway [42–44]. This observation was critical because it eliminated the need for electrode pretreatment or Nafion® coating, although both methods are still used for some applications.

The third key development was the introduction of in vitro voltammetric monitoring of neuroactive substances, initially ascorbate, in viable brain slices [45,46]. This methodology was soon extended to the detection of evoked DA release in acute striatal slices [47–49], and eventually to measurements of spontaneous DA release in organotypic slice culture [50]. A creative advance that followed from these first in vitro studies was to use voltammetric methods, specifically amperometry, to monitor quantal release of catecholamines from isolated chromaffin cells [51,52]. Amperometry was subsequently applied to the study of DA release from cultured CNS neurons, including carotid glomus cells [53] and midbrain DA neurons [54], and later to quantal release of DA from substantia nigra neurons in acute midbrain slices [55].

Together, these developments demonstrated the value and feasibility of using voltammetric methods to study DA release in vitro, as well as in vivo. Reflecting this, the titles of most voltammetric studies published in the last decade confidently state the result, rather than the methods used, indicating that these electrochemical techniques have become an accepted part of state-of-the-art methods in neuroscience.

Advantages of Brain Slices to Study DA Release

Brain slices provide a valuable experimental preparation in which intact, but isolated, brain regions containing ensembles of neurons and glia can be readily identified and studied under controlled conditions. In brain slice preparations, DA release elicited by local stimulation, whether the stimulus is chemical (e.g., high K+) or electrical, is a “population response” such that monitored [DA]o reflects the sum of DA released from the local population of all DA axons, or cell bodies and dendrites, within the stimulated region. Stimulation procedures are discussed in “Stimulating and Recording DA Release in Brain Slices” in this chapter.

Key factors that can be controlled in brain slices include recording position, composition of the extracellular fluid, and the concentration of applied drugs or other agents. The ability to identify a region of interest is of obvious importance. For voltammetric recording in vivo, electrodes are inserted through the skull via burr holes made over the region to be studied, then lowered according to coordinates from a stereotaxic atlas of the brain for the species examined, often a rat (e.g., [56]). Using these procedures, electrode placement in vivo is essentially blind and requires post-experiment confirmation of electrode position. Although electrode placement is usually not a problem for large brain structures like the striatum, accurate placement in smaller, deeper structures can be difficult. For example, voltammetric recording in midbrain slices has been successfully used to study somatodendritic DA release in the SNc, which is a small DA cell body region in the midbrain [65,95,100]; however, comparable in vivo studies in the SNc have not been reported. A related advantage of brain slice preparations is that measurements of DA release can be combined with other methods for which ready visualization is beneficial, like electrophysiological recording [57–59], or essential, like fluorescence imaging [60,61]. Indeed, with appropriate instrumentation, a carbon-fiber microelectrode used for voltammetry can also be used for simultaneous electrophysiological recording in slices [57].

The ability to control the extracellular microenvironment in a brain slice is another advantage of this in vitro preparation. The composition of the artificial cerebrospinal fluid (aCSF) used to maintain brain slice viability (discussed in “aCSF Composition for Healthy Slices” in this chapter) typically mimics the ionic composition of CSF in vivo. However, aCSF composition can be varied in vitro, for example to examine the Ca2+ dependence of DA release [62], whereas normal ion homeostasis in the intact brain would preclude addressing such questions in vivo. Additionally, known concentrations of pharmacological agents can readily be added to aCSF. Such direct application circumvents two major problems with in vivo drug administration. First, with systemic administration of a given drug, actual drug concentration in a brain region of interest is difficult to pinpoint because of peripheral metabolism, variable blood–brain barrier permeability, and/or other pharmacokinetic complications. Second, the site(s) of action is often unclear with systemic drug administration. For example, if a decrease in evoked DA release in striatum were seen following systemic administration of a DA auto-receptor agonist in vivo, it would not be obvious, without further experimentation, whether this reflected a direct inhibitory effect within the striatum, an indirect effect due to autoreceptor-dependent suppression of DA neuron excitability in the SNc, or some combination of both. On the other hand, the question of whether DA autoreceptor activation has a direct, local effect on DA release in the striatum can be readily addressed using striatal slices [63–69].

It is important to consider the limitations of brain slice preparations, as well. Most obviously, the use of isolated brain tissue in vitro precludes evaluation of DA release induced by animal behavior and how this might be affected by systemic drug administration. Additionally, some questions are best addressed by considering the influence of distant structures connected by long pathways that cannot easily be preserved in slice preparations. Even then, however, slices can provide important mechanistic information to complement in vivo studies. For example, Phillips et al. [70] examined DA release and uptake in brain slices from rats that had received prenatal exposure to cocaine in order to assess long-term consequences on DA regulation when these animals reached adulthood.

Overall, the strengths of brain slice preparations outweigh the weaknesses for mechanistic studies of DA release in discrete brain regions without interference from systemic interactions that can complicate data interpretation in vivo. Although cultured or freshly dissociated neurons can also be used for such studies at the single cell level, slices provide a reduced preparation in which the geometric structure of brain tissue is preserved, including intact local circuitry and maintained neuron–glial interactions. This structural preservation facilitates the study of the regulation of DA release by other endogenous neurotransmitters and modulators, including those discussed later in “Regulation of Striatal DA Release in Hydroden Peroxide”. Lastly, direct, in situ voltammetric detection of [DA]o in slice preparations offers clear advantages over other methods that require efflux into the medium outside a slice (for review see [71]).

Voltammetric Methods

Comparison of Voltammetric Techniques

A range of voltammetric methods have been used to monitor DA release in vitro, as well as in vivo; the three most commonly used are amperometry, chronoamperometry, and FCV. Of these, FCV has been most widely used in brain slice studies; however, discussion of these methods will begin with the conceptually simplest, which is constant potential amperometry.

Constant Potential Amperometry

For amperometric detection of DA, a potential sufficient for DA oxidation is applied constantly to the working electrode. The potential required depends on the electrode used and must be determined experimentally; a range of +0.3 to +0.8 V versus Ag/AgCl can be found in the literature [55,59,69,72]. Given that the electrode potential is always sufficient to oxidize all molecules of DA (or other electroactive substances oxidizing at the same or lower potential) in contact with the electrode surface, the integrated current monitored during a release event is directly proportional to the number of molecules oxidized. An advantage of this method is that continual DA oxidation curtails release signals by enhancing DA “clearance” as DA molecules are electrochemically consumed, which can improve the resolution of two signals separated by <1 s. Therefore, amperometry is well-suited for detection and quantification of the release of DA from single vesicles, i.e., quantal release [54,55]. Additionally, because of the constant applied potential, amperometric measurements have little contribution from residual non-Faradaic current (capacitative or charging current), and are not subject to interference from pH or divalent ions like Ca2+ (J. Patel and M. Harrell, unpublished observations), which can be a problem with FCV, as discussed below.

On the other hand, there are several aspects of amperometric monitoring that limit its usefulness for the study of population-based DA release. First, amperometry alone provides little information about signal identity. Other electroactive substance with oxidation potentials similar to that of DA include ascorbate, the DA metabolite DOPAC, and the related monoamine transmitters, norepinephrine (NE) and serotonin (5-hydroxytryptamine; 5-HT). Consequently, amperometric measurements require the use of an independent method, like FCV [69,72], to confirm that the monitored substance is DA. Without this, the identity of the monitored substance(s) remains uncertain. Second, this technique cannot readily be used to estimate [DA]o after a release event because electrode calibration factors obtained using amperometry are flow-rate dependent [72]; again, however, FCV has been used as an adjunct method for quantitative assessment of a release response. Third, because constant-potential amperometry irreversibly consumes DA, this complicates assessment of DA uptake from release records obtained using this method [73,74]. Thus, although amperometry is conceptually simple, data interpretation can be complex.

Chronoamperometry

The earliest voltammetric method used for DA detection in brain slices was chronoamperometry [47], in which the potential is stepped briefly from a non-oxidizing holding potential to a voltage sufficient for DA oxidation, then returned to the holding potential. When first introduced, the duration of the voltage step for chronoamperometry was 1 s delivered at regular intervals of 20–30 s. However, more recent studies have used shorter pulses of 100 ms applied at 200 ms intervals, leading to the term “high-speed” chronoamperometry [75–78]. Like the closely related technique of amperometry, chronoamperometry permits DA detection with minimal interference from pH and Ca2+ [77], which offers particular advantages for studies of the ionic dependence of DA release, e.g., the Ca2+ dependence [76]. A typical voltage step in chronoamperometry is from 0 to +0.55 V versus Ag/AgCl and back, which allows for reduction of oxidized DA after the pulse. The signal-to-noise ratio is usually optimized by integrating the current detected during the last 70 ms of the oxidation step, and again during the reduction step. The reduction step not only minimizes sampling-induced DA depletion, but also provides assistance in the identification of a monitored substance, since the ratio of reduction/oxidation current (or S2/S1) for a given signal tends to be substance-specific. For example, the S2/S1 ratio for DA is typically >0.5 in calibration, albeit >1.0 in slices. By contrast, the ratio for 5-HT is <0.1 [75,76], which permits these substances to be distinguished when only one or the other is released. The variability of the S2/S1 ratio for DA in calibration versus in slices precludes 100% certainty of signal identification, such that the term “DA-like” is sometimes used in reports in which chronoamperometry was used (e.g., [75,76]).

Fast-Scan Cyclic Voltammetry

FCV is more widely used for monitoring DA release than amperometry or chronoamperometry for three main reasons: (1) reliable signal identification; (2) reduction of oxidized DA; and (3) a short sampling time that is commensurate with physiological events. The simplest voltage waveform used for FCV is a triangle wave (Figure 11.2a), with an initial positive-going ramp that starts from a typically negative initial potential, rises to potential that is more positive than the DA oxidation potential, then reverses as a negative-going ramp that returns to the initial potential, which is typically more negative than the potential required for reduction of oxidized DA. Other voltage waveforms have also been used for FCV measurements; indeed, the waveform first introduced began with an initial negative (cathodic) ramp, followed by a conventional triangle wave that returned to the most negative potential, with a final anodic ramp back to the starting point, so that the overall quadraphasic waveform was like a “W” [40]. More dramatically altered waveforms have also been used to facilitate the detection of a substance or multiple substances. For example, to examine DA release and oxygen consumption simultaneously, Kennedy et al. [79] used a scan that rose from a holding potential of 0.0 V versus Ag/AgCl to +1.0 V, reversed to −1.3 V, then returned to the holding potential, which encompassed potentials required for both DA oxidation and oxygen reduction.

FIGURE 11.2. Fast-scan cyclic voltammetry (FCV) voltage waveform and dopamine (DA) voltammograms.

FIGURE 11.2

Fast-scan cyclic voltammetry (FCV) voltage waveform and dopamine (DA) voltammograms. (a) Typical triangular voltage waveform applied to a carbon-fiber electrode with FCV. (b) Background current generated at the carbon-fiber electrode with FCV in the absence (more...)

Regardless of the waveform used, the DA oxidation peak in FCV scans typically occurs around +0.6 V versus Ag/AgCl, with a single reduction peak around −0.2 V (Figure 11.2). Although the voltammogram of 5-HT obtained with FCV has a similar oxidation peak potential to that of DA, the 5-HT voltammogram has two reduction peaks, neither of which coincide with the potential of the single DA reduction peak [49,80,81]. Thus, the characteristic DA voltammogram can be used as a “fingerprint” for signal identification (Figure 11.2c). The reduction of oxidized DA during the cathodic part of the FCV scan allows sampling of [DA]o following a release event, with minimal DA consumption, such that FCV is less destructive than amperometry. This characteristic helps ensure that evaluation of DA uptake and release parameters from release records obtained using FCV reflect physiological characteristics and not consequences of oxidative loss of DA from the detection method used [73,74].

Like high-speed chronoamperometric voltage steps, FCV scans are typically repeated at subsecond intervals, ranging from 250 ms (4 Hz) to 100 ms (10 Hz). However, in the case of FCV, the term “fast” (or sometimes “high-speed”) refers to the scan rate, typically 300–900 V/s [40,49,65,82], rather than the sampling interval. With these scan rates, a forward and backward sweep over a 2 V potential range, e.g., −0.7 to +1.3 V, takes ~5 ms (Figure 11.2), which only slightly exceeds the duration of action potentials in the SNc [11].

Overall, the positive characteristics of FCV have made it the electrochemical method of choice for in vitro studies of endogenous DA release. There are some caveats for this method, however. In contrast to the low non-Faradaic current component of amperometry or chronoamperometry, there is a high non-Faradaic background current in all FCV scans [40,82] (Figure 11.2b). An unfortunate physicochemical fact is that the background current generated with any linear scanning method is directly proportion to scan rate, whereas the Faradaic oxidation current for a given concentration of analyte is proportional to the square root of scan rate [83]. This creates an analytical challenge, since the background current at a carbon-fiber microelectrode is usually ~1 μA with FCV (Figure 11.2b), whereas a physiologically relevant increase in [DA]o of 1 μM at such an electrode would generate a DA oxidation current in the low nA range (Figure 11.2c). One strategy to address this problem is to improve electrode sensitivity for DA by adjusting FCV scan parameters. For example, increasing the anodic limit from +1.0 to +1.4 V versus Ag/AgCl can enhance electrode sensitivity several fold, thereby lowering the detection limit for DA [84].

To monitor the time course of a release response with FCV, current detected at the DA oxidation peak potential can be followed using a sample-and-hold circuit; and the change above background can then be amplified as needed. Signal identification, however, requires the storage of a background scan before a DA increase (Figure 11.2b) that can be digitally subtracted from subsequent scans to provide background-subtracted voltammograms of the monitored substance (Figure 11.2c). Background subtraction is often done off-line; in early studies, this was accomplished using a digital oscilloscope with subtraction capabilities that permitted identification of only the peak DA signal [43,82]. Subsequently, Julian Millar developed an instrument, the Millar Voltammeter, which stores a background scan that is subtracted from subsequent scans, so the Faradaic voltammogram corresponding to changes in [DA]o can be monitored directly in real time. The Millar Voltammeter was, until recently, commercially produced by PD Systems International, U.K., but now is available by special request to Dr Millar (Queen Mary, University of London, U.K.). More recently, improved software for use with 12- to 16-bit D/A boards (e.g., National Instruments, Austin, Texas) has been developed that permits off-line subtraction of all scans [85].

Electrodes

As noted earlier in this chapter, the first electrodes developed to monitor DA release in brain tissue were made using carbon paste, including carbon paste mixed with epoxy to improve electrode stability. These were superseded by carbon-fiber microelectrodes, which are now used in almost all in vitro or in vivo voltammetric studies. A number of methods for the fabrication of carbon-fiber microelectrodes suitable for brain slice recording have been described in the literature [39,55,77,82,86–91]. In all of these, the fundamental design consideration has been to achieve the highest signal-to-noise ratio and lowest detection limit possible. The basic procedure often involves inserting a carbon fiber in a glass capillary tube, then pulling it on a conventional microelectrode puller to provide a seal and insulate the fiber with glass, although other coatings have been used to provide insulation. Every attempt is then made to ensure a tight seal, because small gaps or cracks that draw solution by capillary action can lead to electrode noise.

The design of carbon-fiber microelectrodes to be used for brain slice recording should include an active surface that can be contained completely within a slice, which has a typical thickness of ≤400 μm. Such electrode surfaces can be prepared by beveling or cutting, or by chemical or electrical etching of the carbon-fiber tip. The resultant working surface is either a disk that is flush with surrounding insulation (shown for a beveled electrode in Figure 11.3a), the surface of a cylinder that extends ≤50 μm beyond the insulation (Figure 11.3b), or that of a slightly-extended fiber with a conical shape (Figure 11.3c). Longer electrodes, like the 500 μm cylinders introduced by Gonon et al. [36], work well in vivo, but are not practical for use in slices. Selection of electrode design depends on the application and the carbon fibers available. Beveled and conical electrodes, which have tip diameters of typically 2–4 μm (e.g., [91]), minimize tissue damage at the recording site. Cylindrical and conical electrodes have larger surface areas than disk electrodes, and therefore produce more current for a given DA concentration. This can improve absolute detection limits, with the caveat that background currents are also proportional to surface area, so that signal-to- noise ratio may not necessarily improve. An additional point is that some carbon fibers are “sized,” which means that they have a coating that may prevent electron transfer on the surface of the cylinder; in some cases, this can be removed by soaking fibers in an appropriate solvent, like acetone or dimethyl sulfoxide [91]. Also, isopropanol has also been used to clean electrode surfaces, whether sized or not [89].

FIGURE 11.3. Carbon-fiber microelectrodes used for voltammetric detection of dopamine (DA) release.

FIGURE 11.3

Carbon-fiber microelectrodes used for voltammetric detection of dopamine (DA) release. Carbon-fiber microelectrodes are typically prepared by inserting a carbon fiber into a glass capillary, then pulling in a conventional electrode puller to melt the (more...)

One final issue in making carbon-fiber microelectrodes is creating electrical contact between a 5–10 μm carbon fiber and the instrumentation required for voltammetric recording (Figure 11.3d). Poor contact with the fiber can be a major source of electrical noise. Materials used include Wood’s, metal, silver epoxy, silver paint, carbon paste, and conductive solutions like potassium acetate mixed with KCl, each of which can link the fiber to a larger diameter wire that is then used for connection to an appropriate headstage.

Brain Slice Methodology

The use of brain slices as model CNS preparations were first explored in the 1950s by McIlwain (e.g., [92]). Over time, optimal conditions and incubation media were established to prepare and maintain viable slices; many of the issues evolved, as well as a range of uses for brain slice preparations are discussed in the comprehensive book Brain Slices edited by Dingledine [93]. An important parameter for brain slices is slice thickness; this should not exceed 400 μm, which was determined experimentally to be the maximum thickness at which the core of the slice can receive adequate oxygen and glucose. On the other hand, the minimal thickness of acute slices is ~200 μm, because the cut surfaces of a slice contains a layer of damaged cells that can extend 50 μm into the tissue, such that a 200 μm slice provides at least 100 μm of viable tissue within the slice where DA release can be studied.

Brain slices are typically prepared from deeply anesthetized animals; after decapitation, the brain is carefully removed into ice-cold aCSF (composition is discussed in “aCSF Composition for Healthy Slices”) for 1–2 min to minimize on-going metabolism and possible damage from anoxia. In early studies, McIlwain and others prepared slices using a “chopper,” which is composed of a weighted arm with attached blade to cut slices quickly; McIlwain tissue choppers are commercially available (Brinkmann Instruments, Westbury, New York). Later designs for tissue slicing instrumentation incorporated a vibrating blade to cut slices submerged in ice-cold aCSF; these instruments include the now classic Vibratome (Ted Pella, Inc., Redding, California) and the more recently introduced Leica vibrating microtome (Leica Microsystems, Bannockburn, Illinois). The idea behind a vibrating microtome is that a plane of solution and foam generated at the edge of the vibrating blade precedes the blade and separates the slice from the tissue block, which minimizes compression injury to the tissue. Once slices are prepared, they are usually allowed to recover for at least an hour in appropriate media at room temperature before experimentation. The slices can be held in individual vials or in a larger group holding chamber (e.g., a large Petri dish); with continuous oxygenation, slices can remain viable for eight to ten hours at room temperature. The number of slices prepared from a given animal depends on the region of interest, with more slices possible from the striatum than from smaller structures like the SNc. If DA release is to be examined in more than one region from a given animal, optimal viability will be obtained if more than one slicer is available for tissue preparation; otherwise, the quality of the second region may be degraded from lack of oxygenation during slicing of the first tissue block.

For voltammetric recording, it is preferable to use a submersion chamber in which slices (and electrode tips) are continuously immersed in aCSF, rather than an interface chamber in which the slice surface is exposed to a humidified atmosphere of 95% O2/5% CO2. Removal of electrodes from the lipid environment of brain tissue into air can alter electrode sensitivity and response time. The use of a superfusion chamber also helps ensure the accuracy of electrode calibration data used to calculate evoked [DA]o, because electrodes can be readily calibrated immediately after recording by raising the electrode from the tissue into the superfusing aCSF, then adding a known concentration of DA. A related technical point is that electrodes calibrated in the recording chamber should be positioned near the superfusion inlet to avoid dilution of the calibration solution with DA-free aCSF in the surrounding chamber; such dilution can give an erroneously low calibration factor, thereby causing overestimation of evoked [DA]o in a tissue slice.

aCSF Composition for Healthy Slices

The composition of modified aCSF solutions used for brain slice studies is critical for slice viability. These solutions differ from in vivo CSF in several ways. First, aCSF is hyperglycemic: the glucose concentration of CSF is 1–3 mM, whereas aCSF typically contains 10 mM glucose. Second, aCSF is hyperoxic: the medium surrounding a brain slice is continually saturated with 95% O2/5% CO2 to provide normal O2 levels in the interior of the slice. Third, the superfusing aCSF is hypothermic: the temperature used in most studies of DA release in brain slices is 31–32°C, such that the tissue is somewhat below normal core body temperature of 37°C. This mild hypothermia helps preserve slice viability during recording periods that can last for several hours. In initial studies of electrically- evoked DA release in brain slices at 37°C, Bull and colleagues [49] found that evoked [DA]o in striatal slices was below detection limits; however DA release was readily detected when the temperature was lowered to 32°C. This reflected both enhanced temperature-dependent DA uptake via the DAT at 37°C, as well as poorer slice viability: even in the presence of a DAT inhibitor, stable DA release is difficult to maintain at 37°C [49]. A further decrease in temperature is required for brain slice preparation; as noted in the introduction to this section, slices prepared on a vibrating microtome are submerged in ice-cold aCSF to preserve slice viability during cutting.

A variety of media changes have also been introduced for slice preparation and for use in a recovery or holding chamber. Many of these are indeed beneficial, with the caveat that some changes like decreasing Ca2+ levels or adding glutamate receptor antagonists can have lasting effects on cell excitability. One particularly helpful approach is to use a HEPES-bicarbonate-buffered aCSF for slice preparation, which improves slice quality [94] without obvious effects on tissue physiology. This solution contains (in mM): NaCl (120); NaHCO3 (20); glucose (10); HEPES acid (6.7); KCl (5); HEPES sodium salt (3.3); CaCl2 (2); MgSO4 (2), equilibrated with 95% O2/5% CO2 [81,95,96]. Slices are then maintained in this solution at room temperature until they are used (at least 1 h, but up to 10 h). HEPES was originally included to enhance buffering capacity when the medium was not bubbled with 95% O2/5% CO2 during slicing; it was later found that inclusion of HEPES improves slice quality by preventing excessive water gain (edema) during the holding period [94].

In addition to being hyperglycemic, hyperoxic, and hypothermic, the aCSF used for brain slice studies also often has slightly elevated concentrations of K+ and/or Ca2+ compared to normal levels in CSF (3 mM for K+ and 1.2 mM for Ca2+) to enhance slice excitability and/or elevate Ca2+-dependent transmitter release. An example of aCSF composition is (in mM): NaCl (124); KCl (3.7); NaHCO3 (26); CaCl2 (2.4); MgSO4 (1.3); KH2PO4 (1.3); and glucose (10), equilibrated with 95% O2/5% CO2 [62,81,95]. An advantage of in vitro studies is that the aCSF composition can be readily altered to suit the experiment; for example, Ca2+ concentration can be varied to examine the Ca2+ dependence of DA release [62,76], oxygen and glucose can be lowered to study the effects of ischemia on DA release [96], or Mg2+ can be omitted to enhance glutamatergic NMDA (N-methyl-D-aspartate) receptor activation [97].

Species Selection

Selection of the species to be studied, like the other methodological options already discussed, depends on the questions asked. Many fundamental properties of DA release regulation have been examined in rats, which are often a natural choice simply because of the large body of literature available on many aspects of DA physiology, anatomy, biochemistry, and pharmacology in this species. On the other hand, the increasing availability of transgenic mice developed to address aspects of DA regulation, like DAT or D2 receptor knockouts, make the mouse the species of choice for some applications. The use of other species can offer specific advantages, as well. For example, the study of somatodendritic DA release in the SNc of small rodents is hindered by significant 5-HT innervation [98], such that 5-HT rather than DA can be the primary substance detected in rat and mouse midbrain slices [57,99–102]. The pattern of 5-HT innervation in guinea pig SNc differs, however, such that pure DA release can be detected [62,95,100]. Thus, guinea pigs are the species of choice for studies of somatodendritic DA release in the SNc, although rats or mice also offer advantages for studying 5-HT in this region.

The use of brain slices also permits species comparisons of DA release regulation in animals for which in vivo recording is technically challenging, including non-human primates and birds. For example, Cragg and colleagues [103,104] examined axonal DA release probability and uptake in motor versus limbic regions of striatal slices from the marmoset; these studies revealed a different and more extensive pattern of DA regulation and plasticity in primate brain than seen in rodent striatum. Gale and Perkel [105] recently examined DA release in slices containing the songbird striatal analog, Area X, to assess the extent to which avian Area X is like mammalian striatum. Intriguingly, regulation of evoked [DA]o by uptake and by DA autoreceptors in Area X is surprisingly similar to that seen in rodent striatum.

Plane of Slicing

Brain slices prepared from either midbrain or forebrain are most commonly cut in the coronal plane (Figure 11.4a). In the midbrain, the lateral projecting dendrites of DA neurons in the SNc lie in this plane (Figure 11.4b, left), such that the use of coronal slices avoids having to sever DA cell dendrites, which can compromise cell viability. In axon terminal regions like dorsal striatum (CPu) or NAc, DA axons fan out after leaving the MFB, such that most axons will be perpendicular to the coronal plane (Figure 11.4b, right). When local electrical or chemical stimulation is used, axon orientation should not matter significantly, because the stimulus has a field effect that elicits DA release from all sites for which the field strength is sufficient to cause action potential generation and Ca2+ entry. Such stimulation also causes the release of other local transmitters, which can be an advantage or disadvantage depending on the goals of a given experiment (see “Regulation of Striatal DA Release in H2O2” in this chapter). It is sometimes desirable to elicit DA release by pathway stimulation. Because of the non-planar path of the MFB, only a few millimeters can be contained in a given slice, even when cut in a parasagittal plane. Nonetheless, we have found that preservation of 1–2 mm of DA axons in guinea pig striatum cut in the parasagittal plane is sufficient to permit examination of pure axonal DA release [61,106]. With the smaller mouse brain, horizontal slices can be prepared in which the MFB connections between midbrain and ventral striatum are sufficiently intact to generate spontaneous axonal DA release in ventral striatum arising from action potentials generated in midbrain DA neurons [107].

FIGURE 11.4. (See color insert following page 272.

FIGURE 11.4

(See color insert following page 272.) Rat brain sections showing major midbrain and forebrain dopamine (DA) regions. (a) Sagittal section of rat brain, stained with cresyl violet, showing main dopaminergic midbrain (somatodendritic) and forebrain (axon (more...)

Stimulating and Recording Da Release in Brain Slices

Release can be elicited by various means, including chemical (e.g., high K+), electrical, or pharmacological (e.g., amphetamine) stimulation. In this section, we will focus on stimulation paradigms that are used to elicit exocytotic release, especially high K+ and local electrical stimulation. It should be noted that local electrical stimulation can also be used in combination with other releasing agents to address specific questions. For example, the effect of amphetamine on DA release via DAT reversal and its consequence on impulse-dependent DA release has been used to examine mechanisms of vesicular storage [72,108–111].

Regardless of the stimulation method used, recording electrodes are typically inserted ~50–100 μm into the slice [49,62,95]. When local electrical stimulation is used to evoke DA release, the carbon-fiber electrode is typically positioned within 100 μm of the stimulating electrode. A technical note about recording in striatal slices is that a number of myelinated fiber bundles pass through the striatum, giving it a striated appearance (hence its name). Recording (and stimulating) electrodes should be placed in gray matter, away from these non-dopaminergic fibers [49].

The optimal stimulation paradigm for a given experiment, once again, depends on the question addressed. When the question is about factors that regulate DA release (or DA uptake), a physiological, or depolarizing, stimulus is often preferable to a pharmacological stimulus. Neuronal membranes are normally polarized to a negative intracellular potential (often −60 to −70 mV with respect to a grounded reference electrode) (discussed in [112]). Transmitter release occurs when a Na+-dependent action potential propagates to a synapse (or other release sites) to cause Ca2+ entry through voltage-sensitive Ca2+ channels, vesicle fusion, and diffusion of transmitter into the synaptic cleft and/or extracellular space (see “Physiology of DA Release” in this chapter). This process can be initiated artificially by a variety of methods that cause membrane depolarization. The simplest method is to superfuse a slice with aCSF made with a high K+ concentration (e.g., 30 mM) for a brief period of time [76] or to apply veratridine, which is a pharmacological agent that opens Na+ channels directly [81,113]. The disadvantage of these methods is that they can produce non-physiological DA release signals that last several minutes and are accompanied by large ion shifts that can be problematic for measurements with FCV [112]. Moreover, application of a depolarizing agent to the entire slice causes the uncontrolled release of many other transmitters, as well as DA. An additional concern about veratridine is that it can lead to Na+ overload and consequent DA release via reversal of the DAT, rather than by Ca2+-dependent exocytosis [113].

Better protocols to elicit DA release involve more discrete stimulation. Local electrical stimulation using insulated bipolar electrodes is used most widely [49,62,66,67,69,79,85,95,103,111, 114–117] although local application of high K+ via pressure ejection has also been reported [76]. Both single electrical pulses and trains of multiple-pulses have been used to elicit DA release. Each offers advantages for specific experiments. For example, single pulses are used to indicate whether or not a regulatory factor affects DA axons directly, whereas indirect effects involving local micro-circuitry can be revealed using pulse-train stimulation (see “Regulation of Striatal DA Release in H2O2” in this chapter). It is also sometimes informative to vary the frequency and number of applied electrical pulses to mimic tonic versus phasic firing patterns of DA neurons [66,106,115–117].

Stimulating electrodes are typically bipolar and can be made from tungsten, platinum, or platinum–iridium, and can be made locally or purchased commercially. The stimulating electrode is usually placed on the slice surface, rather than implanted, with both poles of the electrode in the same plane. In general, narrow gauge wire (e.g., 75 μm diameter bare) is recommended for construction of twisted bipolar electrodes; tip separation is usually ~100 μm. Commercially available electrodes should also be selected on the basis of size, based on the balance between small size and adequate field of depolarization. The parameters required to elicit DA release depend on the specific electrode used, as well as the brain region examined. Usually, stimulation pulse duration is 100–1000 μs, which is shorter than a typical action potential, with similar parameters used for both single- and multiple-pulse stimulation. Although large gauge electrodes used with high currents can elicit release that is not action potential dependent (i.e., that cannot be prevented by the Na+ channel blocker tetrodotoxin (TTX) [95]), this is not ideal. Rather, stimulation parameters should be optimized to produce consistent release that is TTX sensitive [62,103].

Signal versus Interference

The earliest voltammetric studies designed to monitor DA release in vivo were hindered by over-whelming interference from ascorbate and from the anionic DA metabolite, DOPAC, as discussed earlier. In vivo, extracellular ascorbate concentration is ~400 μM and DOPAC is 20–30 μM [118,119]. These levels are initially preserved in acute slices made with a tissue chopper and recorded in an interface chamber (see “Brain Slice Methodology” in this chapter) [46]. After only thirty minutes of in vitro incubation, ascorbate is lost from its intracellular stores in the absence of added ascorbate in the incubation medium, resulting in intra-and extracellular depletion [46]. Striatal DOPAC is lost even more quickly, since it is predominantly extracellular and readily washed out of slices [46,81]. Thus, neither of these electroactive species interfere with DA detection in superfused brain slices [81].

Slice recording is not devoid of interference, however, as several electroactive and non-electroactive substances can interfere with DA detection. The most commonly reported electroactive interferents are 5-HT and NE. The degree of interference depends on the relative innervation of each transmitter versus DA in a given region. For example, in dorsal striatum, DA is the only electroactive substance detected because of the high density of DA axons, as well as the low level of 5-HT axons arising from 5-HT neurons in midbrain. By contrast, in the SNc and the SN pars reticulata, DA content is ~10-fold lower than in striatum [81] with specific 5-HT innervation of DA dendrites, such that 5-HT can be the predominant substance identified, especially in rat and mouse midbrain [57,99–102], as noted in “Species Selection.” The extent of 5-HT interference is often amplified by the typically 2-4-fold higher sensitivity of carbon-fiber microelectrodes for 5-HT than for DA. Consequently, even if DA were 4-fold higher than 5-HT, both substances would contribute equally to the current monitored. When FCV is used, 5-HT interference is immediately indicated by the characteristic 5-HT voltammogram, which can be readily distinguished from that of DA (see “Species Selection” in this chapter) [57,80,81,100]. With amperometry or chronoamperometry, however, pharmacological methods (e.g., uptake blockade) must be used to confirm signal identity. In contrast to the cyclic voltammogram for 5-HT, the voltammogram for NE is largely indistinguishable from that for DA, especially in tissue, since the redox characteristics of DA and NE are defined primarily by their identical catechol ring structures; electrode sensitivity for NE is typically similar to or less than that for DA. Consequently, in regions with possible NE interference, additional pharmacological and/or anatomical evidence must be obtained to confirm that DA is the substance monitored, regardless of the electrochemical method used. For example, both anatomical and pharmacological data were used to confirm that DA was the primary substance detected in the VTA (A10 DA cell body region) and in the subthalamic nucleus, both of which have particularly low DA innervation, as well as NE fibers passing through [120,121].

The most important non-electroactive interferents are Ca2+ and pH (H+). These factors are most problematic for FCV, which has a large non-Faradaic background that is sensitive to the ionic microenvironment (as discussed in “Fast-Scan Cyclic Voltammetry” in this chapter). Possible interference can arise from ion-dependent background shifts [82,114] and from the fact that changes in ionic composition, especially in the divalent cations Ca2+ and Mg2+, can alter electrode sensitivity to DA [122]. Understanding these interferents is important because the ionic microenvironment of brain cells is dynamic, with decreases in extracellular Ca2+ concentration and activity-dependent alkaline and acid shifts in the extracellular space accompanying all depolarizing stimuli (see [112] for review).

The sensitivity of electrode background in FCV to changes in ionic composition was first shown for pH, with a decrease in pH (increase in acidity) reflected in an anodic surface wave with a peak around 0.3 V versus Ag/AgCl [82,112,114]. The potential of this background wave can readily be distinguished from the DA oxidation potential (~+0.6 V versus Ag/AgCl in FCV at most electrodes); however, this can interfere with DA quantitation in brain regions with low DA innervation like the amygdala [111]. Stimulated Ca2+ entry (required for exocytotic release of DA) and the associated transient decrease in extracellular Ca2+ concentration is reflected in a negative deflection in current output at roughly the same potential as the pH shift [114]. Heien, Wightman and colleagues recently reported the use of principal component regression coupled with appropriate calibration data to resolve contributions from substances that give overlapping voltammetric wave-forms, including DA and pH [85].

A less obvious source of ionic interference arises from interactions of divalent cations with an electrode surface, which can alter electrode sensitivity to DA [122]. The calibration factor for DA using FCV can be two-fold lower in media containing Ca2+ and Mg2+ than in divalent-ion free solutions, presumably because divalent ions associate strongly with anionic functionalities on electrode surfaces, thereby decreasing DA adsorption and, thus, DA oxidation current. This is relevant for two practical reasons. First, electrode calibration in many previously reported studies was done using HEPES- or phosphate-buffered saline that lack divalent ions; this means that when these data are used to calculate evoked [DA]o for tissue bathed in normal aCSF (see “aCSF Composition for Healthy Slices” in this chapter), [DA]o can be underestimated by at least 50%. Second, studies of the Ca2+- or Mg2+-dependence of DA release will require that electrode sensitivity to DA be determined in all media tested to ensure quantitative accuracy (e.g., [62]).

One final set of potentially interfering substances is the agents used to manipulate DA release. We have found that a number of such substances are electroactive, including the DA-receptor blocker haloperidol, the DAT inhibitor nomifensine, and the catalase inhibitor 3-aminotriazole. An advantage of in vitro methods is that testing whether a drug agent is electroactive, and calibrating electrodes in the presence and absence of the agent, can be done readily in the slice chamber at the same temperature used for DA release studies.

Regulation Of Striatal DA Release By H2O2

Research in our laboratory over the past decade has focused on two converging areas of interest. The first is the neuroprotective role of low molecular weight antioxidants, including ascorbate; the second is DA release regulation by endogenous neurotransmitters and neuromodulators. These topics converged when we began to examine DA release regulation under conditions of oxidative stress. In our initial studies, we examined the effect of exogenously applied H2O2 on electrically evoked [DA]o in guinea pig striatal slices, monitored using carbon-fiber microelectrodes and FCV. Transient exposure to exogenous H2O2 (15 min, 1.5 mM) causes a reversible 30%–40% decrease in pulse-train evoked [DA]o in dorsolateral striatum and is not accompanied by a change in tissue DA content or evidence of oxidative damage [123]. In the course of these studies, we also discovered that endogenous H2O2 plays an important role in the physiological regulation of axonal DA release in striatum. This finding shifted our focus from viewing H2O2 as a potential mediator of oxidative damage, to viewing it as a neuromodulator. In this section we summarize studies that have been done to determine the source of modulatory H2O2, the neuronal circuitry involved, and the mechanism of DA release suppression. This body of work not only provides a good example of the power of voltammetric detection of DA release, but also demonstrates an advantage of in vitro brain slice preparation studies, which is that they facilitate the use of companion techniques, including whole-cell physiological recording and fluorescence imaging.

Endogenous H2O2 Inhibits Axonal DA Release

We first examined a possible regulatory role for endogenously generated H2O2 by manipulating slice peroxidase activity using exogenously applied catalase and/or by inhibiting endogenous glutathione (GSH) peroxidase (as noted in “Signal Versus Interference,” the catalase inhibitor, 3-aminotriazol is electroactive and could not be tested in these experiments). For example, amplification of endogenous H2O2 levels by GSH-peroxidase inhibition with mercaptosuccinate (MCS) causes a similar decrease in pulse-train evoked [DA]o (30 pulses, 10 Hz) in dorsolateral striatum to that seen with exogenously applied H2O2 [9,124]. Again, no change in DA content is seen and DA release suppression is fully reversible upon MCS washout or when the slice is superfused with catalase in the continued presence of MCS [9]. Regulation of DA release by endogenous H2O2 also occurs in the shell of the NAc and in the SNc [124]. Importantly, MCS has no effect on [DA]o evoked by a single stimulus pulse [9], implying that even under conditions of impaired GSH peroxidase activity, basal levels of H2O2 in striatum are insufficient to inhibit DA release. Instead, modulatory H2O2 must be generated dynamically during the first pulse of a stimulus-train and inhibits DA released by subsequent pulses. Additional studies show that H2O2 generation is derived from concomitant glutamate release and AMPA receptor activation (see “Regulation of Axonal DA Release by Glutamate Acting at AMPA Receptors Requires H2O2” in this chapter). The time-scale of H2O2-dependent DA release suppression is rapid, occurring within a few hundred milliseconds after initiation of a 10 Hz pulse train [9].

Potential Sources of H2O2 Generation

The major source of ROS in neurons, as in all cells, is mitochondrial respiration [125], although other superoxide (·O2) generating processes, notably NADPH oxidase [126], also contribute. During mitochondrial respiration, (·O2) is formed from the one electron reduction of O2; H2O2 is generated from (·O2) by superoxide dismutase (SOD), as well as by spontaneous dismutation [4,127] (Figure 11.5). The amount of H2O2 produced by mitochondria is 1%–2% of O2 metabolized [127], such that the concentration of H2O2 within the restricted volume of a dendrite, for example, could transiently reach mM levels [123]. Absolute levels of H2O2 at a given point in time, however, will depend on the balance between H2O2 generation and cellular peroxidase activity [4,11] (Figure 11.5).

FIGURE 11.5. Production of H2O2 by the mitochondrial respiratory chain.

FIGURE 11.5

Production of H2O2 by the mitochondrial respiratory chain. The primary source of H2O2 generation in all cells is mitochondrial respiration during which ·O2 is formed from the single-electron reduction of O2. H2O2 is then generated from (more...)

Is mitochondrial respiration the primary source of modulatory H2O2 in the striatum? Our recent studies indicate that the answer is yes: the effect of MCS on DA release in striatum is lost when mitochondrial complex I is inhibited by rotenone in the presence of the complex II substrate succinate to maintain tissue ATP content [128]. These data also argue against significant involvement of other ·O2 = H2O2 generating pathways, including NADPH oxidase. Additional experiments confirm a lack of involvement of monoamine oxidases (MAOs), which produce one molecule of H2O2 for each molecule of DA metabolized [4]: a cocktail of MAO inhibitors does not alter evoked [DA]o versus control or prevent the effect of MCS [128].

Regulation of Axonal DA Release by Glutamate Acting at AMPA Receptors Requires H2O2

Given the close apposition of mitochondria to presynaptic sites in DA axons [129], we intially assumed that modultory H2O2 generation in the striatum might occur directly in DA axons, such that activity-dependent H2O2 would provide a feedback signal to decrease DA release and thereby “augment the effect of D2-autoreceptor-mediated inhibition” [123]. However, our subsequent studies disprove DA axons as primary sites of generation. The most direct evidence comes from comparisons of the effect of GSH peroxidase inhibition on DA release evoked by local versus distal stimulation in parasagittal slices of striatum, in which DA-axon tracts are preserved for 1–2 mm. In this preparation, as in our usual coronal slices, local stimulation evokes the release of glutamate, GABA, and other transmitters, as well as that of DA, whereas distal pathway stimulation selectively activates DA axons to elicit relatively pure DA release. Under these conditions, inhibiting GSH peroxidase with MCS suppresses [DA]o evoked by local stimulation, as seen previously, but not by distal stimulation [61]. Also arguing against H2O2 generation in DA axons is the finding that H2O2- dependent inhibition of DA release requires glutamatergic AMPA receptors [9], which are apparently absent from DA axons [130,131].

Thus, H2O2-mediated inhibition of DA release in striatum is not an auto-regulatory process, but rather represents potent external regulation by glutamatergic input to the striatum. Indeed, when striatal AMPA receptors are blocked by a selective antagonist, GYKI-52466, pulse-train evoked [DA]o increases by 100% [9]; this effect is completely prevented by catalase, confirming that H2O2 is required (Figure 11.6a and Figure 11.6b). Thus, glutamate released under physiological conditions inhibits action potential-dependent DA release in striatum via H2O2. Moreover, H2O2 amplification by MCS has no effect on DA release in the presence of GYKI-52466, indicating that glutamate acting at AMPA receptors is required for generation of modulatory H2O2 in striatum [9]. Consistent with the non-DA axon origin of AMPA-receptor dependent H2O2 generation, blockade of AMPA receptors with GYKI-52466 enhanced [DA]o evoked locally, but had no effect on that evoked distally [61]. The question of how, or even if, glutamate regulates striatal DA release has been a long-standing source of controversy. Our discovery that glutamate inhibits DA release via H2O2 generation resolves this conundrum.

FIGURE 11.6. Regulation of striatal DA release by glutamate and GABA requires H2O2.

FIGURE 11.6

Regulation of striatal DA release by glutamate and GABA requires H2O2. (a) AMPA-receptor blockade by GYKI-52466 (GYKI; 50 μM) causes a ~100% increase in evoked [DA]o in striatum (p<0.001, n = 6). (b) The effect of AMPA-receptor blockade (more...)

Our studies show that regulation of DA release in the dorsolateral striatum by GABAergic input is also unconventional: blockade of GABAA receptors by picrotoxin causes a ~50% decrease in evoked [DA]o (Figure 11.6c), whereas GABAB-receptor blockade with saclofen has no effect [9], indicating that GABA, acting at GABAA receptors, normally enhances DA release. The influence of GABA on DA release, like that of glutamate, must be indirect, since DA axons in striatum do not

Activity-Dependent H2O2 Generation in Striatal Medium Spiny Neurons

Increasing evidence from our laboratory implicates striatal medium spiny neurons as the primary cellular source of modulatory H2O2 in the striatum. Not only are these the most abundant striatal neurons (90%–95%; [133]), but the pattern of sensitivity of DA release to glutamate and GABA antagonists [9] mirrors the electrophysiological responsiveness of medium spiny neurons to these agents [134,135]. Moreover, glutamate synapses are closely apposed to DA synapses on medium spiny neuron dendrites [130,131,136], such that they are ideally positioned to modulate DA release via postsynaptically generated H2O2. Medium spiny neurons also express GABAA receptors at dendritic sites near spines [132], which would facilitate GABAergic opposition of glutamatergic excitation and H2O2 generation.

Consistent with this indirect evidence, we have recently confirmed that medium spiny neurons are a key cellular source of modulatory H2O2 in striatum. In these studies, we used a method for H2O2 detection that we developed for simultaneous whole-cell recording and fluorescence imaging using the H2O2-sensitive dye H2DCF (dihydro-dichlorofluorescein) in brain slices; this dye becomes fluorescent DCF when oxidized by ROS [11]. Under control conditions, basal DCF fluorescence is seen in all medium spiny neurons, reflecting a basal ROS tone (Figure 11.7). Local stimulation (with the same 30 pulse, 10 Hz pulse trains used to elicit DA release) causes a ~30% increase in DCF fluorescence in medium spiny neurons, which is further enhanced by inhibition of GSH peroxidase with MCS [61]. Importantly, AMPA-receptor blockade by GYKI- 52466 prevented stimulus-induced action potentials, as well as activity-dependent H2O2 generation in MSNs [61].

FIGURE 11.7. Activity-dependent H2O2 generation in a striatal medium spiny neuron.

FIGURE 11.7

Activity-dependent H2O2 generation in a striatal medium spiny neuron. H2O2-dependent DCF fluorescence intensity (FI) under control conditions and after local stimulation (10 Hz, 30 pulses) with simultaneously recorded membrane voltage (Vmemb); FI plateau (more...)

H2O2 Acts via KATP Channels to Inhibit DA Release

The mechanism by which H2O2 regulates the nigrostriatal pathway is H2O2-dependent opening of KATP channels [9,10]. These channels are multimeric proteins composed of four inwardly rectifying pore-forming units, typically Kir 6.2 in neurons [137], and four sulfonylurea-binding subunits (SUR1 or SUR2) [138]. Previous physiological studies demonstrated that exogenous H2O2 can cause membrane hyperpolarization by activating a K+ conductance in a variety of cell types, including pancreatic β-cells [139] and CA1 hippocampal neurons [140]. Our studies of striatal DA release provided the first evidence that endogenous H2O2 can activate KATP channels. Blocking KATP channels with tolbutamide or glibenclamide leads to a significant increase in evoked [DA]o in dorsolateral striatum, indicating that during normal stimulation, KATP-channel activation causes the inhibition of DA release; these sulfonylureas also prevent the usual pattern of H2O2-dependent modulation by MCS, GYKI-52466, and picrotoxin [9,10], demonstrating that KATP channels are required for modulation of DA release by H2O2, glutamate, and GABA. Conversely, striatal DA release is decreased by ~40% by either diazoxide, a SUR1-selective KATP-channel opener, or cromakalim, a SUR2-selective opener. However, diazoxide prevents the effects of MCS, GYKI-52466, and picrotoxin, whereas cromakalim does not, demonstrating that glutamate-dependent H2O2 preferentially opens SUR1-based KATP channels (Avshalumov and Rice 2003) [10].

As mentioned in the introduction to this chapter, KATP channels are expressed abundantly throughout the nigrostriatal pathway [12–14]. We recently reported that DA neurons in the SNc are also responsive to endogenously generated H2O2: 50% of these neurons respond to moderate H2O2 elevation during GSH peroxidase inhibition with membrane hyperpolarization and loss of spontaneous firing, whereas all cells respond in this way to greater H2O2 elevation when catalase is inhibited [11]. These effects are reversible and are blocked by glibenclamide, confirming a role for KATP channels. However, the more sensitive “responders” are those that express SUR1-containing KATP channels, which is the channel subtype involved in mediating AMPA-receptor/H2O2-dependent DA release suppression in the striatum [10]. Whether regulation of KATP channels by H2O2 is direct or indirect in DA neurons remains to be elucidated, although previous studies using inside-out membrane patches from cardiac cells have shown a direct, effect of H2O2 on KATP channel opening by decreasing channel sensitivity to ATP [141].

Summary and Future Directions for studies of H2O2 as a Neuromodulator

In this chapter, we have presented rationale, methods, and results for the study of DA release in brain slices. This background should give readers a general understanding of the strengths and weaknesses of brain slices as experimental preparations, as well as an overview of the voltammetric methods and carbon-fiber microelectrodes currently used to monitor DA release. There is much left to discover about the regulation and function of DA in the CNS. Future studies are likely to exploit the ease of combining other methods with voltammetry in brain slices. Fluorescence imaging and whole-cell recording of single neuron activity are especially promising companion techniques, as illustrated in “Regulation of Striatal DA Release by H2O2” in this chapter.

Our exploration of H2O2 as an endogenous neuromodulator indicates that activity-dependent H2O2 generation is predominantly mitochondrial. In dorsolateral striatum, H2O2 is generated down-stream from AMPA-receptor activation in medium spiny neurons, rather than DA axons, and must therefore diffuse to DA synapses where it inhibits DA release via opening of SUR1-based KATP channels. Our working model of H2O2-dependent regulation of striatal DA release involves a triad of DA, glutamate, and GABA synapses, which are separated by a few micrometers on the dendrites of medium spiny neurons, but bound together functionally by diffusible H2O2 [9,142].

A noted in the beginning of this chapter, H2O2 and other ROS can be neurotoxic. Thus, neuromodulation by H2O2 can be a double-edged sword: oxidative stress resulting from an imbalance between H2O2 generation and regulation could lead to cellular damage. Indeed, oxidative stress has been implicated in nigrostriatal degeneration in Parkinson’s disease, which selectively targets the nigrostriatal pathway [1,2,4]. Thus, loss of normal H2O2 regulation could contribute to nigrostriatal pathophysiology and to the motor deficits of Parkinson’s disease. Further exploration of H2O2 as a diffusible messenger in the migrostriatal pathway should provide new insights into both normal and pathophysiological roles of this novel neuromodulators.

Acknowledgments

The authors are grateful for support from NIH grants NS-36362 and NS-45325 and the National Parkinson Foundation.

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Copyright © 2007, Taylor & Francis Group, LLC.
Bookshelf ID: NBK2562PMID: 21204377

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