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Ernest Gallo Clinic and Research Center, Department of Neurology, University of California, San Francisco, Emeryville, California
Submitted 24 July 2007; accepted in final form 13 August 2007
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ABSTRACT |
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INTRODUCTION |
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; Diana et al. 1992; Mereu et al. 1984) and oral self-administration of ethanol increases DA release in a VTA target region, the nucleus accumbens (Gonzales and Weiss 1998). In contrast, withdrawal after chronic ethanol exposure decreases VTA cell firing and DA release in the nucleus accumbens for several days after removal from ethanol (Bailey et al. 1998, 2001; Diana et al. 1993; Shen 2003; Weiss et al. 1996), and reexposure to ethanol during withdrawal restores nucleus accumbens DA release (Weiss et al. 1996). Behavioral studies have also shown that responses to ethanol are modified by pharmacological manipulations of DA transmission. For example, quinpirole microinjection into the VTA, which inhibits DA cell activity through activation of DA D2/D3 receptors, decreases the number of lever presses an animal will exert to obtain ethanol (Hodge et al. 1993). Several lines of evidence also suggest that the mesolimbic system is involved in ethanol seeking and relapse (Gonzales et al. 2004; Katner and Weiss 1999; McBride et al. 2002). Taken together, these results suggest that VTA DA cell activity contributes to the motivation to self-administer ethanol.
From a physiological perspective, midbrain DA neurons present two patterns of activity—regular, pacemaker firing and burst firing (reviewed in Cooper 2002; Komendantov et al. 2004)—that are considered important for control of different physiological responses. Tonic, pacemaker-like firing may be altered under conditions ranging from withdrawal from addictive drugs to schizophrenia (Grace 2000; Weiss et al. 1992). In addition, burstlike firing in VTA DA neurons, which may produce phasic increases in DA in the different target areas of the VTA (reviewed in Overton and Clark 1997), is thought to encode salience of important stimuli (Robinson and Berridge 2001) and may represent a teaching signal (Schultz 2002).
A number of channels, including small conductance, calciumactivated potassium channels (SK) and inwardly rectifying hyperpolarization-activated cation channels (HCN, responsible for generating the Ih current), can regulate pacemaker and burst firing (Johnson and Seutin 1997; Neuhoff et al. 2002; Seutin et al. 1993, 2001). However, little is known about the long-term effects of chronic ethanol treatment on the firing properties of VTA DA neurons. Furthermore, intermittent ethanol exposure can enhance the locomotor-activating effects of cocaine (Itzhak and Martin 1999), but the cellular mechanisms underlying hyperlocomotion following this pattern of ethanol exposure are poorly understood. In this study, we evaluated the function of different channels involved in control of firing in VTA DA neurons 7 days after the end of a 5-day repeated, intermittent ethanol treatment, and also the ability of repeated ethanol exposure and withdrawal to enhance locomotor activation by cocaine.
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METHODS |
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Male Sprague–Dawley rats were singly housed in a temperature-controlled (21 ± 1°C) room on a 12-h light/dark cycle (lights on at 07:00). Food and water were available without restriction. Rats were acclimated to handling procedures starting 3 days before experiments began. All ethanol injections and behavioral testing were performed during the light cycle. Rats were injected with ethanol [2 g/kg, administered intraperitoneally (ip), twice a day at 10:00 and 18:00, from a 20% vol/vol solution] or an equivalent volume of saline for 5 days, with injections starting at postnatal day (P) 21.
Slice preparation
All electrophysiological recordings were performed in rats 7 days after the end of repeated treatment with ethanol or saline (described earlier). To prepare VTA slices, animals were anesthetized with halothane and perfused transcardially with about 10 ml of nearly frozen (
0°C) perfusion solution at a rate of about 10 ml/min. This perfusion solution was saturated with 95% O2-5% CO2 and contained (in mM): 119 NaCl, 2.5 KCl, 1.0 NaH2PO4, 6.1 MgCl2, 0.1 CaCl2, 26.2 NaHCO3, 1.25 glucose, 50 sucrose, 1 ascorbic acid, and 3 kynurenic acid. Horizontal slices of 230 µm containing the VTA were prepared with a VT1000S vibratome (Leica, Nussloch, Germany) in the same perfusion solution at 4°C. Slices were then placed in a holding chamber at 31–32°C containing artificial cerebrospinal fluid (aCSF) saturated with 95% O2-5% CO2 and composed of (in mM): 119 NaCl, 2.5 KCl, 1.0 NaH2PO4, 1.3 MgCl2, 2.5 CaCl2, 26.2 NaHCO3, and 11 glucose. Ascorbic acid (1 mM) was added about 15 min before slices were placed in the recovery chamber. Slices were allowed to recover for 
45 min before being placed in the recording chamber and superfused with oxygenated aCSF at 31–32°C with picrotoxin (100 µM) added to block 
-aminobutyric acid type A (GABAA)–receptor-mediated inhibitory postsynaptic currents.
Electrophysiology
Cells were visualized using infrared differential interference contrast video microscopy. Whole cell current- and voltage-clamp recordings were made using a Multiclamp 700A or 700B amplifier and Clampex 9.2 (Axon Instruments, Foster City, CA). Electrodes (2.8–4.0 M
) generally contained: 130 mM KOH, 105 mM methanesulfonic acid, 17 mM hydrochloric acid, 20 mM HEPES, 0.2 mM EGTA, 2.8 mM NaCl, 2.5 mg/ml MgATP, and 0.25 mg/ml GTP (pH 7.2–7.4, 280–290 mOsm). A low level of the calcium buffer EGTA was included in the pipette solution to preserve calcium-dependent potassium currents during whole cell current- and voltage-clamp recordings (Wolfart et al. 2001). For experiments examining brief (30 s) bath application of N-methyl-D-aspartate (NMDA) under voltage clamp (Vholding = +40 mV), the internal solution was (in mM): 117 Cs-methanesulfonic acid, 20 HEPES, 0.4 EGTA, 2.8 NaCl, 5 TEA-Cl, 2.5 Mg2ATP, and 0.25 Mg2GTP (pH 7.2–7.4, 280–290 mOsm). The reversal potential of NMDA currents was not determined.
Current-clamp data were acquired at 20 kHz and filtered at 2 kHz and voltage-clamp data were acquired at 10 kHz and filtered at 2 kHz. VTA DA neurons were identified by the presence of a large Ih current (Johnson and North 1992). Because Ih is present in both principal and tertiary VTA neurons (Margolis et al. 2006), we recognize that its presence does not unequivocally identify DA neurons in midbrain slices. However, in previous work (Ungless et al. 2001) and in the present study, this criterion was enough to obtain differences between control and experimental treatments. In addition, prior results from our laboratory (Borgland et al. 2006 and F. Sarti, unpublished observations) suggest a higher (75%) correlation between the presence of Ih and tyrosine hydroxylase, indicative of DA neurons, in neurons just medial to the medial terminal nucleus of the accessory optic tract (MT), where nearly all the recordings in the present study were performed. Also, the decrease in Ih amplitude in ethanol animals was only about 40% (Fig. 7) and thus was not sufficient to prevent the use of Ih to identify putative DA neurons relative to GABA neurons, which do not have an Ih current (Johnson and North 1992; Margolis et al. 2006).
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10 sequential APs with a coefficient of variation of the interspike interval (ISI) of <20%. The average basal instantaneous firing frequency and ISI during periods of pacemaker firing were then determined across 30 successive APs just after breaking into a neuron, before any dialysis of the neuron with the intracellular pipette solution could occur. However, prolonged dialysis with the internal solution did not alter the firing pattern or AP waveform of neurons (data not shown). In addition, due to variability among neurons in the basal pacemaker firing rate, neurons were brought to about 1 Hz by injecting DC current through the patch amplifier before experiments examining the effects of NMDA, 4-(N-ethyl-N-phenylamino)-1,2-dimethyl-6-(methylamino) pyridinium chloride (ZD7288), or recovery from hyperpolarization.
All voltage values were corrected for the liquid junction potential, estimated to be 10 mV using the Junction Null Calculator in Clampex 9.2, and also by direct measurement of the potential difference between internal and external solutions present after zeroing the pipette current. Bridge balance was used to compensate 60–80% of the series resistance. Also, because the average series resistance and instantaneous currents evoked by hyperpolarization (taken as a measure of input resistance) were the same between ethanol and saline animals (see RESULTS), any errors in compensation would be similar in both groups and thus would not explain the differences in channel function observed here. Further, to estimate whether repeated ethanol exposure and withdrawal might have altered the size of VTA neurons, we estimated the whole cell capacitance using a method adapted from Gentet et al. (2000). In Ih experiments, the voltage step from –60 to –70 mV was used for analysis of capacitance, where the series resistance was determined from the peak of the capacitive transient; the input resistance was determined 40 ms into plateau of the voltage response; the capacitive current transient just after application of the voltage step was best fit with two exponentials; and the fast exponential was used in the determination of cell capacitance. For SK experiments, the voltage step from –70 to –60 mV was used for capacitance analysis. Both methods gave equivalent measures of capacitance (data not shown).
To investigate SK tail currents, neurons were held at –70 mV, and depolarizing current steps (400 ms, from –60 to –20 mV in 10-mV steps) were applied, with 1 s between successive steps. On returning to –70 mV, a tail current was evident (see Fig. 3A). The peak magnitude of the SK tail current was determined for the steps to –20, –30, and –40 mV. In addition, the charge transfer for SK (in picocoulombs) was calculated by integrating (nanoamperes x milliseconds) the tail current evoked after the depolarizing pulse, beginning from the initiation of the tail current to 550 ms into the tail current because the apamin-sensitive component of the tail current should be <5% by 550 ms into the tail current (Abel et al. 2004). The area under the curve was determined using GraphPad Prism (San Diego, CA). Finally, the rate of inactivation of the SK current was estimated in the tail current evoked after a –20-mV depolarization by fitting a single exponential from about 55 ms after the peak of the tail current to 550 ms after the peak of the tail current (Abel et al. 2004).
To examine Ih, neurons were held at –60 mV and a series of voltage steps (500 ms, from –40 to –150 mV in 10-mV steps, with 4 s between steps) was applied. In some Ih voltage-clamp experiments, to reduce contamination from other currents and to better isolate Ih, we performed experiments under conditions where many currents other than Ih were inhibited using a modified internal solution (3 mM BAPTA added) and external solution (aCSF containing 2 mM tetraethylammonium, 20 mM MgCl2, 0.5 µM tetrodotoxin, and 2 mM 4-aminopyridine, with NaCl reduced in an equimolar fashion) (Liu et al. 2003).
Ih was determined from the difference between the peak current response at the end of the pulse and the instantaneous current response about 30 ms after the onset of the hyperpolarizing pulse (see Fig. 7A; Watts et al. 1996). In addition, we determined the Ih charge transfer using methods similar to those described earlier for SK, except that Ih charge transfer was determined by integrating the slowly activating inward current beginning from 30 ms after the initiation of the –140-mV hyperpolarizing voltage pulse (Neuhoff et al. 2002). We also determined the magnitude and charge transfer of the instantaneous current, which could represent inwardly rectifying potassium channels (IRK) and/or potassium leak channels (Watts et al. 1996; Wilson 2005). We did not perform experiments to determine the relative contribution of these currents to the instantaneous current. The voltage dependence of activation of Ih was determined as described (Liu et al. 2003) using the equation G = I/ (E – Erev), where –55 mV was used for the Erev for Ih because it was not different between saline and ethanol animals (Fig. 8F). Data were fit with a Boltzmann equation (Liu et al. 2003) using GraphPad Prism to determine the half-activation voltage (V1/2) and slope factor. The Ih reversal potential was determined using described methods (Cathala and Paupardin-Tritsch 1997; Mayer and Westbrook 1983), where the reversal potential of Ih was estimated by comparing the instantaneous current evoked from a voltage of –60 mV, where Ih was not active, and the instantaneous current evoked from a voltage of –80 mV, where Ih was activated (see Fig. 8E). Because instantaneous currents evoked from –60 mV likely represent IRK/leak channels (Watts et al. 1996; Wilson 2005), whereas currents evoked from –80 mV represent Ih in addition to IRK/leak, one can estimate the reversal potential of Ih by determining the point where the two current–voltage (I–V) curves intersect, which essentially represents the voltage where the Ih current is zero after the IRK/leak current is subtracted out.
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; Seutin et al. 1993). However, this applied current (values given for 20 µM NMDA) was not significantly different between ethanol (–31.1 ± 18.9 pA) and saline (–41.6 ± 14.4 pA) animals. In addition, the amount of holding current applied during NMDA exposure did not correlate with the development of bursting, measured as the coefficient of variance of the ISI (see following text), in either saline or ethanol animals (r2 = 0.197 and 0.070 for ethanol and saline, respectively; n.s.). Motor activity assay
Locomotor activity was measured in a 17 x 17-in. open-field chamber lined with three 16-beam infrared arrays. Distance traveled (in centimeters) was measured using Open Field Activity software (MED Associates, St. Albans, VT). Animals were habituated to the locomotor chamber during the 4th through 6th days after the last ethanol injection. Each day, the animals were allowed 1 h of free exploration, then were given an ip injection of saline (1 ml/kg) and then returned to the chamber for 30 min. This was followed by a second ip injection of saline and a final 30 min in the locomotor chamber. On the 7th day after ethanol injection, animals were allowed 1 h of free exploration, followed by an injection of saline and then 30 min in the locomotor chamber. The animals then received an ip injection of cocaine (15 mg/kg) and were returned to the chamber for a final 30 min.
One concern with repeated injections is the possibility of stress-induced changes. However, as subsequently described, saline and naïve neurons exhibited a similar afterhyperpolarization (AHP) and voltage-clamp measures of SK and Ih, and saline animals exhibited a locomotor response to cocaine very similar to that in naïve animals. These results suggest that any stress related to multiple injections or handling procedures did not alter channel function or enhance the locomotor response to cocaine. As an additional indicator of possible stress due to injections, we examined the weight of the animals across the period of injection. Saline and ethanol animals showed a similar weight before injections (saline: 51.6 ± 1.4 g; ethanol: 52.2 ± 1.2 g, n = 25 both for saline and ethanol), but by the 5th day the weight of ethanol animals was slightly but significantly reduced (10%) relative to saline animals (saline: 74.9 ± 1.6 g; ethanol: 68.4 ± 1.7 g, P < 0.05). The cause of this difference is unclear, but may relate to differences in activity of the animals, with two periods of intoxication and lethargy each day in ethanol but not saline animals. However, the weight of saline and ethanol animals was not significantly different after the 7-day withdrawal period (saline: 129.6 ± 3.4 g; ethanol: 121.0 ± 5.2 g), suggesting that repeated ethanol injection did not result in persisting weight loss due to stress associated with injection.
Reagents
All reagents were purchased from Sigma (St. Louis, MO), except for ZD7288, which was purchased from Tocris Cookson (Ellisville, MO). Reagents were prepared in a 1:1,000 stock that was frozen in aliquots at –20°C, except for cocaine, which was made fresh on the day of experimentation. Apamin was prepared in distilled water and ZD7288 in DMSO.
Statistical analysis
All data are expressed as means ± SE. Statistical analysis was performed with a two-tailed, unpaired t-test, except, where indicated, a two-way, repeated-measures ANOVA was used. Significance was determined using GraphPad Prism with confidence intervals of 95%. All exponentials were fit using Clampfit. During experiments aimed at examining changes in firing during NMDA application, the coefficient of variance of the interspike interval (CV-ISI) was determined from 100 successive action potentials. CV-ISI was calculated as the SD divided by the mean and expressed as a percentage (Zhang et al. 1994).
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RESULTS |
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Pacemaker activity was not altered after repeated ethanol treatment followed by 7 days withdrawal
We first investigated the basal pacemaker activity in VTA DA neurons. Here, neurons were considered able to fire in pacemaker mode if they were able to fire 10 sequential APs (see METHODS). We recorded neurons meeting this criterion from 18 saline and 19 ethanol animals. The average basal instantaneous firing frequency and ISI during periods of pacemaker firing were determined across 30 successive action potentials just after breaking into a neuron. When pacemaker firing occurred, there were no significant differences in the average instantaneous firing frequency (Fig. 1B; saline, 1.19 ± 0.07 Hz; ethanol, 1.27 ± 0.12 Hz; not significant, P > 0.05 (n.s.)) or the average ISI (Fig. 1C; saline, 958 ± 69 ms; ethanol, 967 ± 109 ms; n.s.) of neurons from saline and ethanol rats, with firing rates similar to those in several other studies using whole cell recording (Margolis et al. 2006; Marinelli et al. 2005). In addition, although a resting membrane potential was difficult to determine because neurons were undergoing pacemaker firing, the average membrane potential during pacemaker firing was not different between saline and ethanol animals (saline, –59.1 ± 1.2 mV; ethanol, –59.3 ± 0.6 mV, n.s.), in agreement with no change in many aspects of the action potential waveform (see following text).
SK function was reduced in ethanol animals
To further characterize the firing properties of VTA DA neurons, we examined the action potential waveform of neurons from saline and ethanol animals (n = 21 and 20, respectively). A fundamental feature of neuronal firing is the generation of action potentials, and analysis of specific components of the action potential waveform can often provide evidence that the function of a particular channel has been altered. For example, decreased Na+ channel function could be reflected by an increase in action potential threshold and a decrease in the action potential peak (Zhang et al. 1998). However, neither the threshold nor the peak of the action potential were different between saline and ethanol animals (Fig. 2, A and B; threshold: saline, –43.5 ± 0.5 mV; ethanol, –44.1 ± 0.7 mV; peak: saline, 17.6 ± 1.4 mV; ethanol, 16.3 ± 1.5 mV; n.s.), suggesting no changes in Na+ channel function. Further, a reduction in the peak value of the AHP is often observed during inhibition of fast-repolarizing potassium channels (such as BK and Kv3 delayed rectifier channels; Appel et al. 2003; Grillner and Mercuri 2002). However, the peak value of the AHP was not altered in ethanol animals (Fig. 2, A and B; saline, –69.0 ± 1.7 mV; ethanol, –69.8 ± 0.8 mV; n.s.), suggesting no change in function of the fast-repolarizing channels. In addition, the action potential width, determined halfway between the action potential threshold and peak, was not different between saline and ethanol animals (saline, 1.27 ± 0.05 ms; ethanol, 1.29 ± 0.05 ms; n.s.).
These data suggest that some of the basic electrophysiological properties of VTA DA neurons were not modified by withdrawal after repeated ethanol treatment. However, the time-to-peak of the AHP (TTP-AHP), determined relative to the action potential threshold, was significantly reduced in ethanol animals (Fig. 2C shows results from neurons treated with apamin: saline, 70.9 ± 8.7 ms; ethanol, 46.6 ± 7.0 ms; n = 5 for saline and 10 for ethanol, P < 0.05; significant differences were also apparent in an additional set of neurons from 17 saline and 14 ethanol animals in which apamin was not tested: saline, 56.0 ± 4.8 ms; ethanol, 37.5 ± 4.6 ms; P < 0.05). Although a number of potassium channels contribute to the AHP, the small-conductance, calcium-dependent potassium current (SK) is of particular interest because it is hypothesized to regulate the transition from pacemaker to burst firing (Komendantov et al. 2004). Neurons from saline animals exhibited a very broad AHP with a greatly delayed TTP-AHP, typical of naïve neurons with a strong SK (age-matched naïve, 61.2 ± 11.1 ms, n = 7; and see Ping and Shepard 1996; Shepard and Bunney 1988). Further, the SK-selective antagonist apamin (200 nM) significantly reduced the TTP-AHP in both saline and ethanol animals (Fig. 2, D and F; P < 0.05; n = 5 for saline and n = 10 for ethanol), with a significantly greater change in the TTP-AHP in saline animals (Fig. 2, E and F; saline, 60.6 ± 9.9 ms; ethanol, 35.8 ± 6.3 ms; P < 0.05), and resulted in a similar TTP-AHP in both groups (Fig. 2D; saline, 10.4 ± 1.5 ms; ethanol, 11.8 ± 1.1 ms; n.s.). Because SK was a dominant contributor to the TTP-AHP (see also Ping and Shepard 1996; Shepard and Bunney 1991), the significantly reduced TTP-AHP in ethanol animals suggests that withdrawal after repeated ethanol exposure reduced SK function.
To more directly investigate the effects of ethanol withdrawal on SK function, we performed voltage-clamp experiments to better isolate the apamin-sensitive SK current. Neurons were held at –70 mV, then depolarized for 400 ms with pulses ranging from –20 to –60 mV in 10-mV steps. On returning to –70 mV, a tail current due to slow deactivation of a channel was evident (see Fig. 3A; see also Paul et al. 2003). Both the peak amplitudes of the tail currents evoked after depolarizing steps to –20, –30, and –40 mV and the tail current charge transfer, calculated by integrating the tail current evoked after the depolarizing pulse (see METHODS), were significantly reduced in ethanol animals (Fig. 3, A–D), with saline similar to age-matched naïve animals (naïve, 240 ± 40 pA for step to –20 mV, n = 7; compared with 239 ± 12 pA for saline and 178 ± 13 pA for ethanol). In addition, the tail current was inhibited by >90% by apamin in both saline and ethanol animals (analyzed for the depolarizing step to –20 mV; saline, 92.9 ± 2.5% block; ethanol, 92.8 ± 2.5% block; n = 6 and 9 for saline and ethanol, respectively), indicating that SK is the primary channel active during the tail current following depolarization (see also Paul et al. 2003). In this regard, the apamin-sensitive component of the peak tail current was significantly reduced in ethanol relative to saline animals (Fig. 3E; saline, 60.6 ± 9.9 pA; ethanol, 34.8 ± 5.8 pA, P < 0.05), strongly suggesting that SK function was reduced in ethanol animals.
Differences in SK function could result from changes in basic membrane properties. However, neither the series resistance (saline, 14.7 ± 0.9 M; ethanol, 16.1 ± 0.8 M; n.s.) nor the instantaneous currents evoked by hyperpolarization (see Fig. 7, E and F, taken as a measure of input resistance) were altered in ethanol animals. In addition, we estimated whether cell capacitance might be different in neurons from saline and ethanol animals, perhaps indicating differences in cell size (see Neuhoff et al. 2002), following a method adapted from Gentet et al. (2000). Although there could be considerable error using this method due to space-clamp issues associated with low-pass filtering and voltage attenuation during patch clamp of neurons in brain slice with intact dendrites (Major 1993; Spruston et al. 1993), as well as a lack of direct measurement of the cell surface area, we observed no differences in the fast exponential capacitive transient after a voltage step from –70 to –60 mV (saline, 0.99 ± 0.06 ms; ethanol, 1.08 ± 0.05 ms; n.s.) or the cell capacitance (saline, 76.7 ± 3.4 pF; ethanol, 77.6 ± 4.0 pF; n.s.). Further, differences between saline and ethanol animals in integrated currents of SK (Fig. 3C) and Ih (see following text) persisted when normalized to the cell capacitance estimated in this way (data not shown). Taken together with a lack of differences in series resistance or the instantaneous currents evoked by hyperpolarization, these results suggest that apparent changes in SK function did not result from differences in basic membrane parameters.
Reduced SK function in ethanol animals could be due to several factors, including altered kinetics or reversal potential. However, the time constant of inactivation (fit from 55 to 550 ms into the tail current; Abel et al. 2004) was not different between ethanol and saline animals (saline, 139 ± 6 ms; ethanol, 140 ± 7 ms; n.s.). Also, the reversal potential of SK (determined by depolarizing to –20 mV for 400 ms, then stepping the cell to voltages ranging from –70 to –120 mV in 5-mV increments, in the presence of the Ih blocker ZD7288, 30 µM) was not different between groups (saline, –89.0 ± 3.2 mV; ethanol, –87.7 ± 1.9 mV; n = 4 for each group; n.s.). The deviation of the SK reversal potential relative to the EK+ derived from the Nerntz equation ( –100 mV) could be due to small space-clamp errors while patch-clamping neurons in a brain slice and/or localization of SK channels in the dendrites (Abel et al. 2004). Additionally, smaller SK amplitudes in ethanol animals might impair accurate determination of the reversal potential in those neurons. However, because the whole cell capacitance, input resistance, and series resistance were similar between saline and ethanol animals, any such errors should be equivalent in both groups.
NMDA burst firing was enhanced in ethanol animals
In addition to shaping the AHP, SK can regulate the firing pattern of VTA DA neurons. In particular, studies have demonstrated that SK inhibition facilitates the transition to bursting after application of NMDA (Johnson and Seutin 1997; Seutin et al. 1993). Entry of Na+ and Ca2+ after activation of NMDA receptors produces a depolarization and an increase in intracellular Ca2+ that, in addition to the inactivation of SK (which reduces the AHP), may facilitate the transition from pacemaker firing to burst firing (Johnson and Seutin 1997; Komendantov et al. 2004; Seutin et al. 1993). Because our action potential waveform analysis and voltage-clamp data suggest that SK function was reduced in VTA DA neurons from ethanol animals, we studied whether reduced SK function in ethanol animals would facilitate the ability of NMDA to induce bursting activity. To more precisely quantify bursting, we determined the coefficient of variation of the interspike interval (CV-ISI), a measure of spike train irregularity, before and after NMDA application. CV-ISI was defined as the SD divided by the mean of the ISI distribution (Zhang et al. 1994) and was calculated across 100 action potentials. The change from pacemaker to bursting activity increases the CV-ISI, where the CV-ISI is small for very regular spike trains (like pacemaker) and larger in irregular or bursting spike trains.
Bath application of NMDA (20 µM) for 10 min increased the firing frequency in neurons from saline animals, but generally did not induce bursting activity, indicated by no difference in the CV-ISI before and after application of 20 µM NMDA (Fig. 4, A and D; pre-NMDA, 23.8 ± 4.8%; post-NMDA, 31.1 ± 9.6%; n = 10; n.s.). In contrast, the same dose of NMDA (20 µM; n = 11) induced bursting activity in ethanol animals, indicated by a significant increase in CV-ISI (Fig. 4, A and D; pre-NMDA, 27.5 ± 5.5%; post-NMDA, 79.4 ± 16.1%; P < 0.05), with a significant difference in the CV-ISI between saline and ethanol after 20 µM NMDA (P < 0.05). Example histograms of the ISI distribution before and after 20 µM NMDA are shown in Fig. 4, B and C, and demonstrate that NMDA enhanced firing rate but maintained a regular firing pattern in saline animals, whereas NMDA produced an ISI distribution with both short ISIs (during a burst) and longer ISIs (between bursts) in ethanol animals.
A lower dose of NMDA (10 µM) increased firing rate but did not produce bursting or alter the CV-ISI in either group (Fig. 4D; saline, 20.2 ± 7.5% post-NMDA; ethanol, 33.0 ± 10.7% post-NMDA; n = 7 and 6 for saline and ethanol, respectively). A lack of bursting in saline animals was not due to insufficient NMDA receptor activation because 50 µM NMDA also did not significantly increase the CV-ISI in saline animals (Fig. 4D; 34.1 ± 14.1% post-NMDA; n = 5). However, neurons from saline animals exhibited bursting and a significantly elevated CV-ISI when 20 µM NMDA was combined with SK inhibition by apamin (Figs. 4D and 5B; 78.8 ± 6.2% post-NMDA; n = 5; P < 0.05), in agreement with studies showing bursting after NMDA and apamin in combination (Johnson and Seutin 1997; Seutin et al. 1993).
Although NMDA alone generally did not elicit bursting in saline neurons, bursting was occasionally observed (an example is shown in Fig. 5B). Thus to further define whether bursting was induced within a particular neuron, a criterion was defined whereby individual neurons would be considered bursting if they had a CV-ISI greater than the mean + 2 SDs of the CV-ISI values taken from neurons from saline animals before NMDA addition, yielding a threshold of a 68.5% CV-ISI (Fig. 5A). With this criterion, bursting was observed in saline animals in 2 of 10 neurons exposed to 20 µM NMDA, 1 of 5 neurons exposed to 50 µM NMDA, but 4 of 5 neurons exposed to 20 µM NMDA combined with apamin (Fig. 5A). Further, bursting was observed in 8 of 11 neurons from ethanol animals exposed to 20 µM NMDA (Fig. 5A). Thus NMDA alone can induce bursting in some neurons from saline animals, but the incidence of bursting in 20 µM NMDA was significantly greater in ethanol animals (P < 0.05, 
2 test). Further, inhibition of SK at the same time as depolarization with NMDA strongly facilitated the transition to bursting in saline animals and generated bursting that was very similar to that observed in ethanol animals exposed only to NMDA (Figs. 4D and 5B).
Because there was variability among neurons regarding the pre-NMDA baseline (Fig. 5A), we also calculated a CV-ISI ratio, defined as the ratio of the CV-ISI after NMDA to the CV-ISI before NMDA. The CV-ISI ratio was significantly greater in ethanol relative to saline neurons (Fig. 5C; saline, 1.3 ± 0.3; ethanol, 3.0 ± 0.6; P < 0.05), in agreement with a greater increase in variability in ISIs that typifies burst firing (see Fig. 4, A–D). In addition, we observed that the CV ratio was significantly negatively correlated with the SK tail current (Fig. 5D; r2 = 0.387; P < 0.05). Because neurons from ethanol animals that exhibited smaller SK function showed a significantly larger increase in the CV ratio, this suggests that reduced SK function might be responsible for the enhanced NMDA-induced transition to bursting in ethanol animals.
Finally, differences in NMDA-induced bursting could result from different NMDA-dependent receptor activation. To examine this possibility, we determined the current response to a 30-s exposure to NMDA (Vholding = +40 mV) under voltage clamp with a Cs+-based internal solution (Borgland et al. 2006). Bath application of 10 or 20 µM NMDA produced a dose-dependent increase in evoked current that was similar in neurons from saline and ethanol animals (Fig. 6; n = 6 for all groups and doses), suggesting that NMDA-receptor activation was not different between the two groups. Thus bursting induced by NMDA was significantly facilitated in ethanol animals, with no change in NMDA-receptor activity, whereas neurons from saline animals could exhibit bursting with 20 µM NMDA combined with apamin, but generally did not exhibit bursting after NMDA alone. Taken together, these results strongly suggest that the transition to burst firing was facilitated in ethanol animals and that the reduced SK function in ethanol animals might play a key role in this enhanced transition to bursting.
Ih function was reduced and recovery from hyperpolarization impaired in ethanol animals, but reduced Ih did not account for enhanced transition to bursting
The Ih current can also regulate the activity of VTA DA neurons (Neuhoff et al. 2002; Okamoto et al. 2006; Seutin et al. 2001). To measure Ih, we performed voltage-clamp experiments where a series of hyperpolarizing voltage steps (500 ms, ranging from –40 to –150 mV in 10- mV steps) was applied to VTA neurons from a holding potential of –60 mV, and the magnitude of Ih determined from the slowly developing current sag (Cathala and Paupardin-Tritsch 1997; Liu et al. 2003; Neuhoff et al. 2002). We also determined the Ih charge transfer by integrating the area under the slowly activating inward current in the –140-mV hyperpolarizing voltage pulse (see METHODS). Finally, to better isolate Ih, we performed some voltage-clamp experiments under conditions where many currents other than Ih were inhibited using modified internal and external solutions (see METHODS and Liu et al. 2003).
Figure 7, A and B shows examples of the current response to a step from –60 to –140 mV in both experimental groups. The Ih current was significantly decreased in ethanol compared with saline animals, measured both in terms of peak current (Fig. 7, A–D; P < 0.01) and Ih charge transfer (Fig. 7G, measured for step to –140 mV; P < 0.05), and using either standard intracellular and extracellular solutions (Fig. 7, A, C, and E; saline, n = 11; ethanol, n = 9) or solutions that provide better isolation of Ih (Fig. 7, B, D, and F; saline, n = 6; ethanol, n = 8). Also, the Ih current in standard solutions in saline neurons was similar to that in neurons from age-matched naïve rats (measured for step to –140 mV, naïve, 625 ± 74 pA, n = 7; compared with 670 ± 59 pA from saline and 476 ± 61 pA from ethanol). In addition, Ih was determined in neurons from the additional 18 saline and 16 ethanol animals using a current step from –80 to –140 mV. Although the Ih measured in this manner was significantly reduced in ethanol relative to saline animals (saline, 382 ± 20 pA; ethanol, 321 ± 18 pA; P < 0.05), Ih is partially active at –80 mV (Cathala and Paupardin-Tritsch 1997; Liu et al. 2003; Neuhoff et al. 2002), and thus these experiments were not used for kinetic analyses. Finally, instantaneous currents were not altered in ethanol relative to saline animals, measured either as peak currents or charge transfer during a hyperpolarizing step from –60 mV (Fig. 7, E, F, and H), perhaps suggesting no changes in IRK and/or potassium leak channels (Watts et al. 1996; Wilson 2005).
Although total Ih currents were reduced in ethanol animals, several kinetic parameters of Ih were not altered. First, the voltage dependence of activation of Ih, fit with a Boltzmann equation (see METHODS), was not different between saline and ethanol animals (Fig. 8, A and B), with no differences in V1/2 (Fig. 8C) or slope factor (Fig. 8D). Thus ethanol reduced the total Ih current without altering the voltage dependence of activation of Ih (see also Okamoto et al. 2006). Repeated ethanol exposure also did not alter the reversal potential of Ih (Fig. 8, E and F; n = 6 for saline and n = 9 for ethanol), which was estimated, as described (Cathala and Paupardin-Tritsch 1997; Mayer and Westbrook 1983), by comparing the instantaneous currents evoked from a holding voltage of –60 mV, where Ih is not active, and the instantaneous currents evoked from a holding voltage of –80 mV, where Ih is activated (Cathala and Paupardin-Tritsch 1997; Liu et al. 2003; Neuhoff et al. 2002). In particular, one can estimate the reversal potential of Ih from the point where the two I–V curves intersect; because currents from a –60-mV holding potential contain IRK/leak but little Ih, whereas currents from a –80-mV holding potential contain both IRK/leak and Ih due to activation of Ih at –80 mV, the intersection of the I–V curves from –60 and –80 mV essentially represents the voltage where the Ih current is zero after the IRK/leak current is subtracted out.
Ih has been reported to alter burst firing in other types of neurons (Wilson 2005). Thus we examined the effects of the Ih antagonist ZD7288 (30 µM) on firing or NMDA modulation of bursting after 10-min exposure to ZD7288, after which time >90% of Ih should be inhibited (Fig. 9C; Gasparini and DiFrancesco 1997; Gu et al. 2007; Okamoto et al. 2006). In saline animals, ZD7288 did not facilitate a transition to bursting when NMDA was applied subsequent to ZD7288, indicated by no significant increase in the CV-ISI (Fig. 9A; baseline: 31.1 ± 6.6%; after ZD7288 addition: 25.1 ± 5.5%; after ZD7288 and NMDA: 32.3 ± 10.8%; n.s.; n = 5). Thus the reduced Ih in ethanol animals cannot account for the facilitated transition to bursting by NMDA in ethanol animals. In addition, ZD7288 did not alter pacemaker firing, measured by determining the instantaneous firing frequency, in either saline animals (Fig. 9B; before ZD7288: 0.75 ± 0.06 Hz; after ZD7288: 0.72 ± 0.14 Hz; n = 8; n.s.) or ethanol animals (before ZD7288: 0.86 ± 0.05 Hz; after ZD7288: 0.98 ± 0.10 Hz; n = 8; n.s.), in agreement with no differences in pacemaker firing (Fig. 1).
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; Puopolo et al. 2007; Satoh and Yamada 2000; Wilson 2005). Therefore to further investigate the effects of ethanol withdrawal on Ih, a set of hyperpolarizing steps (300 ms, ranging from –50 to –300 pA) was applied to the neurons during pacemaker firing, and the time to recover the firing activity was measured. Recovery from hyperpolarization was significantly delayed in ethanol animals relative to saline animals (examples shown in Fig. 9D; n = 10 for saline and n = 9 for ethanol). Using a two-way, repeated- measures ANOVA, we found a significant effect of treatment (ethanol vs. saline; F(1,17) = 8.344; P < 0.05), the hyperpolarizing step (F(5,85) = 28.690; P < 0.001), and the interaction between the two factors (treatment by hyperpolarizing steps; F(5,85) = 4.365; P < 0.01). These results support the suggestion that the reduced Ih function, apparent in voltage-clamp experiments (Fig. 7), also impaired the ability of VTA DA neurons from ethanol animals to recover from hyperpolarizing events. In addition, as shown in Fig. 9, E and F, ZD7288 significantly delayed the recovery from hyperpolarization in saline animals (F(1,14) = 7.073; P < 0.05; two-way, repeated-measures ANOVA; n = 8). In strong contrast, ZD7288 had no effect on the recovery from hyperpolarization in ethanol animals (Fig. 9, E and G; F(1,14) = 0.227; n.s.; two-way, repeated-measures ANOVA; n = 8). These results suggest that the reduction in Ih may impair recovery from hyperpolarization in ethanol animals, in agreement with studies suggesting a critical role for Ih in DA neurons in recovery from hyperpolarization (Neuhoff et al. 2002; Puopolo et al. 2007). Withdrawal from repeated ethanol treatment enhanced the locomotor response to cocaine
Finally, we wanted to establish whether the repeated ethanol treatment used here could be sufficient to produce physiological changes in ethanol animals. In this regard, drug-induced locomotor sensitization has been associated with some aspects of drug reward, dependence, and relapse (Robinson and Berridge 2001), and repeated ethanol exposure has been shown to produce a cross-sensitization to the locomotor-activating effects of cocaine (Itzhak and Martin 1999). Thus we examined whether the ethanol exposure protocol used here enhanced the motor response to cocaine, examined after 7 days of withdrawal.
Animals withdrawn after repeated ethanol or saline were habituated to the open-field locomotor chamber for 3 days (see METHODS) before administration of cocaine on the 7th day of withdrawal. Ethanol animals exhibited a significantly greater locomotor response to an acute injection of cocaine (15 mg/kg, ip) relative to saline animals or naïve animals (Fig. 10; n = 16, 13, and 11 for saline, ethanol, and naïve animals, respectively; P < 0.05). No differences were observed between groups in the locomotor activity during a 60-min period of habituation or in response to an acute injection of saline (Fig. 10). Further, saline animals exhibited a locomotor response to cocaine very similar to that in naïve animals, suggesting that any stress related to multiple injections or handling procedures did not enhance the locomotor response to cocaine. Taken together, these results indicate that the repeated ethanol exposure and withdrawal paradigm used here altered channel function and also produced a cross-sensitization to the acute locomotor-activating effects of cocaine.
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DISCUSSION |
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Pacemaker activity was not altered in VTA DA neurons from ethanol animals
Pacemaker firing is under strong control of a number of currents including Ca2+ channels and A-type potassium channels (see Marinelli et al. 2006; Puopolo et al. 2007) and the Na+/K+-ATPase (Johnson et al. 1992). However, studies of SK and Ih regulation of pacemaker activity in midbrain DA neurons have yielded mixed results. When pacemaker frequencies are more similar to those observed here, SK inhibition with apamin did not alter pacemaker firing (Brodie et al. 1999; Johnson and Wu 2004; Seutin et al. 1993; Waroux et al. 2005), although, with more rapid firing, apamin can increase firing rates (Ping and Shepard 1996; Shepard and Bunney 1988; Waroux et al. 2005; Wolfart et al. 2001). Further, Ih-dependent depolarization enhanced firing in some studies (Okamoto et al. 2006; Seutin et al. 2001), but other results suggest a less-consistent relationship between Ih and pacemaker activity (Liu et al. 2003; Neuhoff et al. 2002), and Ih can reduce firing by decreasing the input resistance (Fan et al. 2005). Thus the currents responsible for the strong oscillatory drive in VTA neurons could override the contribution of Ih and SK during pacemaker firing, perhaps also explaining the lack of difference in pacemaker activity between saline and ethanol animals despite functional changes in several currents.
Previous in vivo and in vitro observations during withdrawal after chronic ethanol exposure found reduced firing rates in adult midbrain DA neurons after 1–3 days (Bailey et al. 1998; Diana et al. 1993, 1996; Shen 2003; but see Brodie 1999) or 6 days of withdrawal (Bailey et al. 2001), with no changes in firing rates of active DA neurons 3 wk after repeated ethanol exposure (Bailey et al. 2001; Shen et al. 2007), although the number of neurons firing was reduced (Shen et al. 2007). Here, we observed no changes in pacemaker activity 7 days after repeated ethanol treatment. Taken together, these observations suggest that withdrawal from chronic ethanol treatment decreases pacemaker activity for several days (but see Brodie 1999), but that this effect is absent after 7 days of withdrawal. However, it should be noted that our studies used juvenile animals, and thus our results might not fully reflect those collected from adult animals, although differences in species (mice vs. rat) and route of ethanol administration (liquid diet vs. intermittent ip injections) could also explain apparent discrepancies between our studies and those of Bailey and colleagues (2001).
SK function was reduced and burst firing enhanced in ethanol animals
Several lines of evidence suggest that repeated ethanol exposure decreased SK function in VTA neurons. The time-to-peak of the AHP (TTP-AHP) was significantly reduced in ethanol animals, with no apparent changes in several other parameters of the action potential waveform. The TTP-AHP was also greatly reduced by the SK antagonist apamin (Ping and Shepard 1996; Shepard and Bunney 1988), suggesting a primary contribution of SK to the TTP-AHP. Further, SK-mediated tail currents (Paul et al. 2003) were significantly reduced in ethanol animals, with a significant correlation between the TTP-AHP and the tail current magnitude in ethanol animals. Taken together, these results strongly suggest that SK function was reduced in ethanol animals. Further experiments will be required to determine whether differences in SK function in ethanol animals reflect changes in number or regulation of SK or of the calcium channels required for SK activation (Bond et al. 2005).
In addition to regulation of the AHP, SK inhibition facilitates NMDA-induced burst firing (Johnson and Seutin 1997; Seutin et al. 1993; Waroux et al. 2005). Here, NMDA (20 µM) increased pacemaker frequency in saline animals, but induced burst firing in ethanol animals, which could be explained by decreased SK activity facilitating NMDA induction of bursting in ethanol animals. Differential NMDA induction of burst firing was not due to differences in NMDA receptor activation because the current response to NMDA, measured under voltage clamp, was similar in saline and ethanol animals. In addition, control neurons were capable of bursting because SK inhibition in combination with NMDA elicited bursting in saline animals, as is widely observed in control neurons (Johnson and Seutin 1997; Seutin et al. 1993). Thus neurons from saline animals could exhibit bursting, especially when SK was inhibited during NMDA application, but NMDA induction of bursting was greatly facilitated in ethanol animals, perhaps due to decreased SK function. In addition, NMDA induction of bursting in ethanol neurons was significantly negatively correlated with the SK tail current magnitude, strongly supporting the possibility that reduced SK function might be responsible for the enhanced NMDA-induced transition to bursting in ethanol animals.
An increased ability of the glutamate system in the VTA to induce bursting activity after repeated drug exposure may contribute to drug seeking and enhance the vulnerability to relapse during ethanol withdrawal. Glutamatergic afferents into the VTA can regulate the shift from pacemaker to burst firing (Chergui et al. 1994; Floresco et al. 2001; Grillner and Mercuri 2002), which increases DA release in midbrain target regions and may contribute to some aspects of drug-induced reinstatement (Grace 2000; Schmidt and Pierce 2006; Tupala and Tiihonen 2004).
Ih function was reduced and recovery from hyperpolarization impaired in ethanol animals
Repeated ethanol exposure and 7 days' withdrawal also significantly reduced hyperpolarization-evoked Ih currents, which could significantly influence firing properties. For example, Ih drives certain aspects of oscillatory activity in striatal cholinergic interneurons (Wilson 2005), and thus decreased Ih might retard bursting by reducing the depolarizing drive (Wilson 2005). However, NMDA-induced bursting was enhanced in ethanol animals and Ih inhibition in saline animals did not enhance NMDA-induced bursting, suggesting that differences in Ih did not account for the increased NMDA-induced transition to bursting in ethanol animals. In addition, Ih can facilitate recovery from hyperpolarization (Neuhoff et al. 2002; Satoh and Yamada 2000; Wilson 2005). Firing is delayed after strong hyperpolarization (see Fig. 9D) due to an A-type potassium current (Koyama and Appel 2006) and Ih activation at hyperpolarized potentials can provide depolarization to drive neurons to voltages permissive for firing. Here, neurons from ethanol animals showed impaired recovery of firing after hyperpolarization, in agreement with voltage-clamp observations suggesting decreased Ih function. Also, the Ih antagonist ZD7288 significantly delayed recovery of firing following hyperpolarization in saline animals, but had no effect in ethanol animals, suggesting that reduced Ih in ethanol animals was responsible for the delayed recovery from hyperpolarization. Reduced Ih was not due to altered voltage dependence of activation, in agreement with a previous study showing a dissociation between changes in total Ih current and voltage dependence of activation of Ih (Okamoto et al. 2006).
Several papers have raised the possibility that ZD7288 exhibits nonspecific effects through targets other than Ih (Chevaleyre and Castillo 2002; Wilson 2005). The concentration of ZD7288 used here is widely used, and has been reported not to affect a number of physiological parameters including instantaneous currents during hyperpolarization (Harris and Constanti 1995; Liu et al. 2003; Okamoto et al. 2006; but see Wilson 2005), AP potential waveform except for the AHP (Gasparini and DiFrancesco 1997; Harris and Constanti 1995; and data not shown), and several forms of synaptic plasticity (Chevaleyre and Castillo 2002; Gasparini and DiFrancesco 1997; but see Mellor et al. 2002). In addition, in several studies, nonspecific effects of ZD7288 were generally evident at higher ZD7288 concentrations and/or after much longer exposure times (Chevaleyre and Castillo 2002; Mellor et al. 2002; Wilson 2005) than those used here. We found that ZD7288 did not alter pacemaker firing or NMDA modulation of firing, but did alter recovery from hyperpolarization in saline but not ethanol animals, suggesting a more selective effect of ZD7288 on VTA DA neuron firing.
We should note that it is unclear whether the Ih modulation of recovery from hyperpolarization is directly relevant in vivo because the VTA in vivo generally does not exhibit such stereotyped pauses in firing, compared, for example, with striatal cholinergic interneurons (see Wilson 2005). The purpose of measuring the recovery from hyperpolarization was not to claim that a pause such as that observed by the strong hyperpolarizing current step actually occurs in vivo in the VTA. Instead, our results suggest that, under conditions of strong hyperpolarization (e.g., with strong activation of IRK), a reduction in Ih will facilitate the potency of hyperpolarization for reducing firing, as has been demonstrated by Liu and colleagues (2005). However, several groups have found a reduction in VTA firing during omission of an expected reward (see Schultz 2002); the underlying cellular mechanism is unclear, and could result from circuit interactions, but neuromodulator activation of IRK might also contribute to this pause in firing (but see Ji and Shepard 2007).
At present, the mechanisms through which repeated ethanol exposure might produce lasting changes in SK and Ih function remain unclear. Expression of both SK and HCN subunits could exhibit persistent plasticity (Chaplain et al. 2003; Kye et al. 2007; Seroussi et al. 2002), and SK subunit expression can be regulated by transcriptional modulators (Kye et al. 2007; see also Sun et al. 2001). Because ethanol can activate a number of transcriptional pathways (Mulligan et al. 2006), we speculate that protracted changes in channel function after repeated ethanol could reflect altered channel expression. In addition, it is possible that decreased Ih function after repeated ethanol exposure and withdrawal might represent a compensatory mechanism opposing the actions of reduced SK channel function on firing; for example, substantia nigra neurons exhibit greater function of both Ih and SK relative to VTA (Neuhoff et al. 2002; Wolfart et al. 2001), suggesting a possible counterbalancing role of Ih and SK in midbrain DA neuron firing.
Repeated ethanol treatment enhanced locomotor activation by cocaine
Here, repeated ethanol exposure enhanced the locomotor activation produced when cocaine was acutely administered after 7 days of withdrawal, in agreement with a previously described cross-sensitization to the locomotor-activating effects of cocaine after repeated ethanol exposure (Itzhak and Martin 1999). We should note that not all studies examining repeated ethanol injection have observed subsequent enhanced locomotor activation by cocaine (Lessov and Phillips 2003; Wise et al. 1996). However, as noted by Lessov and Phillips (2003), the drug treatment schedules and time of testing for sensitization vary widely across these studies, making comparisons difficult. In addition, because all these studies were performed in mice, ours is the first to demonstrate in rats that repeated ethanol exposure and 7 days' withdrawal can significantly enhance the locomotor-activating effects of cocaine, perhaps representing a form of cross-sensitization.
It has been proposed that sensitization represents an increase in the positive value attributed to a drug or drug-associated stimulus, and sensitization has thus been considered a model of altered drug seeking (Robinson and Berridge 2001). In addition, sensitization may result from persisting neuroadaptations in mesolimbic brain regions (Nestler 2001; Vanderschuren and Kalivas 2000; Vezina 2004), including the VTA (e.g., Liu et al. 2005; Ungless et al. 2001). Although the VTA is critical for the development of sensitization for many abused drugs, the direct role of the VTA in expression of sensitization may depend on the particular drug of abuse, where a clearer role for opiates but a less clear role for psychostimulants has been described (Leite-Morris et al. 2004; Vanderschuren and Kalivas 2000). In addition, a recent study showed that ethanol-related sensitization in mice can be apparent without increased dopamine release, perhaps dissociating the dopaminergic system from enhanced locomotor activation by ethanol after repeated ethanol exposure in mice (Zapata et al. 2006). Although little is known about the cellular mechanisms underlying enhanced locomotor activation after repeated ethanol exposure, reduced SK and facilitation of NMDA-induced bursting might contribute to the enhanced locomotor activation by cocaine after repeated ethanol exposure that we and others (Itzhak and Martin 1999) have observed.
In conclusion, we have shown here that repeated ethanol exposure and 7 days' withdrawal led to several alterations in regulation of firing of VTA DA neurons. In particular, several lines of evidence indicate that SK function was reduced in ethanol relative to saline animals. Interestingly, neurons from ethanol animals showed an increased propensity for bursting during exposure to NMDA, with no change in the amount of NMDA-dependent depolarization measured under voltage-clamp conditions, suggesting that the reduced SK function could be responsible for the enhanced bursting in ethanol animals. Repeated ethanol exposure also led to reduced Ih function, which impaired the ability of VTA neurons to recover from hyperpolarization, but likely did not account for the enhanced bursting in ethanol animals. Also, repeated ethanol exposure increased the locomotor activation by acute cocaine, confirming that the ethanol exposure procedure used here was able to produce functional physiological changes. Thus repeated ethanol exposure and withdrawal produced enhanced bursting of VTA DA neurons during withdrawal from repeated ethanol exposure, which may contribute to other addiction-related behaviors through increasing the salience of drug-related stimuli.
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GRANTS |
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ACKNOWLEDGMENTS |
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Present address of M. Martin: Unit of Neuropharmacology, Pompeu Fabra University, Barcelona, Catalonia, Spain.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Address for reprint requests and other correspondence: A. Bonci, Ernest Gallo Clinic and Research Center, University of California, San Francisco, Department of Neurology, 5858 Horton St., Suite 200, Emeryville, CA 94608 (E-mail: Antonello.Bonci{at}ucsf.edu)
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REFERENCES |
|---|
|
Appel SB, Liu Z, McElvain MA, Brodie MS. Ethanol excitation of dopaminergic ventral tegmental area neurons is blocked by quinidine. J Pharmacol Exp Ther 306: 437–446, 2003.
Bailey CP, Manley SJ, Watson WP, Wonnacott S, Molleman A, Little HJ. Chronic ethanol administration alters activity in ventral tegmental area neurons after cessation of withdrawal hyperexcitability. Brain Res 803: 144–152, 1998.[CrossRef][Web of Science][Medline]
Bailey CP, O'Callaghan MJ, Croft AP, Manley SJ, Little HJ. Alterations in mesolimbic dopamine function during the abstinence period following chronic ethanol consumption. Neuropharmacology 41: 989–999, 2001.[CrossRef][Web of Science][Medline]
Bond CT, Maylie J, Adelman JP. SK channels in excitability, pacemaking and synaptic integration. Curr Opin Neurobiol 15: 305–311, 2005.[CrossRef][Web of Science][Medline]
Borgland SL, Taha SA, Sarti F, Fields HL, Bonci A. Orexin A in the VTA is critical for the induction of synaptic plasticity and behavioral sensitization to cocaine. Neuron 49: 589–601, 2006.[CrossRef][Web of Science][Medline]
Brodie MS. Increased ethanol excitation of dopaminergic neurons of the ventral tegmental area after chronic ethanol treatment. Alcohol Clin Exp Res 26: 1024–1030, 2002.[CrossRef][Web of Science][Medline]
Brodie MS, McElvain MA, Bunney EB, Appel SB. Pharmacological reduction of small conductance calcium-activated potassium current (SK) potentiates the excitatory effect of ethanol on ventral tegmental area dopamine neurons. J Pharmacol Exp Ther 290: 325–333, 1999.
Brodie MS, Shefner SA, Dunwiddie TV. Ethanol increases the firing rate of dopamine neurons of the rat ventral tegmental area in vitro. Brain Res 508: 65–69, 1990.[CrossRef][Web of Science][Medline]
Cathala L, Paupardin-Tritsch D. Neurotensin inhibition of the hyperpolarization-activated cation current (Ih) in the rat substantia nigra pars compacta implicates the protein kinase C pathway. J Physiol 503: 87–97, 1997.
Chaplan SR, Guo HQ, Lee DH, Luo L, Liu C, Kuei C, Velumian AA, Butler MP, Brown SM, Dubin AE. Neuronal hyperpolarization-activated pacemaker channels drive neuropathic pain. J Neurosci 23: 1169–1178, 2003.
Chergui K, Akaoka H, Charlety PJ, Saunier CF, Buda M, Chouvet G. Subthalamic nucleus modulates burst firing of nigral dopamine neurones via NMDA receptors. Neuroreport 5: 1185–1188, 1994.[Web of Science][Medline]
Chevaleyre V, Castillo PE. Assessing the role of Ih channels in synaptic transmission and mossy fiber LTP. Proc Natl Acad Sci USA 99: 9538–9543, 2002.
Cooper DC. The significance of action potential bursting in the brain reward circuit. Neurochem Int 41: 333–340, 2002.[CrossRef][Web of Science][Medline]
Diana M, Gessa GL, Rossetti ZL. Lack of tolerance to ethanol-induced stimulation of mesolimbic dopamine system. Alcohol 27: 329–333, 1992.[Web of Science]
Diana M, Pistis M, Carboni S, Gessa GL, Rossetti ZL. Profound decrement of mesolimbic dopaminergic neuronal activity during ethanol withdrawal syndrome in rats: electrophysiological and biochemical evidence. Proc Natl Acad Sci USA 90: 7966–7969, 1993.
Diana M, Pistis M, Muntoni A, Gessa G. Mesolimbic dopaminergic reduction outlasts ethanol withdrawal syndrome: evidence of protracted abstinence. Neuroscience 71: 411–415, 1996.[CrossRef][Web of Science][Medline]
Fan Y, Fricker D, Brager DH, Chen X, Lu HC, Chitwood RA, Johnston D. Activity- dependent decrease of excitability in rat hippocampal neurons through increases in I(h). Nat Neurosci 8: 1542–1551, 2005.[CrossRef][Web of Science][Medline]
Floresco SB, Todd CL, Grace AA. Glutamatergic afferents from the hippocampus to the nucleus accumbens regulate activity of ventral tegmental area dopamine neurons. J Neurosci 21: 4915–4922, 2001.
Gasparini S, DiFrancesco D. Action of the hyperpolarization-activated current (Ih) blocker ZD 7288 in hippocampal CA1 neurons. Pfluegers Arch 435: 99–106, 1997.[CrossRef][Web of Science][Medline]
Gentet LJ, Stuart GJ, Clements JD. Direct measurement of specific membrane capacitance in neurons. Biophys J 79: 314–320, 2000.[Web of Science][Medline]
Gonzales RA, Job MO, Doyon WM. The role of mesolimbic dopamine in the development and maintenance of ethanol reinforcement. Pharmacol Ther 103: 121–146, 2004.[CrossRef][Web of Science][Medline]
Gonzales RA, Weiss F. Suppression of ethanol-reinforced behavior by naltrexone is associated with attenuation of the ethanol-induced increase in dialysate dopamine levels in the nucleus accumbens. J Neurosci 18: 10663–10671, 1998.
Grace AA. The tonic/phasic model of dopamine system regulation and its implications for understanding alcohol and psychostimulant craving. Addiction 95, Suppl. 2: S119–S128, 2000.
Grillner P, Mercuri NB. Intrinsic membrane properties and synaptic inputs regulating the firing activity of the dopamine neurons. Behav Brain Res 130: 149–169, 2002.[CrossRef][Web of Science][Medline]
Gu N, Vervaeke K, Hu H, Storm JF. Kv7/KCNQ/M and HCN/h, but not KCa2/SK channels, contribute to the somatic medium after-hyperpolarization and excitability control in CA1 hippocampal pyramidal cells. J Physiol 566: 689–715, 2005.
Harris NC, Constanti A. Mechanism of block by ZD 7288 of the hyperpolarization-activated inward rectifying current in guinea pig substantia nigra neurons in vitro. J Neurophysiol 74: 2366–2378, 1995.
Hodge CW, Haraguchi M, Erickson H, Samson HH. Ventral tegmental microinjections of quinpirole decrease ethanol and sucrose-reinforced responding. Alcohol Clin Exp Res 17: 370–375, 1993.[CrossRef][Web of Science][Medline]
Itzhak Y, Martin JL. Effects of cocaine, nicotine, dizocipline and alcohol on mice locomotor activity: cocaine-alcohol cross-sensitization involves upregulation of striatal dopamine transporter binding sites. Brain Res 818: 204–211, 1999.[CrossRef][Web of Science][Medline]
Ji H, Shepard PD. Lateral habenula stimulation inhibits rat midbrain dopamine neurons through a GABA(A) receptor-mediated mechanism. J Neurosci 27: 6923–6930, 2007.
Johnson SW, North RA. Two types of neurones in the rat ventral tegmental area and their synaptic inputs. J Physiol 450: 455–468, 1992.
Johnson SW, Seutin V. Bicuculline methiodide potentiates NMDA-dependent burst firing in rat dopamine neurons by blocking apamin-sensitive Ca2+-activated K+ currents. Neurosci Lett 231: 13–16, 1997.[CrossRef][Web of Science][Medline]
Johnson SW, Seutin V, North RA. Burst firing in dopamine neurons induced by N-methyl-D-aspartate: role of electrogenic sodium pump. Science 258: 665–667, 1992.
Johnson SW, Wu YN. Multiple mechanisms underlie burst firing in rat midbrain dopamine neurons in vitro. Brain Res 1019: 293–296, 2004.[CrossRef][Web of Science][Medline]
Katner SN, Weiss F. Ethanol-associated olfactory stimuli reinstate ethanol-seeking behavior after extinction and modify extracellular dopamine levels in the nucleus accumbens. Alcohol Clin Exp Res 23: 1751–1760, 1999.[CrossRef][Web of Science][Medline]
Komendantov AO, Komendantova OG, Johnson SW, Canavier CC. A modeling study suggests complementary roles for GABAA and NMDA receptors and the SK channel in regulating the firing pattern in midbrain dopamine neurons. J Neurophysiol 91: 346–357, 2004.
Koyama S, Appel SB. A-type K+ current of dopamine and GABA neurons in the ventral tegmental area. J Neurophysiol 96: 544–554, 2006.
Kye MJ, Spiess J, Blank T. Transcriptional regulation of intronic calcium-activated potassium channel SK2 promoters by nuclear factor-kappa B and glucocorticoids. Mol Cell Biochem 300: 9–17, 2007.[CrossRef][Web of Science][Medline]
Leite-Morris KA, Fukudome EY, Shoeb MH, Kaplan GB. GABA(B) receptor activation in the ventral tegmental area inhibits the acquisition and expression of opiate-induced motor sensitization. J Pharmacol Exp Ther 308: 667–678, 2004.
Lessov CN, Phillips TJ. Cross-sensitization between the locomotor stimulant effects of ethanol and those of morphine and cocaine in mice. Alcohol Clin Exp Res 27: 616–627, 2003.[Web of Science][Medline]
Liu QS, Pu L, Poo MM. Repeated cocaine exposure in vivo facilitates LTP induction in midbrain dopamine neurons. Nature 437: 1027–1031, 2005.[CrossRef][Medline]
Liu Z, Bunney EB, Appel SB, Brodie MS. Serotonin reduces the hyperpolarization-activated current (Ih) in ventral tegmental area dopamine neurons: involvement of 5-HT2 receptors and protein kinase C. J Neurophysiol 90: 3201–3212, 2003.
Major G. Solutions for transients in arbitrarily branching cables: III. Voltage clamp problems. Biophys J 65: 469–491, 1993.[Web of Science][Medline]
Margolis EB, Lock H, Hjelmstad GO, Fields HL. The ventral tegmental area revisited: is there an electrophysiological marker for dopaminergic neurons? J Physiol 577: 907–924, 2006.
Marinelli M, Rudick CN, Hu XT, White FJ. Excitability of dopamine neurons: modulation and physiological consequences. CNS Neurol Disord Drug Targets 5: 79–97, 2006.[Medline]
Marinelli S, Pascucci T, Bernardi G, Puglisi-Allegra S, Mercuri NB. Activation of TRPV1 in the VTA excites dopaminergic neurons and increases chemical- and noxious-induced dopamine release in the nucleus accumbens. Neuropsychopharmacology 30: 864–870, 2005.[CrossRef][Web of Science][Medline]
Mayer ML, Westbrook GL. A voltage-clamp analysis of inward (anomalous) rectification in mouse spinal sensory ganglion neurones. J Physiol 340: 19–45, 1983.
McBride WJ, Le AD, Noronha A. Central nervous system mechanisms in alcohol relapse. Alcohol Clin Exp Res 26: 280–286, 2002.[CrossRef][Web of Science][Medline]
Mellor J, Nicoll RA, Schmitz D. Mediation of hippocampal mossy fiber long-term potentiation by presynaptic Ih channels. Science 295: 143–147, 2002.
Mereu G, Fadda F, Gessa GL. Ethanol stimulates the firing rate of nigral dopaminergic neurons in unanesthetized rats. Brain Res 292: 63–69, 1984.[CrossRef][Web of Science][Medline]
Mulligan MK, Ponomarev I, Hitzemann RJ, Belknap JK, Tabakoff B, Harris RA, Crabbe JC, Blednov YA, Grahame NJ, Phillips TJ, Finn DA, Hoffman PL, Iyer VR, Koob GF, Bergeson SE. Toward understanding the genetics of alcohol drinking through transcriptome meta-analysis. Proc Natl Acad Sci USA 103: 6368–6373, 2006.
Nestler EJ. Molecular basis of long-term plasticity underlying addiction. Nat Rev Neurosci 2: 119–128, 2001.[CrossRef][Web of Science][Medline]
Neuhoff H, Neu A, Liss B, Roeper J. I(h) channels contribute to the different functional properties of identified dopaminergic subpopulations in the midbrain. J Neurosci 22: 1290–1302, 2002.
Okamoto T, Harnett MT, Morikawa H. Hyperpolarization-activated cation current (Ih) is an ethanol target in midbrain dopamine neurons of mice. J Neurophysiol 95: 619–626, 2006.
Overton PG, Clark D. Burst firing in midbrain dopaminergic neurons. Brain Res Brain Res Rev 25: 312–334, 1997.[CrossRef][Medline]
Paul K, Keith DJ, Johnson SW. Modulation of calcium-activated potassium small conductance (SK) current in rat dopamine neurons of the ventral tegmental area. Neurosci Lett 348: 180–184, 2003.[CrossRef][Web of Science][Medline]
Ping HX, Shepard PD. Apamin-sensitive Ca(2+)-activated K+ channels regulate pacemaker activity in nigral dopamine neurons. Neuroreport 7: 809–814, 1996.[Web of Science][Medline]
Puopolo M, Raviola E, Bean BP. Roles of subthreshold calcium current and sodium current in spontaneous firing of mouse midbrain dopamine neurons. J Neurosci 27: 645–656, 2007.
Robinson TE, Berridge KC. Incentive-sensitization and addiction. Addiction 96: 103–114, 2001.[CrossRef][Web of Science][Medline]
Satoh TO, Yamada M. A bradycardiac agent ZD7288 blocks the hyperpolarization-activated current (I(h)) in retinal rod photoreceptors. Neuropharmacology 39: 1284–1291, 2000.[CrossRef][Web of Science][Medline]
Schmidt HD, Pierce RC. Cooperative activation of D1-like and D2-like dopamine receptors in the nucleus accumbens shell is required for the reinstatement of cocaine-seeking behavior in the rat. Neuroscience 142: 451–461, 2006.[CrossRef][Web of Science][Medline]
Schultz W. Getting formal with dopamine and reward. Neuron 36: 241–263, 2002.[CrossRef][Web of Science][Medline]
Seroussi Y, Brosh I, Barkai E. Learning-induced reduction in post-burst after-hyperpolarization (AHP) is mediated by activation of PKC. Eur J Neurosci 16: 965–969, 2002.[CrossRef][Web of Science][Medline]
Seutin V, Johnson SW, North RA. Apamin increases NMDA-induced burst-firing of rat mesencephalic dopamine neurons. Brain Res 630: 341–344, 1993.[CrossRef][Web of Science][Medline]
Seutin V, Massotte L, Renette MF, Dresse A. Evidence for a modulatory role of Ih on the firing of a subgroup of midbrain dopamine neurons. Neuroreport 12: 255–258, 2001.[CrossRef][Web of Science][Medline]
Shen RY. Ethanol withdrawal reduces the number of spontaneously active ventral tegmental area dopamine neurons in conscious animals. J Pharmacol Exp Ther 307: 566–572, 2003.
Shen RY, Choong KC, Thompson AC. Long-term reduction in ventral tegmental area dopamine neuron population activity following repeated stimulant or ethanol treatment. Biol Psychiatry 61: 93–100, 2007.[CrossRef][Web of Science][Medline]
Shepard PD, Bunney BS. Effects of apamin on the discharge properties of putative dopamine-containing neurons in vitro. Brain Res 463: 380–384, 1988.[CrossRef][Web of Science][Medline]
Shepard PD, Bunney BS. Repetitive firing properties of putative dopamine-containing neurons in vitro: regulation by an apamin-sensitive Ca(2+)-activated K+ conductance. Exp Brain Res 86: 141–150, 1991.[Web of Science][Medline]
Spruston N, Jaffe DB, Williams SH, Johnston D. Voltage- and space-clamp errors associated with the measurement of electrotonically remote synaptic events. J Neurophysiol 70: 781–802, 1993.
Sun G, Tomita H, Shakkottai VG, Gargus JJ. Genomic organization and promoter analysis of human KCNN3 gene. J Hum Genet 46: 463–470, 2001.[CrossRef][Web of Science][Medline]
Tupala E, Tiihonen J. Dopamine and alcoholism: neurobiological basis of ethanol abuse. Prog Neuropsychopharmacol Biol Psychiatry 28: 1221–1247, 2004.[CrossRef][Medline]
Ungless MA, Whistler JL, Malenka RC, Bonci A. Single cocaine exposure in vivo induces long-term potentiation in dopamine neurons. Nature 411: 583–587, 2001.[CrossRef][Medline]
Vanderschuren LJ, Kalivas PW. Alterations in dopaminergic and glutamatergic transmission in the induction and expression of behavioral sensitization: a critical review of preclinical studies. Psychopharmacology 151: 99–120, 2000.[CrossRef][Medline]
Vezina P. Sensitization of midbrain dopamine neuron reactivity and the self-administration of psychomotor stimulant drugs. Neurosci Biobehav Rev 27: 827–839, 2004.[CrossRef][Web of Science][Medline]
Waroux O, Massotte L, Alleva L, Graulich A, Thomas E, Liégeois JF, Scuvée-Moreau J, Seutin V. SK channels control the firing pattern of midbrain dopaminergic neurons in vivo. Eur J Neurosci 22: 3111–3121, 2005.[CrossRef][Web of Science][Medline]
Watts AE, Williams JT, Henderson G. Baclofen inhibition of the hyperpolarization-activated cation current, Ih, in rat substantia nigra zona compacta neurons may be secondary to potassium current activation. J Neurophysiol 76: 2262–2270, 1996.
Weiss F, Markou A, Lorang MT, Koob GF. Basal extracellular dopamine levels in the nucleus accumbens are decreased during cocaine withdrawal after unlimited-access self-administration. Brain Res 593: 314–318, 1992.[CrossRef][Web of Science][Medline]
Weiss F, Parsons LH, Schulteis G, Hyytia P, Lorang MT, Bloom FE, Koob GF. Ethanol self-administration restores withdrawal-associated deficiencies in accumbal dopamine and 5-hydroxytryptamine release in dependent rats. J Neurosci 16: 3474–3485, 1996.
Wilson CJ. The mechanism of intrinsic amplification of hyperpolarizations and spontaneous bursting in striatal cholinergic interneurons. Neuron 45: 575–585, 2005.[CrossRef][Web of Science][Medline]
Wise RA, Gingras MA, Amit Z. Influence of novel and habituated testing conditions on cocaine sensitization. Eur J Pharmacol 307: 15–19, 1996.[CrossRef][Web of Science][Medline]
Wolfart J, Neuhoff H, Franz O, Roeper J. Differential expression of the small-conductance, calcium-activated potassium channel SK3 is critical for pacemaker control in dopaminergic midbrain neurons. J Neurosci 21: 3443–3456, 2001.
Zapata A, Gonzales RA, Shippenberg TS. Repeated ethanol intoxication induces behavioral sensitization in the absence of a sensitized accumbens dopamine response in C57BL/6J and DBA/2J mice. Neuropsychopharmacology 31: 396–405, 2006.[CrossRef][Web of Science][Medline]
Zhang J, Chiodo LA, Freeman AS. Influence of excitatory amino acid receptor subtypes on the electrophysiological activity of dopaminergic and nondopaminergic neurons in rat substantia nigra. J Pharmacol Exp Ther 269: 313–321, 1994.
Zhang XF, Hu XT, White FJ. Whole-cell plasticity in cocaine withdrawal: reduced sodium currents in nucleus accumbens neurons. J Neurosci 18: 488–498, 1998.
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