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1Neurobiology, Department of Neurology and 4Institutes of Anatomy and 5Physiology, University of Rostock, Rostock, Germany; 2Center for Anatomy, Charite CCM, Berlin, Germany, 3Department of Pharmacology, University of Tennessee Health Science Center, Knoxville, Tennessee; and 6Research Unit Behavioural Physiology, Research Institute for the Biology of Farm Animals, Dummerstorf, Germany
Submitted 10 April 2007; accepted in final form 26 March 2008
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ABSTRACT |
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INTRODUCTION |
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). STN neurons have two main modes of functioning: they discharge continuously and repetitively in resting animals and in bursts of high-frequency spikes prior to, during, and after movement (Wichmann et al. 1994). Hence STN is central in movement control, driving the major output structures of the basal ganglia (Hamani et al. 2004). Studies performed on parkinsonian states (Bergman et al. 1994; Hassani et al. 1996; Obeso et al. 1997) have shown an increase in neuronal activity and a change in the discharge pattern of STN neurons. The central role of altered STN activity is evidenced by an alleviation of parkinsonism through lesioning and high-frequency stimulation of the STN in patients with Parkinson's disease (PD) (Dostrovsky and Lozano 2002; Vitek 2002) and in animals with experimental parkinsonism (Benazzouz et al. 1993; Bergman et al. 1994; Chang et al. 2003). Moreover, hyperactivity of STN may contribute to the degeneration of dopaminergic neurons in PD in humans and in a variety of animal models of PD (Hassani et al. 2001).
An interplay of different network-based and intrinsic mechanisms may be involved in controlling the activity of STN neurons (Blandini et al. 2000; Wichmann and DeLong 1996). GABAergic inputs from the globus pallidus exert a major influence on the rate and pattern of STN neuron activity (Bevan et al. 2002a; Hallworth and Bevan 2005). The loss of dopamine in other brain regions, such as the cerebral cortex and thalamus, may also contribute to altered STN activity (Hassani et al. 1996; Orieux et al. 2002; Wilson et al. 2006; Zhu et al. 2002a). However, firing patterns seen in vivo are maintained in brain slices (Nakanishi et al. 1987) and persist even when fast synaptic transmission is blocked (Beurrier et al. 2000; Bevan and Wilson 1999). This autonomous activity also occurs in dissociated STN neuron cultures (Do and Bean 2003) and is caused by subthreshold sodium currents (Do and Bean 2003). Additionally, intrinsic mechanisms modulate autonomous firing, using voltage-gated calcium channels and small-conductance calcium-activated potassium (SK) channels (Hallworth et al. 2003), which shape activity by activating nonspecific cation channels (Beurrier et al. 1999; Bevan and Wilson 1999). Furthermore, N-methyl-D-aspartate (NMDA)-induced burst firing in STN neurons requires activation of either a Ca2+-activated nonselective cation channel or a Na+-Ca2+ exchanger (Zhu et al. 2004). Taken together, a modulation of intrinsic properties may be pathophysiologically significant in altered STN activity in PD as a response or in addition to abnormal synaptic transmission in the disturbed basal ganglia motor loop.
Generally, changes in potassium (K+) homeostasis result in neuronal hyperexcitability and abnormal synchronization as shown in various pathological conditions, i.e., during or even before periods of seizure activity (D'Ambrosio et al. 1999) or with hypoxic/ischemic or traumatic injury in vivo (Hansen 1985; Schröder et al. 1999). The present study addresses the influence of different [K+]o on the in vitro firing characteristics of STN neurons from nonlesioned and 6-OHDA-lesioned rats.
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METHODS |
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All experimental procedures were conducted in accordance with the regulations and licensing of the local authorities. Adult male Wistar rats (Charles River, Sulzfeld, Germany) with a body weight of 280–330 g were used and housed under a 12-h light/dark cycle with free access to water and standard diet. Stereotaxic surgery was performed under pentobarbital anesthesia (45 mg/kg body wt ip). At 15 min prior to the injection of 6-OHDA, the animals were administered desipramine (25 mg/kg body wt ip) to prevent uptake of the toxin by other catecholaminergic systems. Lesions were made in the right medial forebrain bundle (MFB) by injection of 26 µg 6-OHDA (Sigma-Aldrich, Steinheim, Germany) dissolved in 4 µl of solution comprising 1 ml saline and 1 mg ascorbic acid, which was delivered over 4 min via a 26-gauge 5-µl Hamilton syringe. The coordinates relative to the bregma were A = –2.3 mm, L = 1.5 mm, V = –8.5 mm (Paxinos and Watson 1998). Two weeks after 6-OHDA lesion, the animals were tested for apomorphine-induced rotations (0.25 mg/kg body wt in saline sc) using a self-constructed, automated rotometry device. Apomorphine-induced rotation provides a sensitive and rapid behavioral correlate of dopaminergic deafferentation of the caudate putamen (Beal 2001; Ungerstedt and Arbuthnott 1970). The rotations, measured over 40 min, were defined as complete 360° contralateral turns and reported as net differences between the two directions per minute. Only rats that exhibited at least four full turns per minute, indicating a lesion of 
95% of the dopaminergic neurons in the SNc, were selected for further use (Ungerstedt and Arbuthnott 1970). Effectiveness of 6-OHDA lesions was also tested by means of the limb-use asymmetry test (Schallert and Tillerson 1999) 2 wk after surgery. The lesioned rats showed a 100% preference of ipsilateral forepaw use.
Tyrosine hydroxylase immunohistochemistry
To demonstrate dopaminergic cell loss, immunohistochemical staining of tyrosine hydroxylase (TH) was performed. The rats were injected with an overdose of pentobarbital (60 mg/kg) and transcardially perfused with ice-cold 0.9% sodium chloride (50 ml), followed by 300 ml of 3.7% paraformaldehyde (dissolved in 0.1 M PBS, pH 7.4). The brains were immediately removed, postfixed for 4 h, and transferred into PBS (pH 7.4) containing 20% sucrose (overnight, 4°C). The cryoprotected brains were frozen in isopentane (–50°C) and stored at –80°C until further processing. The brains were cut with a cryostat (30 µm thickness). Free-floating sections were pretreated for 15 min with 3% H2O2, washed three times in PBS, blocked with 3% bovine serum albumin (BSA), normal horse serum (1:67, polyclonal, Vector Laboratories, Burlingame, CA), and 0.05% Triton-X/100 for 1 h and subsequently incubated with mouse anti-TH (1:1,000, monoclonal, Sigma) overnight at 4°C. The sections then were washed three times in PBS and incubated with biotinylated horse anti-mouse secondary antibody (1:200, Vector) overnight at 4°C. After three rinses in PBS, the sections were incubated with peroxidase-conjugated avidin-biotin complex (1:50, Vector Laboratories) for 2 h at room temperature (RT), washed three times in PBS, followed by incubation with 0.02% DAB for 8 min at RT. The mounted sections were dehydrated in graded ethanol solutions and embedded in DePeX mounting medium (Serva, Heidelberg, Germany). Dopaminergic cell and fiber loss was observed in substantia nigra and caudate putamen in the hemisphere ipsilateral to the lesion (Fig. 1).
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Simultaneous field potential recordings and measurements of [K+]o were taken in vivo by using double-barreled ion-sensitive microelectrodes based on the valinomycin exchanger Fluka 60031. The ion-selective microelectrodes had tip diameters of 2–6 µm. The reference channel was backfilled with 150 mM NaC1 and the ion-selective channel with 100 mM KCl. The electrodes showed a potential change of 55.24 ± 1.3 mV (n = 20) in response to a 10-fold increase in [K+]o in calibration solutions. Electrodes were inserted at the position of the STN (coordinates relative to bregma A = –3.6 mm, L = –2.5 mm, V = –7.6 mm) through a drill hole in the skull and dura of the anesthetized rats (9.25 mg ketamine hydrochloride (Pfizer, Germany) and 0.8 mg xylazine hydrochloride (Bayer, Leverkusen, Germany) (100 g body wt ip) in 2-mm steps until the STN was reached. The [K+]o was estimated by the Nernst equation from the potential of the ion-sensitive channel subtracted by the field potential channel, taking individual electrode slopes into account.
Experimental groups of neurons
The STN neurons were investigated in three experimental groups: neurons of nonlesioned rats ("nonlesioned"), neurons of the contralateral hemisphere of unilaterally 6-OHDA-lesioned rats ("contralateral"), and neurons of the lesioned hemisphere of unilaterally 6-OHDA-lesioned rats ("ipsilateral"). A total of 71 STN neurons in the nonlesioned group, 30 STN neurons from the contralateral group and 38 STN neurons from the ipsilateral group were analyzed.
Slice preparation
Parasagittal slices were prepared from adult male Wistar rats (280–380 g). The animals were deeply anesthetized with ether and decapitated. The brains were vertically cut at the rhinal fissure and at the inferior colliculus, the hemispheres separated, quickly removed, and transferred to cold (4–5°C), oxygenated artificial cerebrospinal fluid (ACSF). Each hemisphere was glued with cyanoacrylate to a mounting block, which was then mounted onto the stage of a Vibratome 1000 (Technical Products International, St Louis, MO). Each glued block was immersed in ACSF (4–5°C), and three parasagittal slices of 400 µm nominal thickness, containing the subthalamic nucleus (STN), were cut at 2.90, 2.50, and 2.10 mm lateral of the midline, respectively, adapted from the rat brain atlas of Paxinos and Watson (1998). The slices were transferred to a storage chamber containing ACSF bubbled constantly with 95% O2-5% CO2 and incubated for 2 h in ACSF maintained at RT (20–23°C) before commencing electrophysiological recording. The ACSF used was composed of (in mM) 124 NaCl, 5.0 KCl, 1.25 NaH2PO4, 2.0 MgSO4, 26 NaHCO3, 2.0 CaCl2, and 10 D-glucose. The concentration of potassium was altered by substituting part of the KCl with equimolar NaCl; i.e., 127.5, 125.5, or 116.5 mM NaCl was used for 1.5, 3.5, and 12.5 mM KCl-containing ACSFs, respectively. All ACSFs were continuously bubbled with 95% O2-5% CO2 and had a pH of 7.34–7.40. The osmolarity of ACSFs ranged from 290 to 300 mOsmol as measured with a cryoscopic osmometer (Osmomat 030, Gonotec, Berlin, Germany).
Electrophysiological recording
For conventional intracellular recording, microelectrodes were pulled from thin-wall borosilicate glass with an outer diameter of 1.200 mm, and an inner diameter of 0.982 mm (Hilgenberg, Mansfeld, Germany) on a vertical puller PUL-100 (WPI, Sarasota, FL), yielding a final DC resistance of 70–100 M
when filled with 3 M K+-acetate. For recording, the slices were placed on supporting nets in a submerged-type chamber and continuously superfused with oxygenated ACSF at a constant flow rate (3–5 ml/min) by a fast gravity system, resulting in a dead time to entering the bath of 30 s. The ACSFs were maintained at the constant temperature of 32.0 ± 0.5°C by a temperature control unit (TC10, npi electronic, Tamm). Using a dissection microscope for visual guidance, the STN was identified by its ovoid structure and localization as gray matter 
1.5 mm dorsal to center of the zona incerta and 2 mm posterior to the center of the substantia nigra pars reticulata (Paxinos and Watson 1998). Once the electrode reached the surface of the tissue, it was advanced in 2.0-µm steps through the brain slice with a Piezomanipulator PM500-20 (Frankenberger Biophysikalische Technik, Gilching, Germany). Neurons were impaled in the STN and current clamped. Voltage signals were amplified using a high-impedance amplifier with active bridge circuitry (npi-amplifier SEC-05LX, npi electronic), monitored on an oscilloscope (HM404-2, HAMEG Instruments, Melrose, MA) and stored digitally. In some additional experiments, tonic current was injected to compensate for the membrane potential changes induced by changes in the external K+ concentration.
Data acquisition and analysis
Data were recorded using ISO2 software (MFK, Niederegge, Germany). Signals were low-pass filtered at 2–5 kHz and digitized at 5–20 kHz according to the kinetics of the events under study [AP characteristics at 20 kHz, posttrain afterhyperpolarizations (AHP) and action potential counts at 5 kHz]. Evaluation and analysis were performed with ISO2 and Origin 7 (OriginLab, Northampton, MA). The spike threshold was defined as the point at which the first derivative of voltage with respect to time exceeded 8 V/s and corresponded to a sharp inflection in the voltage trace. The firing rate was determined in a 5-s recording window and counted manually.
Currents ranging from +0.2 to –0.3 nA (300 ms) were injected and the steady-state voltage was measured in the last 50 ms of the pulse to obtain the current-voltage (I-V) relationship. To estimate the input resistance, a linear regression was fitted to the I-V curve at its linear portion (–0.1–0.0 nA), and the slope of the regression was taken as the input resistance of the cell. The amplitude of the postspike AHP was defined as the difference in voltage between the spike threshold and the peak negativity after a spike. The amplitude of posttrain AHP was calculated from the voltage differences from the action potential threshold to the dip voltage of the posttrain AHP. The duration of posttrain AHP was estimated from the termination of the current pulse to the resurgence of the next action potential (Bevan and Wilson 1999). The width of the spikes at half-maximal amplitude (half-width) was measured at the midpoint between the threshold and peak of an action potential.
Statistical analysis
All data are presented as means ± SE. Statistical analysis was performed for between-group comparisons of more than two groups using one-way ANOVA followed by Bonferroni post hoc means comparison and for between-group comparisons of two groups using Student's t-test. For smaller experimental numbers or non-Gaussian distributions, nonparametric tests such as Kruskal-Wallis followed by Dunn post hoc means comparison for comparisons between more than two groups or the Mann-Whitney for comparisons between two groups were used. The numbers and percentage of spontaneously firing and silent STN neurons were calculated using 
2 contingency analysis. Differences were considered statistically significant if P < 0.05 unless stated otherwise.
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RESULTS |
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We studied [K+]o directly, using ion-selective electrodes in the STN of lesioned and nonlesioned rats. As expected due to the known fast diffusion of K+, baseline [K+]o in lesioned rats was 3.05 ± 0.41 mM (n = 9) and therewith not different from that in nonlesioned rats (3.51 ± 0.44 mM, n = 11; P = 0.42). However, the decay of lesion-induced [K+]o increase was remarkably slower in the STN of lesioned rats. To exclude individual differences between the rats, as well as electrode differences, time constants of the exponential decay of [K+]o in STN were normalized to the value in the cortex of the respective rat (Fig. 2A). Whereas in nonlesioned rats, the time constant ratio between the STN and cortex of 1.0 ± 0.19 (n = 11) remained the same throughout the brain (Fig. 2B), the ratio more than doubled in lesioned rats to 2.44 ± 0.42 (n = 9). This difference between the groups was significant (P < 0.002), indicating a disturbed [K+]o clearance in the STN of lesioned rats. Although the ion-sensitive electrode passed through the neocortex, hippocampus, and thalamus on its way to the STN, the latter was the only structure in lesioned rats in which the lesion-induced decay of [K+]o in STN was slowed (Fig. 2C). This suggests that, of the structures studied, the effect is limited to the STN. It should, however, be noted that other basal ganglia structures that suffer from dopamine depletion may also have been affected.
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We hypothesized that a change in the potassium gradient of the STN of lesioned rats could lead to an increase in neuronal excitability. On single-cell level, we examined the impact of a 6-OHDA lesion in the MFB and its putative relation to experimental [K+]o modulation on STN neuronal behavior by intracellular sharp microelectrodes in parasagittal slices. We recorded the membrane voltages of 38 single STN neurons from 21 rats at 24.9 (13–46) days postlesion by current clamping. We began at day 13, as early lesion effects are known to have ended by this stage (Ni et al. 2001). We compared the results with 26 neurons contralateral to the lesion from 17 rats at 28.7 (13–47) days postlesion and with 71 neurons from 34 age-matched nonlesioned rats. The neurons in the ipsilateral group were mostly from the same rats as those in the contralateral group, which is reflected by the lack of difference in the rotational score (ipsilateral: 7.55 ± 3.0, n = 21; contralateral: 8.06 ± 4.03, n = 17; P = 0.55). In the three groups we varied the [K+]o between 1.5 and 12.5 mM; for the visualization of neuronal characteristics and their main comparative analysis, we used the value at 5 mM [K+]o.
We observed a dramatic reduction in the precision of firing in the ipisilateral group and also across all groups with increasing [K+]o. In the nonlesioned group, all STN neurons appeared to have a more regular pattern (Fig. 3A) than in the ipsilateral (Fig. 3, B and C) and contralateral (not shown) groups, which fired in a regular or slightly irregular pattern. We assessed the precision of firing by the ratios of both the maximal to mean and minimal to mean spontaneous firing rates (Zhang et al. 1992). As depicted in Fig. 3D, the deviation of both ratios indicates that ipsilateral STN neurons (n = 26) fired more irregularly than neurons from the nonlesioned (P < 0.0003, minimum to mean; P < 0.022, maximum to mean, n = 29) and contralateral (P < 0.005, minimum to mean; P < 0.04, maximum to mean, n = 14) groups, whereas the ratios were comparable between the nonlesioned rats and the contralateral group (P = 0.75, P = 0.21, respectively). This irregularity culminated in pronounced burst firing in the ipsilateral group (Fig. 3C), where bursting started at 5 mM in a subgroup of neurons (8/38 = 21.1%) but did not expand any further at 12.5 mM [K+]o (4/18 = 22.2%). In the nonlesioned and contralateral group, we observed bursting solely at 12.5 mM [K+]o, in 3 of 17 (17.6%) and in 2 of 14 (14.3%) neurons, respectively. Both changes, irregularity and bursting, occurred in the absence of a change in membrane potential, i.e., a hyperpolarization as reflected in comparable membrane potentials between the three groups (P = 0.14, ipsilateral vs. nonlesioned; P = 0.12, ipsilateral vs. contralateral; P = 0.92, contralateral vs. nonlesioned; Table 2) and especially between the subgroup of bursting neurons (VM = –50.2 ± 0.72 mV) and the nonlesioned group (VM = –51.1 ± 0.5 mV, n = 71, P = 0.55). We additionally observed an increased half-width of action potential with increasing [K+]o (Table 2), but action potential changes did not vary between the three groups, ipsilateral, nonlesioned, and contralateral, in terms of amplitude (P = 0.47 and P = 0.41), threshold (P = 0.23 and 0.34) or half-width of action potential of STN neurons (Fig. 3E and Table 2).
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Although STN neurons exhibited slow spontaneous activity in all three groups, the number of spontaneously firing neurons was generally higher in the ipsilateral than in the other two groups. Comparing activity at different [K+]o, spontaneous firing occurred more frequently in the ipsilateral group than in the nonlesioned (P < 0.01) and the contralateral (P < 0.05) groups at 3.5 mM [K+]o and 5 mM [K+]o (P < 0.05) but was equal at 1.5 [K+]o and 12.5 mM [K+]o (Table 1). The number of spontaneously firing neurons grew steadily with increasing [K+]o in the nonlesioned and the contralateral groups. In the ipsilateral group, that number leapt abruptly from 75% at 1.5 mM [K+]o to 100% at 3.5 mM [K+]o (Table 1). The ipsilateral STN neurons spontaneously fired 9.7 ± 0.76 action potentials/s (n = 38), compared with nonlesioned (7.3 ± 0.74 action potentials/s, n = 60, P < 0.025) and contralateral (7.4 ± 0.65 action potentials/s, n = 26, P < 0.025) STN neurons at 5 mM [K+]o (Fig. 4, A and B). Increasing [K+]o caused more frequent spontaneous firing in STN neurons regardless of the group (Fig. 4, D and E). In terms of the differences between the groups, the spontaneous firing rate in the ipsilateral group was higher than in the nonlesioned and contralateral groups at and above 3.5 mM [K+]o (P 
0.05). In slices from nonlesioned rats, the rate did not differ from that in the contralateral group at any level of [K+]o (P = 0.24 to 0.9583). The firing elicited by DC injection also differed between the three groups (Fig. 4C). We used the slope of the relationship between injected current and firing rate (I-f curve) for a quantitative comparison of the inter- and intragroup differences at the various [K+]o. As described by Bevan and Wilson (1999), the firing frequency of most STN neurons could be divided into two ranges, which have been expressed as 2 slopes (Fig. 4, C and F). Both slopes were steeper in STN neurons ipsilateral to the lesion than in the contralateral group and the nonlesioned group (Table 2, Fig. 5). However, for 20% of STN neurons (8/36 in the ipsilateral, 5/20 in the contralateral, 8/38 in the nonlesioned group), only one linear regression (single range) was exhibited in the I-f curve, which may point to a preexistent excitatory state and therefore indicate that only the second range of firing is relevant. There was no intergroup (nonlesioned, contralateral, and ipsilateral) variation in the slopes of single-linear regression neurons, possibly due to the relatively small number of neurons (Table 2, Fig. 5, P > 0.05).
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The variations in firing following 6-OHDA lesion suggest lasting changes to the intrinsic neuronal properties of STN neurons following dopaminergic denervation. This is more probable than alterations in the basal ganglia circuitry because, in our parasagittal-slice preparations, physiological circuitry is likely to be markedly reduced as a result of the slicing technique.
Increased excitability is accompanied by reduced postspike and posttrain AHPs in ipsilateral STN neurons following 6-OHDA-lesion
POSTSPIKE AHPS. By searching for an underlying cause of the increased excitability in the ipsilateral STN, we studied postspike AHPs because they prevent further phasic firing in STN neurons by following each action potential. We used two different conditions for this: first, untriggered spontaneous and action potentials elicited by long current injections and, second, triggered action potentials.
The amplitudes of untriggered postspike AHPs were decreased in the ipsilateral group and with increasing [K+]o in each group (Fig. 6B) with and without DC injections. In both untriggred conditions, we found a striking difference between the groups for amplitudes of postspike AHPs (Fig. 6A). In the ipsilateral group at 5 mM [K+]o (Fig. 6A), the amplitude of postspike AHP following spontaneous action potentials (–7.1 ± 0.24 mV, n = 37) was smaller than in the nonlesioned (–8.0 ± 0.17 mV, n = 33, P < 0.0006) and contralateral (–8.9 ± 0.25 mV, n = 24, P < 0.004) groups. This was the case for action potentials elicited during 500-ms current injections for the whole current range 0.3 nA. Postspike AHPs following both spontaneous action potentials and action potentials elicited by current injections at 0.3 nA were comparable in the nonlesioned and the contralateral group (Supplemental Table S11 ). Although the effect of [K+]o on action potentials elicited by long current injections was qualitatively similar within each group, [K+]o elevations showed quantitative effects. Thus the amplitude of postspike AHPs in the ipsilateral group was lower than in the nonlesioned rats or contralateral group at 3.5 mM [K+]o. Moreover, the difference was less pronounced at 12.5 mM and absent at 1.5 mM over the whole range of injected current pulses with or without constant DC injection (Supplemental Table S1).
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). The amplitudes of postspike AHPs were strongly sensitive to voltage in all three groups (Fig. 6A, bottom). The average AHP amplitude at –50 mV and 5 mM [K+]o for instance, was –7.0 ± 0.5 mV (n = 6) in the ipsilateral group compared with –8.1 ± 0.4 mV (n = 7) and –8.1 ± 0.4 mV (n = 5) in the nonlesioned and contralateral groups, respectively. This confirms our results from AHP measurements of spontaneously firing neurons. However, postspike AHPs nearly disappeared at potentials below the hypothetical K+-reversal potential of –75 mV, which suggests K+ currents are the determining factor in these AHPs. There was no difference in the voltage necessary to obtain the half-maximal AHP amplitude between the groups (V1/2 = –53.5 ± 2.5 mV, n = 5 for ipsilateral neurons, V1/2 = –54.2 ± 1.6 mV, n = 7 for contralateral neurons, and V1/2 = –55.7 ± 1.5 mV, n = 8 for neurons from nonlesioned rats, P = 0.77). Additionally, the voltage sensitivity itself, represented here as the slope of the curve (k), was comparable among the three groups (k = 2.4 ± 0.9 mV, n = 5 for ipsilateral neurons, k = 5.6 ± 0.8 mV, n = 7 for contralateral neurons, and k = 5.8 ± 0.8 mV, n = 8 for neurons from nonlesioned rats, P = 0.71). Changing [K+]o resulted in shifts in V1/2 from –60.7 ± 0.4 mV to –47.9 ± 5.2 mV, n = 5, P < 0.05 in the ipsilateral group and from –60.3 ± 1.0 to –54.3 ± 1.1 mV, n = 8, P < 0.05 in the nonlesioned group. V1/2 was comparable (–59.9 ± 1.9 mV and –52.4 ± 2.5 mV, n = 5, P = 0.08) in the contralateral group, determined at 1.5 and 12.5 mM [K+]o, respectively. The slope of the voltage-sensitivity curve remained comparable (5.2 ± 1.1 and 5.2 ± 1.8, n = 5, P = 1.0 in the ipsilateral group, 4.7 ± 1.2 and 6.6 ± 1.6, n = 5, P = 0.63 in the contralateral group; 3.8 ± 0.5 and 5.4 ± 0.9, n = 8, P = 0.14 in the nonlesioned group, determined at 1.5 and 12.5 mM [K+]o, respectively; Fig. 6C).
POSTTRAIN AHPS.
We additionally evoked posttrain AHPs by high-frequency action potential firing induced by 1,000-ms-long depolarizing current pulses in the range of 0.02 to 0.3 nA (Fig. 7, A and C). Both increasing the current used to drive the neuron and the frequency during the driving period led to an accumulation in the depth and duration of posttrain AHP in all three groups. The posttrain AHPs were markedly influenced by changes in [K+]o in all three groups (Fig. 7, A and C). Both their amplitude (Fig. 7, B, left, and D, Supplemental Table S2) and duration (Fig. 7, B, right, and E, and Supplemental Table S3) decreased with increasing [K+]o at the same pulse current injection for currents 0.1 nA. A comparison of the groups revealed attenuation in the posttrain AHP for the ipsilateral group. The amplitude of posttrain AHPs in the ipsilateral group for injected currents of 0.1 nA and at a [K+]o of 5 mM was lower than in the nonlesioned and the contralateral group regardless of DC injection (Fig. 7, A and B, left, for comparison between the groups at 5 mM [K+]o; C and D, for comparison within the groups, Supplemental Table S2). Posttrain AHPs at 5 mM were also shorter in the ipsilateral group than in the nonlesioned group and in the contralateral group at injections of 0.02, 0.2 or 0.3 nA (Fig. 7, A and B, right, and Supplemental Table S3). This was similar for 12.5 mM [K+]o with DC injection (Fig. 7, C and E, and Supplemental Table S3). The amplitude and duration of posttrain AHP in the contralateral group was comparable to the nonlesioned group.
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DISCUSSION |
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Postlesion changes in STN neurons were replicated by increasing [K+]o
Our results from the STN of nonlesioned rats resemble those of previous studies with respect to the regular or slightly irregular single-spike firing (Beurrier et al. 1999; Overton and Greenfield 1995), the percentage of spontaneously firing neurons, and the firing rates in vitro (Beurrier et al. 2001; Bevan and Wilson 1999).
Following 6-OHDA-lesion, the great majority of STN neurons ipsilateral to the lesion recorded in this study fired irregularly or in bursts as shown previously (Hassani et al. 1996; Ni et al. 2001). By using brain slices, we provide evidence that single-cell bursting is relatively independent of an intact basal ganglia circuitry as also demonstrated by Wilson et al. (2004). However, the percentage of STN neurons discharging in bursts in our in vitro study is lower than in some in vivo studies where 35–55% of STN neurons exhibit burst firing (Hollerman and Grace 1992; Ni et al. 2001), which also demonstrates that inputs to STN shape firing patterns (Baufreton et al. 2003; Bevan et al. 2002b; Hallworth and Bevan 2005). In our study, the firing rates of STN neurons ipsilateral to the lesion were tonically elevated under basal conditions in accordance with predictions based on prior studies conducted on basal ganglia organization (DeLong 1990) and the majority of studies on dopamine depletion (Hassani et al. 1996; Kreiss et al. 1997; Perier et al. 2000). However, other slice preparations (Zhu et al. 2002a) and animal models (Wilson et al. 2006) revealed a decreased firing rate postlesion.
An experimentally increased [K+]o in nonlesioned rats and contralateral to the lesion mimicked all changes found in STN neurons of lesioned rats, namely changes in the firing rate and pattern, including burst firing, as well as a decrease in AHP and input resistance. Therefore it is tempting to conclude that the disruption of physiological firing is at least in part attributable to changes in K+ conductance. Our findings augment and strengthen previous results showing that the reduction or the absence of K+ conductance mimic the pathophysiological changes in PD: 1) spontaneous bursting was observed in regularly firing STN neurons after a blockade of SK channels (Baufreton et al. 2003; Beurrier et al. 1999, 2001; Hallworth et al. 2003) and 2) increased activity of STN neurons with a marked decline in precision was achieved by reducing K+ conductance (Hallworth et al. 2003; Zhu et al. 2002b). Moreover, dopamine effects are known to be exerted in part via changes in the K+ conductivity. While most neurons in rats show an increase in K+ currents on dopamine binding, mostly to dopamine receptor 2 (for striatum: Greif et al. 1995), some studies instead indicate a reduction in K+ currents by dopamine in neurons (Kitai and Surmeier 1993). It can, however, be stated that K+ current changes are likely following dopamine depletion.
In terms of the channels underlying these effects, members of voltage-gated Kv1 and 3 (Verma-Kurvari et al. 1997; Weiser et al. 1994), as well as SK channels (Stocker and Pedarzani 2000), have been detected on mRNA level in the STN. The existence of Kv3.1 and SK channels has been functionally confirmed (Wigmore and Lacey 2000). Outward rectifying potassium currents, in particular those underlying AHPs, limit excitability (Alger and Nicoll 1980) and contribute to the encoding function of neurons (Stocker et al. 2004). This is also true for STN neurons, where a reduction of AHPs was associated with an increase in their firing rate (Bevan and Wilson 1999) and a change in firing pattern (Baufreton et al. 2003; Beurrier et al. 1999, 2001). The fast component of the postspike AHPs is mediated by Kv3.1 tail currents (Wigmore and Lacey 2000), as also suggested by the strong voltage sensitivity of the AHP amplitude in our study. However, SK-mediated currents are more important for the precision, pace, and pattern of action potential generation in the STN (Hallworth et al. 2003). It is very likely that SK-mediated currents are also required for the oscillatory mechanism underlying regular spontaneous activity (Bevan and Wilson 1999). Furthermore, SK channels strongly curtailed the burst duration in rebound bursting STN neurons (Hallworth et al. 2003). Although posttrain AHPs have been suggested as limiting excitability in many neurons (Kim and McCormick 1998; Lorenzon and Foehring 1992; Tanabe et al. 1998), in STN neurons, they are important for the smooth and uniform resumption of spontaneous activity after high-frequency firing (Hallworth et al. 2003). The alteration of any of the mentioned currents following the lesion could explain the differences between the groups. However, the changes, i.e., the discrepancies between the ipsilateral, contralateral, and nonlesioned groups under regular [K+]o conditions are not clearly attributable to a specific K+ conductance in our experiments. Although our data point to a disturbance of multiple K+ conductance itself as underlying cause of the pathological firing properties in lesioned STN, we cannot exclude that a putative compromised [K+]o regulation in the parkinsonian state alters normal functioning.
Suggested reasons for a transient [K+]o increase in vivo
[K+]o is known to rise during periods of intense neuronal firing (Sykova 1991), but in the intact brain, adjacent astrocytes remove excess K+ sufficiently (Walz 2000). In other pathological conditions, a glial response, primarily due to reactive astrocytes, was observed (Walz and Wuttke 1999). In PD, such a response has been described for substantia nigra (Teismann et al. 2003; Wu et al. 2002). Reactive astrocytosis may also occur in remote but anatomically connected areas, as demonstrated by astrocytosis in the STN following FeCl2 injection in the striatum (Hironishi et al. 1999). The slow decay of [K+]o elevations in vivo found in the present experiments maybe the result of such an astrocytic dysfunction. In our study, [K+]o differentially influenced the firing rates and after-spike AHP of STN neurons of nonlesioned and 6-OHDA-lesioned rats. The relative similarity of firing in STN neurons from lesioned and nonlesioned tissue at low [K+]o where astrocytic clearance is not necessary was contrasted by an unduly increased activity at physiological and elevated [K+]o in lesioned rats. These results point to an impaired K+ clearance in the lesioned rats. This notion is supported by our finding of an increased time constant of K+ clearance in lesioned rats. However, transiently increased [K+]o in vivo might also be a consequence of increased STN neuronal activity in the parkinsonian state even if astrocyte function is unimpaired.
Possible mechanisms of [K+]o effects on firing properties in STN neurons
An increase of [K+]o reduces the driving force of any K+ current present in the membrane by shifting the reversal potential in the positive direction. Alterations in [K+]o also directly influence the membrane resting potential and tilt the balance of key elements for oscillatory bursting such as T-type channels, h-channels and Ca-dependent K+ channels (Beurrier et al. 1999; Bevan and Wilson 1999). The consequence might be the generation of rhythmic bursting by the STN neurons (Beurrier et al. 1999; Hallworth et al. 2003), which was not observed in nonlesioned tissue under physiological [K+]o conditions (Magill et al. 2000). The fact that bursting was observed in the absence of either a hyperpolarization or an NMDA increase in both "lesioned" and elevated [K+]o suggests that the persistent Na+ current (Beurrier et al. 2000; Bevan and Wilson 1999) or a change in Ca2+-channel inactivation (Otsuka et al. 2001) is a burst-driving mechanism. A disturbed balance in intrinsic conductance induced by elevated [K+]o may result in a transition to a burst state without an excitatory (NMDA) (Zhu et al. 2004) or inhibitory hyperpolarizing (GABA) (Bevan et al. 2002a; see also Urbain et al. 2002) drive. The role of a [K+]o elevation—even if transient—might be more complex in vivo because high [K+]o elevates extracellular glutamate (Fujikawa et al. 1996) and reduces extracellular GABA (Engblom et al. 2003), thereby influencing pathways believed to be involved in STN hyperactivity. 6-OHDA lesioning directly reduces monoaminergic inputs to the STN (Francois et al. 2000; Hassani et al. 1997). Even the remaining dopaminergic tone (Delfs et al. 1995) could not exert its full effect as it acts partially via K+ channels controlled by D2 receptors (Zhu et al. 2002b). Finally, [K+]o affects the postsynaptic STN neuron's level of polarization, which in turn is critical in determining response to synaptic inputs (Bevan et al. 2002a). However, the slices used in this study were devoid of any structured input from other STN afferents.
In conclusion, we show a regulation of the firing behavior of STN neurons by [K+]o and that this could be another factor controlling the pathological discharge pattern in PD. Such a control could be mediated by disturbed K conductance i.e., K+ channel impairment or possibly the latter in combination with a temporal increase in [K+]o. Our data cannot be used to conclusively prove either of these hypotheses. The effect of [K+]o variation on STN neurons is of particular interest with regard to the role played by the STN in the basal ganglia, particularly because the GP-STN pathway is a key component of the circuitry involved in the symptoms of PD (DeLong 1990). There is increasing evidence that an altered firing pattern in this circuitry, especially burst firing, may correlate better with parkinsonian symptoms than an enhanced firing rate alone (Bergman et al. 1994; Hollerman and Grace 1992; Levy et al. 2002). The impact of [K+]o variations on increased correlated rhythmic bursting and increases in mean firing frequency of STN activity in PD has been neglected to date (for review, see Bevan et al. 2002b). The results of this study provide the foundation for future analysis of neuronal and glial K+ channels and of glial function in the regulation of STN firing. Further analysis of the ion channels underlying excitability changes and their anatomical location will be critical for our understanding of basal ganglia function and dysfunction. Such research may lead to the development of pharmacological or molecular tools which alter STN firing and serve as a nonsurgical treatment for PD.
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ACKNOWLEDGMENTS |
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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.
1 The online version of this article contains supplemental data.
Address for reprint requests and other correspondence: U. Strauss, Institute for Cell Biology and Neurobiology, Center for Anatomy, Charite CCM, Philippstr. 12, D-10115 Berlin, Germany (E-mail: ulf.strauss{at}charite.de)
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curves) in the 3 groups. Note that the 6-OHDA lesion in the medial forebrain bundle (MFB) was associated with a shift in the I-f curve and an increase in 1st and 2nd gain in STN neurons. All comparative recordings depicted here were obtained under superfusion with artificial cerebrospinal fluid (ACSF) containing 5 mM [K+]o. The color code and symbols are explained for A–C below the figures. For numbers of STN neurons in B, see RESULTS and 
), contralateral to a 6-OHDA lesion (
) and for neurons of nonlesioned rats (
). DC: constant direct-current. Note that DC injection did not change the impact on the input resistance, indicating that a change in membrane potential alone did not account for the effect. *P < 0.05; numbers of STN neurons displayed above columns.