Neurotensin modulation of acetylcholine, GABA, and aspartate release from rat prefrontal cortex studied in vivo with microdialysis

Brain Research Bulletin 77 (2008) 129–135

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Neurotensin modulation of acetylcholine, GABA, and aspartate release from rat prefrontal cortex studied in vivo with microdialysis Polina Petkova-Kirova a,1 , Angelina Rakovska b,∗ , Laura Della Corte c,2 , Galina Zaekova b,3 , Radomir Radomirov b,4 , Aliz Mayer d,5 a

Institute of Biophysics, Bulgarian Academy of Sciences, Acad. G. Bonchev Street, bl. 21, 1113 Sofia, Bulgaria Institute of Neurobiology, Bulgarian Academy of Sciences, Acad. G. Bonchev Street, bl. 23, 1113 Sofia, Bulgaria Department of Preclinical and Clinical Pharmacology, University of Florence, Viale Pieraccini 6, 50139 Florence, Italy d Institute of Experimental Medicine, Hungarian Academy of Sciences, H-1450 Budapest, P.O. Box 67, Hungary b c

a r t i c l e

i n f o

Article history: Received 9 April 2008 Accepted 14 April 2008 Available online 7 May 2008 Keywords: Prefrontal cortex Neurotensin Amino acids Acetylcholine Microdialysis

a b s t r a c t The effects of the peptide transmitter neurotensin (NT) on the release of acetylcholine (ACh), ␥aminobutyric acid (GABA), glutamate (Glu), aspartate (Asp), and taurine from the prefrontal cortex (PFC) of freely moving rats were studied by transversal microdialysis. Neurotensin (0.2 and 1 ␮M) administered locally in the PFC produced a concentration-dependent increase in the extracellular levels of ACh, GABA, and Asp, but not of Glu or taurine. The increase produced by 1 ␮M NT reached a maximum of about 240% for ACh, 370% for GABA, and 380% for Asp. Lower doses of NT (0.05 ␮M) did not cause a significant change in ACh, GABA, or Asp output in the PFC. Higher concentrations of NT (2 ␮M) did not induce further increases in the level of neurotransmitters. A high-affinity selective neurotensin receptor (NTR1) antagonist SR 48692 (0.5 ␮M) perfused locally blocked neurotensin (1 ␮M)-evoked ACh, GABA, and Asp release. Local infusion of the sodium channel blocker tetrodotoxin (TTX) (1 ␮M) decreased the release of ACh, had no significant effect on GABA or Asp release, and prevented the 1 ␮M neurotensin-induced increase in ACh, GABA, and Asp output. Removal of calcium from the Ringer’s solution prevented the peptide from having any effects on the neurotransmitters. Thus, in vivo NT plays a modulatory role in the PFC by interacting with cortical neurons releasing GABA and Asp and with ACh-containing neurons projecting to the PFC. The NT effects are of neural origin, as they are TTX-sensitive, and mediated by the NTR1 receptor, as they are antagonized by SR 48692. © 2008 Elsevier Inc. All rights reserved.

1. Introduction Neurotensin (NT) is an endogenous tridecapeptide [10] known to play a role as a neurotransmitter and neuromodulator both in the central and peripheral nervous systems [38,17,27,7]. The peptide and its receptors are widely distributed throughout the brain, including the basal forebrain and cerebral cortex [36,21,7,65]. In

∗ Corresponding author at: Institute of Neurobiology, Lab. ‘Neuropeptides’, Bulgarian Academy of Sciences, Acad. G. Bonchev Street, bl. 23, 1113 Sofia, Bulgaria. Tel.: +359 2 9792305; fax: +359 2 8719109. E-mail addresses: [email protected] (P. Petkova-Kirova), [email protected] (A. Rakovska), laura.dellacorte@unifi.it (L. Della Corte), ashlie @mail.bg (G. Zaekova), [email protected] (R. Radomirov), [email protected] (A. Mayer). 1 Tel.: +359 2 9792130; fax: +359 2 9712493. 2 Tel.: +39 055 4271226; fax: +39 055 410778. 3 Tel.: +359 2 9792387; fax: +359 2 8719109. 4 Tel.: +359 2 9792164; fax: +359 2 8719109. 5 Tel.: +36 1 2109979; fax: +36 1 2109423. 0361-9230/$ – see front matter © 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.brainresbull.2008.04.003

the prefrontal cortex (PFC), the basal concentration of neurotensin measured in awake, freely moving rats is found to be 2–5 pg/sample [45]. There are no neurotensin-containing cell bodies in the PFC, and the only source of NT arises from axons derived from the ventral tegmental area (VTA) [25,56,60] that contain tyrosine hydroxylase and the dopamine synthetic enzyme [60,15]. In vivo release of coexistent NT and dopamine was measured from the prefrontal cortex after electrical stimulation of mesocortical axons [5]. Thus, neurotensin in the PFC is exclusively localized to dopamine axons. Dopamine synapses are found on PFC pyramidal cells as well as interneurons [58], making neurotensin a likely regulator of the cognitive functions of the prefrontal cortex via an effect on glutamatergic and GABAergic function. Cholinergic neurons of the basal forebrain constitute the primary source of ACh to the cerebral cortex [50,18,33] and are thought to play a major role in mediating cortical activation and plasticity. Autoradiographic studies combined with acetylcholinesterase histochemistry have shown an association of NT receptors with cholinergic neurons in the rat basal forebrain [62]. Furthermore,

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electrophysiological data have demonstrated that NT not only binds to, but is also internalized in cholinergic cells of the basal forebrain and directly affects multiple electrophysiological features of these neurons [2]. In vitro experiments indicate that NT enhanced spontaneous, electrically stimulated, potassium-evoked ACh release in the striatum [29,64] and potassium-evoked ACh release in the parietal cortex [30]. In frontal cortical slices, NT significantly inhibited evoked ACh release via a tetrodotoxin (TTX)-insensitive mechanism and had no effect on spontaneous release [30]. In our previous studies using in vivo microdialysis, we found that neurotensin increases spontaneous ACh release from the rat hippocampus [46]. Such a stimulant effect of neurotensin was not found to occur for in vivo striatal ACh levels [40,59], although it was established that NT regulates acetylcholine release when D2 -dopaminergic inhibitory input is suppressed [59]. So far, the role of neurotensin in the regulation of cholinergic neuronal activity in vivo in the cortex remains unclear. The present study sought to characterize the neuromodulatory role of neurotensin in the prefrontal cortex. We examined the effects of neurotensin on PFC Glu, Asp, GABA, and ACh release in vivo in awake and freely moving rats using transversal microdialysis, i.e. we measured their concentrations in the extracellular space [61] where they exert their actions [69] on nonsynaptic receptors [67,68,48,55,35,34] and extrasynaptic transporters involved in their uptake [61,28]. 2. Materials and methods 2.1. Animals and surgery Male adult Wistar rats weighing 250–300 g were used. Prior to surgery, they were housed in groups in macrolon cages with ad libitum access to food and water. Rats were housed on a 12 h light/12 h dark cycle. On the day of surgery, the rats were anaesthetized with chloral hydrate (400 mg/kg, i.p.) and placed in a stereotaxic apparatus (Stoeling, Stellar, Wood Dale, Il, USA). Microdialysis tubes (AN 69 membrane; Dasco, Bologna, Italy, 220 ␮m i.d. and 310 ␮m o.d., molecular weight cut-off 15,000 Da) were inserted transversally into the prefrontal cortex following the procedure described by Giovannini et al. [19]. The microdialysis tubes were covered with super-epoxy glue along their entire length except for the region corresponding to the cortex (8 mm) (see scheme). The coordinates used for the implantation of the microdialysis tubing in the cortex were as follows: anterior–posterior +1.0 mm and −1.8 mm, according to Paxinos and Watson [41]. All coordinates refer to Bregma, with bregma and lambda on a horizontal plane. The day after surgery, each rat was placed in a Plexiglas cage with free access to food and water. All animal manipulations were carried out according to the European Community guidelines for animal care (DL 116/92, application of the European Community Council Directive 86/609/EEC). All efforts were made to minimize animal suffering and to use only the number of animals necessary to produce reliable scientific data. Procedures involving animals and their care were conducted in conformity with the institutional guidelines that are in compliance with national and international laws and policies (EEC Council Directive 86/609, OJ L 358, 1 Dec. 12. 1987; NIH Guide for the Care and Use of Laboratory animals, NIH publication No. 85-23, 1985).

Mod. CMA/100, Sweden), and the PFC was perfused with Ringer’s solution (NaCl 147 mM, CaCl2 1.2 mM, KCl 4.0 mM) containing 0.1 ␮M physostigmine sulfate at a constant flow rate of 2 ␮l/min. After a 1-h recovery period, during which the PFC was perfused without collecting the dialysate, samples were collected at 20 min intervals. After collecting the first three samples to measure the basal outflow, drugs dissolved in Ringer’s solution were administered locally through the dialysis membrane for 80 min (4th–7th samples) or until the end of the experiment (TTX or TTX and NT). Thereafter, the drugs were withdrawn and the membranes were perfused with normal Ringer’s solution until the end of experiment. The content of ACh, GABA, Glu, Asp, and taurine in the dialysate was analyzed by HPLC as described below. In order to assay all neurotransmitters simultaneously when necessary, samples from the same animal were split into two halves. 2.3. Histological control At the end of the experiments, rats were anaesthetized with urethane (1.2 ␮g/kg, i.p.) and sacrificed by decapitation. The brain was rapidly removed and placed in a vial containing 9% formaldehyde solution in phosphate buffer. Two or three days later, the brains were frozen with liquid CO2 and coronal slices (50 ␮m) cut using a freezing microtome. The slices were examined by light microscopy (Nikon, Labophot-2) to verify the position of the dialysis membrane. Data obtained from rats in which the dialysis membrane was positioned outside the defined structure were discarded (<5%). The condition of the cortical neurons around the probe was checked at the end of the experiment by staining coronal sections of formaldehyde-fixed brains with Richardson’s solution (modified Nisal’s staining). 2.4. Assay of ACh in the dialysate ACh was directly assayed in the dialysate using an HPLC method with a postcolumn enzyme reactor and an electrochemical detector as previously described [12,19]. ACh was separated on a cation-exchange column, prepared by loading a reverse-phase column (Chromspher 5 C18, Chrompack, Middelburg, Netherlands) with sodium lauryl sulfate (0.5 mg/ml). The mobile phase consisted of 0.2 M phosphate buffer (pH 8.0) containing 5 mM KCl, 1 mM tetramethylammonium (TMA), and 0.3 mM EDTA. The flow rate was 0.7 ml/min. ACh was hydrolyzed by acetylcholine esterase (AChE) to acetate and choline in a post-column enzyme reactor; choline was oxidized by choline oxidase to produce betaine and hydrogen peroxide. Hydrogen peroxide was electrochemically detected by a platinum-working electrode at +500 mV with Ag/AgCl reference electrode. For the quantitative analysis of ACh, we constructed a calibration curve by spiking the Ringer’s solution with standard ACh in the concentration range we expected to find in the dialysates. Three or four concentrations for each calibration curve were then injected at the beginning and end of the analysis, and the heights of the recorded peaks were then plotted against the concentrations. A regression line was calculated, and quantification of unknown samples was carried out by the method of inverse prediction. Under these experimental conditions, the sensitivity limit for ACh was 100 fmol (s/n ratio > 3/1). 2.5. Assay of excitatory amino acids GABA, Glu, Asp, and taurine in the dialysate GABA, Glu, Asp, and taurine analyses were carried out as previously described [6]. We performed HPLC with fluorimetric detection at the excitation and emission wavelengths of 340 and 455 nm, respectively, after derivatization of the amino acids with o-phthalaldehyde (OPA). A 5-␮m nucleosil C18 column (200 mm × 4 mm i.d., Macherey-Nagel, Duren, Germany) was used. The mobile phase consisted of methanol–potassium acetate (0.1 M) adjusted to pH 5.52 with glacial acetic acid. A gradient (flow rate 0.9 ml/min) of three linear steps – from 25 to 43% methanol (1 min), 43 to 70% methanol (10 min), and 70 to 90% (1 min) – followed by an isocratic hold at 90% methanol (1 min) was used. One volume (10 ␮l) of the dialysate was mixed with 1 ␮l of the OPA derivatization reagent [6] in a glass capillary tube and injected after 1.5 min. Standard curves were linear over the concentration range of 25–1000 fmol/␮l. The minimum detectable concentration of the neurotransmitters was 2 fmol/␮l. 2.6. Chemicals All reagents were of analytical grade. The following chemicals, purchased from Sigma Chemical Co. (St. Louis, MO, USA), were used: physostigmine, OPA, homoserine, neurotensin, tetrodotoxin (TTX), acetylcholine esterase, AChE (EC 3.1.1.7., Grade VI-S), choline oxidase (EC 1.1.3.17), and p-formaldehyde. The neurotensin antagonist 2-(1-(7-chloro-4-quinolinyl)-5-(2,6-dimethoxyphenyl)pyrazol3yl)carbonylamino tricyclo (3.3.1.1.3.7 )decan-2-carboxylic acid (SR 48692) was kindly supplied by Sanofi Recherche, France.

Schematic diagram showing the positioning of the microdialysis membrane in the prefrontal cortex. For stereotaxic coordinates, see Section 2. 2.2. Microdialysis procedure One day after surgery, each rat was placed in a Plexiglas cage. The inlet of the microdialysis probe was connected to a microperfusion pump (Carnegie Medicine,

2.7. Statistical analysis ACh, GABA, Asp, and Glu release rates are expressed as pmol/40 ␮l or presented as the percent variation over the basal output (average of the first three samples taken before drug administration). All values are given as the mean ± S.E.M. Differences among experimental groups are evaluated by comparing areas under

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Fig. 1. (A) Effects of 1 ␮M neurotensin (NT) on ACh release from the prefrontal cortex (PFC) in freely moving rats in the absence and presence of TTX and 2-(1-(7-chloro4-quinolinyl)-5-(2,6-dimethoxyphenyl)pyrazol-3yl)carbonylamino tricyclo (3.3.1.1.3.7 )decan-2-carboxylic acid (SR 48692). NT and SR 48692 were administered for 80 min locally to the PFC via the dialysis membrane after collection of three samples (horizontal bar). TTX was introduced into the perfusion medium after collection of the three samples and maintained in the medium until the end of the experiment. ACh output is expressed as the percentage change over the mean of the first three control samples. The means ± S.E.M. of at least six different rats in each group are presented. Dialysate samples were collected for 20 min (2 ␮l/min). (B) Effects of 1 ␮M NT on ACh release from the PFC in freely moving rats in the absence and presence of TTX and 2-(1-(7-chloro-4-quinolinyl)-5-(2,6-dimethoxyphenyl)pyrazol-3yl)carbonylamino tricyclo (3.3.1.1.3.7 )decan2-carboxylic acid (SR 48692). Values are expressed as the mean percentage change calculated for the 80–200 min period. Significant differences among the four experimental groups were evaluated by comparing the area under curve (AUC) via one-way analysis of variance (ANOVA) (F3,14 = 13.93; P < 0.0002) followed by a Newman–Keuls multiple comparison test (# P < 0.05 NT vs. basal release; ***P < 0.001 NT + TTX vs. NT; **P < 0.01 NT + SR 48692 vs. NT).

the curves (AUC) via a one-way analysis of variance (ANOVA) followed by the Newman–Keuls multiple comparison test. Differences were considered significant at P < 0.05.

hemisphere by running under the frontal and through prefrontal cortices. These areas are involved in sensorimotor and cognitive functions.

3. Results

3.2. Effects of neurotensin, SR 48692, and tetrodotoxin on the basal release of ACh, GABA, and Asp

3.1. Neuroanatomical localization of microdialysis probes According to the maps of Zilles and Wree [71], the transverse microdialysis membrane in our experiments was inserted through the parietal region of one hemisphere and reached the opposite

After a 60 min recovery period, ACh, GABA, Glu, and Asp levels were measured in the microdialysis perfusates collected from the PFC of control rats not treated with any chemicals. These levels remained relatively constant from one collection period to the

Fig. 2. (A) Effects of 1 ␮M neurotensin (NT) on GABA release from the PFC in freely moving rats in the absence and presence of TTX and 2-(1-(7-chloro-4-quinolinyl)-5-(2,6dimethoxyphenyl)pyrazol-3yl)carbonylamino tricyclo (3.3.1.1.3.7 )decan-2-carboxylic acid (SR 48692). NT and SR 48692 were administered for 80 min locally to the PFC via the dialysis membrane after collection of three samples (horizontal bar). TTX was introduced into the perfusion medium after collection of the three samples and maintained in the medium until the end of the experiment. GABA output is expressed as the percentage change over the mean of the first three control samples. The means ± S.E.M. of at least six different rats in each group are presented. Dialysate samples were collected for 20 min (2 ␮l/min). (B) Effects of 1 ␮M NT on GABA release from the PFC in freely moving rats in the absence and presence of TTX and 2-(1-(7-chloro-4-quinolinyl)-5-(2,6-dimethoxyphenyl)pyrazol-3yl)carbonylamino tricyclo (3.3.1.1.3.7 )decan-2-carboxylic acid (SR 48692). Values are expressed as the mean percentage change calculated for the 80–200 min period. Significant differences among the four experimental groups were evaluated by comparing the area under curve (AUC) via one-way ANOVA (F3,11 = 30.3; P < 0.0001) followed by a Newman–Keuls multiple comparison test (### P < 0.001 NT vs. basal release; ***P < 0.001 NT + TTX vs. NT; ***P < 0.001 NT + SR 48692 vs. NT).

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Fig. 3. (A) Effects of 1 ␮M neurotensin (NT) on aspartate (Asp) release from the PFC in freely moving rats in the absence and presence of TTX and 2-(1-(7-chloro-4-quinolinyl)-5(2,6-dimethoxyphenyl)pyrazol-3yl)carbonylamino tricyclo (3.3.1.1.3.7 )decan-2-carboxylic acid (SR 48692). NT and SR 48692 were administered for 80 min locally to the PFC via the dialysis membrane after collection of three samples (horizontal bar). TTX was introduced into the perfusion medium after collection of the three samples and maintained in the medium until the end of the experiment. Asp output is expressed as the percentage change over the mean of the first three control samples. The means ± S.E.M. of at least six different rats in each group are presented. Dialysate samples were collected for 20 min (2 ␮l/min). (B) Effects of 1 ␮М NT on aspartate (Asp) release from the PFC in freely moving rats in the absence and presence of TTX and 2-(1-(7-chloro-4-quinolinyl)-5-(2,6-dimethoxyphenyl)pyrazol-3yl)carbonylamino tricyclo (3.3.1.1.3.7 )decan-2-carboxylic acid (SR 48692). Values are expressed as the mean percentage change calculated for 80–220 min. Significant differences among the four experimental groups were evaluated by comparing the area under curve (AUC) via one-way ANOVA (F3,16 = 106.4; P < 0.0001) followed by a Newman–Keuls multiple comparison test (### P < 0.001 NT vs. basal release; ***P < 0.001 NT + TTX vs. NT; ***P < 0.001 NT + SR 48692 vs. NT).

next throughout each experiment (for up to 5 h; range of variation ±10–15% n.s.; Figs. 1A, 2A, 3A, and 4). Table 1 shows the extracellular ACh, GABA, Glu, and Asp levels found in the microdialysis perfusates in control rats. Neurotensin applied for 80 min locally in the PFC induced an increase in extracellular ACh, GABA, and Asp levels (Figs. 1A, 2A, and 3A). Two concentrations of NT (0.2 and 1 ␮M) were used to study neurotensin’s effects on ACh, GABA, and Asp release. The stimulant effects on the three neurotransmitters were concentration-dependent and reached a maximum at 1 ␮M neurotensin. The peak values of the effects are shown in Table 2. At a concentration of 1 ␮M, the stimulant effects of NT on the release of ACh, GABA, and Asp began roughly 15–20 min after the administration of the peptide. They lasted for 40 min (ACh and GABA) or 140 min (Asp) and then returned to basal levels (Figs. 1A, 2A, and 3A). A second application of 1 ␮M NT given 180 min after the first application failed to increase ACh, GABA, or Asp release, suggesting that receptor desensitization occurs. A lower dose of NT (0.1 ␮M) did not cause a change in ACh, GABA, or Asp output in the PFC. Higher concentrations of NT (2 ␮M) did not induce further increase in any of the neurotransmitters measured. NT had no effect on Glu (Fig. 4) or taurine outflow. After perfusion of neurotensin, changes in the behavior of the rats studied during the experiments were not observed. Removal of calcium from Ringer’s solution for a period of 120 min before 1 ␮M NT administration reduced baseline levels of the neurotransmitters and prevented the peptide from having any significant effects on the levels of the studied neurotransmitters.

Baseline levels of the studied neurotransmitters returned to normal after Ringer’s solution containing calcium was reinfused. Experiments with tetrodotoxin (TTX), a sodium channel blocker, were also performed to determine the contribution of neuronal depolarization to the changes in extracellular levels of the studied neurotransmitters after introduction of NT. After collection of the three basal samples, 1 ␮M TTX (dissolved in Ringer’s solution) was added locally in the PFC via a dialysis probe and maintained until the end of the experiment. Behavioral activation of the rats was not observed during this in vivo perfusion of TTX. At a concentration of 1 ␮M, TTX caused a significant decrease in basal ACh output in the PFC to a minimum of 55 ± 4% (n = 3) and had no significant effect on basal GABA or Asp levels (data not shown). The significant decrease in basal ACh release in the presence of TTX reveals the

Table 1 Basal release of acetylcholine (ACh), GABA, glutamate (Glu), and aspartate (Asp) measured by transversal microdialysis from the PFC of freely moving rats Neurotransmitters

Basal concentration (pmol/40 ␮l)

ACh GABA Glu Asp

12.39 0.40 65.50 7.60

± ± ± ±

1.26 0.08 12.60 1.46

Number of animals (n) 38 20 20 20

Dialysate outputs are expressed as the mean ± S.E.M. of the first three basal samples.

Fig. 4. Effect of 1 ␮M neurotensin (NT) on glutamate (Glu) release from the PFC in freely moving rats. NT was administered locally for 80 min to PFC via the dialysis membrane after collection of three samples (horizontal bar and arrow). Glu output is expressed as the percentage change over the mean of the first three control samples. The means ± S.E.M. of six different rats are presented. Dialysate samples were collected for 20 min (2 ␮l/min).

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Table 2 Peak values of the effects of neurotensin on the extracellular levels of ACh, GABA, and Asp measured with transversal microdialysis from the PFC of freely moving rats Parameters

1. Basal release (control) 2. Neurotensin 1 ␮M 3. Neurotensin 0.2 ␮M

Neurotransmitter release (% of basal) ACh

GABA

Asp

98 ± 3.80 (n = 9) 240.56 ± 48.50 (n = 9), P < 0.001, 2 vs. 1 160.81 ± 10.12 (n = 6), P < 0.001, 3 vs. 1

110 ± 4.20 (n = 6) 370.00 ± 28.70 (n = 6), P < 0.001, 2 vs. 1 175.20 ± 12.80 (n = 6), P < 0.001, 3 vs. 1

112 ± 3.40 (n = 6) 382.00 ± 70.93 (n = 6), P < 0.001, 2 vs. 1 189.01 ± 18.12 (n = 6), P < 0.001, 3 vs. 1

Neurotransmitter output is expressed as the percentage change over the mean of the first three control samples.

endogenous tonic excitatory control of the cholinergic system in the PFC. Co-administration of TTX (1 ␮M) and NT (1 ␮M) prevented the stimulant effect of neurotensin on the release of the three neurotransmitters (Figs. 1A, B; 2A, B, and 3A, B). The extracellular level of ACh in the presence of NT and TTX was 58.24 ± 3.9% (n = 6); this can be compared with levels of 140 ± 8.1% (n = 9) when NT was introduced in non-treated rats, P < 0.001 (Fig. 1B). The data for GABA and Asp, respectively, are as follows: 81.6 ± 1.2% (n = 6) for NT + TTX versus 150.7 ± 1.7% (n = 6) for NT alone (P < 0.001, Fig. 2B) and 96.09 ± 3.45% (n = 6) for NT + TTX versus 278.78 ± 12.11% (n = 6) for NT (P < 0.001, Fig. 3B). Differences among experimental groups were evaluated by comparing the areas under the curves (AUC) via oneway analysis of variance (ANOVA) followed by the Newman–Keuls multiple comparison test. All three NT receptor subtypes (NTR1, NTR2, and NTR3) are expressed in the cortex [1,52–54]. However, NTR2 has a substantially lower affinity for neurotensin [63,11,65], and neurotensin acts as an NTR2 antagonist in heterologous expression systems [66]. In contrast, NTR3 is located in intracellular vesicles of neurons and glia and appears to be involved in NT inactivation, cell trafficking, and trophism in cancer cells [39,31,37]. In fact, many of the known central and peripheral effects of NT are exerted mostly through NTR1, since they are blocked by the selective non-peptide neurotensin receptor 1 (NTR1) antagonist SR 48692 [47]. The effect of SR 48692, a non-peptide antagonist specific for the NT receptor type 1 [24] was examined in order to investigate whether the NT-induced increase in the release of ACh, GABA, and Asp in the PFC is mediated by NTR1. Local administration of SR 48692 (0.5 ␮M) for 80 min was followed by an insignificant decrease in the basal output of ACh, GABA, and Asp compared to control basal release and returned upon washout (not shown). When SR 48692 (0.5 ␮M) was administered together with NT (1 ␮M), no increase in ACh, GABA, or Asp levels was observed (Figs. 1A, B; 2A, B, and 3A, B). The extracellular level of ACh in the presence of NT and SR 48692 was 89.00 ± 9.40% (n = 9) versus 140 ± 8.10% (n = 9) when NT was introduced in rats not treated with the antagonist (P < 0.01, Fig. 1B). The data for GABA and Asp, respectively, are as follows: 109.18 ± 1.80% (n = 6) for NT and SR 48692 versus 150.7 ± 1.7% (n = 6) for NT alone (P < 0.001, Fig. 2B) and 117.00 ± 9.80% (n = 6) for NT and SR 48692 versus 278.78 ± 12.11% (n = 6) for NT alone (P < 0.001, Fig. 3B). Differences were evaluated by comparing the area under the curve (AUC) using a one-way ANOVA followed by the Newman–Keuls multiple comparison test. 4. Discussion The aim of the present research was to study whether NT modulates the neuronal input and output of cortical neurons in vivo. Neurotensin was administered locally in the PFC by transversal microdialysis of awake and freely moving rats concurrent with HPLC monitoring of the release of ACh, GABA, Glu, Asp, and taurine. We found that NT stimulated ACh, GABA, and Asp release, but had no effect on Glu or taurine. In our experiments, the main finding was that local administration of neurotensin in the PFC increased ACh outflow. The

demonstrated stimulant effect of neurotensin was TTX-sensitive and was antagonized by the NTR1 neurotensin receptor antagonist SR 48692. To our knowledge, the presented result is the first demonstration of a stimulant effect of neurotensin on ACh release in the cortex of freely moving rats. Supporting our data is the study of Prus et al. [44], which shows that NT69L, a synthetic non-peptide analog of the biologically active 8–13 fragment of neurotensin, stimulates the release of ACh in the medial prefrontal cortex in vivo. The main cholinergic input to the cerebral cortex is provided by the basal forebrain cholinergic complex, including the medial septum, diagonal band of Broca, substantia innominata, ventral pallidum, and nucleus basalis magnocellularis [33,18]. Intrinsic cholinergic neurons have also been reported to exist in the cerebral cortex, where they are often colocalized with vasoactive intestinal polypeptide and lie close to blood vessels [14]. It remains unclear whether neurotensin increases cortical ACh release through an effect on afferent or local cholinergic neurons. Lapchak et al. [30] reported that quinolinic acid lesions of basal forebrain cell bodies abolish the regulatory action of neurotensin on evoked ACh release in cortical slices. Additionally, Wenk et al. [70] observed that ibotenic acid lesions of basal forebrain neurons produce a decrease in the density of NT receptors in the frontoparietal cortex. The loss of NT-mediated regulation of ACh release in cortical regions in lesioned animals may be a reflection of the loss of NT receptor binding sites in these regions and suggests an effect of neurotensin on afferent cholinergic neurons. This is supported by the fact that cortical cholinergic interneurons represent a small population of cells whose contribution to the total amount of releasable cortical ACh should be minor. 4.1. Aspartate and glutamate In the present study, we report a NT-induced increase in extracellular Asp levels. Using a push–pull cannulae system, a similar increase was observed by Sanz et al. [51]. We further show that the stimulant effect of neurotensin is TTX- and calcium-sensitive and is antagonized by the NT receptor antagonist SR 48692. Taken together, these data demonstrate that the NT-stimulated Asp increase is of neuronal origin and is mediated through NTR1. The rise in extracellular Asp in the PFC is not surprising, given the high levels of the amino acid in the cortex. Although large numbers of Asp-immunoreactive neurons are seen in the diagonal band of Broca, globus pallidus, medial habenular nucleus, hippocampus, pons, and brainstem, these neurons are most abundant in the cerebral cortex [9]. Within the cerebral cortex of the rat, significantly higher concentrations of Asp have been reported in the frontal cortex [42]. The NT-induced increase in extracellular Asp is most likely due to a direct effect of the peptide on cortical pyramidal neurons. This is supported by the fact that the majority of dopamine synapses (in the PFC, NT is exclusively localized to dopamine axons) in all prefrontal cortex layers are on pyramidal cells [22,54]. Moreover, pyramidal cells are the major cortical cell type in which NTR1 is expressed [43]. Next, immunocytochemical localization of Asp-like immunoreactivity has shown that many pyramidal cells, ranging from the large pyramidal neurons in layer

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V to the smaller ones in layers VI, III, and II, are Asp-positive [9,13]. Autoradiographic studies using 3 [H]-d-Asp have also demonstrated that many pyramidal neurons in layers V, VI, and III of the cortex use Glu/Asp as transmitters [49,4]. This finding has been further confirmed by autoradiography combined with immunocytochemistry [20]. As mesocortical projections from the VTA innervate deep cortical layers V and VI of the PFC [8] and neurotensin is colocalized with dopamine in all of the mesocortical projections to the deep cortical layers [15], a possible target for neurotensin involves the efferent fibers to subcortical areas (cortical layers V and VI). Although they project to other brain areas, efferent cells may also contribute to the cortical Asp/Glu pool measurable by microdialysis. Almost all efferent cells have axon collaterals projecting to intracortical neurons [26], and these collaterals allow them to participate in the intracortical circuitry. Our data show that glutamate levels in the PFC are not increased after NT perfusion. Since a number of studies demonstrate colocalization of Asp with Glu [32], the selective increase in Asp produced by neurotensin is intriguing. It is possible that Glu- and Asp-immunoreactive neurons represent in part two distinct populations. This is supported by Giuffrida and Rustioni [20], who show that a large fraction of cortico-cortical neurons in the cortex of rat are immunoreactive for either Asp or Glu, but not both. Dori et al. [13] further detect morphological differences between the two groups of neurons, reporting that aspartatestained neurons are larger than glutamate-positive cells in every layer in rat visual cortex. Our results are consistent with the findings of Petrie et al. [43], which show that neurotensin does not have an effect on glutamate release in vivo in conscious rats. While neurotensin was perfused through a vertical probe in their experiments, we administered the neuropeptide transversally so that it could activate cortical neurons in a larger area (8 mm). The question about the effect of NT on Glu levels in the PFC remains controversial, as two other groups report NT-induced rises in glutamate [3,17]. However, one of the groups studies the effect of NT in vitro in rat primary cultures of cortical neurons and cortical slices [3,16]. In the latter, NT (1–13) in the range of 100–1000 nM only slightly increased spontaneously released Glu; neurotensin’s effect was mainly on potassium-evoked Glu release [16]. 4.2. GABA Interneurons play a central role in cortical function, and a number of neuropathological studies have indicated a dysfunction of cortical GABAergic neurons in schizophrenia and bipolar disorder (for a review, see [23]). Ultrastructural studies of the cortex show that dopamine axons synapse on both the dendritic shafts and spines of pyramidal neurons and also the dendrites of local circuit neurons [58] that are immunoreactive for GABA [57]. These studies estimate that ∼40% of the dopamine synapses in the PFC arrive onto GABA-containing local circuit neurons. Although pyramidal cells are the major cortical cell type in which NTR1 is expressed, many interneurons defined on the basis of calcium-binding protein expression also display NTR1 immunoreactivity [43]. Therefore, NT might be a modulator of interneurons that is implicated in both normal and symptomatic cortical function. In our experiments, NT increased GABA release in the PFC. This effect was of neural origin, as it was TTX-sensitive, and mediated by the NTR1 receptor, as it was antagonized by SR 48692. The effect of NT is most likely exerted directly on cortical interneurons, because neurotensin failed to increase Glu that can further activate GABAergic interneurons. The possibility exists, however, that part of this effect is secondary to a neurotensin increase in aspartate levels.

Conflict of interest The authors have no conflicting professional and personal interests. Acknowledgements This work was supported financialy by Ente CRF (Firenze, Italy) and Fondazione MPS (Siena, Italy). The authors are grateful to CNR (Rome, Italy) and the EU COST D34 for supporting our international cooperation and to Dr. Maffrand, Sanofi Recherche, France for the generous supply of the neurotensin antagonist SR 48692. For excellent assistance in measuring amino acid transmitters release by HPLC we are grateful to Ms. Alesandra Colivicchi (Department of Pharmacology, University of Florence). References [1] M.J. Alexander, S.E. Leeman, Widespread expression in adult rat forebrain of mRNA encoding high-affinity neurotensin receptor, J. Comp. Neurol. 402 (1998) 475–500. [2] A. Alonso, M.P. Faure, A. Beaudet, Neurotensin promotes oscillatory bursting behavior and is internalized in basal forebrain cholinergic neurons, J. Neurosci. 14 (1994) 5778–5792. [3] T. Antonelli, L. Ferraro, K. Fuxe, S. Finetti, J. Fournier, S. Tanganelli, M. De Mattei, M.C. Tomasini, Neurotensin enhances endogenous extracellular glutamate levels in primary cultures of rat cortical neurons: involvement of neurotensin receptor in NMDA induced excitotoxicity, Cereb. Cortex 14 (2004) 466–473. [4] P. Barbaresi, M. Fabri, F. Conti, T. Manzo, d-[3H]aspartate retrograde labelling of callosal and association neurones of somatosensory areas I and II of cats, J. Comp. Neurol. 236 (1987) 159–178. [5] A.J. Bean, M.J. During, R.H. Roth, Stimulation-induced release of coexistent transmitters in the prefrontal cortex: an in vivo microdialysis study of dopamine and neurotensin release, J. Neurochem. 53 (1989) 655–657. [6] L. Bianchi, L. Della Corte, K.F. Tipton, Simultaneous determination of basal and evoked output levels of aspartate, glutamate, taurine and 4-aminobutyric acid during microdialysis and from superfused brain slises, J. Chromatogr. Biol. 723 (1999) 47–59. [7] E.B. Binder, B. Kinkead, M.J. Owens, C.B. Nemeroff, Neurotensin and dopamine interactions, Pharmacol. Rev. 53 (2001) 453–486. [8] A. Bjorklund, O. Lindvall, Dopamine containing systems in the CNS, in: A. Bjorklund, T. Hokfelt (Eds.), Handbook of Chemical Neuroanatomy: Classical Transmitters in the CNS, vol. 2, Elsevier, Amsterdam, 1984, pp. 55–122. [9] G. Campistron, R.M. Buijs, M. Geffard, Specific antibodies against aspartate and their immunocytochemical application in the rat brain, Brain Res. 365 (1986) 179–184. [10] R. Carraway, S.E. Leeman, The isolation of a new hypotensive peptide, neurotensin from bovine hypothalamic, J. Biol. Chem. 248 (1973) 6854–6861. [11] P. Chalon, N. Vita, M. Kaghad, M. Guillemot, J. Bonnin, B. Delpech, G. Le Fur, P. Ferrara, D. Caput, Molecular cloning of a levocabastine-sensitive neurotensin binding site, FEBS Lett. 386 (1996) 91–94. [12] G. Damsma, D. Lammerts Van Bueren, B.H.C. Westering, A.S. Horn, Determination of acetylcholine in the femtomole range by means of HPLC, a post-column enzyme reactor, and electrochemical detection, Chromatographia 24 (1987) 827–831. [13] I. Dori, M. Petrou, J.G. Parnavelas, Excitatory transmitter amino acid-containing neurons in the rat visual cortex: a light and electron microscopic immunocytochemical study, J. Comp. Neurol. 290 (1989) 169–184. [14] F. Eskenstein, R.W. Baughman, Two types of cholinergic innervation in cortex, one co-localized with vasoactive interstinal polypeptide, Nature 309 (1984) 153–155. [15] A. Febvret, B. Berger, P. Gaspar, C. Verney, Further indication that distinct dopaminergic subsets project to the rat cerebral cortex: lack of colocalization with neurotensin in the superficial dopaminergic fields of the anterior cingulate, motor, retrosplenial and visual cortices, Brain Res. 547 (1991) 37–52. [16] L. Ferraro, M.C. Tomasini, A. Siniscalchi, K. Fuxe, S. Tanganelli, T. Antonelli, Neurotensin increases endogenous glutamate release in rat cortical slices, Life Sci. 66 (2000) 927–936. [17] C.F. Ferris, in: G.M. Marklouf (Ed.), The Gastrointestinal System: Neural and Endocrine Biology, Betseda, 1989, pp. 559–586. [18] R.P. Gaykema, P.G. Luiten, C. Nyakas, J. Traber, Cortical projection patterns of the medial septum-diagonal band complex, J. Comp. Neurol. 293 (1990) 103–124. [19] M.G. Giovannini, F. Camilli, A. Mundula, G. Pepeu, Glutamatergic regulation of acetylcholine output in different brain regions: a microdialysis study in the rat, Neurochem. Int. 25 (1994) 23–26. [20] R. Giuffrida, A. Rustioni, Glutamate and aspartate immunoreactivity in corticocortical neurons of the sensorimotor cortex of rats, Exp. Brain Res. 74 (1989) 41–46.

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