α6β2*-subtype nicotinic acetylcholine receptors are more sensitive than α4β2*-subtype receptors to regulation by chronic nicotine administration

Authors


Abstract

Nicotinic acetylcholine receptors (nAChR) of the α6β2* subtype (where *indicates the possible presence of additional subunits) are prominently expressed on dopaminergic neurons. Because of this, their role in tobacco use and nicotine dependence has received much attention. Previous studies have demonstrated that α6β2*-nAChR are down-regulated following chronic nicotine exposure (unlike other subtypes that have been investigated – most prominently α4β2* nAChR). This study examines, for the first time, effects across a comprehensive chronic nicotine dose range. Chronic nicotine dose–responses and quantitative ligand-binding autoradiography were used to define nicotine sensitivity of changes in α4β2*-nAChR and α6β2*-nAChR expression. α6β2*-nAChR down-regulation by chronic nicotine exposure in dopaminergic and optic-tract nuclei was ≈three-fold more sensitive than up-regulation of α4β2*-nAChR. In contrast, nAChR-mediated [3H]-dopamine release from dopamine-terminal region synaptosomal preparations changed only in response to chronic treatment with high nicotine doses, whereas dopaminergic parameters (transporter expression and activity, dopamine receptor expression) were largely unchanged. Functional measures in olfactory tubercle preparations were made for the first time; both nAChR expression levels and nAChR-mediated functional measures changed differently between striatum and olfactory tubercles. These results show that functional changes measured using synaptosomal [3H]-DA release are primarily owing to changes in nAChR, rather than in dopaminergic, function.

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This study examined dose–response relationships for murine α6β2*-nicotinic acetylcholine receptor (nAChR) down-regulation by chronic nicotine treatment. The ID50 value for α6β2* down-regulation (35 nM) is ≈ 3x lower than the ED50 value for α4β2* nAChR up-regulation (95 nM), both well within the range reached by human smokers. Chronic nicotine treatment altered α6β2*- and α4β2*-nAChR-mediated [3H]-dopamine release from striatal and olfactory tubercle synaptosomes, but dopaminergic parameters were largely unaffected. We conclude that functional changes are primarily driven by altered nAChR activity.

Abbreviations used
α-Ctx

α-conotoxin

A85380

3-((2S)-azetidinylmethoxy)pyridine

DA

dopamine

DAT

dopamine transporter

DLG

dorsolateral geniculate nucleus

HEPES

2-[4-(2-hydroxyethyl)piperazin-1-yl]ethanesulfonic acid

mHab

medial habunula

nAcc

nucleus accumbens

nAChR

nicotinic acetylcholine receptor

OPTN

olivary pretectal nucleus

opt

optic tract

OT

olfactory tubercle

PMSF

phenylmethylsulfonyl fluoride

SCH23390

7-chloro-3-methyl-1-phenyl-1,2,4,5-tetrahydro-3-benzazepin-8-ol

SC

superior colliculus (superficial layers)

Str

striatum

Th

thalamus

VLGN

ventrolateral geniculate nucleus

VTA

ventral tegmental area

The effects of chronic nicotine exposure on nicotinic acetylcholine receptors (nAChR) have been the subject of many reports. It has been repeatedly demonstrated that chronic exposure to nicotine or tobacco smoke elicits an up-regulation of the number of α4β2-nAChR binding sites for many brain regions and in many species (Marks et al. 1983, 1992, 2011; Schwartz and Kellar 1983; Perry et al. 1999; Govind et al. 2009). However, some brain regions (such as thalamus and medial habenula) are less affected than others (such as cerebral cortex and hippocampus). The up-regulation occurs with no change in mRNA levels (Marks et al. 1992). The cellular processes underlying the up-regulation and the functional consequences of this up-regulation are complex and not fully understood. For example, the function of the α4β2*-nAChR has been shown to increase, decrease, or remain unchanged depending on the measure used (Marks et al. 1993; Jacobs et al. 2002; Grilli et al. 2005). Up-regulation of nAChR expression is not exhibited by every subtype. Specifically, down-regulation has been reported for the α6β2*-nAChR binding sites (Lai et al. 2005; Perry et al. 2007; Doura et al. 2008). Furthermore, the function of α6β2*-nAChR subtypes also appears to decrease or remain unchanged after chronic nicotine exposure (Lai et al. 2005; McCallum et al. 2006; Perry et al. 2007).

Differential nAChR subtype responses to chronic nicotine exposure are of particular importance in dopaminergic systems. Dopaminergic neurons express a variety of nicotinic receptor subtypes that contain α4β2*-nAChR-and/or α6β2*-nAChR-binding sites (Champtiaux et al. 2003; Gotti et al. 2005). Some of the α4β2*-nAChR also include the α5 subunit; the (α4β2)2α5-nAChR subtype seems to be generally resistant to up-regulation (Mao et al. 2008; Moretti et al. 2010). In addition, (α4β2)2β2-nAChR sites located on dopaminergic neurons may not up-regulate (Nashmi et al. 2007). Consequently, up-regulation of α4β2*-nAChR sites in dopaminergic regions may be restricted to other types of neurons, perhaps GABAergic.

The α6β2*-nAChR are diverse and appear to respond differently to nicotine treatment. The subtype that contains both α4 and α6 subunits [(α4β2)(α6β2)β3] may down-regulate more than other α6β2-nAChR subtypes [(α6β2)2β3 and (α6β2)2β2] (Perez et al. 2008; Quik et al. 2011). Given the complexity and variety of nAChR subtypes expressed on dopaminergic neurons, it has been difficult to assess consequences of chronic nicotine exposure on this system. More recently, longer term chronic nicotine treatments by water bottle, minipump, and/or food, with or without cycles of withdrawal in mice, rats, or monkeys have shown changes in reward behavior as well as changes in modulation of dopamine (DA) release by cyclic voltammetry methods (Hilario et al. 2012; Perez et al. 2012; Zhang et al. 2012; Baker et al. 2013; Bordia et al. 2013).

Several smoking cessation aids that target nicotinic acetylcholine receptors (nAChR) are in current use, including nicotine replacement by patch and gum, and varenicline, a partial agonist with high potency at the α4β2*-nAChR subtype. The suboptimal efficacy of these treatments in achieving tobacco abstinence necessitates a search for other therapeutics, perhaps for alternative targets (Pierce et al. 2012; Hurst et al. 2013). Some of the less widely distributed nAChR subtypes have been proposed as targets. One of these is the α6β2*-nAChR with expression restricted mainly to dopaminergic and visual pathways (Brunzell 2012). This subtype regulates function of ventral tegmental area (VTA) dopaminergic projection neurons, a pathway considered important in reward. Pharmacological manipulation of a more selective target, such as α6β2*-nAChR could be effective in aiding smoking cessation attempts with possibly fewer side effects. Availability of more detailed information about a target receptor should aid in designing better pharmacotherapies.

This study was undertaken to compare changes in both α4β2*-nAChR and α6β2*-nAChR binding sites as well as to evaluate functional changes resulting from variation in chronic nicotine treatment dose. We report that the chronic dose required for half-maximal change of α6β2*-nAChR site expression (decrease) was significantly lower than that required for half-maximal change of the α4β2*-nAChR sites (increase). In addition, binding site changes for both α6β2*-nAChR and α4β2*-nAChR-nAChR binding sites occur at lower chronic treatment doses than those leading to functional changes measured by nAChR-mediated [3H]-dopamine release using synaptosomal preparations. In comparison to nAChR binding and functional parameters, measures of dopaminergic receptor expression and DA transporter were largely unchanged; we conclude that nAChR-level changes are likely primarily responsible for the observed effects of chronic nicotine on nAChR-mediated [3H]-dopamine release.

Methods

Materials

[125I]-Epibatidine (2200 Ci/mmol), [3H]SCH23390 (N-methyl [3H], 84.3 Ci/mmol), [3H]mazindol (4'-[3H], 20.6 Ci/mmol), and [3H]raclopride (methoxy-[3H], 20.6 Ci/mmol) were obtained from Perkin-Elmer NEN, Boston, MA, USA. α-Conotoxin MII (α-CtxMII) and [125I]-α-CtxMII were prepared as described previously (Cartier et al. 1996; Whiteaker et al. 2000). Unlabeled 5I-epibatidine was a generous gift from Dr Kenneth Kellar, Georgetown University, Washington, DC, USA. Liquid (-)-nicotine was purchased from Sigma-Aldrich Chemical Company (St. Louis, MO, USA) and was redistilled periodically. All additional chemicals were of reagent grade.

Mice

Animal production methods and experimental procedures using mice were reviewed and approved by the Institutional Animal Care and Utilization Committee at the University of Colorado, Boulder. All mice were bred at the Institute for Behavioral Genetics, University of Colorado, Boulder. β2 null mutant mice (Picciotto et al. 1995) were originally obtained from Marina Picciotto, Yale University, New Haven, CT, USA. Heterozygous mice were bred and pups genotyped from ear or tail clippings taken at weaning (21 days) (Salminen et al. 2004). Mice were housed at 22°C with lights on from 7 AM to 7 PM and allowed free access to food and water.

Surgery

Cannulas were inserted in the right jugular vein of each mouse using previously published procedures (Barr et al. 1979). Briefly, mice were anesthesized with pentobarbital (50 mg/kg) and chloral hydrate (100 mg/kg). An incision exposed the superficial right jugular vein into which a silastic cannula (0.51 mm I.D. 0.94 mm O.D) was inserted. The cannula was passed through the back in the midscapular region and stabilized with surgical thread and a wound clip. The mouse was injected with buprenorphine (0.1 mg/kg), placed in a clean cage and warmed until wakening.

Nicotine treatment

Mice were transferred to an individual infusion chambers and cannulas were connected to a 1 mL syringe mounted on an infusion pump (Harvard Apparatus, Holliston, MA, USA). Sterile saline was continuously infused at a rate of 35 μL/h for 2 days before nicotine treatment was begun. Mice were divided into seven treatment groups that received the following nicotine doses (free base as mg/kg/h, prepared from liquid nicotine neutralized with HCl): 0 (saline-infused control), 0.125, 0.25, 0.5, 1.0, 2.0, or 4.0. This range of doses was chosen to maintain compatibility with our previous chronic nicotine treatment studies and to provide plasma nicotine concentrations that increase linearly with dose (Marks et al. 2004). In addition, as mice metabolize nicotine much faster than humans, plasma nicotine concentrations within the chosen dosing range will also cover the range of doses likely to be experienced by human smokers. The highest mouse dose (4.0 mg/kg/h) will produce an ≈ 1.2 μM plasma concentration; this slightly exceeds the upper end of the range seen in very heavy smokers (Marks et al. 2004; Matta et al. 2007). Following 10 days of treatment with the indicated nicotine dose, nicotine treatment was discontinued and each cannula was checked for free fluid flow. A 2 h interval of withdrawal was allowed before killing to metabolize nicotine (Petersen et al. 1984).

Tissue preparation

For samples to be prepared for sectioning, each mouse was killed by cervical dislocation, its brain was rapidly (< 1 min) removed from the skull and quickly frozen by immersion in isopentane (−35°C) for 10 s. The frozen brain was wrapped in aluminum foil and stored at −70°C until sectioning.

Preparation of tissue sections

Frozen brains were removed from the −70°C freezer and allowed to warm to the temperature of the cryostat (−14°C) and mounted with M-1 Embedding Matrix (Anatomical Pathology, Pittsburgh, PA, USA). Subsequently, 14 μm coronal sections were cut using either a Leica CM 1850 cryostat/microtome (Leica, Nussloch, Germany) or an IEC Minotome (Damon Corp., Needham, MA, USA) and thaw mounted on Fisher Suprafrost/Plus microscope slides. A series of ten sets of slides were prepared from each brain to allow comparison of results for several different experiments on adjacent or near-adjacent sections. Slides containing the brain sections were stored, desiccated at −70°C, until use.

[125I]Epibatidine autoradiography

Slides with tissue sections prepared were warmed to 22°C in a desiccators, transferred to Bel-Art (Wayne, NJ, USA) slide racks that have been modified to hold 50 slides and rehydrated by incubation at 22°C for 15 min in isotonic buffer (NaCl, 144 mM; KCl, 2.2 mM, CaCl2, 2.0 mM, MgSO4, 1.0 mM; HEPES, 25 mM; pH = 7.5). Rehydrated slides were subsequently transferred to isotonic buffer containing 200 pM [125I]epibatidine (specific activity 2200 Ci/mmol mixed with unlabeled 5I-epibatidine to yield a final specific activity of 220 Ci/mmol, a 10-fold dilution). Samples were incubated for 2 h at 22°C, then redistributed to slide racks containing 25 slides and washed as follows (all solutions at 4°C): Twice for 30 s in isotonic buffer, twice for 5 s in hypotonic buffer (0.1 x), and twice for 5 s in 10 mM HEPES, pH = 7.5. Parallel series of sections were used to determine [125I]epibatidine (200 pM) binding levels in the presence of 30 nM cytisine [sufficient to block binding to α4β2* nAChR, while leaving other subtypes unaffected (Whiteaker et al. 2000, 2002)]. Subsequently, slides were air dried and stored desiccated at 22°C under vacuum overnight before exposure initially to Packard Super Resolution Cyclone Storage Phosphor Screens to yield images for quantitation, and subsequently to Kodak MR autoradiography film (both from PerkinElmer, Inc., Waltham, MA, USA) to yield higher resolution images for photography. Each Phosphor Screen was also simultaneously exposed to a series of tissue paste standards containing measured amounts of 125I to allow quantitation of the image intensity. Tissue sections from β2 null mutant mice or samples incubated in the presence of 10 μM nicotine were used to establish blanks that did not differ from film background (Whiteaker et al. 2006).

[125I]-α-CtxMII autoradiography

[125I]-α-CtxMII binding was performed as previously described (Whiteaker et al. 2000). Sections were incubated for 10 min in binding buffer containing 1 mM phenylmethylsulfonyl fluoride (PMSF). Sections were then incubated with 0.5 nM [125I]-α-CtxMII in binding buffer with the addition of the protease inhibitors leupeptin, pepstatin, and aprotinin (10 μg/mL each) and bovine serum albumin (0.1% w/v), for 2 h at 22°C. In parallel series of slides, Nicotine was used to gauge nonspecific binding in the presence of [125I]-α-CtxMII (see also legend to Fig. 2). Slides were subsequently washed in ice-cold protein free binding buffer twice for 30 s followed by two 10 s washes in 0.1× protein free binding buffer. Final rinses (2 × 5 s each) were conducted in ice-cold 5 mM HEPES, pH 7.5. Sections were subsequently air dried and desiccated overnight prior to exposure to Packard Phosphor screens and Kodak MR film as previously described for [125I]epibatidine autoradiography. Tissue sections from β2 null mutant mice were again used to establish blanks.

Quantitation

Tissue paste samples prepared from whole brain homogenates containing measured amounts of 125I were used to construct standard curves. The Phosphor screens produce a linear relationship between signal intensity and tissue radioactivity content over several orders of magnitude. The regression line calculated for the standard curve was used to convert the measured value of pixels/mm2 to the cpm/mg wet weight. Signal intensity in fmol/mg wet weight was estimated from the specific activity of each ligand. Brain regions were identified using a mouse brain atlas (Paxinos and Franklin 2004) as a guide. Multiple (4–6) measurements were made in each brain region of each mouse and the average of these measurements defined the signal intensity for each region.

Synaptosomal superfusion [3H]-Dopamine release assay

Two DA terminal regions, striatum (ST) and olfactory tubercle (OT) were assessed for [3H]-dopamine ([3H]-DA) release from crude synaptosomes as previously described (Salminen et al. 2004, 2007). These two regions were chosen as they are easily distinguished, rich sources of terminals from DA neurons originating in the substantia nigra (SN) and the VTA, respectively. The OT and the nucleus accumbens both receive projections from similar areas of the VTA, and both regions respond similarly in drug-reward assessments (Ikemoto 2007). Nucleus accumbens was not used as sufficient material could only be obtained for functional assays by pooling samples from multiple mice. This was not practical given the demands of the chronic treatment experiments. Nicotine treatment was terminated at least two h before tissue preparation to allow metabolism of nicotine. For uptake of [3H]-DA, crude synaptosomes were incubated at 37°C in uptake buffer (containing the following, in mM, 128 NaCl, 2.4 KCl, 3.2 CaCl2, 1.2 KH2PO4, 1.2 MgSO4, 25 HEPES, pH 7.5, 10 glucose, 1 ascorbic acid, and 0.01 pargyline) for 10 min before addition of 100 nM [3H]-DA (1 μCi for every 0.2 mL of synaptosomes) and diisopropyl fluorophosphate (10 μM). The suspension was incubated for an additional 5 min. Aliquots of crude synaptosomes (80 μL) were distributed onto filters and superfused with buffer (uptake buffer containing 0.1% bovine serum albumin, 1 μM nomifensine, and 1 μM atropine) at 0.7 mL/min for 10 min. For experiments using α-conotoxin MII (α-CtxMII) to inhibit α6β2*-nAChR, crude synaptosomes were exposed to α-CtxMII (50 nM) for the last 5 min of the 10 min buffer superfusion. This concentration of α-CtxMII inhibits all α6β2*-nAChR subtypes present in the mouse striatum (Salminen et al. 2007). [3H]DA release mediated by α6β2*-nAChR was calculated as the difference between total release (in the absence of α-CtxMII) and α4β2*-nAChR-mediated release (retained in the presence of α-CtxMII). After buffer and/or α-CtxMII superfusion, crude synaptosomes were exposed to ACh for 20 s. Basal and ACh-stimulated release was determined by collecting 23 × 10 s fractions into 96-well plates.

Radioligand binding to membrane preparations

Expression levels of DA receptors and transporter were measured by binding of radioligands to membrane fractions isolated from olfactory tubercles or striata of mice treated with saline, 0.25, 1.0, or 4.0 mg/kg/h nicotine.

The expression of D1/D5 DA receptors was measured by membrane binding of [3H]-SCH23390. Membrane preparations were incubated in binding buffer (30 μL volume including in mM: NaCl, 144; KCl, 1.5; CaCl2, 2; MgSO4, 1; HEPES, 20; pH = 7.5). [3H]-SCH23390 was used at 1.7 nM, (Kd ~ 0.1 nM), with 0.5 mM mianserin added to block binding to 5HT2 receptors, and 10 mM flupenthixol added for blank determination. Samples were incubated in 30 μL of binding buffer for 60 min at 22°C. Expression levels of D2/D3 DA receptors were measured with [3H]-raclopride at 17 nM, (Kd ~ 2 nM) with 10 mM sulpiride added for blank determination. Assay volume and incubation time was identical to that used for determination of D1/D5 DA receptor expression, as was the bulk buffer composition.

Dopamine transporter (DAT) expression was measured by binding of [3H]-mazindol (200 nM, Kd ~ 25 nM). Alternatively, for a small number of samples, [125I]-RTI-121 (8 nM, Kd ~ 4 nM) was used. Fluoxetine (1 μM) and desipramine (100 nM) were used to block off-target binding to the serotonin and norepinephrine transporters, respectively. Blanks were determined using 100 mM nomifensine. Samples were incubated in 30 μL of binding buffer for 90 min at 22°C followed by 30 min at 4°C. Assay volumes were the same as for DA receptor assays, previously described.

Reactions were terminated by filtration of samples at 4°C onto a two layer filter consisting of one sheet of GFA/E glass fiber filter (Gelman Sciences, Ann Arbor, MI, USA) and one sheet of GF/B glass fiber filter (Micro Filtration Systems, Dublin, CA, USA) both treated with 0.5% polyethylenimine using an Inotech Cell Harvester (Inotech, Rockville, MD, USA). Samples were subsequently washed six times with ice-cold binding buffer. Radioactivity was measured using a Wallac 1450 Microbeta scintillation counter (PerkinElmer Life and Analytical Sciences, Waltham, MA, USA) after addition of 150 μL of Optiphase Supermix scintillation cocktail (PerkinElmer Life Sciences, Waltham, MA, USA) to each well of a 96-well counting plate.

Dopamine transporter activity assays

Synaptosomes were prepared as for DA release assays. Synaptosomes were suspended in uptake buffer (in mM: NaCl, 128; KCl, 2.4; CaCl2, 3.2; KH2PO4, 1.2; MgSO4, 1.2; HEPES, 25, pH 7.5, glucose, 10; ascorbic acid, 1; and pargyline, 0.01) to a volume of 8 mL per mouse for ST and 4 mL per mouse for OT. Previously, we have determined the Km concentration of DA for uptake into striatal synaptosomes of C57Bl6 mice to be 0.080 ± 0.003 mM (data not shown). Synaptosomes were incubated in uptake buffer (final volume 100 μL) with either 0.05 μM (close-to-Kd) or 1 μM (saturating concentration) of [3H]-DA (40–55 Ci/mmol diluted with unlabeled DA to achieve concentration desired) at 22°C for 5 min. Nomifensine (100 μM) was used for blank determination. The reaction was terminated by filtration onto a two layer filter consisting of one sheet of GFA/E glass fiber filter (Gelman Sciences) and one sheet of GF/B glass fiber filter (Micro Filtration Systems) both soaked in uptake buffer. Samples were washed with cold uptake buffer four times. Radioactivity was measured using a Wallac 1450 Microbeta scintillation counter (PerkinElmer Life and Analytical Sciences) after addition of 150 μL of Optiphase Supermix scintillation cocktail (PerkinElmer Life Sciences) to each well of a 96-well counting plate.

Protein

Protein was measured using the method of Lowry et al. (1951).

Statistical analyses

SigmaPlot V9 (Systat Software, Inc., San Jose, CA, USA) was used to derive assay parameters by non-linear regression curve fitting. The following equations were used to examine the decreases or increases in binding site density after chronic nicotine treatment: For decreases: BN = BS/(1 + nic/ED50) + BR and for increases: BN = BS/(1 + ED50/nic) + BR: where BN is the binding measured following treatment with nicotine, BS is the binding affected by the chronic nicotine treatment at the dosage = nic, ED50 is the nicotine treatment dose eliciting 50% of the maximal change, and BR is the binding unaffected by nicotine treatment.

The SPSS statistical package (IBM Corp., Somers, NY, USA) was used for statistical analyses. Two-way anovas (independent variables were nicotine treatment dose and brain region) were used to evaluate the effects of nicotine treatment on [125I]epibatidine and [125I]α-CtxMII binding. Subsequently, the effects of chronic nicotine treatment on binding site density were analyzed by non-linear regression analysis as described above. One-way anovas were also used to examine the effects of nicotine treatment on binding site density for each ligand in each brain region, and to evaluate the effects of chronic nicotine treatment on the binding of [3H]SCH23390, [3H]mazindol, and [3H]raclopride.

Results

The typical regional distribution of [125I]α-CtxMII and [125I]epibatidine binding is shown in Fig. 1. Regions that were assessed by quantitative autoradiography are labeled. [125I]α-CtxMII binding sites are prominently expressed in a limited set of brain nuclei, typically associated with optic tracts and DA projections originating in the substantia nigra and VTA (Whiteaker et al. 2000). Within these regions, almost all [125I]α-CtxMII binding sites correspond to α6β2* nAChR (Champtiaux et al. 2002; Whiteaker et al. 2002; Gotti et al. 2005).

Figure 1.

Autoradiograms of [125I]α-conotoxinMII ([125I]α-CtxMII) and [125I]epibatidine binding. Representative examples are shown of radioligand binding patterns, recorded on X-ray film, using autoradiography approaches as described in the 'Methods' section. Left column: [125I]α-CtxMII (500 pM) labeling. Right column: [125I]epibatidine (200 pM). Regions of interest are labeled as follows: DLG, dorsolateral geniculate nucleus; mHab, medial habunula; nAcc, nucleus accumbens; opt, optic tract; OPTN, olivary pretectal nucleus; OT, olfactory tubercle; SC, superior colliculus (superficial layers); Str, striatum; Th, thalamus; VLGN, ventrolateral geniculate nucleus.

[125I]α-CtxMII binding sites are highly sensitive to down-regulation by chronic nicotine exposure

[125I]α-CtxMII binding site densities were determined in brains from mice exposed to a range of chronic nicotine doses (0–4 mg/kg/h for 10 days; see 'Methods' for details). Regions with measurable [125I]α-CtxMII binding were quantitated (see Fig. 2). Chronic treatment with even low doses of nicotine produced down-regulation of [125I]α-CtxMII binding site expression in most regions (Fig. 2). Comparison of binding levels by 2-way anova indicated significant differences by Region, and by Dose, but no Dose × Region interaction (statistical analysis outcomes are provided in the legend to Table 1). Results were subsequently analyzed to evaluate the dose-dependence of the down-regulation and calculate the extent of the down-regulation. The mean ID50 value across regions exhibiting significant down-regulation was 0.108 ± 0.040 mg/kg/h (see Table 1). This corresponds to a serum nicotine concentration of 35 nM (Marks et al. 2004). This analysis also provides an estimate of the maximal decrease in expression of [125I]α-CtxMII binding sites and illustrates that chronic nicotine administration modestly, but significantly, reduces α6β2* nAChR expression across multiple regions (mean of five regions analyzed −24.1 ± 1.2% of saline control, see Table 1). Optic tracts and superior colliculus (for which regions no significant effect of nicotine treatment was observed) were not included.

Table 1. Curve-fit parameters for chronic nicotine-induced changes (up- or down-regulation) of nAChR expression
Region[125I]α-CtxMII binding sites (α6β2* nAChR)Cytisine-sensitive [125I]-epibatidine binding sites (α4β2* nAChR)
ED50 (nM)/mg/kg/hMax % changeED50 (nM) /mg/kg/hMax % change
  1. Parameters are derived from curve fits performed in Figs 2 (for α6β2* nAChR; left column) and 3 (for α4β2* nAChR; right column). nd = not determined for reasons of undetectable binding; ns = un-measurable change by treatment. Two-way anova was performed to determine the effects of Region and Dose on α6β2* nAChR down-regulation following chronic nicotine treatment levels. Both factors significantly affected expression (F[7,56] = 591, p < 0.001 and F[6,56] = 6.33, p < 0.001, respectively), confirming that expression of α6β2* nAChR varies across brain regions, and is significantly affected by chronic nicotine treatment. No Region × Dose interaction was seen, however (F[7,42] = 0.86, p < 0.727), demonstrating that the dose dependence of α6β2* nAChR down-regulation is generally similar for each region. Results were further analyzed to calculate the nicotine treatment dose dependence on binding site densities for each brain region as described in the 'Methods'. Significant and similar extents of down-regulation were observed for five of the eight brain regions, and are denoted by *(p < 0.05). The means ± SEM for the maximum percentage decreases shown are significantly less than zero as determined by t-test for that parameter calculated from the non-linear least-squares curve fits. Accordingly, a mean ED50 down-regulation nicotine dose was calculated for those five brain regions. Expression of α4β2* nAChR was generally up-regulated, in contrast to the effects recorded for α6β2* nAChR. Two-way anova showed that both Region (F[12,91] = 104, p < 0.001) and Dose (F[6,91] = 3.08, p = 0.006) significantly affected expression of this receptor population. Again, no Region × Dose interaction was observed (F[12,72] = 0.27, p = 1). Results were further analyzed to calculate the nicotine treatment dose dependence for the increase in binding site densities as described in the 'Methods'. Significant up-regulation was detected in eight of the 14 regions analyzed, and are denoted by *(p < 0.05). The means ± SEM for the maximum percentage decreases shown are significantly greater than zero as determined by t-test for that parameter calculated from the non-linear least-squares curve fits. Mean ED50 values for nicotine-induced up-regulation were also calculated for these regions. By t-test, the ED50 values for α6β2* nAChR down-regulation are significantly different from the ED50 values for the α4β2* nAChR up-regulation (t = 5.23, degrees-of-freedom = 12; p < 0.001). Regions of interest are labeled as follows: Regions of interest are labeled as follows: DLG, dorsolateral geniculate nucleus; fCX, frontal cortex; in CX, inner layers of cortex; nAcc, nucleus accumbens; opt, optic tract; OPTN, olivary pretectal nucleus; orbCx, orbital cortex; outCx, outer layers of cortex; OT, olfactory tubercle; SCdl, superior colliculus (deep layers); SCsg, superior colliculus (superficial gray); Str, striatum; Th, thalamus; VLG, ventrolateral geniculate nucleus.

OT0.128 ± 0.057−28.8 ± 13.4*0.258 ± 0.13071.1 ± 23.5*
nAcc0.136 ± 0.110−23.9 ± 9.6*0.446 ± 0.21289.5 ± 33.3*
Str0.241 ± 0.129−22.0 6.7*0.255 ± 0.17047.2 ± 19.9*
optnsnsnsNs
OPTNnsnsnsNs
DLG0.066 ± 0.045−22.4 ± 6.7*nsns
VLG0.061 ± 0.047−23.2 ± 6.2*nsns
SCsgnsnsnsns
fCXndnd0.339 ± 0.18973.6 ± 25.7*
inCXndnd0.208 ± 0.19393.2 ± 29.9*
orbCXndnd0.218 ± 0.11671.8 ± 22.9*
outCXndnd0.287 ± 0.16061.4 ± 23.1*
SCdlndndnsns
Thndndnsns
Overall for regions with changes0.108 ± 0.040−24.1 ± 1.20.293 ± 0.02772.5 ± 5.9*
Figure 2.

Concentration dependence of decreased [125I]α-CtxMII binding site [α6β2* nicotinic acetylcholine receptor (nAChR)] expression following chronic nicotine administration. Mice were treated for 10 days with continuous infusion of sterile saline (control) or a range of nicotine doses (0.125–4 mg/kg/h). Expression of α6β2* nAChR was assessed using [125I]α-CtxMII autoradiography in sections (14 μm) collected from each brain. Specific labeling was determined as the difference in labeling by quantitation of total binding (500 pM [125I]α-CtxMII alone) and non-specific labeling (500 pM [125I]α-CtxMII in the presence of 100 μM nicotine) between adjacent sections or equivalently by binding to sections of β2 null mutant mice, by reference to known radioactive standards. Please see 'Methods' section for experimental details. n = 9 for each point; points represent mean ± SEM; *denotes a significant difference (p < 0.05) from no-nicotine control binding detected by one-way anova within each region, followed by the Duncan post hoc test. All other statistical analyses and calculated parameters are given in Table 1.

Cytisine-sensitive [125I]Epibatidine binding sites are less sensitive to up-regulation by chronic nicotine exposure

Expression of cytisine-sensitive [125I]epibatidine binding sites (corresponding to α4β2* nAChR) was generally up-regulated by chronic nicotine treatment (Fig. 3) in the same regions that showed down-regulation of [125I]α-CtxMII binding sites (corresponding to α6β2* nAChR). Note that [125I]epibatidine binding data are shown for seven of the eight regions analyzed for [125I]α-CtxMII binding plus two cortical regions. Optic tract [125I]epibatidine binding could not be distinguished from [125I]epibatidine binding to the adjacent thalamic region. Comparison by 2-way anova of the effects of chronic nicotine treatment on [125I]epibatidine binding levels in the brain regions listed in Table 1 indicated significant differences by Region, and by Dose, but no Dose × Region interaction (statistical analysis outcomes are provided in the legend to Table 1). As for [125I]α-CtxMII binding sites, levels of α4β2* nAChR expression vary between regions of interest, and are affected by chronic nicotine administration (although expression changes are opposite in direction between the two nAChR populations). Analysis of the dose dependence for up-regulation of [125I]epibatidine binding by non-linear regression indicated that higher doses of nicotine were required to up-regulate α4β2* nAChR expression than to down-regulate α6β2*-nAChR expression. The mean ED50 value across seven7 regions with significant response to nicotine treatment was 0.293 ± 0.027 mg/kg/h (see Table 1). This corresponds to a serum nicotine concentration of 95 nM (Marks et al. 2004), which is ~3-fold higher than the ID50 value for the [125I]α-CtxMII binding down-regulation. The extent of up-regulation varied from a 47% to a 93% increase for regions with significant changes (mean for seven regions, +72.5 ± 5.9%, see Table 1).

Figure 3.

Concentration dependence of increased cytisine-sensitive [125I]epibatidine binding site [α4β2* nicotinic acetylcholine receptor (nAChR)] expression following chronic nicotine administration. As in Fig. 2, mice were treated for 10 days with continuous infusion of sterile saline (control) or a range of nicotine doses (0.125–4 mg/kg/h). Expression of α4β2* nAChR was assessed using [125I]epibatidine autoradiography in sections (14 μm) collected from each brain. Specific labeling of α4β2* nAChR was determined as the difference in labeling by quantitation of total binding (200 pM [125I]epibatidine alone) and cytisine-sensitive labeling (200 pM [125I]epibatidine in the presence of 50 nM cytisine) between adjacent sections, by reference to known radioactive standards. Non-specific binding measured by including 10 μM nicotine in the incubation did not differ from film background. Please see 'Methods' section for experimental details. n = 5–6 for each point; points represent mean ± SEM; *denotes a significant difference (p < 0.05) from no-nicotine control binding detected by one-way anova within each region, followed by the Duncan post hoc test. All other statistical analyses and calculated parameters are given in Table 1.

Chronic exposure to nicotine doses reduces α6β2*-nAChR-mediated [3H]DA release from striatal synaptosomes

The effects of chronic nicotine treatment on α-CtxMII-sensitive (α6β2*) nAChR function were assessed using [3H]DA release from striatal and OT synaptosomal preparations. Chronic nicotine treatment doses were chosen that corresponded to partial α6β2* down-regulation (0.25 mg/kg/h), partial α4β2 up-regulation (1 mg/kg/h), or full effect on both (4 mg/kg/h). The outcomes of these experiments are illustrated in Fig. 4.

Figure 4.

Concentration–response curves for nicotine-evoked [3H]dopamine release mediated by α6β2* nicotinic acetylcholine receptor (nAChR) following chronic nicotine administration. Crude synaptosomes were produced from striatum and olfactory tubercles of animals treated for 10 days with saline (control) or three different nicotine doses as indicated. Synaptosomes were loaded with [3H]-DA, then superfused for 5 min in the presence or absence of 50 nM α-CtxMII just prior to stimulation with a range of nicotine concentrations (10 nM–3 μM). α-CtxMII-sensitive release (mediated by α6β2* nAChR) of [3H]-DA was calculated by subtracting the resistant fraction from the total (n = 6–11 for each point; points represent mean ± SEM), and is expressed as a multiple of baseline release just before stimulation. The curves for stimulation by nicotine represent the best fits of the data as one saturable component, for each chronic treatment regime, in each region, as described in the 'Methods'. Calculated parameters and statistical analyses are given in Table 2.

In the striatum, only the very highest dose of chronic nicotine treatment produced a significant change in the amount of nicotine-evoked α6β2*-nAChR-mediated [3H]-DA release (see Table 2) [3H]-DA release EC50 values, while trending downward, were not significantly changed with any of the treatments. In contrast, no statistical differences were detected by one-way anova in nicotine-evoked α6β2*-nAChR-mediated release parameters from OT preparations at any chronic nicotine dose. Statistical analysis is presented in the legend to Table 2.

Table 2. Curve-fit parameters for DA release (all Hill fit numbers)
TreatmentEC50 R max n H EC50 R max n H
  STSTSTOTOTOT
  1. Parameters are derived from curve fits performed in Figs 4 (for α6β2* nAChR) and 5 (for α4β2* nAChR). One-way anova was performed comparing each parameter across treatment conditions, for each region. Where significant effects of treatment were detected, Duncan's post hoc test was used to identify which treatment condition(s) produced significantly different parameter values compared to saline controls (*p < 0.05). Significant differences were only seen at the highest nicotine dose used (4.0 mg/kg/h), and were restricted to a subset of Rmax values, as follows:α6β2*-nAChR significant difference seen for striatal Rmax (F3,11 = 10.41, p = 0.004).

    α4β2*-nAChR significant difference seen for striatal Rmax (F3,11 = 7.54, p = 0.010), and for olfactory tubercle Rmax (F3,11 = 9.39, p = 0.005).

α6β2*-nAChR
Saline0.099 ± 0.0262.48 ± 0.191.21 ± 0.0320.110 ± 0.0242.79 ± 0.181.12 ± 0.22
0.25 mg/kg/h nicotine0.077 ± 0.0302.28 ± 0.251.32 ± 0.560.107 ± 0.0272.51 ± 0.181.07 ± 0.23
1.0 mg/kg/h nicotine0.075 ± 0.0232.21 ± 0.201.75 ± 0.770.181 ± 0.1212.47 ± 0.501.00 ± 0.49
4.0 mg/kg/h nicotine0.048 ± 0.0221.57 ± 0.20*1.54 ± 0.870.156 ± 0.0572.68 ± 0.270.84 ± 0.18
α4β2*-nAChR
Saline0.98 ± 0.1813.9 ± 0.60.81 ± 0.081.26 ± 0.4415.3 ± 1.20.71 ± 0.11
0.25 mg/kg/h nicotine1.34 ± 0.3515.4 ± 1.00.77 ± 0.102.10 ± 0.6018.7 ± 1.40.69 ± 0.08
1.0 mg/kg/h nicotine1.14 ± 0.4013.9 ± 1.20.71 ± 0.110.87 ± 0.2415.2 ± 1.00.72 ± 0.10
4.0 mg/kg/h nicotine0.90 ± 0.2211.8 ± 0.8*0.89 ± 0.141.47 ± 0.5613.2 ± 1.2*0.68 ± 0.10

Chronic exposure to nicotine reduces α4β2*-nAChR-mediated [3H]DA release from striatal and olfactory tubercle synaptosomes

The effects of chronic nicotine treatment on α-CtxMII-resistant (α4β2*) nAChR function were also determined, using [3H]DA release from striatal and OT synaptosomal preparations. The same chronic nicotine doses were used as for assessment of α6β2* nAChR function. The results are shown in Fig. 5. Note that the relative errors associated with the measurements are small for α-CtxMII-resistant (α4β2* nAChR) function compared to determinations of α-CtxMII-sensitive (α6β2* nAChR) function as the latter is calculated as the difference between two measures (total and α-CtxMII-resistant function).

Figure 5.

Concentration–response curves for nicotine-stimulated [3H]dopamine release mediated by α4β2* nicotinic acetylcholine receptor (nAChR) following chronic nicotine administration. Crude synaptosomes were produced from striatum and olfactory tubercles of animals treated for 10 days with saline (control) or with the indicated different nicotine doses. Synaptosomes were loaded with [3H]-DA, then superfused for 5 min in the presence of 50 nM α-CtxMII just prior to stimulation with a range of nicotine concentrations (10 nM–3 μM). The residual α-CtxMII-resistant release of [3H]-DA is mediated by α4β2* nAChR (n = 6–11 for each point; points represent mean ± SEM), and is expressed as a multiple of baseline release just before stimulation. The curves for stimulation by nicotine represent the best fits of the data as one saturable component, for each chronic treatment regime, in each region, as described in the 'Methods'. Calculated parameters and statistical analyses are given in Table 2.

For striatal synaptosomes, the two highest chronic nicotine treatment doses elicited significant decreases in the amount of nicotine-evoked α4β2*-nAChR-mediated [3H]-DA release (see Table 2). This contrasts with the effects on striatal α6β2*-nAChR-mediated function, for which function as well as amount of binding was reduced; whereas reductions in α4β2*-nAChR-mediated [3H]-DA release are seen even though binding sites are increased. Furthermore, only the highest dose of nicotine significantly changed either function, even though half-maximal values for changing binding are considerably lower. No significant effect of chronic nicotine treatment was seen on α6β2*- or α4β2*-nAChR-mediated [3H]-DA release EC50 values (see Table 2).

Similar significant decreases were seen for α4β2*-mediated function in OT synaptosomal preparations following treatment with the highest nicotine dose.

Chronic nicotine treatment does not affect dopamine receptor or transporter expression in striatum or olfactory tubercles

As previously described, chronic nicotine treatment elicited changes in nAChR-mediated DA release from striatal and OT synaptosomal preparations. This effect might reflect changes in either nAChR function or be because of changes in dopaminergic parameters induced by chronic nicotine treatment. Accordingly, we assessed expression of D1/D5 dopamine receptors (using [3H]SCH-23390), of D2/D4 dopamine receptors (using [3H]raclopride), and of DAT (using [3H]mazindol) in membrane preparations from these two regions. As shown in Fig. 6, none of these dopaminergic markers were changed by chronic nicotine treatment across the dose range that altered nAChR-mediated synaptosomal [3H]dopamine release.

Figure 6.

Chronic nicotine administration does not affect expression of dopamine system markers. Expression of dopaminergic markers was assessed in striatum and olfactory tubercle membrane preparations of animals treated for 10 days with saline (control) or with the three indicated nicotine doses. Values obtained in striatal membranes are represented as black circles, whereas those measured in olfactory tubercles are shown as gray triangles (n = 6–7 per point; points represent mean ± SEM). Expression of D1/D5 receptors was measured using [3H]-SCH23390 (1.7 nM; top panel), whereas that of D2/D4 receptors was measured using [3H]-raclopride (17 nM; middle panel). Dopamine transporter (DAT) expression was assessed by binding of [3H]-mazindol (200 nM; lower panel). One-way anova was performed for each marker in each region to determine if chronic nicotine treatment significantly affected expression levels. No effect of nicotine dose was seen in any combination of region × marker.

Dopamine transporter function in striatum and olfactory tubercles is generally resistant to the effects of chronic nicotine treatment

In a further examination of possible chronic nicotine treatment effects on dopaminergic parameters, DAT activity was also measured in striatal and OT synaptosomal preparations. As shown in Table 3, almost all measures of DA transporter activity were unaffected by chronic nicotine treatment. The rate of [3H]-DA uptake was measured following 5 min incubations using of [3H]-DA concentrations at approximately Km (50 nM), or at a saturating concentration (1 μM). Rates of uptake were not altered at any chronic nicotine dose. Equilibrium [3H]-DA uptake was also measured at 30 min, using the same substrate concentrations. In striatum, total uptake was unaltered by chronic nicotine for either [3H]-DA concentration. A similar outcome was observed for total uptake of 50 nM [3H]-DA in OT. However, at the highest (4 mg/kg/h) chronic nicotine dose only, OT synaptosomal [3H]-DA uptake was significantly increased compared to the saline-treated control preparation.

Table 3. [3H]-DA Uptake after chronic treatment
Chronic TreatmentRate of uptake using 0.05 μM DA fmol/mg/minRate of uptake using 1.0 μM DA fmol/mg/minTotal uptake using 0.05 μM DA fmol/μgTotal uptake using 1.0 μM DA fmol/μgRatio of high DA/low DA for rateRatio of high DA/low DA for total
  1. Rate of uptake was determined from 5 min incubations. Total equilibrium uptake was determined at 30 min. Both assays were conducted at 22°C, with synaptosomal preparations from mice chronically treated with saline, 0.5 or 4 mg/kg/h nicotine (n = 6,7,6 mice, respectively). One-way anova and Duncan's post hoc test was used to identify any significant differences (*p < 0.05) treatment-induced differences from saline-treated control for each parameter within each region. Only one significant change was identified: an increase in total olfactory tubercle synaptosomal uptake following 4.0 mg/kg/h nicotine treatment (F2,17 = 7.671, p = 0.005). We have previously determined the Km for C57 mouse DA uptake to be 0.080 ± 0.003 μM DA; the ratio of ~ Km conc/ saturating conc gives an estimate of whether the Km has changed. Ratios are not different by region or treatment in two-way anova. A constant ratio of approximately two indicates that our estimate of Km is accurate and does not change under any of the experimental conditions.

Striatum
Saline (control)14.6 ± 0.843.4 ± 7.2181 ± 16378 ± 322.98 ± 0.462.15 ± 0.18
Nic 0.5 mg/kg/h15.2 ± 1.533.9 ± 2.4171 ± 25319 ± 242.47 ± 0.462.15 ± 0.36
Nic 4.0 mg/kg/h16.5 ± 3.439.4 ± 3.6232 ± 50459 ± 583.00 ± 0.852.50 ± 0.56
Olfactory tubercles
Saline (control)9.6 ± 2.326.7 ± 3.2131 ± 24208 ± 333.58 ± 0.971.70 ± 0.31
Nic 0.5 mg/kg/h10.2 ± 1.322.3 ± 3.1123 ± 20211 ± 252.21 ± 0.231.85 ± 0.24
Nic 4.0 mg/kg/h12.6 ± 2.032.9 ± 2.7166 ± 34349 ± 31*2.85 ± 0.342.57 ± 0.55

The use of two different [3H]-DA concentrations allows an estimate to be made of whether the DAT Km for DA uptake has changed. A ratio of ~ Km concentration / saturating concentration uptake should remain at approximately 2. As shown in Table 3, this is indeed the case; no significant changes in uptake ratio were observed for region or treatment for either rate or total amount of [3H]-DA uptake. We conclude that the DAT affinity for [3H]-DA uptake is not altered by chronic nicotine treatment in either region, under any of the conditions tested.

Discussion

This study represents the first comprehensive examination of dose–response relationships performed for down-regulation of murine [125I]α-CtxMII binding sites (corresponding to α6β2*-nAChR) by chronic nicotine treatment. Importantly, the ID50 values for down-regulation and the extent of down-regulation are very similar in the multiple nuclei in those dopaminergic and optic tracts responding to chronic nicotine treatment. Although previous studies have not examined the effect of nicotine dose on α6β2*-nAChR, where points of comparison are available, the current data (summarized in Table 1) generally agree well with those from previous studies. In striatum, down-regulation of α6β2* nAChR expression by chronic nicotine treatment has reliably been observed across multiple animal models. For example, in rat striatum declines of −3% (Nguyen et al. 2003), −33% (Mugnaini et al. 2006), −28% (Perry et al. 2007), and −27% (Doura et al. 2008) versus control were seen in α6β2*-nAChR expression frequently using nicotine administration (6 mg/kg/day) with osmotic minipumps. Owing to differences in metabolism, this treatment is estimated provide plasma nicotine concentrations comparable to those following treatment of mice with a dose of 2.5 mg/kg/h (Matta et al. 2007). The plasma nicotine level measured in one study was 1.91 μM (Doura et al. 2008) almost twice that attained in the plasma of mice treated with the highest nicotine dose (1.2 μM after 4 mg/kg/h) used in this study (Marks et al. 2004). Thus the apparently high doses that we used to treat the mice yield steady-state nicotine concentrations comparable to those achieved with the much lower doses used in rats. Declines have also been reported for striatal binding in mice chronically treated with nicotine: −30%, −28%, and −30% (Quik et al. 2012). Finally in squirrel monkey striatum, chronic nicotine treatment resulted in a −20% decline compared to control (McCallum et al. 2006). As another point of comparison, we observed little change in superior colliculus α6β2* nAChR expression following chronic nicotine administration. This is supported by three previous rat studies which also report modest declines: −2% (Perry et al. 2007), −10% (Mugnaini et al. 2006) and −2.9% (Doura et al. 2008). In this study, we show that relatively low doses elicit declines in α6β2*-nAChR expression (declines plateau after approx 0.5 mg/kg/h). The studies previously cited generally used doses comparable to our higher treatment doses, so decreases in expression can be compared to the maximum decreases observed in this work.

Consistent with numerous previous reports, chronic nicotine treatment up-regulated α4β2* nAChR expression. This phenomenon was first observed 30 years ago (Marks et al. 1983; Schwartz and Kellar 1983), and provides a valuable demonstration that the chronic nicotine treatment protocol worked as intended. The observed ED50 values for α4β2*-nAChR up-regulation (Table 1) also match well with our recent publication (Marks et al. 2011), which demonstrated that this phenomenon is caused by increased expression of nAChR protein. The ED50 value for α4β2* nAChR up-regulation by chronic nicotine treatment (95 nM) was ≈ 3x higher than the ID50 value for α6β2* down-regulation (35 nM).

The down-regulation or α6β2* nAChR sites is thus exceptionally sensitive. For comparison, plasma nicotine concentrations in smokers are typically 150–250 nM post-smoking, and fall as low as 6 nM following overnight abstinence (Jarvik et al. 2000). On this basis, α4β2*-nAChR will be exposed to a ≈ ED50 (for up-regulation) nicotine concentration for significant parts of the day, but well below overnight (although CNS levels will be somewhat higher than plasma levels). α6β2*-nAChR, however, will be at a > ID50 down-regulation concentration for the whole day and much of the night.

Both α6β2*- and α4β2*-nAChR are expressed on striatal and olfactory tubercle DA terminals. This allowed the use of the long-established synaptosomal [3H]-DA release technique to assess the impact of chronic nicotine treatment on α6β2*- and α4β2*-nAChR function. Mice were withdrawn from nicotine treatment for 2 h before tissue preparation, which should allow sufficient time between cessation of treatment and functional assessment for nicotine to be metabolized. Functional effects are thus a consequence of chronic treatment, not of residual nicotine. Intriguingly, DA-terminal α6β2*-nAChR function appeared to be more sensitive to reduction by chronic nicotine in striatum where DA terminals are projections from the SN than in the olfactory tubercles where terminals are from the VTA. Even in striatal samples, only the highest dose (4 mg/kg/h) affected the total nAChR-mediated [3H]-DA release. The decrease in function resembled that seen for high (single) doses tested in previous studies: −35% (Lai et al. 2005) and ≈ 50% (Quik et al. 2012) in mice, and −54% in rats (Perry et al. 2007). However, the chronic nicotine dose required to down-regulate α6β2*-nAChR function was much greater than that required to down-regulate expression. It therefore seems unlikely that loss of receptors per se is solely responsible for loss of function. This disconnect raised the question of whether changes in α6β2* nAChR composition might underlie the observed loss of function [perhaps because of changes in incorporation of α4 and/or β3 subunits that commonly co-assemble into α6β2*-nAChR (Gotti et al. 2005)]. As α6β2* nAChR are a mixed population of related subtypes, it is possible that individual subtypes may be differentially regulated by chronic nicotine treatment. If relevant subunit-specific antibodies were available to allow an immunostaining approach to be deployed, it would be possible to address this question directly. However, in their absence, radioligand autoradiography and functional assays were the best available approaches. It has been shown that α6β2β3* nAChR are more sensitive to down-regulation at high (560 nM) serum nicotine concentrations than are α6β2*(no β3)-nAChR (Perry et al. 2007). In addition, α6α4β2*-nAChR appear to be more sensitive to chronic nicotine down-regulation than are α6β2*(no α4)-nAChR (Perez et al. 2008). In contrast, our results indicated no significant changes EC50 values of nicotine evoked [3H]-DA release for either striatal or OT synaptosomes, at any chronic nicotine dose [as might be expected if subtype composition was changed following chronic treatment (Salminen et al. 2007)]. However, a small change in the ratio of populations with rather similar EC50 values (Salminen et al. 2007) may not be detectable. In addition, the total amount of nicotine evoked [3H]-DA release was unchanged for OT, but decreased for striatum. No previous studies exist to provide points of comparison, but the regional difference is striking. Perhaps the olfactory tubercle α6β2*-nAChR population more-closely resembles that in the treated striatum. However, striatal EC50 values tended to fall with chronic nicotine treatment, moving further away from those measured in OT, rather than converging. Alternatively, α6β2*-nAChR in the two regions may be subject to different regulation mechanisms, in which case the observed differences may be context rather than receptor-dependent.

The overall magnitude of effects on maximal α4β2*-nAChR function in striatum is less than for α6β2*-nAChR. This also corresponds well to results for mouse striatum from previous studies: no change (Lai et al. 2005) or a ≈ 30% decrease in [3H]-DA release at high stimulating agonist concentrations only (Quik et al. 2012). No functional change was seen in rat striatum (Perry et al. 2007). Again, no previous studies are available for comparison to our results in OT.

The changes in α6β2*- and α4β2*-nAChR-mediated [3H]-DA release following chronic nicotine treatment could be because of alterations in nAChR function, or to changes in dopaminergic system parameters. However, no changes in DA receptor or transporter expression levels were seen at any chronic nicotine treatment dose. Further, no evidence was seen for changes in DAT affinity for DA, and only in one set of conditions (total equilibrium uptake of [3H]-DA in olfactory tubercles following chronic treatment at the highest nicotine dose) was transporter activity significantly changed. We note that despite this change in DAT-mediated uptake, α6β2* nAChR-mediated [3H]-DA release from OT synaptosomes remained unchanged under the same chronic treatment regime, whereas [3H]-DA release mediated by α4β2* nAChR decreased similarly to striatum (Table 2). Although equivalent experiments have not been performed previously in OT, previous studies in striatum are in general agreement with our current observations. No changes in striatal DAT binding or overall DA levels were found following chronic nicotine treatment in squirrel monkeys (Perez et al. 2009), or for DAT binding or DA uptake as measured by voltammetry (Perez et al. 2012). A similar lack of effect on DAT binding levels was also observed in mice (Quik et al. 2012).

Taken together, our findings indicate that functional changes (as measured using synaptosomal [3H]-DA release) are due more directly to changes in nAChR function, than in dopaminergic function per se. Our work expands on previous studies by examining effects across a comprehensive chronic nicotine treatment dose range, and by measuring α6β2*- and α4β2*-nAChR function in OT as well as striatal preparations. Intriguingly, nAChR populations in these two regions respond differently (both in terms of changing expression levels and of function) to chronic nicotine treatment. This finding resembles earlier observations of disparate responses in different regions of non-human primate caudate-putamen. The observed significant differences in ED/ID50 values and direction for changes in α4β2* and α6β2* nAChR expression may result in notable changes in system/circuit equilibrium, as α4β2 nAChR are also expressed on GABA terminals that affect function in DA projection areas (English et al. 2011; Luo et al. 2013), whereas α6β2* nAChR are found only on the DA projections themselves (Gotti et al. 2005, 2010; Quik et al. 2005).

Acknowledgments and conflict of interest disclosure

This study was primarily supported by NIH grant R01 DA012242 (to PW). Additional support was provided by NIH grants R01 GM103801 and P01 GM48677 (to JMM).

All experiments were conducted in compliance with the ARRIVE guidelines. The authors have no conflict of interest to declare.

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