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Keywords:

  • β2;
  • EPSC;
  • FrA;
  • Fr2;
  • heteromeric;
  • 5IA85380;
  • M2;
  • nicotinic acetylcholine receptors;
  • premotor;
  • VGLUT1;
  • VGLUT2

ABSTRACT

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. REFERENCES

We studied how nicotinic acetylcholine receptors (nAChRs) regulate glutamate release in the secondary motor area (Fr2) of the dorsomedial murine prefrontal cortex, in the presence of steady agonist levels. Fr2 mediates response to behavioral situations that require immediate attention and is a candidate for generating seizures in the frontal epilepsies caused by mutant nAChRs. Morphological analysis showed a peculiar chemoarchitecture and laminar distribution of pyramidal cells and interneurons. Tonic application of 5 µM nicotine on Layer V pyramidal neurons strongly increased the frequency of spontaneous glutamatergic excitatory postsynaptic currents. The effect was inhibited by 1 µM dihydro-β-erythroidine (which blocks α4-containing nAChRs) but not by 10 nM methyllicaconitine (which blocks α7-containing receptors). Excitatory postsynaptic currents s were also stimulated by 5-iodo-3-[2(S)-azetidinylmethoxy]pyridine, selective for β2-containing receptors, in a dihydro-β-erythroidine -sensitive way. We next studied the association of α4 with different populations of glutamatergic terminals, by using as markers the vesicular glutamate transporter type (VGLUT) 1 for corticocortical synapses and VGLUT2 for thalamocortical projecting fibers. Immunoblots showed higher expression of α4 in Fr2, as compared with the somatosensory cortex. Immunofluorescence showed intense VGLUT1 staining throughout the cortical layers, whereas VGLUT2 immunoreactivity displayed a more distinct laminar distribution. In Layer V, colocalization of α4 nAChR subunit with both VGLUT1 and VGLUT2 was considerably stronger in Fr2 than in somatosensory cortex. Thus, in Fr2, α4β2 nAChRs are expressed in both intrinsic and extrinsic glutamatergic terminals and give a major contribution to control glutamate release in Layer V, in the presence of tonic agonist levels. Synapse 00:000–000, 2013. © 2013 Wiley Periodicals, Inc.


INTRODUCTION

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. REFERENCES

In the mammalian neocortex, acetylcholine (ACh) stimulates arousal and regulates attention and working memory (Jones, 2008; Sarter et al., 2006). ACh exerts its effects through metabotropic and ionotropic (nAChRs) receptors. The prominent role of the latter is increasingly recognized (e.g., Bailey et al., 2010; Guillem et al., 2011; Hahn et al., 2003; Howe et al., 2010), and stimulation of nAChRs is being considered for cognitive enhancing therapy in neurodegenerative and other neurologic diseases (Quik and Wonnacott, 2011; Williams et al., 2011). However, comprehending the physiological and pathological roles of nAChRs in the neocortex is complicated because of the diversity of nAChR subtypes and their expression in different cell types and compartments (Dani and Bertrand, 2007). What is more, different neocortical regions are anatomically and functionally heterogeneous (Franklin and Chudasama, 2012; Palomero-Gallagher and Zilles, 2004).

The nAChR is a pentamer of subunits surrounding a pore permeable to cations. In the mammalian brain, nine gene products concur in forming functional receptors: α2-α7 and β2-β4. In neocortex, common nAChR subtypes are the heteropentamer α4β2 and the homopentamer α7, which are characterized by different permeability to Ca2+, desensitization rate, and sensitivity to the agonists (Dani and Bertrand, 2007). In primates' neocortex, the distribution of homo- and heteromeric nAChRs is broadly similar to that observed in rodents, although a higher contribution of other subunits, such as β4, is observed (Hellström-Lindahl et al., 1998; Quik et al., 2000). Widespread expression of β4 has also been detected in the mouse (Gahring et al., 2004). In prefrontal cortex, homo- and heteromeric nAChRs cooperate in regulating the cognitive effects of nicotine, but the specific roles of different subtypes are still matter of debate (Dani and Bertrand, 2007; Gotti et al., 2009).

The nAChRs can be expressed at synaptic and extrasynaptic sites. Accordingly, different lines of evidence indicate that low tonic ACh levels exert important modulatory roles by diffuse transmission (Lendvai and Vizi, 2008), in agreement with behavioral studies showing that ACh regulates attentive behavior at widely different time scales (Parikh et al., 2007). Heteromeric nAChRs are thought to give a major contribution to such sustained control of neocortical excitability, as they display higher sensitivity to the agonists and slower desensitization, compared with the homomeric forms. Consistently, the large majority of mutations known to be linked to autosomal dominant nocturnal frontal lobe epilepsy and affine pathologies map on genes coding for non-α7 nAChR subunits (Aridon et al., 2006; Becchetti, 2012; De Fusco et al., 2000; Phillips et al., 2001; Steinlein et al., 1995).

Control of neocortical excitability depends on a fine balance between excitatory and inhibitory transmission, and nAChRs are known to regulate both. For coherence with literature, we retain the expression “prefrontal cortex,” although its appropriateness in rodents has been questioned, because of the difficulties of making comparisons with the immensely more complex dorsolateral prefrontal cortex in primates (Uylings et al., 2003). Expression of nAChRs on the soma of different interneuronal populations is established in the prefrontal cortex of rats (Christophe et al., 2002; Porter et al., 1999; Xiang et al., 1998), humans (Alkondon et al., 2000), and mice (Brown et al., 2012; Couey et al., 2007). Evidence of heteromeric nAChR expression in GABAergic terminals is also available, at least in mouse (Aracri et al., 2010; Klaassen et al., 2006).

Knowledge about the nicotinic control of glutamatergic transmission is more fragmentary. In the rat, nicotine stimulates the thalamocortical (TC) fibers projecting to superficial layers in prefrontal (Gioanni et al., 1999; Vidal and Changeux, 1993) and somatosensory (SS) cortex (Gil et al., 1997). Similar results were obtained in Layer V of rats and mice, and the effect was mostly inhibited by blocking heteromeric nAChRs (Lambe et al., 2003). On the other hand, in the visual cortex of monkeys (Disney et al., 2007) and tree shrew (Bhattacharaya et al., 2012), the functional effects of nAChRs are more prominent in Layer IV, the main target of TC projection. Glutamate release is also likely to be modulated by nAChRs regulating local pyramidal cell excitability, as somatic nicotinic currents have been measured in pyramidal neurons from deep layers (Kassam et al., 2008; Poorthuis et al., 2012; Zolles et al., 2009), at least in rodents.

Previous studies in nonsensory regions generally focused on the medial prefrontal cortex (particularly on its subdivision named prelimbic area), whose laminar structure and connectivity are, however, rather heterogeneous (Franklin and Chudasama, 2012). In the mouse, the morphology and connections of the different prefrontal regions are only beginning to be studied in detail (Franklin and Chudasama, 2012; Paulussen et al., 2011; Van de Werd et al., 2010). This is a serious gap in knowledge because physiological and pharmacological studies carried out in different regions may not be directly comparable and may have different functional implications. Such complexity hampers our interpretation of the ever increasing body of evidence obtained using murine models of human neuropathology.

In the present article, we focus on the dorsomedial shoulder of the murine prefrontal cortex, recently indicated as “secondary motor area” (M2; Franklin and Chudasama, 2012). This region is also frequently called frontal area 2 (Fr2), following the criteria applied to the rat prefrontal cortex (Heidbreder and Groenewegen, 2003; Palomero-Gallagher and Zilles, 2004; Uylings et al., 2003) and has been also referred to as precentral cortex or dorsomedial prefrontal cortex. Because it displays features related to the more complex dorsolateral prefrontal cortex in humans, with partial overlap with higher association functions (Uylings et al. 2003), it has been also named FrA. From a functional point of view, Fr2 has been rather neglected until now. It is distinct from other prefrontal areas because it is innervated by sensory and parietal regions of the cortex. This pattern of connections likely enables Fr2 to properly respond to situations that require immediate attention (Hoover and Vertes, 2007). Work in mice indicates that Fr2 is also implicated in reward-guided decision making (Kargo et al., 2007). In addition, this area projects to the motor cortex and dorsolateral striatum (Berendse et al., 1992; Condé et al., 1995). Therefore, we believe detailed studies about the nicotinic regulation of synaptic function in Fr2 could also help defining how expression of mutant nAChRs can cause the typical hypermotor seizures observed in patients with autosomal dominant nocturnal frontal lobe epilepsy (Picard and Brodtkorb, 2008).

We first carried out a broad chemoarchitectonic characterization of this region. Next, we assessed by patch-clamp methods the steady state contribution of α4β2- and α7-containing nAChRs to the regulation of glutamate release in Layer V. This is the main subcortical output layer of the neocortex and is particularly susceptible to seizure development (Richardson et al., 2008). Finally, we studied the distribution of α4-containing nAChRs in Fr2 and their association with the main populations of glutamatergic axon terminals, identified by labeling the vesicular glutamate transporters (VGLUT) type 1 and type 2 (VGLUT2). These mark the two main populations of classical asymmetric glutamatergic synapses in the neocortex (Graziano et al., 2008; Hur and Zaborsky, 2005; Nakamura et al., 2005). VGLUT1 mostly labels the intrinsic glutamatergic terminals or cortical afferents, whereas VGLUT2 tends to label the thalamic afferents (Graziano et al., 2008). In the primary somatosensory cortex, 96% of the VGLUT2 fibers project from the thalamus, whereas the percentage is approximately 90% in the medial prefrontal cortex; the other VGLUT2 fìbers present in the neocortex originate from other subcortical regions, mainly the hypothalamus (Hur and Zaborsky, 2005).

MATERIALS AND METHODS

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. REFERENCES

Experiments were carried out on FVB mice (Harlan, Italy) according to the Principles of Laboratory Animal Care (86/609/EEC Directive). All efforts were made to minimize animal suffering and reduce the number of animals used. Unless otherwise indicated, chemicals and drugs were purchased from Sigma-Aldrich (Italy).

Definition of the neocortical regions

For SS cortex, sections were chosen between +0.50 and +0.02 mm from bregma (primary SS cortex, S1), according to Paxinos and Franklin (2001). For the prefrontal cortex, no general agreement about nomenclature of the different neuroanatomical subdivisions has yet been reached. For both morphological and electrophysiological experiments, we studied the part of the dorsomedial shoulder region that was recently indicated by Franklin and Chudasama (2012) as “secondary motor area” (M2). Our sections were thus prepared between +2.68 and +2.10 mm from bregma. This region is referred to in the text as Fr2. The reader should be aware that, in the atlas of Paxinos and Franklin (2001), Fr2 covers both the medial part of “frontal association cortex” (FrA, rostrally) and the secondary motor area (M2, caudally). Moreover, a recent cytoarchitectonic characterization of mouse prefrontal cortex labels as Fr2 a more medial region than we have selected (Fig. 6A) and considered it as “agranular cortex” (Van de Werd et al., 2010).

Brain slice preparation for electrophysiology

Mice (aged 17–24 days) were deeply anesthetized with ether and decapitated. Brains were removed and placed in ice-cold solution containing (in mM): 87 NaCl, 21 NaHCO3, 1.25 NaH2PO4, 7 MgCl2, 0.5 CaCl2, 2.5 KCl, 25 d-glucose, and 75 sucrose, equilibrated with 95% O2 and 5% CO2 (pH 7.4). Coronal slices (300 µm thick) were cut with a VT1000S vibratome (Leica Microsystems, Germany) and incubated at room temperature for at least 1 h, in the same solution as above, before being transferred to the recording chamber.

Whole-cell recordings and data analysis

Neurons were voltage- or current-clamped with a Multiclamp 700A patch-clamp amplifier (Molecular Devices, CA) at room temperature. Low-resistance borosilicate micropipettes (2–3 MΩ) were pulled with a P-97 Flaming/Brown Micropipette Puller (Sutter Instrument, CA). The cell capacitance and series resistance (up to 75%) were always compensated. Series resistance was usually below 10 MΩ. Input resistance was generally close to 100 MΩ. Synaptic currents were low-pass filtered at 2 kHz and digitized at 5 kHz, with pClamp/Digidata 1322A (Molecular Devices). Pipettes contained (in mM) 135 K-gluconate, 5 KCl, 1 MgCl2, 10 HEPES, 2 MgATP (pH 7.2). During experiments, slices were superfused at 1.8 mL/min with the following solution (in mM): 129 NaCl, 21 NaHCO3, 1.6 CaCl2, 3 KCl, 1.25 NaH2PO4, 1.8 MgSO4, and 10 d-glucose, aerated with 95% O2 and 5% CO2 (pH 7.4). Cells were inspected with an Eclipse E600FN (Nikon Instruments, Italy) equipped with a water immersion differential interference contrast objective and an infrared CCD 100 camera (DAGE-MTI, IN). Resting Vm was measured in open circuit mode, soon after obtaining the whole-cell configuration.

The frequency and peak amplitude of excitatory postsynaptic currents (EPSCs) were analyzed offline with Clampfit 9.2 (Molecular Devices) and OriginPro 8 (OriginLab, MA) software. The spontaneous EPSCs included both smoothly shaped isolated signals and composite signals. Events were inspected one by one, and those not presenting the typical shape of synaptic currents were rejected. The baseline noise (peak to peak) was generally lower than 5 pA. The threshold was generally set around 7–8 pA. Agonists and antagonists were perfused in the bath and their effect calculated at the steady state, which was usually reached within 2 min.

Stock solutions of (−)-nicotine hydrogen tartrate salt, dihydro-β-erythroidine (DHβE) hydrobromide, methyllicaconitine (MLA) citrate hydrate and AP5 [d(−)-2-amino-5-phosphono-pentanoic acid] were prepared in distilled water. Stock solutions of 20mM 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX) were prepared in dimethyl sulfoxide. 5-Iodo-3-[2(S)-azetidinylmethoxy]pyridine (5IA85380; Tocris Bioscience, Bristol, UK) was dissolved in our standard extracellular solution and used at the indicated final concentration.

Primary antisera

A polyclonal antibody made in guinea pig, against the synthetic peptide corresponding to amino acids 568–588 of the rat nAChR α4 protein, (AB5590; Chemicon International, CA; diluted 1:2000) was chosen among different commercially available antibodies. Its suitability for our purposes was previously extensively assessed (Aracri et al., 2010; Nashmi et al., 2003; Zarghooni et al., 2007). In particular, we confirmed that AB5590 does bind to α4-containing nAChRs in human embryonic kidney cells transiently transfected with different combinations of α4, β2, α2, and β4 subunits (Aracri et al., 2010). Two commercial antibodies against VGLUT1 and VGLUT2 (Synaptic Systems, Germany; polyclonal, made in rabbit, dilutions: 1:600 anti-VGLUT1, 1:500 anti-VGLUT2) were used to identify the corresponding glutamatergic synaptic terminals. These antisera were raised against Strep-TagR-fusion proteins containing, respectively, the amino acid residues 456–560 of rat VGLUT1/BNPI (brain-specific Na+-dependent inorganic phosphate transporter) and the residues 510–582 of rat VGLUT2/DNPI (differentiation-associated Na+-dependent inorganic phosphate cotransporter). These antigens were used as control proteins diluted 1:250 in our preadsorption experiments on cortical sections of murine brain. Moreover, these three antibodies have been tested by western blot (WB) on murine cortical tissue (see Results).

To perform both a preliminary neurochemical analysis of Fr2 and double/triple labeling with immunofluorescence in Layer V, we also used two monoclonal mouse antibodies against (i) the nonphosphorylated epitope of the neurofilament heavy subunit (SMI32; Sternberger Monoclonals, MD; diluted 1:1000) to detect pyramidal cells and (ii) the calcium-binding protein parvalbumin (PV; Sigma Aldrich; diluted 1:2000) to mark the main cortical interneuronal population.

Cortical tissue preparation for WB

Mice (n = 4) were anesthetized with diethyl ether and killed by decapitation. Brains were quickly removed and dissected. After removing the ventral noncortical structures, two cortical blocks for SS and Fr2 were prepared following approximate stereotaxic coordinates from bregma (Paxinos and Franklin, 2001). For Fr2, cortical tissue was selected between +3.00 mm and 1.00 mm from bregma; for SS, between +1.00 mm and −1.50 mm. Prefrontal and SS cortices were rapidly separated with fine jeweller's forceps and frozen at −80°C until use. These tissues were washed three times with 50mM Tris buffered solution (TBS; pH 7.5) and then homogenized in the same buffer containing a general-purpose protease inhibitor cocktail (Mini Protease Inhibitor Mixture; Roche Applied Science, Italy) using a Teflon-on-glass homogenizer. Homogenates were centrifuged at 1000g for 5 min at 4°C, and the pellets were resuspended according to Eukaryotic Membrane Protein Extraction Reagent Kit (Pierce Biotechnology, IL) to separate the membrane proteins from the intracellular proteins. Proteins were quantified by the Bradford method, with AppliChem (Gatersleben, Germany) solution. Samples containing 50 mg of total proteins were heated to 100°C for 5 min in sample buffer containing 0.6 g/100 mL Tris, 2 g/100 mL sodium dodecyl sulphate (SDS), 10% glycerol, 1% β-mercaptoethanol, pH 6.8, 0.03% bromophenol blue. Next, they were loaded onto 10% SDS-polyacrylamide gel electrophoresis (PAGE) gel and run at 60 mA until the dye front reached the bottom of the gel, and finally electrotransferred to nitrocellulose membranes (Mini Trans-blot; Bio-Rad Laboratories, CA) in 20% methanol/25 mM Tris/192 mM glycine at 70 V for 2 h, at room temperature. Nitrocellulose membranes were treated for 2 h at room temperature with a blocking solution containing 5% nonfat dry milk in TTBS (TBS supplemented with 0.1% Tween 20), to block nonspecific protein-binding sites. They were then incubated overnight at 4°C with the primary antibodies (see above) diluted in blocking solution. Subsequently, blots were rinsed and incubated for 1 h at room temperature with the following secondary biotin-conjugated antibodies: donkey anti-guinea pig (Sigma Chemical, MO; 1:2000), to identify the anti-α4 nAChR subunit; goat anti-rabbit (Sigma Chemical; 1:2000), to identify the anti-VGLUT1 and anti-VGLUT2. The immunoreactive regions were detected by alkaline phosphatase conjugated streptavidin (Streptavidin-AP; Vector Laboratories, CA; 1:2000), at room temperature for 2 h; washed twice in TTBS and once in TBS and finally developed in AP substrate: 100mM Tris-HCl, 100mM NaCl, 5mM MgCl2, 150 µg/mL 5-bromo-4-chloro-3-indolyl phosphate (Sigma Chemical), 30 µg/mL nitroblue tetrazolium (Sigma Chemical), pH 9.5. The same membranes were subsequently processed with the three primary antibodies to detect the levels of nicotinic subunit or glutamate transporters and actin (using a monoclonal anti-actin antibody; Sigma Aldrich) in the same samples loaded on the gel. To this purpose, primary and secondary antibodies were removed from the membrane by incubation for 15 min at 37°C in Restore WB stripping buffer (Pierce Biotechnology). Membranes were then washed twice with TTBS and processed as described above. The optical intensity of WBs was detected and quantified by densitometric analysis (NIH ImageJ software) on scanned films. Immunoreactivity with anti-α4 nAChR subunit, anti-VGLUT1, and anti-VGLUT2 was normalized to the one obtained with anti-actin, as protein loading control. The following negative controls were carried out: omission of the primary antiserum and preadsorbtion of the primary antiserum with the corresponding 5 mM synthetic peptide, for 12 h at 4°C before use.

Cortical tissue preparation for immunocytochemistry

Five mice (aged 21–40 days) were anesthetized with intraperitoneal 4% chloral hydrate (2 mg/100 g) and sacrificed by intracardiac perfusion with 4% paraformaldehyde, in phosphate buffer (PB) 0.1M (pH 7.2–7.4). Perfusion of paraformaldehyde was preceded by heparin followed by 1% paraformaldehyde in PB. Such a procedure allows to maintain adequate tissue antigenicity. Brains were removed from skulls and immersed in 4% paraformaldehyde in PB, at 4°C, overnight. Coronal sections (50 µm thick) were cut with a VT1000S vibratome (Leica Microsystems) from Fr2 and SS cortex. For each cortical region, three to four sections were selected for immunocytochemistry. Cytoarchitecture controls were performed on alternate sections adjacent to those processed for immunocytochemistry and stained with thionin.

Immunoperoxydase histochemistry

After aldehyde quenching with NH4Cl (0.05M in phosphate buffered saline, PBS) for 30 min and inactivation of endogenous peroxidases with H2O2, sections were simultaneously permeabilized with 0.2% Triton X-100 and treated to block the unspecific interaction sites for 30 min with bovine serum albumine (BSA; 1% in PBS). Next, they were incubated overnight with the primary antibody diluted in 0.1% BSA, at room temperature. This procedure was followed by incubation with either biotinylated donkey anti-guinea pig (for anti-α4 nAChR subunit), horse anti-mouse (for SMI32 and anti-PV), or goat anti-rabbit (for anti-VGLUT1/2) IgG (Vector Laboratories; diluted 1:200), for 75 min. After washing, sections were treated with the avidin-biotinylated complex (ABC kit; Vector; diluted 1:100) and then with a freshly prepared solution (0.075%) of 3-3′-diaminobenzidine tetrahydrochloride and 0.002% H2O2. Finally, sections were mounted, dehydrated, and laid on coverslips. The specificity of primary antibodies was assessed by negative controls: omission of primary antiserum and preadsorbtion of primary antiserum with the corresponding 5 mM synthetic peptide, when available (see Primary antisera). In these cases, no specific staining was ever observed. Specificity for the same antibodies was also tested by immunofluorescence (see later).

Immunofluorescence

Fluorescent labeling was studied with confocal microscopy, which provides the resolution necessary to distinguish both the cell bodies and the synaptic boutons marked by VGLUT expression. After aldehyde quenching with NH4Cl for 30 min, sections were rinsed with PB. They were next treated with BSA (1%) and Triton X-100 (0.2%) in PBS for 30 min. Finally, sections were washed with PBS and incubated for two nights with the following primary antisera (diluted in 0.1% BSA, at 4°C): (A) anti-VGLUT1 and anti-α4 nAChR subunit; (B) anti-VGLUT2 and anti α4 nAChR subunit; (C) anti-VGLUT1 or anti-VGLUT2, anti-α4 nAChR subunit, and SMI32. After washing with PBS, the following mixture of secondary antibodies (diluted in the same solution as the primary) conjugated with appropriate fluorochromes was added for 75 min: (i) GAR- Cy3 (anti-rabbit IgG, made in goat, diluted 1:200, conjugated to the the fluorochrome indocarbocyanine Cy3; Zymed Laboratories, CA); (ii) DAGp-Cy2 (polyclonal, anti-guinea pig IgG, made in donkey; 1:200 dilution; conjugated to the indocarbocyanine Cy2 fluorochrome; Jackson Immunoresearch Laboratories, PA). Only for triple immunolabeling, alternative secondary antibodies were used for monoclonal SMI32 and for anti-VGLUT1/2: namely DAMCy3 (polyclonal, anti-mouse IgG, made in donkey; 1:200 dilution; Cy3 fluorochrome; Jackson Immunoresearch Laboratories) and DAR-Cy5 (polyclonal, donkey anti-rabbit IgG conjugated to the indocarbocyanine Cy5, 1:200, Jackson Immunoresearch Laboratories), respectively. After rinsing, samples were mounted on coverslips with PBS/glycerol and inspected with a TCS-NT (Leica Laserteknik GmbH) laser scanning confocal microscope, to visualize double fluorescent labeling. In Figure 9, the original emission colour of some fluorochromes conjugated to secondary antibodies has been changed to facilitate visual inspection of confocal images (VGLUT1/2 was set to red; SMI32 was set to blue). Immunoreaction specificity was tested by carrying out negative controls as previously explained. In this case, no specific signal was ever observed.

Colocalization analysis

Confocal micrographs were collected and digitized as TIFF files with the Leica confocal scanning microscope (Leica Microsystems) and software package (Leica Confocal Software). Identical parameters were used to acquire images from the Fr2 and SS areas. Digital micrographs were optimized for resolution (final resolution 300 dpi), brightness, and contrast using Adobe Photoshop CS2 9.0 (Adobe Systems, San Jose, CA). Images were not altered in any way, e.g., by removing or adding image details. Nonoverlapping pictures (40× objective, 1.8× zoom) were acquired in Layer V to sample the double immunolabeling in at least two different sections for each cortical region (i.e., three to four cortical fields were analyzed for each cortex and each animal). For two-color analysis, tissues were excited at 488 and 568 nm and fluorescence of Cy2 (green) and Cy3 (red) were detected sequentially to avoid crosstalk. Colocalization of α4 nAChR subunit with either VGLUT1 or VGLUT2 was also determined by Leica Confocal software, by generating the 2D cytofluorograms (Bolte and Cordelières, 2006) shown in Figures 7 and 8. Generation of a binary mask of image data allowed to select in a region of interest certain intensity value pairs in the cytofluorograms and to identify them as white signal in the original images, representing both single and merged antigen localization (Figs. 7-9).

Statistical analysis

Patch-clamp data are given as mean values ± standard error of the mean. Statistical significance of averages from several cells was determined with paired Student t-test (with significance level set to 0.05), which assumes that the differences are sampled from a Gaussian distribution. This assumption was tested with the Kolmogorov-Smirnov (KS) method. The distribution of EPSC frequency (or amplitude) obtained in each cell in different experimental conditions was also evaluated with the KS test (significance level set to 0.01). In the figures, asterisks mark the level of significance; in the text, NS denotes nonsignificant effects.

RESULTS

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. REFERENCES

We studied the Fr2 part of the dorsomedial shoulder region of murine prefrontal cortex (Franklin and Chudasama, 2012). Contrary to the more medial regions, this area presents evident lamination, with distinction of a very thin Layer IV enclosed between Layers III and V (e.g., Fig. 6), as was also previously observed in the rat prefrontal regions (Palomero-Gallagher and Zilles, 2004). In brain slices obtained from Fr2, we first addressed by patch-clamp methods the nicotinic regulation of glutamate release in Layer V.

Nicotine stimulates the EPSCs recorded from pyramidal neurons

Pyramidal neurons were identified based on their electrophysiological and morphological features, as previously reported (Aracri et al., 2010). In brief, the neurons considered in Figures 1, 2, 4 had an average resting Vm of −70.0 ± 0.51 mV (n = 27). When tested with depolarizing current pulses, they displayed the typical low frequency action potential firing with adaptation (Chang and Luebke, 2007; Connors and Gutnick, 1990; Porter et al., 1999).

In pyramidal neurons, we studied the effect of nicotine on spontaneous EPSCs in whole-cell mode. Nicotine was preferred to the physiological agonist ACh, because our main goal was assessing the specific involvement of nAChRs, rather than studying the global ACh effect. Moreover, using nicotine avoids the necessity of applying muscarinic receptor blockers, which are known to also affect nAChRs at relatively low concentrations (Zwart and Vijverberg, 1997). EPSCs were registered as inward events at −70 mV, close to the Vrev of inhibitory postsynaptic currents (Aracri et al., 2010). In this way, we also avoided using blockers of GABAergic currents, thus maintaining as far as possible the physiological conditions in the local microcircuit. After obtaining the whole-cell configuration, we recorded for 4–5 min the spontaneous EPSCs. Next, nicotine was applied in the bath for 5 min and then removed. Typical current traces are shown in Figure 1A. The glutamatergic nature of EPSCs was confirmed by the full inhibition produced by AP5 (50 µM) and CNQX (10 µM), which block the NMDA and the AMPA receptors, respectively. Nicotine was applied at 5 µM, a concentration close to the peak of the “window current” for both α4β2 and α7 nAChRs (Fenster et al., 1997). These experimental conditions favor maximal steady state activation of both hetero- and homomeric nAChRs, which are known to coexist in prefrontal regions (e.g., Dickinson et al., 2008; Marchi et al., 2002). In general, nicotine produced a robust increase of the EPSC frequency, whereas scarce effects were observed on the events' amplitude. Figures 1B and 1C illustrate the cumulative distribution of the EPSC amplitudes and interevent intervals for a typical experiment, respectively. Detailed statistics are given in the figure legend. The time course of a representative experiment is shown in Figure 1D, in which bars represent the number of synaptic events calculated in 1-min intervals for the entire experiment. Stimulation usually peaked within 2–3 min and produced an approximate threefold increase in the EPSC frequency. The average effect of nicotine in a series of similar experiments is illustrated in Figure 1E, which shows the average EPSC frequency calculated for 2 min at the steady state, in the indicated conditions. Stimulation brought the EPSC frequency from 5.06 ± 1.03 Hz (control) to 11.3 ± 2.75 Hz (nicotine; P < 0.05, with t-test; n = 7).

image

Figure 1. Nicotine stimulates EPSCs on pyramidal neurons in Layer V. A: Representative EPSC traces, recorded on pyramidal neurons, at −70 mV. Nicotine (5 µM) increased the EPSC frequency, in a reversible way. All events were abolished by AP5 (50 µM) and CNQX (10 µM), as indicated. B: Amplitude distribution of the EPSCs for the same cell, registered during 2 min, in the indicated conditions, at the steady state. In six of eight neurons tested, 5 µM nicotine produced no increase in the EPSC amplitude (not significant with KS test). In the other experiments, a slight increase (less than 15%) in average EPSC amplitude was observed. C: Distribution of the interevent intervals, for the same experiment. The EPSC frequency measured during 2-min intervals at the steady state was increased by nicotine from 4.302 ± 0.15 to 10.935 ± 0.1 (P < 0.01 with KS test). Similar results were obtained in six of eight neurons. D: Typical time course of the EPSC frequency. Bars give the number of synaptic events measured within consecutive 1-min intervals, from an experiment similar to the one shown in (A), except that AP5 and CNQX were not applied. E: Population effect of nicotine on cells treated as illustrated in the previous panels. Bars give the mean EPSC frequency, measured during 2 min at the steady state, in the indicated conditions. On average, nicotine (5 µM) more than doubled the frequency of synaptic events, bringing it from 5.06 ± 1.03 Hz (control) to 11.3 ± 2.75 Hz (nicotine; P < 0.05, with paired t-test; n = 7). The effect was reversible on wash out.

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The effect of nicotine is inhibited by DHβE but not MLA

The contribution of the main nAChR subtypes expressed in the murine neocortex can be distinguished by using 1 µM DHβE, which blocks α4-containing nAChRs and 10 nM MLA, which blocks α7-containing receptors (Harvey et al., 1996; Alkondon et al., 1999; Murray et al., 2012). We tested the effect of these inhibitors in experiments analogous to those illustrated in Figure 1. Figure 2A shows the current traces from a representative experiment in which we tested 1 µM DHβE. Application of this compound for 3–4 min did not alter the EPSC amplitude or frequency, indicating that DHβE does not produce any nonspecific effect. The time course of a typical experiment is shown in Figure 2D, in which bars represent the number of synaptic events calculated in 1-min intervals for the entire experiment. The average effect of DHβE in a population of neurons treated in the same way is shown in Figure 2E. When 5 µM nicotine was added in the presence of DHβE, the stimulation on EPSCs was largely prevented, as the average EPSC frequency calculated for 2 min at the steady state was not different in the presence or absence of nicotine. In particular, the EPSC frequency was 5.71 ± 1.15 Hz before treatment with nicotine and was 6.64 ± 1.55 Hz in the presence of nicotine plus DHβE (NS with paired t-test; n = 7). Further statistics are given in the figure legend.

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Figure 2. The effects of nicotine are antagonized by DHβE. A: Representative EPSC traces, recorded on pyramidal neurons, at −70 mV. Current traces are shown in control condition, in presence of 1 µM DHβE, in presence of DHβE plus 5 µM nicotine, and after wash out, as indicated. B: EPSC amplitude distribution for the same experiment (calculated for 2 min, at the steady state). In the presence of DHβE, nicotine produced no significant increase of the EPSC amplitude, as tested with the KS test in seven analogous experiments. C: The corresponding interevent interval distribution shows no significant alteration in EPSC frequency in the presence of nicotine plus DHβE compared with treatment with DHβE alone (as tested with the KS test). Similar results were obtained in seven experiments. D: Time course of the EPSC frequency during the entire experiment. Bars give the number of synaptic events measured within consecutive 1-min intervals. Different treatments (DHβE and DHβE plus nicotine) were applied for 4–5 min, as indicated. E: Population effect of nicotine on cells treated with DHβE as illustrated in the previous panels. Bars give the mean EPSC frequency, measured during 2 min at the steady state, in the indicated conditions. In particular, the EPSC frequency was 7.06 ± 1.58 Hz in the controls, 5.71 ± 1.15 Hz in the presence of DHβE (not significantly different with paired t-test; n = 7), and 6.64 ± 1.55 Hz in the presence of nicotine plus DHβE (not significantly different with paired t-test; n = 7). No significant alteration of EPSC frequency was observed after wash out.

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In contrast, MLA (10 nM) turned out to be considerably less effective than DHβE in antagonizing the effect of nicotine. Figure 3A shows typical current traces registered in the indicated conditions. Application of 5 µM nicotine in the presence of MLA produced a reversible increase in the EPSC frequency, which was reflected in the cumulative distribution of the interevent intervals (Fig. 3C). In agreement with the results we obtained with nicotine alone, the EPSC frequency increase in the presence of MLA was not accompanied by a significant effect on EPSC amplitudes (Fig. 3B). The time course of a typical experiment is shown in Figure 3D. The average EPSC frequency in series of similar experiments, measured during 2 min continuous recording at the steady state for each indicated condition, is shown in Figure 3E. In the presence of MLA, nicotine almost doubled the average EPSC frequency. The effect was fully reversible. Hence, DHβE was much more effective than MLA in counteracting the tonic effect of nicotine on the spontaneous EPSCs.

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Figure 3. The effects of nicotine are not inhibited by MLA. A: Representative EPSC traces, recorded on pyramidal neurons, at −70 mV. Current traces are shown in control condition, in presence of 10 nM MLA, in presence of MLA plus 5 µM nicotine, and after wash out, as indicated. B: Amplitude distribution of the EPSCs for the same cell, registered during 2 min, in the indicated conditions, at the steady state. In the presence of MLA, nicotine produced no significant increase of the EPSC amplitude (as tested with the KS test). Similar results were obtained in seven analogous experiments. C: Distribution of the interevent intervals, for the same experiment. In the presence of MLA, the EPSC frequency was significantly increased by nicotine, as shown by the distribution of the interevent intervals (P < 0.01 with the KS test). Analogous results were obtained in six of seven similar experiments. D: Time course of the EPSC frequency in representative experiments. Bars give the number of synaptic events measured within consecutive 1-min intervals. Different treatments (MLA and MLA plus nicotine) were applied for 4–5 min, as indicated. E: Population effect of nicotine on cells treated with MLA as illustrated in the previous panels. Bars give the mean EPSC frequency, measured during 2 min at the steady state, in the indicated conditions. The EPSC frequency was 4.62 ± 0.34 Hz in control condition; 4.36 ± 0.4 Hz in the presence of MLA alone (NS with t-test; n = 7); and 8.35 ± 1.7 Hz in the presence of nicotine plus MLA (P < 0.05 with t-test, compared with the controls; n = 7); after wash out, the residual EPSC frequency was 4.04 ± 0.66.

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Figure 4. 5IA85380 stimulates EPSCs in Fr2. Panel shows the average effect of IA585380 (10 nM), a specific agonist of β2-containing nAChRs, on a population of experiments. Spontaneous EPSCs were registered and analyzed as illustrated in the previous figures. Bars give the mean EPSC frequency, measured during 2 min at the steady state, in the indicated conditions. The effects produced by 10 nM IA585380 were abolished by 1 µM DHβE. The EPSC frequency was 3.67 ± 1.2 Hz in control condition; 6.35 ± 3.1 Hz in the presence of IA585380 (P < 0.05 with t-test, compared with the controls; n = 5); and 2.94 ± 0.5 Hz in the presence of IA585380 plus DHβE.

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Effect of the specific nAChR agonist 5IA85380

The results illustrated so far suggest that heteromeric α4-containing nAChRs regulate glutamate release in Layer V, in the presence of tonic low agonist concentrations. To corroborate this hypothesis and further define the involved nAChR subtypes, we studied the effect on EPSCs of 5IA85380 (Mukhin et al., 2000), which is a highly specific agonist for β2-containing receptors, at concentrations around 20 nM (Mogg et al., 2004). These results are shown in Figure 4. EPSCs were registered and analyzed as discussed above. For brevity, we only show the average effects obtained from an ensemble of neurons. At 10 nM, 5IA85380 approximately doubled the EPSC frequency. This submaximal concentration was chosen to avoid possible interference from β4-containing nAChRs (Mogg et al., 2004). The stimulation produced by 5IA85380 was abolished by adding 1 µM DHβE. 5IA85380 is also known to activate α6β2* receptors (Mogg et al., 2004), which are however not significantly expressed in prefrontal regions (Gotti et al., 2009). Therefore, we attribute the effect of 5IA85380 to activation of α4β2 nAChRs. These are known to comprise two main subtypes: high-affinity α42β23 and low-affinity α43β22 (Nelson et al., 2003). Concentrations of 5IA85380 lower than 100 nM essentially activate the high affinity α42β23 (Li and Steinbach, 2010; Zwart et al., 2006). Considering that the effect of 5IA85380 was comparable with the one produced by 5 µM nicotine (Fig. 1), which is a quasi-saturating concentration for the high-affinity receptors, we conclude that the steady state regulation of glutamate release in Fr2 can be largely attributed to the action of α42β23 nAChRs.

Expression of α4 nAChR subunit in glutamatergic terminals in Fr2 and SS cortex

The glutamatergic terminals in prefrontal areas include both intrinsic fibers and extrinsic TC fibers. We recall that the spontaneous EPSCs comprise miniature events that depend on spontaneous release from presynaptic terminals and larger events that depend on action potential-dependent stimulation of these terminals (Wong et al., 2000). The nicotine-dependent increase of EPSC frequency shown in Figure 1 resembles the one previously observed in medial prefrontal cortex (Couey et al., 2007; Lambe et al., 2003), in which, however, nAChR activation also produces a systematic increase of EPSC amplitude. This effect was mostly attributed to extrasynaptic nAChRs stimulating the action-potential-dependent release of glutamate from TC fibers. In fact, TC fibers are known to generate spontaneous ectopic action potentials, and the effect of nicotine can be abolished by lesion experiments that eliminated these fibers (Lambe et al., 2003).

In contrast, in Fr2 nicotine produced a negligible effect on EPSC amplitude, suggesting that the above mechanism was scarcely operant, at least at the steady state. Because no unequivocal electrophysiological way is available to distinguish the EPSCs produced by release from TC fibers from those produced by release from intrinsic glutamatergic terminals, we turned to immunolocalization methods. The expression of α4-containing nAChRs in glutamatergic terminals was studied by using immunoblots and immunocytochemistry. We focused on α4 subunit, instead of β2, for two reasons. First, we intended to exclude any interference of β4-containing receptors (Gahring et al., 2004), even though our patch-clamp data indicate that their functional contribution in our experimental conditions is minor. Second, several research groups have shown that β2 may associate with α7, in vitro (Khiroug et al., 2002; Murray et al., 2012) and perhaps also in vivo (Azam et al., 2003; Liu et al., 2009). Therefore, labeling α4 instead of β2 produces unequivocal identification of non-α7 nAChRs, independent from the patch-clamp experiments.

We first tested the expression of α4 subunit in glutamatergic terminals by analyzing the electrophoretic profile of neocortex homogenate samples with SDS-PAGE in reducing conditions. Equal amounts of Fr2 and SS homogenates were tested by WB carried out with three different antibodies. A representative experiment is shown in Figure 5A. Colorimetric assay of our samples revealed unique bands of antigen-antibody interaction with sizes comparable with those reported in literature (Liguez-Lecznar and Skangiel-Kramska, 2007; Raju and Smith, 2006). When using the anti-α4 antibody, we observed a unique band with electrophoretic mobility of 116 kDa (Moser et al., 2007). As revealed by the average band intensity normalized to the actin level, the α4 signal was considerably more evident in Fr2 than in SS cortex (Fig. 5B). Next, use of anti-VGLUT1 revealed the typical 60-kDa band, whereas anti-VGLUT2 revealed a single 65-kDa band (Fig. 5). In these cases, the difference between Fr2 and SS was less pronounced (Fig. 5B). Detailed statistics are given in the figure legend. As expected, when our antibodies against VGLUT1 and VGLUT2 were preadsorbed with their specific antigen, no immunoreactive bands were observed in the homogenate samples. These results show conspicuous expression of both VGLUT1 and VGLUT2 in Fr2, which is accompanied by the presence of α4 nicotinic subunits.

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Figure 5. Protein expression of α4 nAChR subunit, VGLUT1, and VGLUT2 in homogenates of murine neocortex. A: Representative immunoblots comparing α4, VGLUT1, and VGLUT2 expression in membrane fractions of Fr2 and SS. The antibody against α4 recognized a specific band with an apparent molecular mass of 116 kDa. The anti-VGLUT1 and anti-VGLUT2 antibodies detected specific bands of 60 and 65 kDa, respectively. Equal protein loading was assayed by evaluating total actin protein using as a control a monoclonal anti-actin antibody, as indicated. Controls carried out with preadsorbed antisera and with omission of primary antibodies were completely negative. B: Comparison of the intensity of the WB bands in the indicated conditions, in three representative experiments. Optical intensity was detected and analyzed as detailed in Methods. Immunoreactivity data obtained with anti-α4 nAChR subunit, anti-VGLUT1, and anti-VGLUT2 was normalized to the one obtained with anti-actin, and the resulting ratios were plotted for both Fr2 and SS, as indicated. In particular, α4 nAChR gave a mean value of 1.43 ± 0.009 for Fr2 and 0.96 ± 0.001 for SS; VGLUT1 gave 1.52 ± 0.01 for Fr2 and 1.38 ± 0.012 for SS; and VGLUT2 gave 1.1 ± 0.002 for Fr2 and 0.88 ± 0.003 for SS. These differences are statistically significant with paired t-test.

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Figure 6. Laminar cytoarchitecture and nAChR distribution in the Fr2 region. By immunocytochemistry, we studied SMI32-containing pyramidal cells, PV-expressing interneurons, VGLUT1- and VGLUT2-expressing glutamatergic fibers, and α4-containing heteromeric nAChRs. A: Thionin staining of a cortical section from the dorsomedial prefrontal cortex. Open rectangle marks the Fr2 area in which we have analyzed the SMI32 (B), PV (C), VGLUT1 (D), VGLUT2 (E), and 4 nAChR subunit (H, I) immunolabeling (brown signals). B: Notice that the SMI32-positive pyramidal neurons are mainly observed in the large Layer V, whereas only scattered SMI32-positive cells are present in the supragranular Layers II and III (arrows). C: PV immunoreactivity labels small neurons homogeneously distributed in all cortical layers. D: VGLUT1-expressing terminals appear intensely and broadly distributed in Fr2 and surround neuronal cell bodies, as shown at high magnification for Layer V (F). E: The overall VGLUT2 immunoreactivity was lower than that displayed by VGLUT1 and produced higher signal in the supragranular layers of the Fr2 neuropil, whereas labeling tended to decrease in Layers V and VI. G: Immunostaining for α4 nAChR subunit was conspicuous in both supragranular (H) and infragranular (I) layers. A slightly denser neuropil can be observed in the most superficial layers of Fr2 (H). Scale bars: 150 µm (A); 100 µm (B, C); 50 µm (D, E, H, I); and 20 µm (F, G).

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Figure 7. Comparison of the distributions of α4 nAChR subunit and VGLUT1 in Layer V of Fr2 (A–D) and SS regions (E–H). Colocalization of α4 nAChR and VGLUT1 was assessed by confocal fluorescence images, showing immunolocalization of α4 (green) and VGLUT1 (red). A, E: Single α4 nAChR signal (green), which is found in neuronal cell bodies (arrows) as well as neuropilar processes (arrowheads); little difference is observed between Fr2 (A) and SS (E). B, F: The VGLUT1 positive terminals (red) are far more concentrated in Fr2 (B) than in SS (F). D, H: The 2D cytofluorograms show the colocalizing puncta inside the region of interest (yellow oval), for Fr2 (D) and SS (H). These puncta are reported as white signals in the single immunolabeling images (A, B, E, F) and in the merged ones (C, G). C, G: Double immunolabeling (merge) for the two antigens are more often colocalized in Fr2 (C) than in SS (G).

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Figure 8. Comparison of the distributions of α4 nAChR subunit and VGLUT2 in Layer V of Fr2 (A–D) and SS regions (E–H). Colocalization of α4 nAChR and VGLUT2 was assessed by confocal fluorescence images, showing immunolocalization of α4 (green) and VGLUT2 (red). A, E: Single α4 nAChR immunolabeling (green). Arrows, α4 nAChR positive neuronal cell bodies; arrowheads, α4 nAChR positive neuropilar processes. B, F: The single VGLUT2 immunolabeling (red) displays a similar degree of VGLUT2-positive innervation in Fr2 (B) and SS (F). However, a higher colocalization of α4 and VGLUT2 (white signal) is observed in Fr2 (B) than in SS (F). White signal in single immunolabeling images (A, B, E, F) and in the merged ones (C, G) corresponds to colocalizing puncta selected inside the region of interest (yellow oval) of the 2D cytofluorograms, as illustrated in (D) and (H). C, G: Double immunostaining (merge) shows how colocalization is present mainly at the level of α4 positive neuropilar processes and rarely nearby cell bodies.

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Figure 9. High magnification confocal images of the colocalization of α4 nAChR subunit and VGLUT1/VGLUT2 in Layer V of Fr2. Immunofluorescence localization is shown for α4 nAChR subunit (green) and either VGLUT1 (red in A, C, C′, D) or VGLUT2 (red in B). For pyramidal cell recognition, immunofluorescence is also shown for SMI32 (blue in C). A, B: Details from Figures 7C and 8C, respectively. A: The α4-VGLUT1 colocalization sites (white signal) tend to be located around the α4-positive cell bodies (arrows) or processes (arrowheads). B: The α4-VGLUT2 colocalization sites are sparser in the neuropil, in correspondence with some VGLUT2 synaptic terminals (arrowheads). C, C′: The soma (asterisk) of the same α4-positive pyramidal neuron, i.e., the SMI32-positive in (C), is juxtaposed with synaptic terminals expressing both α4 subunit and VGLUT1 (white signal; arrows in C'). D: double-labeled boutons (arrows) are in contact with another α4-positive pyramidal cell (asterisk); a synaptic terminal showing colocalization of α4 and VGLUT1 (white signal, arrowhead) is detected at the level of a α4-positive neuropilar process.

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Neuronal distribution in the Fr2 region

As was recently pointed out, little is known about the structural organization of mouse prefrontal cortex, including its morphology and its connections (Van de Werd et al., 2010; Franklin and Chudasama, 2012). Thus, we next performed a morphological and neurochemical study by single immunocytochemistry on Fr2, for comparison with the classical SS cortex. In rostral brain coronal sections stained with thionin, we identified the small Fr2 area (highlighted in Fig. 6A) whose Layer V we studied by electrophysiology. In this region, we stained pyramidal neurons with SMI32 immunoreactivity, which recognizes the nonphosphorylated epitope of the neurofilament heavy subunit (Fig. 6B). Labeled pyramidal neurons showed a peculiar distribution, as signal was mainly concentrated in the large Layer V, whereas scarce labeling was observed in supragranular layers. In contrast, a second band of SMI32 immunoreactivity is usually present in Layers II and III, in the SS cortex (Kirkcaldie et al., 2002; data not shown). Next, we studied the PV-containing interneurons, which are thought to constitute the majority of basket cells (Fig. 6C). PV-positive cells were rather homogeneously scattered in Fr2 layers, at variance with the usual clustering observed in Layer V of other cortical regions (DeFelipe, 1997; Gonchar et al., 2008; Hof et al., 1999; Soriano et al., 1992; data not shown).

Laminar distribution of VGLUT1 and VGLUT2

Previous studies in rat (Minelli et al., 2003) and mouse (Graziano et al., 2008; Hur and Zaborsky, 2005; Nakamura et al., 2005) show that VGLUT1 is mainly expressed in the intrinsic glutamatergic terminals or corticothalamic afferents, whereas VGLUT2 mainly labels the TC projections. No detailed study is however available in the murine prefrontal regions. By labeling anti-VGLUT1, we observed in our mice considerable neuropil staining in the telencephalon and particularly in Layers I–III and V of the adult neocortex. VGLUT1 immunoreactivity was intense in all Fr2 layers (Fig. 6D), as was testified by the numerous immunoreactive puncta that outlined the morphology of different cortical neurons (e.g., Fig. 6F, for Layer V). The observed laminar distribution was quite similar to that described previously in murine SS cortex (Graziano et al., 2008).

In Fr2, the overall immunoreactivity for VGLUT2 was lower (Fig. 6E) than observed for VGLUT1. The density of VGLUT2-positive puncta tended to decrease in infragranular layers (Fig. 6G). Overall, the distribution of VGLUT2 in Fr2 was broadly similar to the one observed in SS cortex, in agreement with the notion that VGLUT2 is mainly expressed in the terminals of the TC projections originating from the ventral posterior thalamic nucleus (Graziano et al., 2008).

Laminar distribution of α4 nAChR subunit

The distribution of α4-containing nAChRs was tested with the anti-α4 AB5590 (Chemicon) that we have previously extensively tested (Aracri et al., 2010). This antibody does recognize α4-containing nAChRs, when they are expressed (which was demonstrated by our patch-clamp results), although some cross-reaction with other nicotinic subunits or other proteins cannot be fully ruled out (Moser et al., 2007). The overall labeling pattern produced by anti-α4 was quite similar to the immunocytochemical localization previously observed in the rat cerebral cortex for α4 and β2 subunits (Bravo and Karten, 1992; Hill et al., 1993; Nakayama et al., 1995; Schröder, 1992; Whiteaker et al., 2006) In brief, the immunoreactive signal was observed in both neuronal somata and processes throughout Fr2 (Figs. 6H and 6I) with little differences between layers. Neuropilar immunoreactivity for α4 nAChR was observed mainly in Layers II–III and V.

Distribution of α4 nAChR subunit in VGLUT1- and VGLUT2-expressing terminals

We next determined by double immunofluorescence whether the α4-containing nAChRs colocalized with the VGLUT1- and VGLUT2-positive terminals in Fr2 and SS cortex (Figs. 7 and 8). For both pyramidal cells (Figs. 9C, 9C′, and 9D) and small interneurons (Figs. 9A and 9B), α4 mainly labeled the cell somata in Layer V (see also Fig. 6I), whereas in the upper layers the neuropilar signal tended to be denser (e.g., Fig. 6H).

In Fr2 and SS sections, confocal microscope images were acquired from Layer V, for direct comparison with our patch-clamp results. Colocalization of α4 nAChR subunit with either VGLUT1 or VGLUT2 (white signal in Figs. 7-9) was obtained by generating the 2D cytofluorograms reported in Figures 7D, 7H, 8D, and 8H. Combining α4 nAChR and VGLUT1 immunocytochemical localization, we observed a higher degree of colocalization in Fr2 (Figs. 7A–7D and 9A), compared with SS (Figs. 7E–7H). This is more evident in Figures 7B and 7F. In these panels, the colocalization puncta were highlighted among the dense VGLUT1-positive innervation of Layer V, by maintaining constant acquisition parameters and omitting single α4 nAChR dense signal. Our results indicate that α4 is highly expressed on intrinsic glutamatergic synaptic terminals. These terminals were often found to contact α4-positive small cell bodies (probably interneurons; Fig. 9A) and pyramidal neurons identified by morphology, SMI32 immunoreactivity, or both (Figs. 9C, C′, and D).

Finally, by combining anti-α4 and anti-VGLUT2 antibodies, we detected considerable colocalization in Fr2 (Figs. 8A–8D and 9B), despite the lower overall density of VGLUT2-positive terminals in Layer V (compare the colocalization in Figs. 7B and 8B). In contrast, the α4/VGLUT2 colocalization was very weak in the SS (Figs. 8E–8H). Moreover, in both Fr2 and SS cortex, the α4/VGLUT2 colocalization was mainly detected at the level of α4-positive neuropilar processes (Figs. 8A, 8C, 8E, 8G and 9B).

DISCUSSION

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. REFERENCES

Comparison between the neurochemical laminar structure of Fr2 and SS cortex

To the best of our knowledge, our results represent the first analytical study of Fr2 in the mouse. The neuronal distribution showed a peculiar concentration of SMI32-labeled pyramidal neurons in Layer V, whereas the available studies indicate that in SS cortex strong immunoreactivity is also present in Layer III (Kirkcaldie et al., 2002; Franklin and Chudasama, 2012). In a recent chemoarchitectonic characterization, the most medial part of murine Fr2 was found to be almost devoid of SMI32 immunoreactivity (Van de Werd et al., 2010). This, however, reappeared in Layer V toward its lateral borders (corresponding to the region we tested), in both mouse (Franklin and Chudasama, 2012) and rat (Kirkcaldie et al., 2002). Hence, a progressive decrease of supragranular layer thickness seems to occur from the lateral to the medial regions of the prefrontal cortex. A better understanding of the morphofunctional differences between SS cortex and prefrontal regions and between different prefrontal portions may bear important physiological implications. One possibility is that in prefrontal regions, and especially in the more medial areas, the increase in cortical thickness and neuropil, accompanied by decreased cell density, makes it more difficult to discern the presence of Layer IV (Shepherd, 2009). However, expression of SMI32 is also correlated with the degree of fiber myelination (Kirkcaldie et al., 2002), so that it is possible that Layer III (and V in medial regions) pyramidal neurons are present but less recognizable with this marker in prefrontal cortex. This would suggest a higher degree of structural plasticity of principal cells, consistent with the flexibility of the prefrontal cognitive functions. Moreover, the PV immunoreactivity suggests that in prefrontal regions the laminar distribution of interneurons is also different from the one observed in SS cortex (DeFelipe, 1997; Hof et al., 1999). Although we did not observe an overall difference in PV-positive cell density between Fr2 and SS (data not shown), in the former, the PV-expressing neurons (mostly basket cells) appeared to be less concentrated in Layer V.

Such differences in neuronal distribution could be interpreted as a cytoarchitectonic peculiarity of Fr2, which mainly concerns the supragranular layers (especially Layer III). Because basket cells appear to be distributed more homogeneously, the balance of pyramidal cells and interneurons seems to be different compared with other regions and may constitute the basis of specific pathological alterations affecting the morphology of this cortical region (Perez-Cruz et al., 2007) and its inhibition pattern (Eyles et al., 2002; Helmeke et al., 2008; Schneider and Koch, 2005).

As for the VGLUT1 and VGLUT2 distribution, no striking differences were observed in Fr2 compared with the previous studies in SS cortex (Fujiyama et al., 2001; Graziano et al., 2008), although our WB approach highlighted a slightly higher expression level for both transporters in Fr2. Moreover, we observed a broader distribution of VGLUT1 terminals in Fr2, whereas the detailed study on the mouse SS system by Graziano et al. (2008) demonstrated a particularly intense labeling in the supragranular layers. These results are in overall agreement with the notion discussed above that the pattern of excitatory and inhibitory transmission in the different layers is more homogeneously distributed in prefrontal areas.

In both mice (Graziano et al., 2008; present article) and rat (Fujiyama et al., 2001; Herzog et al., 2001), VGLUT2 turns out to be far less expressed than VGLUT1 in cerebral cortex, if compared with subcortical structures. Nonetheless, our analysis indicates that the distribution of VGLUT2 presents a more evident laminar pattern than observed for VGLUT1, with Layer V showing the weakest signal, in analogy with what is observed in SS (Graziano et al., 2008).

Distribution of nAChRs

Although we did not perform a detailed quantitative analysis of nAChR distribution in SS and Fr2, our qualitative comparison broadly confirms previous results obtained in rodents' SS (Bravo and Karten, 1992; Hill et al., 1993; Nakayama et al., 1995; Schröder, 1992; Whiteaker et al., 2006). More recent observations aimed to better discriminate the laminar distribution and function of nAChRs in different neocortical regions of mice (Brown et al., 2012; Kawai et al., 2011; Poorthuis et al., 2012). However, the overall picture is still uncertain, because the anatomical studies focused on SS, whereas the electrophysiological approaches were mainly directed toward the prefrontal regions. Moreover, a detailed comparison with other mammalian species is hampered by the scarcity of available data. Recently, the role of nAChRs in enhanced contrast response of granular layer during visual stimulation has been demonstrated in the primary visual cortex of monkeys (Disney et al., 2007) and tree shrew (Bhattacharyya et al., 2012). The cortical organization of tree shrew is more similar to the one observed in primates than in rodents (Remple et al., 2007). These studies highlight a prevalent expression of nAChRs in Layer IV, but no comparison is available with other sensory (SS) or nonsensory cortical regions. Moreover, only fragmentary data are available on the cortical distribution of nAChRs in these closely related species as well as in humans (Bhattacharyya et al., 2012; Quik et al., 2000; Sihver et al., 1998), with no laminar comparison between different regions.

On these basis, we can only compare the distribution of α4-containing nAChRs in Fr2 with the SS of the same brains. In SS, we detected two evident bands of immunoreactivity at the level of Layers III and V, with higher intensity in supragranular layers (data not shown). In contrast, Fr2 showed a higher expression of α4 nAChR subunit, which displayed a homogeneous distribution with little difference between layers. Altogether, the morphological and physiological features of Fr2 are in line with the general shrinkage of Layer IV observed in nonsensory regions. Although uncertainty still lingers about the consistency of Layer IV in different areas (Palomero-Gallagher and Zilles, 2004; Shepherd, 2009; Van de Werd et al., 2010), in motor and premotor regions with thin Layer IV, a dense TC innervation is observed in Layer III and upper Layer V (Shepherd, 2009).

Localization of α4-containing nAChRs in glutamatergic terminal of Fr2 and SS cortex

The higher expression of α4 nAChR throughout Fr2, compared with SS, with barely detectable laminar pattern, agrees with the potent DHβE-sensitive stimulation produced by nicotine on EPSC frequency. Our results suggest that the regulatory role of α4β2 nAChRs on excitatory transmission is particularly potent in dorsomedial prefrontal regions, even with steady state stimulation. In Layer V of both Fr2 and SS, immunolabeling of α4 was detected in neuropilar processes, puncta, and in the cell bodies of pyramidal cells and interneurons, in agreement with the widespread expression of heteromeric nAChR indicated by other lines of evidence (Aracri et al., 2010; Couey et al., 2007; Lambe et al., 2003; Poorthuis et al., 2012).

In Fr2 Layer V, we observed a certain degree of colocalization of α4 subunit with both VGLUT1- and VGLUT2-expressing terminals. These glutamatergic terminals, and especially the VGLUT2-positive, showed scarce nAChR expression in SS cortex. A similar analysis with different methodological approaches was recently performed on Layer IV of murine SS cortex. This demonstrated a low, but significant, degree of colocalization (7–10%) of α4 nAChR with glutamatergic cortical terminals (Brown et al., 2012).

Nicotinic stimulation of glutamate release in Fr2

Activation of α4β2 nAChRs in Fr2 strongly stimulated the EPSC frequency, without a significant increase of EPSC amplitude. This suggests that nicotine was not producing preferential stimulation of the action potential-dependent glutamate release. The latter effect could occur in two ways: (i) stimulation of spontaneous local pyramidal neuron firing and (ii) stimulation of the frequency of spontaneous ectopic action potential in TC fibers. The weak effect of nicotine on EPSC amplitude thus suggests that, in steady state conditions, both of the above mechanisms are poorly active in Fr2. This is consistent with the following two observations. First, we obtained generally negligible whole-cell nicotinic currents by directly stimulating pyramidal neurons with nicotine at Vm around rest (data not shown; see also Couey et al., 2007 for similar results in the medial prefrontal region). Second, immunocytochemistry showed a generally low VGLUT2 fiber density, in Fr2. At the present stage, our interpretation is that tonic stimulation of nAChRs mainly controls glutamate release presynaptically, in both types of glutamatergic terminals.

In terms of nAChR subtypes, the high sensitivity of glutamate release on 5IA85380 indicates that α4β2 receptors account for most of the steady state stimulatory effect of nicotine. In fact, at drug concentrations lower than 100 nM, the effect of this compound on α7-containing receptors is negligible (Mogg et al., 2000; Mukhin et al., 2000), consistent with our results with DHβE and MLA. Moreover, at these concentrations, 5IA85380 is also poorly effective on β4-containing subunit. Finally, the strong efficacy of low concentrations of this drug leads us to attribute most of the nicotinic effect to the α42β23 high-affinity subtype.

Several lines of evidence indicate that in other cerebral regions, α4β2 and α7 receptors cooperate in regulating glutamate release. Most of this evidence was obtained by using synaptosomes extracted from hippocampus (Bancila et al., 2009; Mura et al., 2012; Zappettini et al., 2010) and different neocortical areas (Dickinson et al., 2008; Marchi et al., 2002). An interesting observation is that the contribution of α7-containing receptors often requires a certain degree of synaptosome depolarization. This and other observations (reviewed in Marchi and Grilli, 2010) indicate that the α7 nAChRs cooperate with voltage-gated calcium channels to exert their presynaptic effects. These experimental approaches are more difficult to apply to brain slices. However, the above results are not necessarily in contrast with our data, which suggest that α4β2 nAChRs can stimulate glutamate release (at least in Fr2) at low-tonic levels of ACh (and nicotine, in smokers), irrespective of the general state of network excitation. They do not imply that α7 homomeric receptors are not functional in this region but simply that their contribution to glutamate release in conditions of sustained stimulation with low doses of agonists is negligible. Therefore, we conclude that the α4β2 receptors prevail in regulating the tonic neocortical activation produced by the ascending cholinergic stimulus typically occurring during wakefulness and REM sleep. This is also consistent with the involvement of α4β2 nAChRs in sleep-related epilepsy.

Possible pathophysiological implications

Recent work carried out with two-photon imaging of entire slices in medial prefrontal cortex suggests that the overall effect of tonic nAChR activation is layer specific, with increase of activity in deep layer pyramidal neurons and inhibition in Layers II/III (Poorthuis et al., 2012). Our results suggest that the output of Fr2 could be even more sensitive to nicotinic stimulation, as indicated by (i) the sustained effect of tonic stimulation of α4β2 nAChRs on glutamate release and (ii) the well-developed Layer V, seemingly accompanied by lesser inhibitory weight than is typical in regions such as the SS cortex. Therefore, this region could be particularly prone to develop seizures. Considering also the connectivity of Fr2 that we discussed earlier, we believe further studies on the steady state heteromeric nAChR effects on excitatory transmission and GABA release (Aracri et al., 2010) in this region may offer important insights into the susceptibility to seizures produced by mutant heteromeric nAChRs in the frontal lobes.

REFERENCES

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. REFERENCES
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