The entorhinal cortex (EC) is a part of the hippocampal complex that is essential to learning and memory, and nicotine affects memory by activating nicotinic acetylcholine receptors (nAChRs) in the hippocampal complex. However, it is not clear what types of neurons in the EC are sensitive to nicotine and whether they play a role in nicotine-induced memory functions. Here, we have used voltage-sensitive dye imaging methods to locate the neuronal populations responsive to nicotine in entorhino-hippocampal slices and to clarify which nAChR subtypes are involved. In combination with patch-clamp methods, we found that a concentration of nicotine comparable to exposure during smoking depolarized neurons in layer VI of the EC (ECVI) by acting through the non-α7 subtype of nAChRs. Neurons in the subiculum (Sb; close to the deep EC layers) also contain nicotine-sensitive neurons, and it is known that Sb neurons project to the ECVI. When we recorded evoked EPSCs (eEPSCs) from ECVI neurons while stimulating the Sb near the CA1 region, a low dose of nicotine not only enhanced synaptic transmission (by increasing eEPSC amplitude) but also enhanced plasticity by converting tetanus stimulation-induced short-term potentiation to long-term potentiation; nicotine enhanced synaptic transmission and plasticity of ECVI synapses by acting on both the α7 and non-α7 subtypes of nAChRs. Our data suggest that ECVI neurons are important regulators of hippocampal function and plasticity during smoking.
Nicotine is the major compound in tobacco that affects memory (Peeke and Peeke, 1984; McGehee and Role, 1996) and cognition (Potter and Newhouse, 2008) through its actions on nicotinic acetylcholine receptors (nAChRs) (Dajas-Bailador and Wonnacott, 2004; Davis et al., 2007; Timmermann et al., 2007). Despite the negative health consequences of smoking tobacco, nicotine and related compounds have been proposed as drugs for treating neurological diseases and disorders including Alzheimer's disease (Levin and Rezvani, 2002; Levin et al., 2006), Parkinson's disease (Park et al., 2007; Quik et al., 2007a,b; Villafane et al., 2007), attention-deficit hyperactivity disorder (Potter and Newhouse, 2008), schizophrenia (Tizabi, 2007), and epilepsy (Shin et al., 2007) because of their effects on memory, cognition, and neuronal excitability. However, the mechanisms by which nicotine influences memory and learning are far from understood.
The hippocampal formation (HF) is critical to memory and cognition (Blozovski, 1983; Izquierdo et al., 2008) and mediates the influences of nicotine on memory (Blozovski, 1985; Davis et al., 2007). The HF includes four main subregions (Amaral and Witter, 1995): the dentate gyrus, the hippocampal proper (including CA1, CA2, and CA3 regions), the subicular complex [including subiculum (Sb)], and the entorhinal cortex (EC; including layers I–VI). Each subregion plays distinct roles in memory and cognition (Hunsaker et al., 2008; Li and Chao, 2008). Thus far, investigations regarding the influence of nicotine on hippocampal neurons have been focused on the hippocampal proper and dentate regions (Jones and Yakel, 1997; Frazier et al., 1998a,b; McQuiston and Madison, 1999; Sudweeks and Yakel, 2000; Alkondon and Albuquerque, 2001; Khiroug et al., 2003; Fayuk and Yakel, 2004, 2005; Klein and Yakel, 2005; Welsby et al., 2007); this is likely because of the well studied role of this region in learning and memory (Hasselmo, 2005; Lee et al., 2005; Chen et al., 2006; Hunsaker et al., 2008; Izquierdo et al., 2008; Li and Chao, 2008) and the ability of nicotine to induce synaptic potentiation (Hunter et al., 1994; Sawada et al., 1994; Gray et al., 1996; Fujii et al., 1999, 2000; He et al., 2003; Matsuyama et al., 2000). However, whether nicotine influences neurons in either the subicular complex or EC is still unknown. Because both the EC and Sb play critical roles in memory functions (Blozovski, 1983, 1985; O'Mara et al., 2001; Deadwyler and Hampson, 2004; Burhans and Gabriel, 2007; Martin-Fardon et al., 2007; Harich et al., 2008; Izquierdo et al., 2008; Van Cauter et al., 2008), information about whether and how nicotine influences these structures is important in understanding how nicotine affects memory.
To address this issue, we introduced voltage-sensitive dye imaging (VSDI) techniques to locate the neuronal populations responsive to nicotine in entorhino-hippocampal slices and to clarify which nAChR subtypes are involved. Nicotine was bath applied to emulate systematic administration occurring during smoking, while using nicotine patches, or during subcutaneous injection. Patch-clamp techniques were then used to characterize the receptors and neuronal populations involved and their role in synaptic plasticity.
Materials and Methods
Wistar rats ∼14–21 d of age were anesthetized with halothane and decapitated. Brains were quickly removed; placed into a high-magnesium/low-calcium ice-cold artificial CSF (the dissecting ACSF) containing (in mm) 122 NaCl, 3.5 KCl, 12 MgCl2, 1.3 CaCl2, 1.2 NaH2PO4, 25 NaHCO3, 0.5 ascorbic acid, and 10 glucose; and continuously bubbled with carboxygen (95%O2/5%CO2). Horizontal entorhino-hippocampal slices (∼300–450 μm thick) corresponding to plate 190–200 of the Paxinos rat brain atlas (Paxinos and Watson, 2007) were prepared in icy carboxygenated dissecting ACSF with a vibratome. Slices were then stored in the regular ACSF (same as the dissecting ACSF except with 1.8 mm CaCl2 and 1.2 mm MgCl2) and continuously bubbled with carboxygen at room temperature for >1 h before use.
The VSDI technique was modified from a previous report (Tominaga et al., 2000) to conduct longer-duration imaging sampling (see Fig. 1A). After at least 1 h of recovery from slicing, 400- or 450-μm-thick slices were stained (away from light) for 45–60 min in a oxygenated humidity chamber with staining solution containing either Di-4-ANNEPS or N-(3-triethylammoniumpropyl)-4-(4-(dibutylamino)styryl)pyridinium dibromide (FM 1-43; both 100 μm), 50% FBS, 50% regular ACSF, 0.1% cremophor EL, and 2% ethanol. The stained slice was then placed in an MS3 microscope stage interface chamber (Automate Scientific) over a white FHLC membrane (0.45 μm; Millipore) supported by a nylon mesh (Warner Instruments) that was raised 250 μm above the chamber bottom. Humidified carboxygen was blown over the top of the slice with perfusion medium flowing beneath the FHLC membrane to make sure that the top of the slice was humidified but not covered with liquid; this was a critical step to ensure low-noise imaging while maintaining viability of neurons. Perfusion medium was delivered to the chamber at a rate of ∼1.5–2 ml/min driven by gravity (see Fig. 1A). A THT microscope (BrainVision) was used to acquire VSDI signals and was equipped with epifluorescence optics in a tandem-lens configuration: PLANAPO 1× lens (WD, 55 mm; Leica) as the objective lens and PLANAPO 0.63× lens (WD, 97 mm; Leica) as the projection lens. The final magnification of the system was 1.6. The excitation light was provided by a halogen lamp source (150 W; MHAB-150W; Moritex) passed through an excitation filter (510–550 nm; FF01-531/40-25; Semrock). Light was reflected onto the slice by a dichroic mirror (570 nm), and the fluorescence generated from slices was passed through an emission filter (>590 nm; U-MWG2 filter set; Olympus). Images were projected onto the 6.4 × 4.8 mm2 CCD sensor of the optical imaging system (MiCAM02-HR; BrainVision). Imaging responses to stimulation were acquired at a speed of 5 ms per frame with an actual size of 184 × 124 pixels (spatial resolution of 22 × 24 μm per pixel) for 500 ms. Imaging of nicotine effects was acquired at 1 or 5 s per frame (376 × 252 pixel dimension, 11 × 12 μm per pixel), with 50 ms exposure time for each frame to minimize photobleaching. Imaging acquisition was performed at room temperature in all experiments.
A glass pipette (tip diameter, ∼1 μm), pulled from borosilicate glass capillaries (Garner Glass) with a model P-97 puller (Sutter Instrument Company) and filled with 2 m NaCl (resistance, ∼3–10 MΩ), was placed in regions of interest. Stimulation was delivered through a bipolar platinum/iridium electrode (25 μm thick insulated with formvar; FHC) to specified regions. Field potential signals were amplified 1000 times and bandpass filtered at 0.3–3000 Hz with a P511 AC amplifier (Grass Technologies). Amplified signals were digitized with a Digidata 1322A, acquired, and analyzed with pClamp10 software (Molecular Devices).
Whole-cell patch-clamp recordings.
Slices of ∼300–350 μm thick were used for whole-cell patch-clamp studies in a submerged chamber perfused at a rate of 1 ml/min with regular ACSF. Patch clamp of neurons was performed under guidance of infrared differential interference contrast (IR-DIC) optics using an Axopatch 200B patch amplifier (Molecular Devices) with a glass pipette filled with an internal solution containing (in mm) 120 potassium gluconate (KGluc), 2 NaCl, 5 MgATP, 0.3 Na2GTP, 20 KCl, 10 HEPES, 1 EGTA, and 11.3 d-glucose, with pH ∼7.2–7.3 and osmolarity of ∼270–280 mOsm. Resting membrane potentials (RMPs) were obtained immediately after break-in under current-clamp configuration. Series resistances ranging from 7 to 40 MΩ were not compensated for during recordings but were monitored throughout the experiments. Recordings were discarded when a significant (>20%) change of series resistance was detected. Whole-cell capacitance was obtained by canceling transients when clamping the cell at −70 mV and applying 5 mV hyperpolarizing pulses for 30 ms. After adjusting the membrane potential to −70 mV, firing properties of neurons and h-current were tested with 11 current injections of 500 ms duration at intensities ranging from −150 to +350 pA, with 50 pA increments. The threshold firing potential was determined by the lowest membrane potential needed to elicit the first action potential during this current injection process.
To detect the amount of membrane depolarization by nicotine, recordings were performed under the current-clamp configuration in the presence of tetrodotoxin (TTX; 1 μm) and atropine (10 μm). Current was injected into neurons to maintain a membrane potential of −80 mV at the beginning of the experiment and was kept constant after initiation of recordings. For recordings of postsynaptic events in neurons in layer V of the EC (ECV neurons), KGluc in the internal solution was replaced with the same concentration of KCl [for spontaneous IPSC (sIPSC)] or CsGluc [for spontaneous EPSC (sEPSC)]; glucose was replaced with QX314 (10 mm). All membrane potentials reported here were values after adjustment of liquid junction potentials experimentally tested for each type of internal solution (10 mV for internal solutions with KGluc and CsGluc, 4 mV for internal solution with KCl). Signals were passed through a 1 kHz low-pass filter, digitized with the Digidata 1322A (Molecular Devices), and acquired at 10 kHz with pClamp10 software (MDS Analytical Technologies) in all experiments, except in nicotine bath-application experiments in which recordings were acquired at 10 Hz to reduce data file size.
Determination of functional nAChRs in neurons.
At 5–15 min after break-in during whole-cell recordings, nAChR-mediated responses were evoked by pressure applications (350 ms, 25 psi unless indicated otherwise) of maximal doses of either choline (100 mm) or ACh (4 mm) at a 2 min interval through the tip (0.5–1 μm diameter) of a theta style double-barrel borosilicate glass (outer diameter, 2.0 mm; inner diameter, 1.4 mm; Warner Instruments) located ∼30–50 μm away from the soma of the targeted neuron. Pressure applications used a PPM-2 pneumatic pump (Harvard Apparatus). To minimize desensitization of nAChRs by agonists leaking from the pipette tip between applications, the pipette was manually placed at the targeted position 5–10 s before pressure application of either ACh or choline and was retracted by >200 μm away from the neuron immediately after pressure application [this technique was modified from Pidoplichko and Dani (2005)]. This was done under visual guidance of live IR-DIC imaging, and the targeted position was marked to insure that subsequent pressure applications were at the same location. TTX (1 μm) and atropine (10 μm) were added to the perfusion ACSF in all experiments. Functional α7-containing nAChR (α7 nAChR) current responses were assessed by the amplitude of responses to choline, and the peak of non-α7 current responses were determined from responses to ACh in the presence of methyllycaconitine (MLA) (10 nm) to block α7 nAChR-mediated responses; sometimes MLA was omitted when there was no sign of fast activating and desensitizing α7 responses. Using this protocol, the functional α7 current responses induced by the choline (100 mm) applications shown here were not significantly different from those induced by ACh (4 mm) applications (in the presence of 1 μm dihydro-β-erythroidine (DHβE) in addition to the above-mentioned atropine and TTX) either in amplitude or 10–90% rise time in CA1 interneurons (n = 9; p > 0.1, Student's t test).
Determination of nicotine effects on Sb-ECVI synaptic plasticity.
Synaptic plasticity of Sb input to ECVI pyramidal neurons was tested with whole-cell recordings in ECVI neurons with a KGluc-based internal solution using a stimulating electrode (same as used above) placed in the Sb area near the CA1 region. The stimulation intensity was adjusted to evoke a postsynaptic current of ∼50 pA, and the intensity was usually ∼0.5–5 mA for 0.1 ms. Short-term potentiation (STP), defined as synaptic potentiation for a few seconds or minutes, often 10–15 min (Sajikumar et al., 2009), was induced by a 100 ms train of stimulation at 100 Hz. Local continuous nicotine exposure was achieved by placing a glass puffer pipette 25–50 μm away from the cell body with continuous pressure of 1–2 psi. To elicit EPSCs in CA1 pyramidal neurons, the Schaffer collateral pathway was stimulated with a 0.1-ms-duration pulse, with the stimulation intensity adjusted to 50–200 μA to evoke EPSCs of 50–100 pA in amplitude. The same type of stimulation was used to obtain the paired-pulse ratio (PPR), which was defined as the ratio of responses evoked by the second versus the first stimulation at an interstimulation interval of 20 ms.
Conventional and confocal imaging.
The soma sizes of neurons were determined under IR-DIC optics before patch clamping. The longest axis of each neuron and the axis perpendicular to the longest axis were measured, and the square root of the product of these two values was used to represent its soma diameter. Major dendrites were also counted to help in the determination of whether a neuron was a pyramidal or multipolar neuron. To further visualize the dendritic structures of neurons studied, 5 μm Alexa Fluor 488 hydrazide (Alexa488) was added to the patch pipette solution to replace d-glucose. Fluorescence-labeled neurons were imaged with either a cascade digital camera (Roper Scientific) with Andor iQ 1.8 imaging software (Andor Bioimaging Division) or a confocal imaging system (Radiance 2100; Bio-Rad).
Imaging data were acquired and analyzed with BV-Analyze software (BrainVision). The raw data were filtered with spatial (5 × 5) and cubic (3 × 3) filters, and baseline drift was removed using the built-in, two-dimensional drift-removal function with a fitting period that ensures maximum stability of the control recording. Preliminary experiments found that during 30 min of image acquisition, drift of the fluorescence signal during the first 5 min of imaging did not fit this two-dimensional function and was therefore disregarded before analysis. More than 10 min of control recordings were obtained before drug addition to ensure ≥5 min of control recordings for drift removal.
To compare differences in current or voltage responses mediated by functional nAChRs, values from the different neuronal groups were first tested for a Gaussian distribution [p > 0.1, Kolmogorov–Smirnov (KS) normality test]. One-way ANOVA and the post hoc Dunnet's test were used to compare Gaussian distribution values, whereas Kruskal-Wallis nonparametric one-way ANOVA with post hoc Dunn's test was used for the others. All the above analyses were performed using Prism 4.03 software (GraphPad Software).
sEPSC and sIPSC events acquired with pClamp10 software were analyzed using MiniAnalysis software (version 6.03; Synaptosoft). Events were initially detected by the software with an amplitude threshold of 5 pA and then were manually examined to exclude false events. Kinetics (10–90% rise time and decay constant) of sEPSCs, sIPSCs, and ACh-evoked current responses were also analyzed with MiniAnalysis software. Decay time constants were derived from exponential fits of the 90% decay responses. KS tests were conducted using MiniAnalysis software. One-way ANOVA and post hoc analysis, dose–response curve (EC50) analysis, and normality tests were performed using Prism 4.03 software (GraphPad Software). All values were presented as mean ± SEM unless noted otherwise.
Drugs and chemicals.
TTX was obtained from Tocris. Di-4-ANEPPS, FM 1-43, and Alexa488 were obtained from Invitrogen. All other chemicals or drugs were obtained from Sigma.
Nicotine depolarizes deep layers of the EC and Sb through the activation of non-α7 nAChRs in Wistar rats
VSDI provides high temporal and spatial resolution in monitoring membrane potential changes in neuronal populations in brain slices or under the surface of intact brains (Ebner and Chen, 1995; Airan et al., 2007). Di-4-ANEPPS, a styryl dye that linearly reflects membrane potential changes at a rate of 1% per 10 mV under physiological conditions (Loew et al., 1992), has been found to reliably detect neuronal membrane potential changes within milliseconds (Mann et al., 2005; Airan et al., 2007). The voltage sensitivity of Di-4-ANEPPS was verified in our experimental set-up (Fig. 1) by fluorescent responses to Schaffer collateral stimulation in rat CA1 hippocampal slices stained with this dye, as well as the absence of fluorescent responses in slices stained with the voltage-insensitive dye FM 1-43 (which was used as a control) (Fig. 1B). As reported previously (Tominaga et al., 2000), we also observed significant fluorescence drift (likely caused by photobleaching and other factors) in our slice recordings during 30 min of imaging, which was accounted for and removed (to obtain a stable VSDI signal baseline) by using a two-dimensional drift-removal function in the BV-Analysis software (Fig. 1C).
After adding TTX (1 μm) to remove action potential-dependent synaptic transmission between neurons, we found that bath-applied nicotine (10 μm) significantly depolarized neuronal populations in ECVI and stratum oriens of the Sb (SbSO), whereas other regions were depolarized to a lesser extent (Fig. 2).
The two major subgroups of nAChRs in hippocampal neurons are the α7-containing nAChRs (α7 nAChRs) and a diverse subgroup of non-α7 nAChRs, the most prevalent of which is the α4β2 subtype in the hippocampus and cortex (Alkondon and Albuquerque, 1993; Alkondon et al., 1994; Jones and Yakel, 1997; Sudweeks and Yakel, 2000). We found that the non-α7 nAChR antagonist DHβE (1 μm) blocked nicotine-induced VSDI responses, whereas the α7 nAChR-selective antagonist MLA (10 nm) did not (Fig. 2A,C), which demonstrates that the nicotine-induced neuronal depolarizations were mediated by non-α7 nAChRs. The viability of neurons in slices was verified by the ability of 4 mm KCl to induce significant depolarization (Fig. 1D, 2A) or, in some cases, by Schaffer collateral stimulation-evoked responses recorded through field electrodes (Fig. 1B). Brain slices stained with the voltage-insensitive dye FM 1-43 did not show any fluorescent responses to nicotine (10 μm) (Fig. 2A), which suggests that nicotine-induced fluorescent responses using Di-4-ANEPPS were not attributable to some interaction between nicotine and the fluorescent dye (independent of membrane potential changes), the influence of nicotine on slice optical characteristics, or any potential membrane surface change as a result of synaptic vesicle release triggered by nicotine.
Layer ECVI neurons are the most responsive to nicotine
We used whole-cell patch-clamp techniques to record from individual neurons within various regions and layers of the HF (Fig. 3A) to directly measure the ability of nicotine to induce electrical responses and how this pattern of responsiveness compared with the VSDI results. Under current clamp, we found that all ECVI neurons that we recorded from (n = 26) were depolarized by bath-applied nicotine (10 μm) (Table 1, Fig. 3) and that the amount of depolarization was larger compared with neurons in other regions of the HF (Fig. 3B,C).
To compare the kinetics of the nicotine-induced depolarizations in different regions of the HF, we averaged the membrane potential recordings in response to nicotine (Fig. 3B). We found that the depolarizations induced by the bath application of nicotine for 5 min reached peak within 1 min and then slowly repolarized back to baseline. The repolarizations were caused by the decay of nicotine responses (most likely as a result of desensitization) and not as a result of membrane potential drift since the nicotine responses were completely blocked (i.e., complete recovery of membrane potential to the value before nicotine application) by DHβE (either 1 or 100 μm for complete inhibition). The decay rate (i.e., the percentage of decay after 5 min of nicotine application) in the CA1 stratum radiatum (SR) neurons (17 ± 6%; n = 4) was similar to that in ECVI neurons (15 ± 4%; n = 16), both of which were significantly slower than that in the SbSO neurons (57 ± 6%; n = 9) or ECV neurons (43 ± 8%; n = 8) (p < 0.01, one-way ANOVA followed by the Newman–Keuls multiple comparison test). When DHβE was replaced by the ionotropic glutamatergic receptor blockers 2,3-dioxo-6-nitro-1,2,3,4-tetrahydrobenzo[f]quinoxaline-7- sulfonoamide (NBQX; 20 μm) and APV (50 μm), the nicotine-induced responses in ECVI neurons were not affected (n = 3; data not shown), precluding the contribution of glutamate release to the nicotine-induced depolarization.
The concentration of nicotine required to produce half-maximal depolarizations (i.e., the EC50 value) was 0.6 μm for ECVI neurons (Fig. 3D), which was significantly lower than for either SbSO (1.3 μm) or ECV (5.5 μm) neurons (p < 0.01, two-way ANOVA using concentration and region as variables followed by Bonferroni posttests). The bath application of a low concentration of nicotine (0.1 μm), which is within the range found in the blood of average smokers (i.e., 0.06–0.31 μm) (Moriya and Hashimoto, 2004; Hukkanen et al., 2005; Moriya et al., 2006), still significantly depolarized ECVI neurons, unlike either SbSO or ECV neurons (Fig. 3B).
Functional distribution of nAChR-mediated current responses in the HF
We recorded nAChR-mediated current responses under voltage clamp during the rapid pressure application of ACh to the soma of neurons in different regions of the HF in the presence of TTX (1 μm) and atropine (10 μm). Under these conditions, there will be little to no desensitization of nAChR-mediated responses, which is particularly important for α7 nAChRs that are characterized by rapid desensitization (Jones and Yakel, 1997; McQuiston and Madison, 1999; Alkondon and Albuquerque, 2002; Khiroug et al., 2003). To isolate the non-α7 nAChR-mediated current responses, we activated responses with ACh in the presence of MLA (10 nm), and to isolate the α7 nAChR component, we activated responses with the full α7 nAChR-selective agonist choline (Fig. 4A). Although MLA has previously been reported to block α6-containing nAChRs (Klink et al., 2001), the α6 subunit is not expressed in rat hippocampus and EC (Sudweeks and Yakel, 2000; Mugnaini et al., 2002), and thus this should not affect the interpretation of our results. We found that ECVI and SbSO neurons had the largest non-α7 nAChR-mediated responses (Fig. 4B, Table 1), similar to what we observed for nicotine-induced depolarizations. These data indicate that ECVI and SbSO neurons had the highest levels of functionally expressed non-α7 nAChRs, which is consistent with the previous VSDI results that ECVI and SbSO were the most nicotine-responsive regions in the HF.
After blocking α7 nAChRs with MLA (10 nm), the bath application of DHβE (1 μm) blocked >95% of the nicotine-induced depolarizations in neurons of the ECVI (median, 96.6%; interquartile range, 83.3–99.3%; n = 15), the SbSO (98.3%; 96.0–98.9%; n = 5), and the CA1 SR (98.3%; 94.3–99.1%; n = 11). The α4β2 nAChR subtype, besides being the most prevalent subtype in the hippocampus and cortex, has the highest affinity for nicotine and DHβE (Harvey et al., 1996; Marks et al., 1999). The pharmacological data for our non-α7 nAChR-mediated responses are most consistent with their being composed mostly of the α4β2 subtype, and not consistent with the other common subtypes of non-α7 nAChRs expressed in the brain (e.g., the α3β4 subtype) (Jones and Yakel, 1997; Sudweeks and Yakel, 2000). Neither the α3 nor the α2 nAChR subunits were previously detected in the deep layers of the EC, whereas the α4 and β2 subunits were (Wada et al., 1989), as well as the lack of presence of the α6 nAChR subunit.
The functional distribution of α7 nAChR-mediated current responses (i.e., those responses activated by choline) is shown in Figure 4B and Table 1. Interestingly, most ECVI neurons (89%) had functional α7 responses with a median amplitude not significantly different from CA1 interneurons (p > 0.05, Kruskal-Wallis test and post hoc Dunn's multiple comparison test). Since α7 nAChRs desensitize rapidly, the most likely reason why we did not observe an α7 receptor-mediated depolarizing component in response to bath-applied nicotine in ECVI neurons was because of desensitization. To address this point directly, we used the α7-selective allosteric potentiator PNU-120596 (PNU), which dramatically increases α7 response amplitudes in part by reducing desensitization (Hurst et al., 2005; Poisik et al., 2008). Bath application of nicotine (under current clamp; n = 3) to CA1 SR interneurons that functionally expressed only α7 nAChRs (and not non-α7 nAChRs) failed to induce any membrane depolarization (Fig. 5). However after the application of PNU (10 μm; to other α7-only CA1 SR interneurons), the bath application of nicotine was now able to induce significant membrane depolarizations (Fig. 5) (n = 3). These results demonstrate that desensitization prevented α7 nAChRs from contributing to the neuronal depolarization induced by the bath application of nicotine under physiological conditions.
The α7-only nature of these responses was verified by similar (defined as <5% difference) rise and decay kinetics between responses to ACh and the full and selective α7 nAChR agonist choline, and the significantly slower kinetics of non-α7 nAChRs. The absence of non-α7 nAChR-mediated currents in these α7-only neurons was further confirmed by the lack of changes in the kinetics of responses to ACh after the bath application of DHβE (1 μm). As a final confirmation, 21 α7-only neurons were selected from 68 CA1 SR neurons using these criteria, and ACh-evoked currents in all of these neurons were completely (>95%) blocked by 10 nm MLA.
Characteristics of ECVI neurons
We characterized the morphological and electrical properties of ECVI neurons, which were usually referred to as part of the deep layers of the EC along with ECV neurons (Chrobak and Buzsaki, 1994; Solger et al., 2004; Gnatkovsky and de Curtis, 2006).
Morphological properties of ECVI neurons
As reported previously (Dugladze et al., 2001), the shapes of ECVI neurons may be either pyramidal or multipolar, whereas ECV neurons are primarily pyramidal shaped. When observed under IR-DIC optics, the soma diameters of ECVI neurons were not significantly different from CA1 interneurons but were significantly larger than the diameters of either ECV or CA1 pyramidal neurons (Table 2). The measurements of neuronal size under IR-DIC are consistent with a previous report (Ruan et al., 2007). The membrane capacitance (an indicator of membrane surface area) of ECVI neurons was significantly larger than CA1 neurons, suggesting that ECVI neurons had more dendritic arboration than CA1 neurons since their soma sizes were similar.
Electrical properties of ECVI neurons
The RMPs of ECVI neurons were more negative than those of CA1, Sb, ECII, or ECVIII neurons (Table 2). In addition, the ECVI neurons had the lowest level of h-current (Ih) among all regions of neurons studied. Ih is a current mediated by a hyperpolarization-activated cyclic nucleotide-gated (HCN) channel (Rosenbaum and Gordon, 2004; Siu et al., 2006). Although HCN channels are weakly selective for K+ over Na+ with permeability ratios of ∼4:1, under physiological conditions, Ih is mostly carried by Na+ because of a greater driving force (Siu et al., 2006). Ih has a significant impact on the RMP since inhibition of Ih hyperpolarizes neurons (Lupica et al., 2001; Aponte et al., 2006). Therefore, a low Ih amplitude, such as that found in ECVI neurons, can contribute to the more hyperpolarized RMP of these neurons. The ECVI neurons also had smaller input resistances than CA1 interneurons, spiked at a frequency slower than CA1 interneurons but faster than CA1 pyramidal neurons when depolarized (i.e., medium frequency spiking), and had similar action potential thresholds (Table 2).
ECVI neurons provide both excitatory and inhibitory inputs to ECV neurons
Both ECVI and ECV neurons project mainly to superficial layers of the EC (Kohler, 1986; Dolorfo and Amaral, 1998; Hamam et al., 2000) and a wide range of cortical areas including the temporal pole, medial frontal and orbitofrontal cortices, and the rostral part of the polysensory area of the superior temporal sulcus cortices (Wyss and Van Groen, 1992; Deller et al., 1996; Hamam et al., 2000; Munoz and Insausti, 2005). Through these projections, ECVI and ECV neurons may impact indirectly on hippocampal functions since the neurons in superficial layers of the EC (ECII or ECIII) are the major sources of inputs received by the dentate and the hippocampal proper. Projections from ECVI or ECV were also identified targeting neurons in the Sb and the dentate gyrus directly (Dugladze et al., 2001), suggesting that direct influence on hippocampal function by deep EC neurons is also possible. We found that ECVI neurons, which are mainly excitatory neurons (Hamam et al., 2000), were the most responsive to nicotine (based on the VSDI and patch-clamp experiments described above). Thus far, there are no reports demonstrating the functional synaptic influence of the neuronal activation of ECVI on neurons in other regions. Because the ECV neurons are the closest neuronal population to the ECVI region, any connection between ECV and ECVI neurons may be well preserved in the hippocampal slices that we have studied. In addition, ECV neurons are known to mediate interactions between the hippocampal proper and a wide range of neocortical areas (Insausti et al., 1997; Naber et al., 2001). Therefore, if ECV neurons were influenced directly by ECVI neurons, nicotine may influence the function of various hippocampal regions through the activation of neurons in the ECV, which will further influence ECII/III neurons, the major source of inputs for the dentate gyrus.
To investigate whether ECV neurons are postsynaptically activated by ECVI neurons, we recorded spontaneous glutamatergic (Fig. 6) or GABAergic (Fig. 7) postsynaptic currents (sEPSCs and sIPSCs) in ECV neurons with whole-cell voltage-clamp recordings while bath-perfusing the slices with nicotine (10 μm), or by locally activating non-α7 nAChRs in ECVI neurons and other neighboring regions. Bath-applied nicotine (10 μm) significantly increased the frequency of sEPSCs (n = 5) (Fig. 6A,B) but not that of sIPSCs (n = 5; p > 0.05, KS test) (Fig. 7A) recorded in ECV pyramidal neurons, demonstrating that bath-applied nicotine enhanced excitatory but not inhibitory inputs onto ECV neurons through the activation of non-α7 nAChRs. The brief (350 ms duration) local application of ACh (4 mm; ∼200 μm away from recorded neurons) to ECVI neurons in the presence of antagonists of muscarinic AChRs (atropine, 10 μm) and α7 nAChRs (MLA, 10 nm) also significantly increased the frequency of sEPSCs onto ECV neurons (n = 5; 0.29 ± 0.08 Hz in control condition compared with 7.3 ± 2.0 Hz within 10 s after brief local activation of non-α7 nAChRs in ECVI neurons; p < 0.05, paired t test) (Fig. 6D). This increase did not occur when ACh was locally applied to ECIII or neighboring ECV neurons that were similar distances away from the recorded ECV neuron (n = 5; p > 0.05, paired t test), indicating that ECVI neurons act as a major source of excitatory inputs for ECV.
When we increased the duration of local application of ACh to the ECVI region to 1 min, we could now observe a significant increase in the frequency of sIPSC events in ECV neurons (Fig. 7B), demonstrating that inhibitory synaptic connections from ECVI to ECV neurons exist but they are relatively harder to activate. Increases in the frequency of sIPSCs did not occur when ACh was locally applied to the ECV or ECIII regions similar distances away from the recorded ECV neuron (Fig. 7B), demonstrating that non-α7 nAChR-containing inhibitory neurons that target ECV neurons exist mainly in the ECVI region. This increase in sIPSC frequency recorded from ECV neurons was also blocked by DHβE (1 μm; n = 3) (Fig. 7C), further confirming that these inhibitory ECVI neurons innervating ECV neurons were activated by ACh through the activation of non-α7 nAChRs. The increase in sIPSC frequency in ECV neurons was also prevented by pretreatment of TTX (1 μm; n = 3) (Fig. 7D), indicating that the increase in sIPSC frequency was a result of increased action potential frequency, not a consequence of higher synaptic vesicle release probability (often attributable to increased intracellular calcium in presynaptic terminals). Statistical analysis revealed a significant decrease in sIPSC interval by the local application of ACh to ECVI neurons (n = 5) (Fig. 7E), further suggesting that ECVI GABAergic neurons may provide a major source of inhibitory synaptic influence during systematic non-α7 nAChR activation.
Low concentration of nicotine enhances both amplitude and plasticity in Sb-ECVI synapses
Since we found that both the Sb and ECVI contain nicotine-sensitive neurons, and it is known that Sb neurons project to the ECVI (van Strien et al., 2009), we studied the effect of nicotine on Sb-ECVI synapses by recording evoked EPSCs (eEPSCs) from ECVI neurons while stimulating the Sb near the CA1 region. The bath application of a low concentration of nicotine (0.1 μm) increased the eEPSC amplitude by 40 ± 2% (n = 5) (Fig. 8A). The increase in eEPSC amplitude was also observed when nicotine was applied locally to the recorded ECVI neuron (33 ± 3%; n = 6) (Fig. 8A), and this effect was blocked by either DHβE (0.1 μm) (Fig. 8C) or MLA (10 nm) (Fig. 8D, compare filled symbols), demonstrating that the enhancement of Sb-ECVI synaptic transmission by nicotine is mediated by both α7 and non-α7 nAChRs at a nicotine concentration related to smoking. The PPR, which is inversely related to the presynaptic neurotransmitter release probability (Dobrunz and Stevens, 1997; Lagostena et al., 2008), was 1.70 ± 0.05 (n = 6) under control conditions and decreased significantly to 1.22 ± 0.08 (n = 6) after the local application of nicotine, suggesting a presynaptic mechanism of action (Fig. 8E).
Next we examined whether nicotine had any effect on synaptic plasticity in ECVI neurons. A tetanus stimulation (100 Hz, 100 ms) delivered to the Sb region elicited STP (<20 min) in the Sb-ECVI synapses (Fig. 8B). When the tetanus was preceded by the local perfusion of nicotine (0.1 μm) for 10 min, the STP was boosted to long-term potentiation (LTP; >50 min), an effect that was blocked by the non-α7 nAChR antagonist DHβE (1 μm) (Fig. 8C). Interestingly, the PPR during STP and LTP (1.72 ± 0.11 and 1.58 ± 0.07, respectively; n = 3) was not significantly different from control (1.69 ± 0.15 and 1.65 ± 0.1, respectively; n = 3), suggesting that this enhancement is attributable to a postsynaptic mechanism of action.
The bath application of the α7-selective antagonist MLA (10 nm) appeared to prevent the presynaptic action of nicotine since under these conditions, nicotine failed to enhance the amplitude of the eEPSC and alter the PPR in the presence of MLA (Fig. 8D,E). However, STP was still boosted to LTP (Fig. 8D). These data suggest that nicotine (at a concentration similar to the blood level in cigarette smokers) is acting presynaptically to enhance glutamate release via the activation of α7 nAChRs on presynaptic terminals and, in addition, is acting postsynaptically via non-α7 nAChRs to convert STP to LTP. A further indication of a postsynaptic mechanism of action for LTP is that dialyzing ECVI neurons with the calcium chelator BAPTA (10 mm) reduced the extent of LTP (30 min after the tetanus) by 70% (from 82 to 25%) (Fig. 8D).
We found that ECVI, a region in the HF not previously known to participate in the effects of nicotine, contains neurons that are highly responsive to nicotine under conditions that emulate the delivery of nicotine during smoking. These ECVI neurons were primarily glutamatergic with high RMPs and low h-currents and were capable of influencing synaptic activity to ECV neurons (i.e., by enhancing glutamatergic and GABAergic synaptic responses). In addition, the occurrence of LTP at Sb-ECVI synapses was facilitated by nicotine at a low concentration related to cigarette smoking. Therefore, future investigations focusing on the deep layers of the EC region are critical for our understanding of the cellular mechanisms of nicotine action and the potential role of nicotine in regulating memory.
Identification of nicotine-sensitive neuronal populations in brain slices using VSDI
In this study, we used the VSDI method to map the functional distribution of nicotine receptors to facilitate localization of nicotine-sensitive neuronal populations in hippocampal slices. VSDI is a relatively new technology in neuroscience (Takashima et al., 1999) that is a functional assay that offers high spatial and temporal resolution for imaging neocortical functions in living brain (Grinvald and Hildesheim, 2004). Although patch-clamp functional assays have superior sensitivity and reliability, the time-consuming nature of this single-cell approach does not easily lend itself to recordings from all cellular types in all brain regions. Thus, we used the VSDI technique to locate large areas of functional nAChRs that were activated by nicotine in the HF and used patch-clamp techniques to identify neuronal types in known locations of brain slices to characterize the functional properties of these responses. Future improvements in the sensitivity of the VSDI technique (e.g., more stable and sensitive dyes and better cameras) will undoubtedly increase the usefulness of this method for the functional mapping of other ion channels.
The α4β2 and α7 are the most likely nAChR subtypes responsible for nicotine effects in ECVI neurons
Thus far, six α and three β nAChR subunits (α2-7, β2-4) have been identified in neurons of the mammalian brain, with the most prevalent nAChRs in the hippocampus and cortex composed of the α7 and α4β2 subtypes (Wada et al., 1989; Sargent, 1993; Alkondon and Albuquerque, 2004). We have demonstrated the existence of functional α7 subunit-containing nAChRs in ECVI neurons based on their kinetics and pharmacological properties (Figs. 3⇑–5).
For the non-α7 nAChR-mediated responses, the α2, α3, and α6 subunits have not previously been found in deep EC layers, whereas both the α4 and β2 subunits have (Wada et al., 1989; Boyd, 1997; Sudweeks and Yakel, 2000; Mugnaini et al., 2002). In addition the α5 and β3 subunits are auxiliary subunits that carry no agonist binding sites (Ramirez-Latorre et al., 1996; Wang et al., 1996). Therefore, the most likely subunit composition of the non-α7 nAChRs mediating the nicotine-induced effects in the ECVI neurons are either the α4β2 or α4β4 subtypes. This composition is consistent with the fact that the nAChRs in ECVI neurons were almost completely inhibited (>90%) by a low concentration of DHβE (1 μm) (Harvey et al., 1996; Chavez-Noriega et al., 1997; Marks et al., 1999). Since α4β2 nAChRs are the most prevalent non-α7 nAChRs in the hippocampus and cortex (both of these subunits have previously been found to be expressed in the deep layers of the EC) and the nicotine sensitivity of α4β2 nAChRs (Briggs et al., 2006) is close to what we have observed in ECVI neurons (Fig. 3D), the α4β2 subtype is the most likely non-α7 nAChR subtype responsible for nicotine effects in the EC. However, we cannot completely rule out that other nAChR subunits, including the α5, β3, and β4 subunits, might be contributing. For example, the differences in nicotine sensitivity and decay rates among responses in CA1, SbSO, and ECVI neurons (Fig. 3) may be attributable to several possible mechanisms, including differences in the molecular makeup of the nAChRs (see above).
ECVI neurons may be important in mediating the effects of nicotine on memory
Our results suggest that ECVI neurons may be an important target of nicotine action in the HF. Studies regarding the mechanisms of nicotine influence on the HF have been primarily focused on the dentate gyrus and hippocampal proper regions (Jones and Yakel, 1997; Dani and Bertrand, 2007) where nicotine-sensitive neurons are mainly inhibitory GABAergic interneurons that express both α7 and non-α7 nAChRs (Alkondon and Albuquerque, 1993; Jones and Yakel, 1997; McQuiston and Madison, 1999; Frazier et al., 2003). Our data from both VSDI and patch-clamp recordings indicate that neurons from the deep EC layers (in particular ECVI) and SbSO, the portion of the Sb close to the deep EC layers, had larger responses to bath-applied nicotine (through the activation of non-α7 nAChRs) compared with the CA1 interneurons. All neurons that were studied in the ECVI, as well as most in the ECV, expressed functional non-α7 nAChRs, more so than the percentage of CA1 interneurons expressing non-α7 nAChRs. Because the bath application of low concentrations of nicotine (0.1 μm) to slices emulates the systematic introduction of nicotine during smoking (0.06–0.31 μm), or nicotine injection (Moriya and Hashimoto, 2004; Hukkanen et al., 2005; Moriya et al., 2006), the present study suggests that ECVI neurons may be an important site of nicotine action in the HF; this will have to be verified in future studies in vivo. These data are also consistent with a previous report in which nicotine-induced seizures upregulated immediate-early genes more in the EC regions that in other areas of the HF (Bastlund et al., 2005). Although most ECVI neurons had functional α7 nAChRs that were activated when the agonist was locally applied, these α7 receptors were not activated with bath-applied nicotine because of rapid desensitization. However, the expression of these receptors in the ECVI neurons also may be very important because of their influences on intracellular calcium concentration and potential role in influencing learning and memory in human and animal models (Welsby et al., 2006, 2007).
To investigate possible influences that the activation of ECVI neurons may have on other regions of the HF, we studied possible synaptic inputs to ECV neurons. These were chosen because their dendrites extend deep into the ECVI layer (Hamam et al., 2000; Egorov et al., 2003) and are in close proximity to each other. Direct connections from the ECVI to ECII/III, or more remotely to CA1 and dentate gyrus (Dugladze et al., 2001), are also likely but were not examined in this study. We found that the bath application of nicotine (10 μm) significantly enhanced glutamatergic but not GABAergic postsynaptic events recorded in the ECV neurons. In addition, brief (350 ms) local application of high concentrations of ACh (4 mm) to neurons in the ECVI region increase sEPSC events within 10 s of application, which did not occur when ACh application was applied to other regions of the EC. These results indicate that ECVI neurons provide an important source of excitatory synaptic influence to ECV neurons.
Although brief local application of ACh to ECVI neurons failed to increase sIPSC events in ECV neurons, more prolonged (1 min) local application of ACh to ECVI neurons significantly enhanced the frequency of GABAergic postsynaptic currents (sIPSC) in ECV neurons. This demonstrates that ECVI neurons also provide inhibitory synaptic influences to ECV neurons; however, these inhibitory effects were harder to activate and/or involved fewer ECVI neurons. These results suggest that under our experimental conditions, when the non-α7 nAChRs on ECVI neurons are activated, the enhancement of excitatory glutamatergic neuronal inputs to ECV neurons overcomes any possible inhibition caused by increases in neuronal GABAergic activities. This is consistent with our findings that the nicotine-responsive ECVI neurons are primarily pyramidal- or multipolar-shaped neurons, which are mainly excitatory in nature (Hamam et al., 2000).
We also demonstrated that a low concentration of nicotine (0.1 μm) enhances synaptic transmission (through presynaptic α7 and non-α7 nAChRs) and converts STP to LTP at Sb-ECVI synapses (through postsynaptic non-α7 nAChRs) in a cytoplasmic calcium-dependent pathway. A nicotine-dependent enhancement in plasticity was previously reported in synapses in CA1 and dentate regions through activation of α7 nAChRs (Fujii et al., 1999; Ji et al., 2001; Mann and Greenfield, 2003; Welsby et al., 2006, 2007), but it required a much higher concentration of nicotine (i.e., 1–5 μm). Our finding that non-α7 nAChRs also contributes to the facilitation of LTP occurrence is consistent with a recent report that α4-containing nAChRs contribute to LTP facilitation (Nashmi et al., 2007). These data further suggest that ECVI neurons may play a significant role in learning and memory during smoking.
Based on these results, we postulate that during smoking, ECVI neurons influence hippocampal excitability through their projection to ECV and maybe other regions (including ECII/III, dentate, and CA1 regions) and also potentiate synaptic transmission to ECVI neurons from Sb neurons. Additional studies are needed to better understand the role of ECVI neurons in the influence of nicotine on memory.
This work was supported by the Intramural Research Program of the National Institutes of Health–National Institute of Environmental Health Sciences. We are grateful to Drs. David Armstrong, J. Victor Nadler, Serena Dudek, and Christian Erxleben for providing valuable suggestions in experiments and in the writing of this manuscript.
- Correspondence should be addressed to Jerrel L. Yakel, National Institute of Environmental Health Sciences, F2-08, P.O. Box 12233, 111 T. W. Alexander Drive, Research Triangle Park, NC 27709.