Axonal α7* nicotinic acetylcholine receptors modulate glutamatergic signaling and synaptic vesicle organization in ventral hippocampal projections

Abstract

Modulation of the release of glutamate by activation of presynaptic nicotinic acetylcholine receptors (nAChRs) is one of the most prevalent mechanism of nicotinic facilitation of glutamatergic transmission in cortico-limbic circuits. By imaging gene chimeric co-cultures from mouse, we examined the role of α7* nAChRs mediated cholinergic modulation of glutamate release and synaptic vesicle organization in ventral hippocampal projections. We directly visualized exogenous and endogenous cholinergic facilitation of glutamate release in this specialized preparation of circuits in vitro. Disrupting α7* nAChRs mediated cholinergic signaling genetically or pharmacologically diminished cholinergic facilitation of glutamate release at presynaptic terminals. Alteration of α7* nAChRs mediated cholinergic signaling along glutamatergic axons also decreased functional synaptic vesicle clustering to presynaptic terminals. These findings suggest that presynaptic α7* nAChRs contribute to cholinergic modulation of glutamate release and synaptic vesicle organization.

Introduction

Cholinergic signaling plays critical functions in the mammalian brain, contributing to attention, wakefulness and the formation and recall of memories (Ballinger et al., 2016). The importance of these functions is underscored by the connections between disruption in the cholinergic system and conditions ranging from thought disorders to cognitive impairment and dementia. Understanding how cholinergic inputs influence various forebrain circuits is essential to defining the role of acetylcholine in these conditions (Freedman et al., 2001; Mathew et al., 2007; Barreto et al., 2015; Lim et al., 2015; Alber et al., 2020; Bohnen et al., 2022; Polli and Kohlmeier, 2022).

Cholinergic projection neurons of the basal forebrain and brainstem send axons to virtually all regions of the brain where they branch extensively allowing a relatively small number of neurons to contribute to an array of critical functions. Acetylcholine acts via two general classes of receptors, the G-protein coupled muscarinic receptors (mAChRs) and the ligand gated cation channel nicotinic receptors (nAChRs). The most abundant nAChRs in the brain are the α4β2* and α7* nAChRs (the nicotinic ACh receptors are pentamers comprised of all alpha or combinations of alpha and beta subunits. We use the convention of αβ* and α* to indicate that the subunit stoichiometry is not known) (Kutlu and Gould, 2015; Wittenberg et al., 2020). α4β2* and α7* nAChRs are found at pre-synaptic as well as post-/extra-synaptic sites (Jones and Wonnacott, 2004; Zhong et al., 2008; Cheng and Yakel, 2014). Activation of axonal and/or pre-synaptic nAChRs affects the release of many neurotransmitters including glutamate, dopamine, GABA and even ACh (McGehee et al., 1995; Gray et al., 1996; Kramer et al., 2022; Liu et al., 2022). nAChR activity also is linked to synapse development and maturation (Berg and Conroy, 2002; Lozada et al., 2012). Taken together, there is strong evidence that nAChRs play multiple roles in the formation and function of brain circuits.

Axonal α7*nAChRs can increase glutamate release probabilities by elevating intracellular Ca²⁺-signaling via multiple mechanisms. First, α7*nAChR activation leads to local depolarization and opening of voltage dependent calcium channels. Second, α7* nAChRs are calcium permeable, allowing rapid and local increases in intracellular calcium. When localized close enough to neurotransmitter release sites, α7*nAChR gated calcium transients contribute to synaptic vesicle exocytosis, increasing the probability of transmitter release (Fucile, 2004; Turner, 2004; Fayuk and Yakel, 2005; Shen and Yakel, 2009; Cheng and Yakel, 2015a,b; Cheng et al., 2021).

Activation of axonal α7*nAChRs also leads to sustained increases in intracellular calcium signaling; a response linked to the coupling of ligand-activated α7*nAChRs to heterotrimeric G proteins (Zhong et al., 2013; Nordman and Kabbani, 2014; King et al., 2015). The ability of α7*nAChRs to activate second messenger signaling allows these receptors to regulate axonal actions ranging from growth cone dynamics to sustained glutamate release (Nordman and Kabbani, 2012; Kabbani and Nichols, 2018). Prior studies have predominantly used pharmacological agents to probe axonal α7*nAChR function leaving open the question to the extent to which axo-axonal interactions between cholinergic axons and target axons elicit similar α7*nAChR dependent signaling.

Glutamatergic synaptic transmission requires calcium dependent release of glutamate from presynaptic terminals. Although glutamatergic transmission, and its modulation by pre-synaptic nAChRs, has been studied in detail at individual synapses electrophysiologically, optical reporters of synaptic vesicle recycling and neurotransmitter release allow direct measurement of presynaptic phenomena, allowing us to address questions related to the effects of α7* nAChR signaling on the dynamics of synaptic vesicular cycling (Liu and Tsien, 1995; Moulder et al., 2006; Alabi and Tsien, 2012; Marvin et al., 2013, 2018; Zhong et al., 2015, 2017; Helassa et al., 2018).

In this study we used a variety of imaging modalities to examine exogenous and endogenous cholinergic modulation of glutamate release and synaptic vesicle recycling in ventral hippocampal glutamatergic projections using gene-chimeric micro-slice co-cultures. We provide evidence for two distinct functions of axonal α7* nAChR signaling. The first involves sustained increases in glutamate release in response to both levels of nicotine approximating those delivered to the CNS by smoking and by levels of acetylcholine released from basal forebrain cholinergic neurons. The second function for axonal α7* nAChRs is in the formation of functional glutamate release sites by contributing to the organization of synaptic vesicles.

Materials and methods

Materials

Mice

C57Bl6/J (000668), B6.129S7-Chrna7^TM1Bay/J (003232) and B6;129S6-Chat^TM2(cre)Lowl/J (006410) mice were obtained from Jackson Labs, the Chat-tau-eGFP BAC transgenic line was a gift from Dr. S. Vijayarhagavan, U. Colorado (Grybko et al., 2011). All mice were maintained on 12 h light: 12 h dark cycles with constant temperature (72°F) and humidity (40%). Mice were allowed ad libitum access to food and water. All animal handling was done in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals (NIH Publications No. 80-23, revised 2012) and was approved by either the Stony Brook University Institutional Animal Care and Use Committee (#1618) or the NINDS ACUC (ASP#1490).

Viruses

Packaged preparations of AAV₉.hSyn.iGluSnFr.WPRE. SV40 (gift from Loren Looger) and AAV₉-EF1a-DIO-C1V1(E122T/E162T)-TS-mCherry (gift from Karl Deisseroth) were obtained from Addgene (Addgene viral prep #98929-AAV9; Addgene_98929) or the University of North Carolina viral vector core, respectively.

Antibodies and other reagents

vGluT1 (Synaptic systems, #135 303), pan-axonal neurofilament marker (SMI312, Sigma-Aldrich, # NE1022), Synapsin 1 (EMD Millipore, #1543), phospho (Ser62, Ser67) Synapsin 1 (EMD Millipore, #AB9848), alpha-bungarotoxin (Sigma-Aldrich, #B137), alpha-bungarotoxin-594 (Thermo Fisher #B-13423), FM1-43 (Thermo Fisher #T35356), Fluo4 (Thermo Fisher #F14217), tetrodotoxin (Tocris #1078), bicuculline (Tocris #0130), RuBi-nicotine (Tocris, #3855), D-AP-5 (Tocris #0106), and CNQX (Tocris #1045).

Methods

Micro-slice cultures

The cultures were prepared as described previously (Zhong et al., 2015). Briefly, for ventral hippocampal micro-slice cultures, the ventral CA1 and subiculum area (vHipp) from WT (+/+) or α7 knockout (−/−) mice (postnatal days 0–3, P0–P3) was dissected, sliced into ∼150 μm × 150 μm pieces, and then plated onto poly-D-lysine/laminin-coated glass coverslips (12 mm, BD Sciences) in 24-well plates with a minimal volume (50 μl) of culture media [Neurobasal, 2% B-27 (GIBCO) and 20 ng/ml of brain-derived neurotrophic factor (Invitrogen)]. After the micro-slices attached to the substrate, 100 μl of additional culture media was added. For medial septum/diagonal band (MS/DB) cultures, the MS/DB area (Bakker et al., 2015; Staib et al., 2018) from ChAT-tau-eGFP or ChAT-IRS-Cre mice (P0–P3) was dissected, sliced and plated on coverslips as described before (Zhong et al., 2015) and above for vHipp cultures.

For vHipp-nucleus accumbens (nAcc) synaptic co-cultures, nAcc (ED18-P1) from WT mice (C57BL/6J) were dissected out and dispersed with 0.25% trypsin (GIBCO) for 15 min at 37°C, followed by gentle trituration in culture media. Dispersed nAcc neurons were plated onto poly-D-lysine/laminin-coated glass coverslips at 0.25 ml/coverslip. The next day vHipp micro-explants [prepared as described above and in Zhong et al. (2008)] were added to the same coverslips.

AAV infections

The genetically encoded glutamate indicator, iGluSnFr or the red-shifted opsin, C1V1, were expressed by adding AAV-hSyn-iGluSnFr-WPRE-SV40 (10¹³ vg/ml) or AAV-Ef1a-DIO C1V1 (t/t)-TS-mCherry (2 × 10¹² vg/ml) directly to the coverslip (0.25 μl/well). Twenty-four hours later the culture media was changed and replaced with virus-free media. All cultures were maintained in a humidified 37°C, 5% CO₂ incubator 7–10 days post-infection before performing imaging experiments. For co-culture experiments involving vHipp micro-slices and dispersed nAcc neurons, the nAcc neurons were plated and infected first and vHipp micro-slices were added to the same coverslips 24 h later following virus wash-out. For vHipp – MS/DB co-cultures, vHipp explants from C57Bl6/J mice were plated and infected with AAV-hSyn-iGluSnFr-WPRE-SV40 on day 1 and MS/DB micro-slices from Chat-IRES-Cre mice were plated on the same coverslips 24 h later. Following plating of the MS/DB micro-slices, AAV-Ef1a-DIO C1V1 (t/t)-TS-mCherry was added to the coverslips and infection allowed to proceed for 24 h following which the culture media was replaced. In this way, iGluSnFr and C1V1 expression were limited to vHipp projections or MS cholinergic neurons, respectively.

FM1-43-based live imaging and analysis

After 7–10 days in vitro, coverslips with cultures were transferred and sealed to the bottom of an imaging Ludin chamber (Live Imaging Services, Switzerland) and then mounted on an Olympus IX81 DSU microscope (spinning disk confocal, Olympus America) under continuous superfusion (1 ml/min) with HEPES buffered saline (HBS, 135 mM NaCl, 5 mM KCl, 1 mM MgCl₂, 2 mM CaCl₂, 10 mM HEPES, and 10 mM glucose, pH 7.35) containing 2 μM tetrodotoxin (TTX), 10 μM bicuculline, 50 μM D-AP-5, and 20 μM CNQX. KCl induced depolarization (Amaral et al., 2011) was used for loading FM1-43 into axons. Cultures were loaded with 10 μM FM1-43 in 56 mM KCl ACSF for 90 s, external dye was washed away in Ca²⁺-free HBS, containing ADVASEP-7 (0.1 mM; Sigma, # A3723, to scavenge membrane-bound FM1-43) for 15 min. Fluorescence images of vHipp and/or MS/DB axons were collected by a Plan-Apochromat objective (60× oil with 1.4 NA, excitation 488 nm, emission 530 nm) and captured with a CCD camera (Hamamatsu) every 1.5 s for 5 min. Image acquisition was performed using Slidebook software (version 5; Olympus). After 1 min of baseline data collection, 56 mM KCl in HBS without FM1-43 was applied for 120 s to confirm that high K⁺ depolarization induced de-staining of FM1-43 dye–filled vesicles. The total amount of releasable FM1-43 fluorescence at each synaptic bouton was calculated from the difference between fluorescence intensity after staining (but before de-staining) and after nicotine (1 μM) application (applied by rapid perfusion (2 ml/min) for 1 min), KCl depolarization, or opto-stimulation (100 total 5 ms pulses at 10 Hz) induced de-staining (ΔF = F_staining − F_{de–staining}). The fraction of fluorescence intensity decrease after depolarization was calculated as F_decrease% = ΔF/F_staining. The number and size of FM1-43 positive puncta before and after nicotine application, or KCl depolarization was measured and compared along vHipp axons from α7+/+ wild type vs. α7−/− knockout mice using MetaMorph software. The lengths of axonal projections were also measured by tracing vHipp and/or MS projections in MetaMorph.

For analyzing the time course of nicotine induced FM1-43 de-staining, all frames of the raw FM1-43 fluorescence images were saved as Slidebook files and then exported as a series of TIF format images that were then imported to MetaMorph software and transferred as Z-stack images for further analyses. After setting the threshold of the FM1-43 fluorescence, the integrated intensity of the FM1-43 signals along vHipp axons before and after nicotine application was calculated. FM1-43 fluorescence data are displayed as a normalized integrated intensity: ΔF/F₀ = (F − F₀)/F₀, where F₀ is the background-corrected pre-nicotine FM1-43 fluorescence. Data were analyzed further using Excel and GraphPad Prism 9 software.

iGluSnFr-based live imaging and analysis

After 7–10 days in vitro, cultures expressing iGluSnFr were transferred to an imaging chamber and continuously superfused with HBS cocktail as described above for FM1-43 live imaging. Fluorescence images of iGluSnFr under various culture conditions were collected every 2 s for 20 min. After 1–2 min of baseline data collection, glutamate (100 μM), or nicotine (1 μM) was applied by rapid perfusion (2 ml/min) for 1 min. For testing the effect of endogenous ACh release on glutamate release, C1V1 expressing MS/DB projections were stimulated with a 594 nm LED light (5 ms pulse duration, 10 Hz for 10 s) after 2 min baseline data collection.

All frames of the raw iGluSnFr fluorescence images were saved as Slidebook files and then exported as a series of TIF format images that were then imported to MetaMorph software and transferred as Z-stack images for further analyses. After setting the threshold of the iGluSnFr fluorescence, the integrated intensity of the fluorescence signals before and after glutamate or nicotine application or following opto-stimulation were collected and calculated. Fluorescence data are displayed as a normalized integrated intensity: [ΔF/F₀ = (F −F₀)/F₀], where F₀ is the background-corrected pre-stimulation iGluSnFr fluorescence. Data were analyzed further using Excel software. The iGluSnFr data from vHipp-nAcc and MS/DB-vHipp synaptic co-cultures are displayed in box plots (Statview and GraphPad Prism 9 software) where the boxes include data points between the twenty-fifth percentile (bottom line) and the seventy-fifth percentile (top line). The middle line indicates the fiftieth percentile (median). Vertical lines mark the fifth and ninety-fifth percentiles.

Electron microscopy

After 7–10 days in vitro, vHipp micro-slices grown on Thermanox plastic coverslips (NUNC/Electron Microscopy Sciences) were fixed for 30 min in a mixture of cold 2% paraformaldehyde and 2% glutaraldehyde in 0.1 M phosphate buffer (PB), pH 7.4. After several washes in PB, explants were postfixed with 2% osmium tetroxide for 30 min, en bloc stained with aqueous 1% uranyl acetate for 30 min, dehydrated through an ascending series of ethanol concentrations, followed by acetonitrile, and embedded in Embed 812 resin (Electron Microscopy Sciences) for 48 h at 60°C. Ultrathin sections (60–90 nm) were cut and then stained with 1% methanolic uranyl acetate and 0.3% aqueous lead citrate, and observed with a JEOL 1200EX transmission electron microscope.

Immunocytochemistry and reagents

After 7–10 days in vitro, cultures were fixed in 4% paraformaldehyde/4% sucrose/PBS [20 min, room temperature (RT)], permeabilized with 0.25% Triton X-100/PBS (5 min, RT), blocked with 10% normal donkey serum in PBS (30 min, RT), and then incubated in primary antibodies overnight at 4°C. The following primary antibodies were used: anti-vesicular glutamate transporter 1 (1:250), anti-pan-axonal neurofilament marker (1:1000), anti-synapsin1 (1:500), and anti-phospho-Synapsin1 (1:500). Cultures were washed and incubated in secondary antibodies conjugated to Alexa Fluor 488 (1:500) or Alexa Fluor 594 (1:500) for 1 h at RT. Slips were mounted using VectaShield (with DAPI, Vector Laboratories), and images were captured using a microscope (Axio Imager A1; Carl Zeiss) equipped with Plan-Apochromat objectives (63× oil with 1.4 NA), a CCD camera (Hamamatsu), and MetaMorph software (version 7.1, MetaMorph Microscopy Automation and Image Analysis Software, Molecular Devices). To label surface α7* nAChRs after live imaging of nicotine-induced FM1-43 de-staining, vHipp cultures were incubated in αBgTx conjugated to Alexa 594 (1:1000; Molecular Probe) for 45 min at RT prior to fixation.

Statistical analysis

All data were tested for normality with the D’Agostino-Pearson test and expressed as mean ± SEM unless otherwise indicated and analyzed with StatView (SAS Institute), Microsoft Excel (Microsoft) and Prism 9 (GraphPad by Dotmatics) software. Data sampling units and p-values correspond to the statistical tests for all data collected are shown in Table 1.

Results

α7* nicotinic acetylcholine receptors are required for nicotine-induced vesicular release along vHipp axons

We used FM1-43 de-staining to investigate the dynamics of nicotine induced exocytosis from individual synaptic vesicle clusters along vHipp axons. We used KCl induced depolarization to load vHipp axons from either WT or α7 knockout mice with FM1-43 and then quantified the time course of nicotine-induced FM1-43 exocytosis from individual synaptic vesicle clusters (three pixels, ∼1 μm) by live imaging (every 1.5 s for 5 min) (Figures 1A,B). We plotted the fraction decrease of normalized integrated intensity at single clusters after nicotine application and then calculated the time constants for fluorescence decay along WT (trace in red) and α7 knockout (trace in black) vHipp axons (Figure 1C; traces are averages of recordings along multiple axon segments; the shaded areas around each trace represent standard deviations). Following nicotine application, FM1-43 fluorescence in vesicle clusters along WT vHipp axons decreased by ∼80% with a decay time constant of ∼4 s. In contrast, nicotine only reduced FM1-43 fluorescence by ∼20% with a time constant of ∼16 s from vesicle clusters along α7 knockout axons (Figures 1D,E; WT vs. Knockout, p < 0.001; see Table 1 for additional details regarding n values and statistical analyses).

FIGURE 1

To assess the spatial relationship between α7* nAChRs and sites of nicotine induced FM1-43 de-staining, we used αBgTx conjugated to Alexa 594 to visualize surface α7* nAChRs following nicotine-induced FM1-43 de-staining (Figures 2A,B). Over 80% of individual puncta along vHipp axons at which nicotine induced robust FM1-43 de-staining also stained with αBgTx-Alexa 594 indicating the presence of surface α7* nAChRs (Figure 2C). These double labeled vesicle clusters showed rapid and nearly complete (time constant ∼4 s, 80% decrease in FM1-43; red trace in Figure 2D) nicotine induced FM1-43 exocytosis. In contrast, vesicle clusters that were not stained with αBgTx –Alexa 594 showed significantly slower (time constant of ∼14.0) and only partial (<25% intensity decrease) de-staining (trace in black) (Figures 2D–F); values that were very similar to those recorded from α7 knock-out axons. The effect of nicotine on neurotransmitter vesicle release showed two distinct responses – an α7* nAChR dependent response that was associated with close apposition of surface α7* nAChRs to the sites of release and that was kinetically distinct from the second non-α7* nAChR, presumably α4β2* nAChR (Zhong et al., 2013), mediated response.

FIGURE 2

A dynamic synapsin 1 phosphorylation/dephosphorylation cycle is essential for maintaining synaptic vesicle organization and abundance in axon terminals (Bykhovskaia, 2011; Shupliakov et al., 2011; Patzke et al., 2019). To determine if synapsin 1 phosphorylation is a target for α7* nAChRs signaling, we examined the effect of acute nicotine or chronic αBgTx treatment on axonal levels of phospho-synapsin 1. vHipp micro-slices from α7+/+ or α7+/+ treated for 7-days with αBgTx, were exposed to nicotine (1 μM, 1 min), fixed, permeabilized, and stained with antibodies recognizing phospho-Synapsin 1 (serines 62 and 67; Figure 3A, green) or total synapsin 1 (Figure 3B, green), as well as pan-axonal (SMI312; red) antibodies. Levels of phospho-synapsin 1 increased following nicotine, in an α7* nAChR-dependent manner (blocked by αBgTx; Figures 3A,C; p < 0.01). Neither nicotine nor αBgTx affected the total level of axonal synapsin 1 (Figures 3B,D). These data are consistent with α7* nAChR mediated signaling via synapsin 1 phosphorylation, contributing to local modulation of synaptic vesicle cycling.

FIGURE 3

α7* nicotinic acetylcholine receptors are required for sustained nicotine induced glutamate release

Modulation of the release of glutamate by activation of presynaptic nAChRs is a prevalent mechanism of nicotinic facilitation of glutamatergic transmission in the CNS. To visualize α7* nAChRs modulation of glutamate release, we utilized the genetically encoded fluorescent glutamate indicator, iGluSnFr. Dispersed medium spiny neurons (MSNs) from wild type nAcc were plated and transfected with iGluSnFr before the addition of vHipp micro-slices prepared from either wild type or α7 knock-out pups. We monitored changes in fluorescence of iGluSnFr expressed on the disperse nAcc MSNs following application of nicotine to the co-cultures (Figure 4A). A 1 min nicotine (1 μM) application resulted in sustained increase of iGluSnFr fluorescence intensities around nAcc MSNs soma co-cultured with WT (α7+/+) vHipp (Figures 4B–D; p < 0.01). No significant increase in iGluSnFr fluorescence intensity was seen in WT MSNs cultured with α7 knockout vHipp (Figures 4B–D). These data, consistent with our previous electrophysiological results (Zhong et al., 2008), demonstrate that presynaptic α7* nAChRs mediate the sustained responses to nicotine.

FIGURE 4

Next, we took advantage of the ability of iGluSnFr to sum glutamate signals (Helassa et al., 2018) to image nicotine modulated release sites along vHipp axons. We infected vHipp micro-explants with AAV.hSyn.iGluSnFr and confirmed axonal iGluSnFr expression by imaging responses to glutamate (100 μM, 1 min) added to the cultures. iGluSnFr ΔF/F_o increased from ∼0 to 0.07 during the glutamate applications and returned to baseline following washout (Figures 5A,B). Having established that we could record changes in iGluSnFr fluorescence along vHipp projections, we used iGluSnFr to assay the effect of nicotine application on glutamate release. Nicotine application (1 μM, 1 min) increased iGluSnFr fluorescence intensities (ΔF/F_o ∼4%). After washout of nicotine, iGluSnFr fluorescent intensity continued to increase and then remained elevated for at least 20 min (baseline vs. 1 min post-nicotine: p < 0.01; baseline vs. 20 min post-nicotine, p < 0.01). This robust and sustained response was abolished by preincubation with alpha-bungarotoxin (αBgTx), an α7* nAChRs selective antagonist (Figures 5C,D). Although we are plotting the integrated fluorescence intensity along vHipp axons in Figure 5C, the increase in signal was focal (Figure 5C, inset), consistent with stimulation of glutamate release from discrete sites.

FIGURE 5

Endogenous acetylcholine induces sustained glutamate release along vHipp axons

We next sought to determine whether endogenously released ACh would also stimulate glutamate release from vHipp axons. To test this, we co-cultured vHipp explants with micro-slices of the medial septum/diagonal band of broca (MS/DB). First, we stained MS/DB micro-slices from ChAT-tau-GFP mice cultured alone with vGluT1 antibodies to confirm that these MS/DB micro-slices extend cholinergic (GFP⁺) but not glutamatergic (vGluT1^–) projections (Figure 6A). Next, we infected MS/DB micro-slices from ChAT-IRES-Cre mice with AAV-Ef1a-DIO.C1V1.(t/t)-TS-mCherry to express the red-shifted channel rhodopsin variant, C1V1, only in cholinergic neurons (Figure 6B, pan-axonal marker in green, C1V1 in red). We monitored the effects of focal optogenetic stimulation (594 nm LED light, 5 ms pulses at 10 Hz for 10 s) of cholinergic fibers in three ways: (1) axonal Ca²⁺ dynamics following loading of cultures with Fluo 4; (2) FM1-43 vesicle de-staining; and (3) we looked for evidence of glutamate release by co-expressing iGluSnFr (Figures 6C,D). Photo-stimulation of C1V1 expressed along MS/DB cholinergic axons increased axonal calcium levels (p < 0.001) and induced vesicular release of FM1-43 (p < 0.001). We did not detect any release of glutamate from these explants following C1V1 stimulation (iGluSnFr fluorescence intensity did not change, Figure 6D).

FIGURE 6

Next, we plated co-cultures of MS/DB (from ChAT-tau-GFP pups) and vHipp (from C57Bl6 pups) micro-slices and stained for either vGluT1 or surface α7* nAChRs. Cholinergic projections (in green) from MS/DB micro-slices intermingled with glutamatergic projections (in red) from WT vHipp micro-slices (Figure 7A, top). vGluT1 and α7* nAChRs positive staining were detected along vHipp projections (Figure 7A, bottom). We did not detect any α7* nAChRs along MS/DB projections using this micro-slice preparation.

FIGURE 7

Having established that in this co-culture preparation cholinergic and glutamatergic neurons were physically intermingled we next plated and infected WT vHipp micro-slices with AAV₉-hSyn.iGluSnFr and then 1 day later added MS/DB micro-slices from ChAT-Cre animals to the same coverslips and infected the co-culture with AAV-EF1a.DIO.C1V1-mCherry (Figure 7B). This allowed us to quantify glutamate release along the vHipp axons (expressing iGluSnFr) following optostimulation of C1V1 (expressed in cholinergic neurons). We stimulated C1V1 (594 nm, 5 ms, 10 Hz for 10 s) and measured iGluSnFr fluorescence. Optogenetic stimulation of ACh release resulted in a sustained increase in iGluSnFr fluorescence along vHipp axons with close contacts with C1V1 expressed cholinergic axons (∼10-fold increase in ΔF/F_o for at least 15 min following light stimulation; Figures 7C,D, red trace). There were no changes in iGluSnFr fluorescence detected along vHipp axons without close contacts with C1V1 expressed cholinergic axons in the same cultures (Figure 7D, black trace). The vHipp axon iGluSnFr response following optostimulation of cholinergic axons was blocked by the selective nicotinic receptor antagonist mecamylamine, but not by the muscarinic receptor antagonist atropine (Figure 7E). These results demonstrate that brief stimulation of ACh release (10 s) from septal-cholinergic neurons can facilitate sustained (15 min) vHipp glutamate release via presynaptic nAChRs.

α7* nicotinic acetylcholine receptor signaling regulates synaptic vesicle clustering along vHipp axons

Previously we reported that FM1-43 labeled puncta along α7−/− vHipp axons were more numerous but smaller than FM1-43 positive puncta along WT vHipp axons (Zhong et al., 2013). We found similar differences in our current study. Although FM1-43-labeling was equivalent in WT (+/+) (Figure 8A, left top) and α7 knockout (−/−) (Figure 8A, right top) vHipp axons, and showed no differences in the initial FM1-43 fluorescence intensities after K⁺-dependent loading (Figure 8B), the α7 knockout (−/−) vHipp axons had more (KO: 38.4 ± 4.2 clusters/100 μm vs. WT 27.8 ± 3.1 clusters/100 μm; p < 0.01) and smaller FM1-43 positive puncta (measured in fluorescence pixels) than the WT vHipp axons (KO: 0.4 ± 0.3 vs. WT: 1.78 ± 0.8; p < 0.01) (Figures 8C,D). Despite the differences in FM1-43 labeled puncta number, size and nicotine responsiveness, FM1-43 loading, and de-staining was equivalent in α7 knock out and WT axons. vHipp micro-slices from either WT or α7 knockout mice were loaded with FM1-43 and then KCl –induced de-staining of the readily releasable pool of vesicles was quantified. After high K⁺ (56 mM) application, the fluorescence intensity of all puncta along WT (+/+) vHipp axons (Figure 8A, left bottom) decreased by ∼85% (Figure 8E), reflecting depolarization-induced exocytosis of synaptic vesicles. The decrease in depolarization-induced fluorescence intensity of FM1-43 along α7 knockout (−/−) vHipp axons was the same as WT (Figure 8A, right bottom; Figure 8E).

FIGURE 8

To examine further the effect of α7* nAChRs on vesicle organization we used transmission electron microscopic imaging of ultrathin sections of vHipp micro-slice cultures with extensive axonal arbors. Varicosities containing clusters of synaptic vesicles (≥15 vesicles of 30–60 nm diameter, within less than a vesicle diameter of one another) were clearly seen along vHipp axons (Figure 9A). There were fewer clusters per 100 μm of α7 knockout (−/−) vHipp axon compared with WT (+/+) control (Figure 9B): 8 ± 5 per 100 μm of α7 knockout vHipp axon compared with 29 ± 8 per 100 μm of WT vHipp axon (p < 0.01). When we quantified the total number of vesicles, we found more per μm² of α7 knockout (−/−) vHipp axon compared with WT (+/+) control (Figure 9C): 242 ± 8 per μm² of α7 knockout vHipp axon compared with 108 ± 5 per μm² of WT vHipp axon (p < 0.01). To confirm that these differences were directly related to loss of α7* nAChRs signaling, we prepared WT vHipp micro-slices and maintained them in the presence of αBgTx for 7 days prior to processing for electron microscopy. αBgTx blockade of α7* nAChRs decreased the number of vesicle clusters (Figure 9A, middle, Figure 9B) (12 ± 8 per 100 μm of vHipp axon, p < 0.01) and increased the total number of vesicles (Figure 9C) (218 ± 8 per μm² of vHipp axon, p < 0.01) compared to WT vHipp axon. Taken together, these findings demonstrate that α7* nAChRs signaling regulates synaptic vesicle abundance and the organization of synaptic vesicles into distinct clusters.

FIGURE 9

Discussion

In this report we demonstrate that α7* nAChRs localize to glutamatergic release sites along vHipp axons where they mediate glutamate release in response to either applied nicotine or ACh released from MS/DB cholinergic axons. We also demonstrate that the absence of functional α7* nAChRs (genetically deleted or chronically inhibited with αBgTx) alters the organization of synaptic vesicles along vHipp axons. These results provide evidence for a dual role of axonal α7* nAChRs during both the maturation of presynaptic-like specializations and in modulating glutamate release in mature projections.

The presence and potential functional role of pre-synaptic nAChRs in the regulation of glutamate release has been demonstrated with a variety of approaches from synaptosome preparations to in vitro, ex vivo and in vivo opto-physiology (Vidal and Changeux, 1989; McGehee et al., 1995; Gray et al., 1996; Threlfell et al., 2012; Kosillo et al., 2016). In the BLA, glutamatergic transmission is enhanced by nicotine or acetylcholine released following optogenetic stimulation of cholinergic terminals and is thought to underlie sustained changes in synaptic potentiation associated with fear learning (Jiang and Role, 2008; Jiang et al., 2013, 2016). In prior in vitro and ex vivo studies we have defined two distinct responses, transient increases in glutamate release mediated by non-α7* nAChRs and sustained changes in release that require α7*nAChRs. A limitation of these prior studies was our inability to visualize axonal nAChRs relative to sites of glutamate release. In this study we leveraged the power of micro-explant co-cultures to directly image aspects of nAChRs-mediated axonal modulation of glutamatergic glutamate release, both by using FM1-43 destaining and using an axonally expressed glutamate sensor, iGluSnFr (Marvin et al., 2013, 2018; Helassa et al., 2018).

Contribution of non-α7* nicotinic acetylcholine receptors to nicotine induced glutamate release

α7* nAChRs localized to synaptic vesicle clusters mediate nicotine induced glutamate release. Both α4β2* and α7* nAChRs are associated with axo-axonic regulation of the release of many transmitters with studies of glutamate, GABA and dopamine being the most common (Dickinson et al., 2008; Livingstone et al., 2009; Threlfell et al., 2012). Non-α7*nAChRs (α4β2*nAChRs) mediate rapid axonal responses to nicotine (Zhong et al., 2008, 2013) in synaptic co-culture preparations as well as in response to ACh release in acute striatal slice (Kramer et al., 2022; Liu et al., 2022). Here we show that although nicotine treatment leads to general FM1-43 de-staining along vHipp axons, closer examination reveals two populations of FM1-43 labeled clusters that are distinguished both by the rate and degree of destaining, and by the proximity of surface α7* nAChRs. Clusters that destained rapidly and completely were associated with α7* nAChRs, whereas those that showed slower and incomplete de-staining were not (Figure 2). It is important to note that these latter release sites were not inherently dysfunctional – all FM1-43 sites rapidly and completely de-stained in response to K⁺ depolarization. In some instances, we found the two classes intermingled along the same axon segments (the two examples shown in Figure 2C were within ∼5 microns of each other). The α7* nAChR signals that lead to sustained responses are apparently spatially constrained and do not significantly spread within the axon. It is not clear what factors spatially restrain the α7* nAChR signals, but these findings underscore the importance of appropriate precise targeting of axonal α7* nAChRs.

FM1-43 marks sites of synaptic vesicle cycling but does not distinguish between different types of synaptic vesicles. We used the genetically encoded glutamate reporter, iGluSnFr, to probe α7*nAChR contribution more directly to glutamate release per se along vHipp axons. A brief exposure to nicotine increased iGluSnFr fluorescence along vHipp axons for tens of minutes; a response that required functional α7* nAChRs in the vHipp neurons (Figure 5). In contrast, glutamate application (100 μM, 1 min) increased iGluSnFr transiently with fluorescence returning to baseline levels with a t_1/2 of 2–3 min. Given the reversibility of the iGluSnFr signal following glutamate wash-out, the continued rise in fluorescence after nicotine washout likely reflects continuing, elevated release of glutamate from the vHipp projections involving downstream intracellular signaling cascades.

The minimal non-α7*nAChR mediated response to nicotine, either in terms of FM1-43 destaining or iGluSnFr fluorescence, is difficult to interpret. Previously, using patch clamp recordings in vHipp – nAcc co-cultures, we demonstrated that nicotine increased mEPSP frequency via both a sustained α7* nAChR-dependent and a transient, α7 nAChR-independent mechanism (Zhong et al., 2008). We have also seen clusters of α4* nAChR on vHipp axons that were similar in frequency (cluster/100 μm axon) to α7* nAChR clusters (Zhong et al., 2013). One possibility is that transient increases in synaptic vesicle fusion and glutamate release, as measured by mEPSP frequency (Zhong et al., 2008) are insufficient to give a detectable FM1-43 or iGluSnFr signal under our imaging conditions. Alternatively, the full maturation of a non-α7* nAChR responsive pre-synaptic site might require interactions with post-synaptic targets. This would be an intriguing distinction but requires further study. In sum, our results are consistent with prior studies showing that transient nicotine activation of pre-synaptically localized α7* nAChRs results in sustained glutamatergic synaptic transmission (Jiang and Role, 2008; Zhong et al., 2008, 2013; Jiang et al., 2013, 2016).

Release of acetylcholine from medial septum/diagonal band axons elicits sustained glutamate release

Much of the literature exploring cholinergic modulation of synaptic transmission, especially with a focus on presynaptic mechanisms, has utilized pharmacological approaches – either nicotinic or muscarinic specific drugs, or applied ACh. We took advantage of this co-culture system to demonstrate that ACh released from cholinergic axons could directly activate presynaptic/axonal nAChRs leading to sustained facilitation of glutamate release (Figure 7). These responses were only seen in vHipp axons in close proximity to C1V1 expressing cholinergic axons – vHipp axons from the same explants but not in apposition to C1V1⁺ axons did not visibly respond to ACh release (red vs. black traces in Figure 7D). This is consistent with the formation of specialized axo-axonic sites of cholinergic signaling; utilization of this chimeric co-culture preparation opens the door for more in depth structural and functional studies of these axo-axonic interactions.

Stimulation of ACh release from cholinergic terminals within the basal lateral amygdala ex vivo (i.e., in acute slice preparations) results in a sustained increase probability of evoked glutamate release at cortical-amygdala synapses (Jiang et al., 2016). Optogenetic stimulation of ACh release in the BLA also results in increased BLA principal neuron firing, both in slice and in vivo, and is critical for the formation of associative memories (Jiang et al., 2016). Whether and how ACh modulation of pre-synaptic inputs affects firing is not clear; it is clear that ACh is able to modulate glutamate transmission in a behaviorally relevant manner (Jiang et al., 2016).

Dysfunction of the basal forebrain cholinergic system is an early feature of Alzheimer’s disease and is likely to contribute to impairments in memory formation and recall. To date there is little information about the extent to which presynaptic α7* nAChR signaling occurs in the human brain but both α7* and non-α7* nAChRs have been identified as potential targets for pathological Aβ oligomers (Wang et al., 2000; Liu et al., 2012; Lombardo et al., 2016). Aβ acts as both an agonist and antagonist of α7* nAChRs, depending on Aβ concentration (Liu et al., 2001; Chin et al., 2007; Cecon et al., 2019). If sustained axonal signaling is a critical feature of glutamatergic transmission in vivo, ACh-independent disruption in the timing and/or location of α7* nAChR-dependent axonal signaling as a result of changing Aβ levels, might contribute to cognitive impairment.

Axonal α7* nicotinic acetylcholine receptor signaling contributes to glutamatergic axon maturation

Somato-dendritic α7* nAChRs have been implicated in development of glutamatergic synapses and of adult born glutamatergic neurons (Campbell et al., 2010; Lozada et al., 2012; Fernandes et al., 2014). We demonstrate in this report that axonal α7*nAChR function contributes to both the establishment of glutamatergic release sites and in modulating glutamate release. Taken together, these findings indicate that these receptors act in multiple compartments to regulate the development of some glutamatergic circuits. The contribution of α7* nAChR to neuron development raises several questions. First, what aspects of α7* nAChR functions are responsible for the effects on neuronal development? α7* nAChRs have the interesting feature of operating on two distinct levels: they are ligand-gated cation channels that contribute to depolarization on rapid time scales (milliseconds) and they activate metabotropic-like signaling by direct coupling to heterotrimeric G proteins (Nordman and Kabbani, 2014; Nordman et al., 2014; Kabbani and Nichols, 2018; King et al., 2018). Both mechanisms result in increased intracellular Ca²⁺ levels, and it is likely that calcium signaling is the major driver of α7* nAChR contributions to development. A second outstanding question is the identity of the developmentally relevant α7* nAChR agonist. In this study we demonstrate that adding the α7* nAChR antagonist, αBgTx, to the culture media mimics the effect of genetic deletion of α7. This points to some degree of on-going signaling in the absence of ACh. Whether this signaling is driven by choline, Lynx prototoxins (Miwa et al., 2019) or other agents remains to be determined.

The localized effects of axonal α7* nAChR on glutamate release underscore the importance of accurate targeting of these receptors to specific compartments. It is noteworthy that axonal/pre-synaptic targeting of α7*nAChRs is regulated by at least two signaling molecules – Nrg1 and Wnt – that play important roles in neurodevelopment (Farías et al., 2007; Hancock et al., 2008; Zhong et al., 2017). Variants in both CHRNA7 and NRG1 genes are associated with schizophrenia and related endophenotypes (Leonard et al., 2000; Munafò et al., 2006; Chen et al., 2009) and α7*nAChR levels are associated with NRG1 risk variants. The dual role of α7*nAChR in both glutamatergic axonal maturation and modulation of glutamatergic circuits, raises the possibility that the α7*nAChRs contribution to schizophrenia associated endophenotypes might reflect alteration in both functions.

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