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. Author manuscript; available in PMC: 2026 Jul 25.
Published in final edited form as: Biol Psychiatry. 2026 Feb 17;100(3):237–246. doi: 10.1016/j.biopsych.2026.02.006

Perineuronal Net and Inhibitory Synapse Remodeling on Striatal Fast-Spiking Interneurons by Chronic Alcohol Exposure

Michael S Patton 1, Samuel H Sheats 1, Andreas B Wulff 1, Paige N McKeon 1, Jonathan W VanRyzin 1, Mary H Patton 1, Morgan Heckman 1, Allison N Siclair 1, Phillip H Iffland II 1, Brian N Mathur 1
PMCID: PMC13397362  NIHMSID: NIHMS2194343  PMID: 41713796

Abstract

BACKGROUND:

Alcohol use disorder is characterized by persistent drinking in the face of negative consequences. Such inflexible drinking requires dorsolateral striatum fast-spiking interneurons, which comprise approximately 1% of all striatal neurons. How chronic ethanol exposure affects fast-spiking interneuron physiology is poorly understood.

METHODS:

We exposed adult male and female mice to chronic intermittent ethanol and examined synaptic transmission onto dorsolateral striatum fast-spiking interneurons.

RESULTS:

We discovered that chronic ethanol exposure induced a dramatic loss of GABAergic (gamma-amino-butyric acidergic), but not glutamatergic, synapses onto dorsolateral striatum fast-spiking interneuron somata and proximal dendrites where perineuronal nets (PNNs), a subdivision of the extracellular matrix, are enriched. We found that chronic ethanol exposure degraded these PNNs and that enzymatically degrading PNNs similarly reduced GABAergic transmission onto dorsolateral striatum fast-spiking interneurons. Modeling the effect of alcohol, we found that silencing extrinsic GABAergic projections to the dorsolateral striatum increased voluntary ethanol consumption.

CONCLUSIONS:

These data suggest that chronic alcohol exposure remodels PNNs and inhibitory synapses on fast-spiking interneurons to facilitate alcohol drinking.

Inflexible drinking is a prominent feature of alcohol use disorder that is not adequately addressed by current therapies (1). The dorsolateral striatum encodes inflexible actions that are thought to underlie inflexible drinking in alcohol use disorder (2–6). Ethanol exposure promotes activity in the dorsolateral striatum and biases behavior from goal-directed to inflexible, stimulus response–based strategies (7–12). This includes inflexible consumption of both natural rewards (11) and alcohol (12,13).

Parvalbumin-expressing fast-spiking interneurons (FSIs) are enriched in the dorsolateral striatum and provide feed-forward inhibition onto striatal projection neurons (14–17). Habitual responding for natural rewards requires FSIs (18), and FSIs are necessary for the expression of organized, aversion-resistant alcohol-drinking behavior (13). Thus, FSIs are positioned to shape dorsolateral striatal medium spiny neuron encoding of stereotyped behaviors (19–22). Exploring how alcohol may facilitate this FSI-mediated process, several studies have identified that acute alcohol targets FSIs (23–26). What is not known, however, is how chronic alcohol exposure, modeling the human experience in alcohol use disorder, affects FSI synaptic physiology and related behavior.

Dorsolateral striatum FSIs receive excitatory innervation primarily from the somatomotor cortices and receive inhibitory innervation primarily from the reticular thalamic nucleus (RTN) and the globus pallidus (GP) (27–31). Projections from the GP to the dorsolateral striatum are implicated in the transition from goal-directed to habitual behavior (32), and acute alcohol potentiates GABAergic (gamma-aminobutyric acidergic) synaptic transmission at GP and RTN synapses onto FSIs (23). This suggests that chronic alcohol exposure may result in lasting changes at these synapses to promote drinking. In the following work, we demonstrate that chronic ethanol vapor exposure reduced GABAergic, but not glutamatergic, synaptic transmission onto dorsolateral striatum FSIs from both the GP and the RTN. This reduction in GABAergic transmission was mediated by a decrease in the number of GABAergic synapses, primarily on somata and proximal dendrites of FSIs where perineuronal nets (PNNs) uniquely surround FSIs. Following this observation, we discovered that chronic ethanol exposure reduced the number of PNN-positive FSIs as well as striatal PNN protein expression, while enzymatic digestion of PNNs reduced the number of GABAergic synapses on FSIs. Silencing inhibitory synaptic transmission to the dorsolateral striatum, mirroring what is observed following chronic ethanol exposure, increased voluntary ethanol consumption. These data suggest that a remodeling of inhibitory control of FSIs in the dorsolateral striatum following chronic alcohol exposure contributes to drinking phenotypes.

METHODS AND MATERIALS

Animals

All experiments were performed in accordance with the National Institutes of Health guidelines and were approved by the Institutional Animal Care and Use Committee of the University of Maryland, Baltimore, and the National Institute on Alcohol Abuse and Alcoholism. Randomly sampled 2- to 4-month-old male and female wild-type C57BL/6J mice or mice expressing Cre-recombinase under the parvalbumin promotor (33) crossed with a tdTomato reporter mouse line (PV-tdT) were housed with littermates (2–5 per cage) under a normal 12-hour light/dark cycle (lights on at 9 AM, off at 9 PM) with ad libitum access to food and water.

Surgical Procedures

All stereotaxic injections were performed under isoflurane anesthesia (5% induction; 2%–3% maintenance), and viruses were injected at a rate of 30 nL/min with a 25G syringe (Hamilton Company). To isolate GABAergic projections from the GP or the RTN to the dorsolateral striatum, PV-tdT mice on a C57BL/6J background (33,34) were stereotaxically injected with a virus expressing ChR2 (channelrhodopsin-2) (AAV5-hSyn-ChR2-eYFP) in the GP (anteroposterior [AP] −0.4 mm, mediolateral [ML] ±2.0 mm, dorsoventral [DV] −3.7 mm; 200 nL/side) or the RTN (AP −0.58 mm, ML ±1.25 mm, DV −3.5 mm; 200 nL/side). Further information can be found in Supplemental Methods.

Chronic Intermittent Ethanol Exposure

Chronic intermittent ethanol (CIE) exposure was conducted following our previously established methods (23). Randomly sampled male and female mice were placed in plexiglass inhalation chambers (60 × 36 × 60 cm) (35) and exposed to ethanol vapor or air 16 hours/day, 4 days a week for a maximum of 5 weeks. After 4 days in the inhalation chambers, mice underwent a 72-hour forced abstinence period from ethanol. Vapor chamber ethanol concentrations were monitored daily, and airflow was adjusted to produce ethanol concentrations within 1.8% to 2.0% ethanol content measured by a digital alcohol breath tester (FFtopu). These conditions produce stable blood ethanol concentrations in C57BL/6J mice ranging from 150 to 200 mg/dL (36). To counteract the emergence of metabolic tolerance that increases after 4 weeks of ethanol exposure (37), mice were injected with the alcohol dehydrogenase inhibitor pyrazole (1 mmol/kg, intraperitoneally) 30 minutes prior to each session (weeks 4–5) as this stabilizes blood ethanol concentrations (23,36,38). Air control mice were also injected with pyrazole 30 minutes prior to each session (weeks 4–5).

Whole-Cell Patch-Clamp Electrophysiology

Mice were deeply anesthetized with isoflurane before brain removal. Coronal sections (250 μm) were collected in ice-cold modified artificial cerebrospinal fluid (aCSF) (194 mM sucrose, 30 mM NaCl, 4.5 mM KCl, 1 mM MgCl2, 26 mM NaHCO3, 1.2 mM NaH2PO4, and 10 mM D-glucose bubbled with 95% oxygen, 5% carbon dioxide). Brain sections were then transferred to regular aCSF (124 mM NaCl, 4.5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 26 mM NaHCO3, 1.2 mM NaH2PO4, and 10 mM D-glucose bubbled with 95% oxygen, 5% carbon dioxide) and incubated at 32 °C for 30 minutes, followed by room temperature until recording. Slices were hemisected, placed into a recording chamber, and perfused with aCSF (29–31 °C) during recording (aCSF: 124 mM NaCl, 4.5 mM KCl, 1 mM MgCl2, 26 mM NaHCO3, 1.2 mM NaH2PO4, 10 mMD-glucose, and 2 mM CaCl2 bubbled with 95% oxygen, 5% carbon dioxide). Dorsolateral striatum FSIs were fluorescently visualized using a Nikon Eclipse FN.1 microscope and Intenslight C-HGFIE (Nikon). Images were digitally rendered using a Hamamatsu ORCA-ER digital camera and the associated HCImage Live 4.8.0 software. FSIs were voltage clamped at −60 mV using a MultiClamp 700B Amplifier (Molecular Devices). Electrically evoked postsynaptic currents were generated using a concentric bipolar stimulating electrode (World Precision Instruments) located approximately 150 μm from the recording electrode. Optogenetically evoked currents were generated from an LED driver (LEDD1B T-Cube; Thorlabs) and a 470-nm LED delivering blue light (2–4 ms pulse duration) through an optical fiber located approximately 150 μm from the recording electrode. Excitatory postsynaptic currents (EPSCs) were pharmacologically isolated by adding picrotoxin (50 μM) to the recording aCSF. EPSCs were recorded using a borosilicate glass pipette (3–5 MΩ resistance) filled with cesium-based internal solution (120 mM CsMeSO3, 10 mM HEPES, 5 mM NaCl, 10 mM TEA-Cl, 1.1 mM EGTA, 0.3 mM Na-GTP, 5 mM QX-314, and 4 mM Mg-ATP). Evoked inhibitory postsynaptic currents (IPSCs) and spontaneous IPSCs (sIPSCs) were pharmacologically isolated by adding DL-AP5 (50 μM) and CNQX (5 μM) to the recording aCSF. IPSCs were recorded using a borosilicate glass pipette (3–5 MΩ resistance) filled with a chloride-based internal solution (150 mM CsCl, 10 mM HEPES, 2 mM MgCl2, 0.3 mM Na-GTP, 5 mM QX-314, 3 mM Mg-ATP, and 0.2 mM BAPTA). For optically evoked asynchronous currents, calcium was replaced with strontium (2 mM) in the recording aCSF. To photo uncage GABA, a 2-ms pulse of bluelight (470 nm) was delivered to an acute slice preparation bath perfused with 250 μM ruthenium-bipyridine-triphenylphosphine–caged GABA (RuBi-GABA) (#4709; Toc-ris) (39). For recordings of excitable membrane properties, borosilicate glass recording pipettes (3–7 MΩ resistance) were filled with a potassium gluconate–based solution (126 mM potassium gluconate, 4 mM KCl, 10 mM Hepes, 4 mM ATP-Mg, 0.3 mM GTP-Na, and 10 mM phosphocreatine). Signals were filtered at 2 kHz, digitized at 10 kHz, and acquired using Clampex 10.4.1.4 software (Molecular Devices). Data were analyzed using Clampfit software (Molecular Devices) and Mini Analysis Program (Synaptosoft).

Drinking in the Dark

The drinking in the dark (DID) protocol was adapted from Rhodes et al. (40), following our previously established methods (13). Briefly, mice were randomly assigned to sucrose or ethanol drinking groups. Mice were given 2 hours of access to the drinking bottle during the first 3 days, and on day 4, they were given 4 hours of access. After 5 days of drinking, mice underwent 2 days of forced abstinence, and this cycle was repeated for 2 consecutive weeks. Bottle weights were recorded to determine total consumption and normalized to body weight (g/kg).

Statistical Analysis

All statistical analyses were performed in GraphPad Prism 9.0. Data are represented as mean + SEM. Comparisons between 2 groups were analyzed using Student’s t tests while comparisons among 3 or more groups were analyzed with 2-way analysis of variance (ANOVA). Multiple comparisons were corrected with Sidak’s test or Dunnett’s test when comparing multiple time points to a single baseline. For the Sholl analysis, a mixed-effects model was performed (41). ns are specifically defined in the legend of each figure. For data from CIE mice, n is defined as an individual animal. Each individual animal value represents an average of 2 or 3 cell recordings per animal or an average fluorescence quantification of 5 to 10 cells per animal. For the chondroitinase ABC (ChABC) treatment, n was defined as an individual animal, with 2 or 3 cell recordings per animal. Preliminary analyses revealed no sex differences, so data were pooled across sexes. Individual male and female data points are shown graphically.

RESULTS

We performed whole-cell patch-clamp recordings from dorsolateral striatum FSIs to determine whether CIE modulated FSI excitable membrane properties or synaptic transmission onto FSIs (Figure 1A, B). Recording 4 to 7 days into abstinence, we found that CIE decreased FSI resting membrane potential (mean control = −66.28 mV, mean ethanol = −69.3 mV; t14 = 2.48, p < 05; Student’s t test) (Figure S1A). However, we observed no effect of CIE on input resistance (t14 = 1.41, p = .18; Student’s t test) (Figure S1B), action potential threshold (t14 = 1.28, p = .22; Student’s t test) (Figure S1C), or maximum firing rate in response to depolarizing current injection (t14 = 0.31, p = .76; Student’s t test) (Figure S1D).

Figure 1.

Figure 1.

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CIE vapor exposure specifically reduced inhibitory synaptic transmission onto dorsolateral striatum FSIs. (A) Schematic and timeline of CIE vapor exposure paradigm. (B) Schematic of whole-cell patch-clamp recording from an FSI. (C) CIE dramatically reduced inhibitory synaptic transmission onto FSIs (n = 6 mice per group, 3 F and 3 M). Inset: representative electrically evoked IPSCs from control (black) and CIE-treated (blue) mice (scale bars = 500 pA, 100 ms). (D) However, we observed no change in electrically evoked EPSCs onto FSIs (n = 6 mice per group, 3 F and 3 M). (E, F) sIPSC events onto FSIs (n = 10 mice per group, 5 F and 5 M). Inset: representative traces from control (black) and CIE-treated (blue) mice (scale bar = 100 pA, 2 s). CIE reduced the frequency of spontaneous inhibitory events (E, F) but did not impact event amplitude (G). (H) CIE did not impact FSI dendrite branch complexity (n = 10 cells, 5 mice per group, 2 F and 3 M). Representative dye-filled FSIs from control (left) and CIE-treated (right) mice (Sholl ring radius = 10-μm increments). All data are represented as mean + SEM. **p < .01. (Portions of this figure were created in BioRender.) eIPSC, evoked inhibitory postsynaptic current; EPSC, excitatory postsynaptic current; EtOH, ethanol; F, female; M, male; sIPSC, spontaneous IPSC.

Then we recorded electrically evoked synaptic transmission and found that CIE dramatically reduced inhibitory transmission onto dorsolateral striatum FSIs (F1,10 = 19.59, p = .001; 2-way repeated-measures ANOVA) (Figure 1C). However, we found no difference in the amplitude of electrically evoked excitatory synaptic currents across increasing stimulus intensities (F1,10 = 0.058, p = .82; 2-way repeated-measures ANOVA) (Figure 1D). Following these results, we recorded sIPSC events onto dorsolateral striatum FSIs to determine whether CIE-induced depression of these synapses was expressed pre- or postsynaptically. FSIs from CIE-exposed mice exhibited less frequent sIPSCs (increased interevent interval) compared with FSIs from control mice (t18 = 3.12, p < .001; Student’s t test) (Figure 1E, F), but there was no change in the amplitude of sIPSCs (t18 = 1.47, p = .16; Student’s t test) (Figure 1G). Given the significant change in inhibitory synaptic transmission, we assessed whether CIE impacted the cellular morphology of dorsolateral striatum FSIs. Sholl analysis revealed no statistically significant differences in dendritic complexity between control and CIE mice (F1,18 = 0.037, p = .85; mixed-effects model) (41) (Figure 1H). Considering that the data thus far suggest a suppression of presynaptic GABA release, next we examined the 2 major extrinsic sources of GABA to the dorsal striatum, the GP and the RTN, which synapse onto striatal FSIs (27,29). To investigate which distinct GABAergic projections were specifically impacted by CIE, we virally expressed ChR2 into the GP (Figure 2A) or the RTN (Figure 2F) and recorded optically evoked IPSCs onto dorsolateral striatum FSIs. GABAergic transmission from the GP was dramatically reduced by CIE (F1,70 = 42.45, p < .0001; 2-way repeated-measures ANOVA) (Figure 2B), but there was no change in the paired-pulse ratio (PPR) from the GP (t14 = 1.07, p = .30; Student’s t test) (Figure 2C). Because we found a decrease in the frequency of sIPSCs (Figure 1E, F) but saw no difference in PPR (Figure 2C), next we conducted optically evoked recordings in which we replaced calcium with strontium in the external aCSF recording solution to elicit asynchronous synaptic release events from the GP. CIE reduced the frequency of asynchronous release events from the GP (t10 = 4.14, p < .005; Student’s t test) (Figure 2D) but had no impact on event amplitude (t10 = 0.09, p = .93; Student’s t test) (Figure 2E). Similar to GP inputs, CIE reduced the amplitude of optically evoked IPSCs from the RTN onto dorsolateral striatum FSIs (F1,14 = 10.36, p < .01; 2-way repeated-measures ANOVA) (Figure 2G). Additionally, CIE did not impact PPR from the RTN (t14 = 0.58, p = .57; Student’s t test) (Figure 2H). CIE reduced the frequency of optically evoked strontium-enabled asynchronous inhibitory synaptic release events from the RTN (t10 = 6.20, p < .001; Student’s t test) (Figure 2I) but had no impact on event amplitude (t10 = 0.41, p = .69; Student’s t test) (Figure 2J).

Figure 2.

Figure 2.

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CIE reduced inhibitory synaptic transmission from the GP and the RTN, and silencing inhibitory transmission from these inputs increased EtOH consumption. (A) Schematic of injection of a virus expressing ChR2 into the GP. (B) CIE reduced optically evoked inhibitory currents from the GP onto FSIs but (C) had no impact on release probability (n = 8 mice per group, 4 F and 4 M). Inset: representative traces from control (black) and CIE-treated (blue) mice (scale bar = 500 pA, 100 ms). CIE reduced (D) the frequency but not (E) the amplitude of asynchronous inhibitory current events from the GP. Inset: representative traces from control (black) and CIE-treated (blue) mice (scale bar = 200 pA, 10 ms). (F) Schematic of injection of virus expressing ChR2 into the RTN. (G) CIE reduced oIPSCs from the RTN but (H) had no impact on release probability (n = 8 mice per group, 4 F and 4 M; scale bar = 500 pA, 100 ms). (I) CIE reduced the frequency of asynchronous inhibitory current events but (J) did not impact asynchronous current event amplitude (n = 6 mice per group, 3 F and 3 M; scale bar = 200 pA, 20 ms). (K) Schematic of injection of Cre-dependent virus expressing TeLC into the GP and RTN and retrograde AAV expressing Cre-recombinase in the dorsal striatum. (L) Experimental timeline of drinking in the dark paradigm with a schematic of a mouse drinking EtOH (20%) or sucrose (2%). (M) TeLC silencing of inhibitory transmission from the GP and RTN did not impact sucrose consumption. (N) However, TeLC silencing of inhibitory transmission increased voluntary EtOH consumption. All data are represented as mean + SEM. *p < .05, **p < .01, ***p < .001. (Portions of this figure were created in BioRender.) AAV, adeno-associated virus; aCSF, artificial cerebrospinal fluid; ChR2, channelrhodopsin-2; CIE, chronic intermittent ethanol; EtOH, ethanol; FSI, fast-spiking interneuron; IEI, interevent interval; IPSC, inhibitory postsynaptic current; osIPSC, optically evoked, strontium-enabled IPSC; TeLC, tetanus light chain toxin.

Next, we investigated whether virally silencing inhibitory projections to the dorsolateral striatum, mirroring what is observed following CIE, affects voluntary ethanol consumption in C57BL/6J mice. We microinjected a Cre-dependent tetanus light chain toxin (TeLC)–expressing AAV (adeno-associated virus) into the GP and RTN and a retrograde AAV expressing Cre-recombinase into the dorsolateral striatum (Figure 2K) to silence inhibitory synaptic transmission from the GP and RTN to the dorsolateral striatum (42). This reduced the frequency of spontaneous inhibitory transmission onto dorsolateral striatum FSIs (t9 = 2.4, p < .05; Student’s t test) but had no impact on the amplitude of these events (t9 = 0.5, p = .63; Student’s t test) (Figure S2A, B). Following a 4-week recovery period, mice underwent a 2-week DID paradigm (Figure 2L). Silencing inhibitory projections from the GP and RTN to the dorsolateral striatum did not affect sucrose drinking (Figure 2M) but increased voluntary ethanol consumption (F1,9 = 5.56, p < .05; 2-way repeated-measures ANOVA) (Figure 2N).

To examine the mechanism underlying CIE-induced suppression of GABAergic transmission onto FSIs, we virally expressed a GFP (green fluorescent protein)–tagged intrabody against the postsynaptic GABAergic synapse marker gephyrin in dorsolateral striatum FSIs (Figure 3A) and immunostained for the presynaptic GABAergic marker VGAT (vesicular GABA transporter). We quantified colocalized VGAT- and gephyrin-positive puncta to identify putative GABAergic synapses (Figure 3B). CIE mice exhibited fewer GABAergic synapses onto dorsolateral striatum FSIs compared with control mice (Figure 3B–E). Notably, the loss of synapses was more pronounced on the somata (t14 = 4.49, p < .001; Student’s t test) (Figure 3C) and proximal branches of FSIs (t14 = 6.58, p < .0001; Student’s t test) (Figure 3D) but was still present on distal processes (t14 = 2.16, p < .05; Student’s t test) (Figure 3E). To further investigate whether CIE resulted in postsynaptic silencing of inhibitory synapses or merely a loss of presynaptic elements that leave postsynaptic sites intact (and unsilenced), we photo uncaged RuBi-GABA in a slice preparation and recorded from FSIs using whole-cell patch-clamp electrophysiology in voltage clamp mode (Figure 3F). We found that photo-uncaged GABA IPSCs (uIPSCs) onto FSIs from control and CIE mice were not significantly different, suggesting that postsynaptic inhibitory synapses were not silenced (F1,14 = 0.19, p = .67) (Figure 3G, H).

Figure 3.

Figure 3.

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CIE delocalizes GABAergic synaptic contacts onto FSIs. (A) Schematic of injection of virus expressing a Cre-dependent GFP-tagged gephyrin intrabody into PV-Cre mice. (B) Representative image of immunostaining for gephyrin (green) and VGAT puncta (purple). VGAT-positive puncta were masked onto the gephyrin signal, and colocalized signals (white dots) were quantified (scale bar = 10 μm) (right). Representative fluorescence image of a proximal neuronal branch from control and CIE-treated mice depicting colocalized presynaptic VGAT (purple) puncta onto gephyrin (green)-positive postsynaptic elements (scale bar = 1 μm). CIE reduced the density of colocalized VGAT puncta onto (C) somata, (D) proximal dendrites, (E) and distal dendrites (n = 8 mice per group, 4 F and 4 M). (F) Schematic of a whole-cell patch-clamp recording from an FSI during photo uncaging of RuBi-GABA. (G) Representative traces of uIPSCs from FSIs of control (left) and CIE-treated (right) mice at increasing stimulus intensities (scale bar = 200 pA, 100 ms). (H) CIE did not impact uIPSCs compared with control mice (n = 8 mice per group, 4 F and 4 M). All data are represented as mean + SEM. *p < .05, **p < .01, ***p < .001, ****p < .0001. (Portions of this figure were created in BioRender.) CIE, chronic intermittent ethanol; GABA, gamma-aminobutyric acid; GFP, green fluorescent protein; EtOH, ethanol; F, female; FSI, fast-spiking interneuron; IPSC, inhibitory postsynaptic current; M, male; PV, parvalbumin; uIPSC, photo-uncaged GABA IPSC; VGAT, vesicular GABA transporter.

As the reduction in putative GABAergic synapses was more prominent around the somata and proximal processes of FSIs, we hypothesized that a perisomatic mechanism accounts for cellular subregion–specific synaptic disruption. Because PNNs condense around the somata and proximal branches of FSIs and are known to regulate synaptic transmission onto FSIs (43–45), we investigated whether CIE modulated PNNs. First, we confirmed that PNNs are present in the dorsal striatum of adult C57BL/6J mice using Wisteria floribunda agglutinin (WFA) staining (46). We identified WFA-positive PNNs surrounding FSIs (Figure 4A), consistent with previous literature (47). However, we noticed that some FSIs were enriched with PNNs (PNN+) and that a subpopulation was PNN poor (PNN−) based on a fluorescence signal intensity threshold for WFA staining (Figure 4A–C). Next, we examined the effect of CIE on PNN presence surrounding FSIs. CIE reduced the number of PNN+ FSIs (t14 = 12.83, p < .0001; Student’s t test) (Figure 4D) but had no impact on the total number of FSIs (t14 = 1.01, p = .33; Student’s t test) (Figure 4E). The reduction in PNN+ FSIs was not evident after 2 weeks of CIE (t10 = 0.85, p = .41; Student’s t test) (Figure S2C), indicating that loss of PNN expression was specific to long-term ethanol exposure. We followed these results by Western blotting for 2 PNN proteins: the proteoglycan aggrecan and the link protein HAPLN1 (45,48). Consistent with our population analysis, aggrecan (t14 = 3.87, p < .01; Student’s t test) (Figure 4F) and HAPLN1 (t14 = 2.91, p, .05; Student’s t test) (Figure 4G) expression was reduced in dorsolateral striatum punches from CIE compared with control mice.

Figure 4.

Figure 4.

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CIE degrades PNNs in the dorsolateral striatum. (A) Immunostaining for PV (red) and WFA staining for PNNs (green) depicting PNN-enriched (PNN+) and PNN-poor (PNN−) FSIs (scale bar = 100 μm). (B, C) Fluorescence intensity profile of PNN+ and PNN− FSIs. (D) CIE reduced the percent-age of PNN+ FSIs but (E) did not reduce the number of FSIs (n = 10 mice per group, 5 F and 5 M). CIE reduced the expression of PNN proteins (F) aggrecan and (G) HAPLN1 in the dorsolateral striatum (n = 8 mice per group, 4 F and 4 M). (H) Representative confocal fluorescence images of PV (black) and WFA signal (green) from control (top) and CIE-treated (bottom) mice (scale bar = 15 μm). (I) Top: 3-dimensional volume reconstruction and filament model of WFA fluorescence signal from control (top) and CIE-treated (bottom) mice. (J) CIE reduced the volume of WFA signal (top) surrounding FSI somata and reduced the length of WFA signal (bottom) surrounding FSI proximal branches (n = 10 mice per group, 5 F and 5 M). All data are represented as mean + SEM. *p < .05, **p, .01. ****p < .0001. CIE, chronic intermittent ethanol; EtOH, ethanol; F, female; FSI, fast-spiking interneuron; M, male; PNN, perineuronal net; PV, parvalbumin; WFA, Wisteria floribunda agglutinin.

To determine whether specific cellular subcompartments were impacted by CIE, we collected confocal images of WFA PNN+ signal from control and CIE mice (Figure 4H) and generated 3-dimensional surface expression models surrounding FSI somata and branches (Figure 4I). Analysis of these models revealed a significant reduction in WFA signal surrounding FSI somata (t18 = 6.24, p < .0001; Student’s t test) (Figure 4J) and a reduction in the length of WFA signal extending along FSI branches (t18 = 8.23, p < .0001; Student’s t test) (Figure 4J). However, there was no difference in the branch length of FSIs between groups (t18 = 0.03, p = .98; Student’s t test) (Figure S2D).

During brain development, PNNs regulate the formation of synaptic connections (49). Therefore, we investigated whether PNNs regulated GABAergic transmission onto FSIs in the dorsolateral striatum. First, we recorded electrically evoked GABAergic transmission from FSIs and later identified them as PNN+ or PNN− by dye filling the recorded cell followed by subsequent WFA staining (Figure 5A). PNN+ FSIs exhibited larger electrically evoked IPSC amplitudes compared with PNN− FSIs (F1,15 = 8.1, p < .05; 2-way repeated-measures ANOVA) (Figure 5B). This difference was not due to a change in presynaptic release probability, as no mean difference in the PPR was observed (t15 = 0.74, p = .47; Student’s t test) (Figure 5C). Furthermore, PNN+ FSIs exhibited a greater frequency of sIPSC events compared with PNN− FSIs (t13 = 2.53, p < .05; Student’s t test) (Figure 5D, E), but there was no change in sIPSC amplitude (t13 = 0.56, p = .58; Student’s t test) (Figure 5F).

Figure 5.

Figure 5.

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PNN enzymatic digestion suppresses GABAergic synaptic transmission onto FSIs. (A) Schematic of a whole-cell patch-clamp recording from a PNN+ or PNN− FSI identified by Wisteria floribunda agglutinin staining after recordings. (B) PNN+ FSIs exhibited larger eIPSC amplitudes compared with PNN− FSIs (n = 11 PNN+ cells from 6 F and 5 M; n = 6 PNN− cells from 3 F and 3 M). Inset: representative traces from PNN+ (pink) and PNN− (black) FSIs (scale bar = 1 nA, 100 ms). (C) The presence of PNNs did not affect the probability of GABA release onto FSIs. (D–F) sIPSCs recorded from FSIs (n = 11 PNN+ cells from 6 F and 5 M; 6 PNN− cells from 3 F and 3 M). Inset: representative traces from PNN+ (pink) and PNN− (black) FSIs (scale bar = 100 pA, 1 s). (D, E) PNN+ FSIs exhibited a greater frequency of sIPSC events but (F) no change in sIPSC amplitude compared with PNN− FSIs. (G) Schematic of unilateral microinjection of ChABC into the dorsolateral striatum to enzymatically degrade PNNs. (H) Degrading PNNs with ChABC reduced eIPSC event amplitude onto FSIs but (I) did not impact the probability of GABA release (n = 10 mice, 5 F and 5 M; scale bar = 1 nA, 100 ms). (J, K) Degrading PNNs with ChABC reduced the frequency of sIPSC events onto FSIs but (L) did not impact event amplitude (n = 8 mice, 4 F and 4 M; scale bar = 100 pA, 1 s). (M) Schematic of viral injection of Cre-dependent gephyrin intrabody into PV-Cre mice followed by ChABC treatment. (N) Degrading PNNs with ChABC did not reduce inhibitory synapses onto somata but (O) did reduce inhibitory synapse number onto proximal dendrites (n = 13 mice, 7 F and 6 M). (P) Schematic of a whole-cell patch-clamp recording during photo uncaging of RuBi-GABA onto FSIs treated with ChABC. (Q) Representative traces of uIPSCs in control (left) and ChABC-treated (right) FSIs at increasing stimulus intensities (scale bar = 200 pA, 100 ms). (R) There was no change in uIPSC event amplitude between control (black) and ChABC-treated (pink) FSIs (n = 8 mice per group, 4 F and 4 M). All data are represented as mean ± SEM. *p < .05, **p < .01. (Portions of this figure were created in BioRender.) ChABC, chondroitinase ABC; eIPSC, evoked inhibitory postsynaptic current; F, female; FSI, fast-spiking interneuron; GABA, gamma-aminobutyric acid; M, male; PNN, perineuronal net; sIPSC, spontaneous IPSC; uIPSC, photo-uncaged GABA IPSC.

Next, we examined whether PNN degradation decreased GABAergic transmission to FSIs. To do this, we enzymatically degraded PNNs with ChABC, which reduced the number of PNN+ FSIs at 3 to 4 days following stereotaxic microinjection into the dorsolateral striatum (F2,34 = 105, p < .0001; Sidak’s multiple comparison test) (Figure S3A, B) compared with the vehicle-injected control side. This effect was still evident at 14 days postinjection (Sidak’s multiple comparison test) (Figure S3B). Notably, degrading PNNs had no impact on the number of FSIs (F2,34 = 1.9, p = .15) (Figure S3C). Next, we recorded from dorsolateral striatum FSIs and compared GABAergic transmission from control and ChABC-treated hemispheres (Figure 5G). Enzymatically degrading PNNs reduced electrically evoked IPSCs (F1,18 = 14.89, p < .01; 2-way repeated-measures ANOVA) (Figure 5H). This reduction in GABAergic transmission was not associated with a change in PPR (t18 = 0.27, p = .79; Student’s t test) (Figure 5I) but reduced the frequency of sIPSC events (t14 = 3.0, p < .01; Student’s t test) (Figure 5J, K) without impacting sIPSC event amplitude (t14 = 1.0, p = .33; Student’s t test) (Figure 5L).

Next, we determined whether ChABC treatment delocalized GABAergic synaptic contacts onto FSIs, as we observed following CIE. Again, we virally expressed the GFP-labeled intrabody against gephyrin in dorsolateral striatum FSIs (Figure 5M). After 4 weeks of viral expression, mice received a unilateral microinjection of ChABC or 5% BSA vehicle control into the dorsolateral striatum. ChABC treatment reduced colocalized GABAergic synapses onto proximal dendrites (t12 = 2.66, p < .05; paired t test) (Figure 5O) but not somata (t12 = 1.57, p = .14; paired t test) (Figure 5N) of dorsolateral striatum FSIs compared with control mice. However, uIPSC amplitudes onto FSIs from control and ChABC-treated mice were not significantly different, suggesting that postsynaptic inhibitory synapses remain unsilenced (F1,7 = 0.016, p = .9) (Figure 5Q, R).

DISCUSSION

CIE produced only subtle effects on FSI intrinsic excitability but dramatically reduced GABAergic synaptic transmission onto dorsolateral striatum FSIs, including inputs from the GP and RTN. Modeling this effect by silencing GP and RTN inputs to the dorsolateral striatum increased alcohol drinking. Exploring a possible mechanism, we found that CIE degraded FSI PNNs and that degrading FSI PNNs enzymatically also delocalized FSI GABA synapses and reduced synaptic transmission onto FSIs. These data are consistent with the notion that chronic alcohol exposure suppresses inhibitory control of dorsolateral striatum FSIs to promote alcohol-drinking behavior through a PNN degradation-dependent process.

The dorsolateral striatum encodes habitual actions underlying compulsive behavior seen in substance use disorder (3,4). Dorsal striatum FSIs play a critical role in the expression of habitual behavior (18), including compulsive-like ethanol consumption (13). Here, we demonstrate that chronic ethanol exposure reduces GABAergic, but not glutamatergic, input onto dorsolateral striatum FSIs. In response to acute ethanol, FSI intrinsic excitability is increased (26), and GABA synaptic transmission onto FSIs is increased (23), but GABA release from FSIs onto postsynaptic spiny projection neurons is decreased (25). In response to CIE, we observed here that FSI membrane potential is slightly hyperpolarized, which may be a compensatory response to the observed reduction in GABAergic transmission. Given the critical role of striatal FSIs in neuronal ensemble formation (50), understanding the complex interactions between FSI acute and chronic alcohol exposure effects on the development of striatal ensembles, including those recruited for drug acquisition (20,22,51), is positioned to provide deeper insight into the development of learned, inflexible behaviors in alcohol use disorder.

The effect of CIE on GABAergic transmission was evident in 2 major GABAergic projections to the dorsolateral striatum, the GP and the RTN, suggesting a cell-wide modification of the FSI GABAergic input. While the effect of alcohol on the FSI-FSI synapse has not yet been investigated (52), GABAergic transmission from both GP and RTN inputs is increased in response to acute ethanol exposure (23). How such multiple acute alcohol exposures then give rise to the GABA synapse delocalization that suppresses inhibitory control of FSIs is a major question arising from the present findings. Given the evidence for PNNs in this process, multiple cellular mechanisms could underlie this phenomenon (53), including synaptic pruning (54), possibly mediated by microglia (55–58). Another possible mechanism is ethanol’s modulation of PNN-degrading enzymes released by neurons and glia, including the metalloproteinase-9 (MMP-9) enzyme (45). In humans, serum MMP-9 levels increase during alcohol intoxication (59) and in individuals with a history of alcohol abuse (60). Similarly, in rats, MMP-9 expression is elevated in the nucleus accumbens of alcohol-dependent rats (61). Notably, intracerebroventricular administration of a broad-spectrum MMP inhibitor reduces the escalation of ethanol self-administration in rats (62).

PNNs condense around the soma and proximal processes of FSIs to stabilize synaptic contacts and regulate synaptic transmission (45,63–65). We demonstrated that chronic ethanol exposure decreases the number of FSIs containing PNNs and decreases PNN protein expression in the dorsolateral striatum. This result is consistent with the ethanol-induced PNN loss reported in the retrosplenial cortex (66) and hippocampus (67). However, in the insular (68) and prefrontal cortex (69), repeated ethanol exposure increases PNN expression. Regional differences in PNN expression are also observed following cocaine exposure (70), suggesting that local microenvironments differentially respond to drugs of abuse, resulting in increased or decreased PNN expression. Determining the mediators of these regional effects may reveal important biology underlying neurobiological responses to drug exposure and resulting maladaptive behavioral changes.

Important limitations of this study include that TeLC silencing of GP and RTN projections may impact not only transmission onto FSIs but also other striatal cell populations, including medium spiny neurons (71,72). Although WFA is the most widely used stain for PNNs, PNNs may be differentially detected with aggrecan antibodies (73), which may reveal additional effects. While ChABC degrades PNNs, it also likely disrupts the diffuse extracellular matrix (74) and may not precisely model the changes occurring in response to CIE. Thus, establishing whether PNN remodeling itself directly contributes to increases in inflexible drinking requires further investigation.

Despite these limitations, the study supports the notion that drug-induced PNN modification reopens a critical period of synaptic plasticity, enabling the formation of drug memories and drug-associated behaviors (75,76). More specifically, chronic ethanol exposure may reopen a critical period for dorsolateral striatal–mediated learning that may contribute to alcohol consumption. The discovery that chronic alcohol exposure alters inhibitory control of FSIs, which are required for aversion-resistant alcohol consumption, suggests that future therapeutic strategies aimed at FSI PNN remodeling could present a promising approach for treating alcohol use disorder.

Supplementary Material

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Supplementary material cited in this article is available online at https://doi.org/10.1016/j.biopsych.2026.02.006.

ACKNOWLEDGMENTS AND DISCLOSURES

This work was supported by the National Institute on Alcohol Abuse and Alcoholism (Grant Nos. R01AA028070 and R01AA024845 [to BNM] and F31AA029264 [to MSP]).

MSP and BNM conceived experiments and wrote the article. MSP, PNM, JWV, MHP, and MH performed experiments.

The authors report no biomedical financial interests or potential conflicts of interest.

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Supplementary Materials

1
NIHMS2194343-supplement-1.pdf (2.7MB, pdf)
2
NIHMS2194343-supplement-2.xlsx (22KB, xlsx)

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