Chronic Neuronal Inactivity Utilizes the mTOR-TFEB Pathway to Drive Transcription-Dependent Autophagy for Homeostatic Up-Scaling

Yang Wang; Jingran Lin; Jiarui Li; Lu Yan; Wenwen Li; Xingzhi He; Huan Ma

doi:10.1523/JNEUROSCI.0146-23.2023

Abstract

Activity-dependent changes in protein expression are critical for neuronal plasticity, a fundamental process for the processing and storage of information in the brain. Among the various forms of plasticity, homeostatic synaptic up-scaling is unique in that it is induced primarily by neuronal inactivity. However, precisely how the turnover of synaptic proteins occurs in this homeostatic process remains unclear. Here, we report that chronically inhibiting neuronal activity in primary cortical neurons prepared from embryonic day (E)18 Sprague Dawley rats (both sexes) induces autophagy, thereby regulating key synaptic proteins for up-scaling. Mechanistically, chronic neuronal inactivity causes dephosphorylation of ERK and mTOR, which induces transcription factor EB (TFEB)-mediated cytonuclear signaling and drives transcription-dependent autophagy to regulate αCaMKII and PSD95 during synaptic up-scaling. Together, these findings suggest that mTOR-dependent autophagy, which is often triggered by metabolic stressors such as starvation, is recruited and sustained during neuronal inactivity to maintain synaptic homeostasis, a process that ensures proper brain function and if impaired can cause neuropsychiatric disorders such as autism.

SIGNIFICANCE STATEMENT In the mammalian brain, protein turnover is tightly controlled by neuronal activation to ensure key neuronal functions during long-lasting synaptic plasticity. However, a long-standing question is how this process occurs during synaptic up-scaling, a process that requires protein turnover but is induced by neuronal inactivation. Here, we report that mTOR-dependent signaling, which is often triggered by metabolic stressors such as starvation, is “hijacked” by chronic neuronal inactivation, which then serves as a nucleation point for transcription factor EB (TFEB) cytonuclear signaling that drives transcription-dependent autophagy for up-scaling. These results provide the first evidence of a physiological role of mTOR-dependent autophagy in enduing neuronal plasticity, thereby connecting major themes in cell biology and neuroscience via a servo loop that mediates autoregulation in the brain.

Introduction

With its vast number of synapses, one of the brain's major challenges is optimizing cellular resources for synaptic remodeling and plasticity to process and store information. Recruited by chronic changes in neuronal activity, homeostatic synaptic plasticity (i.e., synaptic scaling) regulates synaptic strength via a negative feedback system and is believed to be essential for ensuring that various brain activities operate within a specific range (Abbott and Nelson, 2000; Davis, 2006; Marder and Goaillard, 2006; Turrigiano, 2008). In recent decades, several studies have yielded valuable insights into synaptic scaling in response to chronic changes in activity (Burrone and Murthy, 2003; Turrigiano and Nelson, 2004; Pozo and Goda, 2010), for example during sleep (Diering et al., 2016; Hengen et al., 2016), sensory deprivation (Desai et al., 2002; Goel and Lee, 2007), and in the context of various neuropsychiatric conditions such as autism spectrum disorder (ASD; Ramocki and Zoghbi, 2008; Toro et al., 2010; Mullins et al., 2016). The reorganization of cellular resources for synaptic remodeling requires de novo protein synthesis driven by activity-dependent gene transcription (Majdan and Shatz, 2006; Tropea et al., 2006; Ibata et al., 2008; Goold and Nicoll, 2010; Meadows et al., 2015; Steinmetz et al., 2016; Schaukowitch et al., 2017; Garay et al., 2020), translation (Aoto et al., 2008; Penney et al., 2012; Schanzenbächer et al., 2016), and alternative splicing (Li et al., 2020), thereby regulating the levels of synaptic proteins. On the other hand, activity-dependent protein degradation plays an essential role in regulating protein levels for synaptic scaling (Ehlers, 2003; Djakovic et al., 2009; Jakawich et al., 2010; Shin et al., 2012; Dorrbaum et al., 2020). However, several questions still remain with respect to how this process occurs.

In neurons, the ubiquitin-proteasome system (UPS) plays a central role in mediating protein turnover (Ehlers, 2003; Bingol and Schuman, 2004; Lee et al., 2008). Triggered by chronically increased neuronal activity, ubiquitin conjugates accumulate in the postsynaptic density, leading to the proteasomal degradation of synaptic proteins and homeostatic down-scaling (Ehlers, 2003; Djakovic et al., 2009; Jakawich et al., 2010; Shin et al., 2012; Dorrbaum et al., 2020). Another major mechanism that target synaptic components (Rowland et al., 2006; Shehata et al., 2012; Tang et al., 2014; Nikoletopoulou et al., 2017) for protein degradation is macroautophagy (hereafter referred to as autophagy), which has drawn considerable attention because of its apparent role in multiple dimensions of neuronal function (Cai et al., 2012; Hernandez et al., 2012; Vijayan and Verstreken, 2017; Griffey and Yamamoto, 2022), including long-term potentiation (LTP; Glatigny et al., 2019; Pandey et al., 2020), long-term depression (LTD; Shehata et al., 2012, 2018; Shen et al., 2020; Compans et al., 2021; Pan et al., 2021; Kallergi et al., 2022), learning and memory (Nikoletopoulou et al., 2017; Hylin et al., 2018; Glatigny et al., 2019; Pandey et al., 2020; Shen et al., 2020), and various neurologic diseases (Yue et al., 2009; Rubinsztein et al., 2011; Nixon, 2013). However, whether and how autophagy plays a role in synaptic scaling, a fundamental process believed to counterbalance LTP and LTD, is currently unknown.

Here, we report that autophagy regulated by mTOR (mechanistic target of rapamycin) is recruited for synaptic homeostasis in response to chronic suppression of neuronal activity. In addition, we show that this autophagy pathway is also triggered by chronic inhibition of UPS; moreover, both neuronal inactivity and chronic UPS inhibition lead to mTOR dephosphorylation and drive an increase in surface AMPA receptors (AMPARs). By targeting proteins such as αCaMKII and PSD95, inactivity-driven autophagy plays an important role in maintaining neuronal homeostasis. During neuronal inactivity, dephosphorylated mTOR drives the cytonuclear translocation of transcription factor EB (TFEB), which drives the expression of autophagy genes leading to long-lasting synaptic up-scaling. Together, our results suggest that mTOR-dependent autophagy is important for linking neuronal inactivity to protein turnover to maintain neuronal homeostasis.

Materials and Methods

Animals

Sprague Dawley rats (both sexes) were purchased from Shanghai SLAC Laboratory Animal Co, Ltd. C57BL/6 male mice were purchased from Zhejiang Academy of Medical Sciences. GFP-LC3 transgenic mice (both sexes, BRC No. RBRC00806) were kindly provided by Prof. Zhen-Ge Luo (ShanghaiTech University). Animals were housed at 22°C under a 12/12 h light/dark cycle with free access to food and water. All animal experiments were approved by the Animal Care and Use Committee at Zhejiang University and were conducted in accordance with institutional guidelines regarding the care and use of laboratory animals.

Plasmid construction

The original pEGFP-LC3 plasmid was kindly provided by Prof. Wei Liu (Zhejiang University School of Medicine). The truncated rat αCaMKII promoter (0.4 kb) was synthesized by Generay Biotech. To selectively transfect excitatory cortical neurons, the truncated αCaMKII promoter was introduced into the pCDH construct (Wheeler et al., 2012). mCherry and eGFP-LC3 were cloned into pCDH-tCaMKIIP to generate the construct expressing mCherry-eGFP-LC3. The H1 promoter was amplified from pLVTHM and then inserted into pCDH-tCaMKIIP-mCherry to knock down expression of the genes of interest. All constructs generated in this project were confirmed using Sanger sequencing.

The short-hairpin RNA (shRNA) sequences are listed below:

ATG7 shRNA #1: 5′-GCAGTGATGACCGCATGAATG-3′

ATG7 shRNA #2: 5′-GCTGGTCTCCTTGCTCAAACA-3′

ATG7 shRNA #3: 5′-GGGTGTAATGTGGCTAGAACA-3′

TFEB shRNA #1: 5′-CTGCTACACATCAGCTCCAATC-3′

CRTC1 shRNA #1: 5′-CGAACAATCCGCGGAAATTTA-3′

Scramble shRNA: 5′-TCGCTTGGGCGAGAGTAAG-3′

Primary neuron culture and cell treatment

Primary cortical neurons were cultured from embryonic day (E)18 Sprague Dawley rats as previously described (Pan et al., 2021). In brief, after the mother was anesthetized, the embryos were quickly removed and placed in a 100-mm culture dish containing DMEM/high glucose (HyClone, SH30243.01) with penicillin-streptomycin (HyClone, SH40003.01) on ice. The cortex was then isolated and washed with ice-cold DMEM containing penicillin-streptomycin. The tissues were then transferred to a clean 1.5-ml tube and digested in 0.25% trypsin-EDTA (Invitrogen, 25200056) at 37°C for 12 min, with gentle shaking every 3 min. After digestion, prewarmed DMEM containing 10% fetal bovine serum (Vistech, SE200-ES) was added to stop the reaction. After washing three times, the tissue was carefully dissociated by trituration through Pasteur pipettes of decreasing tip diameter. The debris was removed by centrifugation, and the cell suspension was filtered through a 40-μm nylon strainer. For immunocytochemistry, electrophysiology, and electron microscopy experiments, approximately 0.5 × 10⁵ neurons were plated on poly-D-lysine-coated coverslips (Sigma-Aldrich, P0899); for Western blot analysis and real-time PCR experiments, 1.0 × 10⁵ neurons were plated in each well of a 12-well plate. All neurons were cultured at 37°C in 5% CO₂ in Neurobasal medium (Invitrogen, 21103049) supplemented with GlutaMAX (Invitrogen, 35050061), B27 (Invitrogen, 17504044), and penicillin-streptomycin. Every 6 d, half of the culture medium was replaced with fresh medium. Unless stated otherwise, all neurons were used 14–18 d after plating.

To induce synaptic up-scaling, 2 μm tetrodotoxin (TTX; Alomone, T-550) was added to cultured cortical neurons for 24 h. For drugs that were dissolved in dimethylsulfoxide (DMSO), control samples contained 0.1% DMSO. Unless stated otherwise, all drugs were applied for 24 h.

Other chemicals were as follows: actinomycin D (ActD; Sigma, A1410), anisomycin (Anis; APExBio, B6674), bicuculline (Bic; MedChemExpress, HY-N0219), bortezomib (BTZ; MedChemExpress, HY-10 227), chloroquine (CQ; Sigma, C6628), D-AP5 (Abcam, ab120003), MG132 (Selleck, S2619), MHY1485 (MedChemExpress, HY-B0795), all-trans retinoic acid (RA; MedChemExpress, HY-14649), SBI-0206965 (SBI; MedChemExpress, HY-16966), STO-609 (MedChemExpress, HY-19805), and wortmannin (Wort; MedChemExpress, HY-10197).

Calcium phosphate transfection

The Ca²⁺-phosphate transfection method was used to transfect cortical neurons as described previously (Ma et al., 2014).

Lentivirus production and infection

To obtain lentivirus, the pLVTHM shRNA construct was transfected into HEK-293T cells (ATCC, CRL-3216) with the envelope plasmid pMD2.G and the packaging plasmid psPAX2. After 12 h, the medium was replaced with fresh medium, and the supernatant was collected three times at 24 h intervals. Cell debris was removed using a 0.45-μm filter, and virus particles were concentrated by centrifugation in a Beckman SW28 rotor at 70,000 × g at 4°C; the particles were then resuspended in PBS, aliquoted, and stored at –80°C. Lentivirus particles (1 μl of viral stock diluted in 20 μl of PBS per coverslip) were added to the cortical neurons 7–9 d after plating; 24 h later, the cultures were fed with 1 ml of fresh Neurobasal medium and used for experiments 7 d later.

Immunocytochemistry

Neurons were fixed in ice-cold 4% (w/v) paraformaldehyde (PFA) in 4% (w/v) sucrose containing 20 mm EGTA for >10 min, and then permeabilized in 0.1% Triton X-100. The cells were then blocked in 10% normal donkey serum (Jackson ImmunoResearch, 017-000-121) for 20 min at room temperature (RT), washed with PBS, and incubated with primary antibodies (e.g., p62, αCaMKII or PSD95) for 16–24 h at 4°C. After washing twice in PBS, the cells were incubated with secondary antibodies (1:1000) for 40–60 min at RT. Finally, the coverslips were washed with PBS for 30 min and mounted with Fluoromount-G+DAPI (Yeasen Biotechnology, 36308ES20).

For surface staining of the GluA1 and GluA2 AMPAR subunits, cortical neurons were cultured with a primary antibody against the N terminus of GluA1 (mouse monoclonal, Millipore, MAB2263, RRID: AB_11212678, 1:500) or GluA2 (mouse monoclonal, Millipore, MAB397, RRID: AB_2113875, 10 μg/ml) for 20 min at 37°C. The neurons were then immediately fixed in ice-cold 4% PFA/4% sucrose containing 20 mm EGTA, and surface-bound antibody-labeled GluA1 or GluA2 was saturated by incubation with secondary antibody for 40–60 min at RT. The neurons were then washed with PBS for 30 min at RT, permeabilized with methanol at −20°C for 1 min, washed with PBS for 5 min, and incubated with MAP2 (Sigma, M3696, RRID: AB_1840999, 1:1000) primary antibody for 16 h at 4°C. The cells were then washed with PBS, incubated with secondary antibody for 40–60 min at RT, washed in PBS for 30 min, and mounted with Fluoromount-G+DAPI.

Other primary antibodies (1:1000) were as follows: c-fos (Synaptic System, 226004, RRID: AB_2619946), CRTC1 (Abcam, ab92477, RRID: AB_10563847), MAP2 (Sigma-Aldrich, M3696, RRID: AB_1840999), MAP2 (Sigma-Aldrich, M4403, RRID: AB_477193), p62/SQSTM1 (MBL, PM045, RRID: AB_1279301), pCaMKII (Thr286; Cell Signaling Technology, 12716, RRID: AB_2713889), pCREB (Ser133; Cell Signaling Technology, 9198s, RRID: AB_2561044), LC3B (Abcam, ab48394, RRID: AB_881433), αCaMKII (Abcam, ab92332, RRID: AB_2049246), PSD95 (Cell Signaling Technology, 3450s, RRID: AB_2292883), mTOR (Cell Signaling Technology, 2983s, RRID: AB_2105622), p70 S6 Kinase (Cell Signaling Technology, 9202s, RRID: AB_331676), TFEB (Proteintech, 13 372–1-AP, RRID: AB_2199611), and H3 (Abcam, Ab176842). Secondary antibodies (1:1000) were Alexa Fluor 488 donkey anti-mouse antibody (Thermo Fisher Scientific, 1796361), Alexa Fluor 488 donkey anti-rabbit antibody (Thermo Fisher Scientific, 1796375), Alexa Fluor 555 donkey anti-mouse antibody (Thermo Fisher Scientific, 1774719), Alexa Fluor 555 donkey anti-rabbit antibody (Thermo Fisher Scientific, 1806149), Alexa Fluor 647 donkey anti-mouse antibody (Thermo Fisher Scientific, 1757130), and Alexa Fluor 647 donkey anti-rabbit antibody (Thermo Fisher Scientific, 1826679).

SDS-PAGE and Western blot analysis

For whole-cell protein extraction, cultured cortical neurons were lysed in RIPA buffer (Yeasen, 20101ES60) containing phosphatase inhibitor cocktail (Roche, 04906837001) and protease inhibitor cocktail (Roche, 04693132001). The lysates were then sonicated and centrifuged at 14,000 × g at 4°C for 15 min to collect the proteins in the supernatant. For subcellular fraction samples, cytoplasmic and nuclear fractions were isolated using the CelLytic NuCLEAR Extraction kit (Sigma, NXTRACT-1KT) in accordance with the manufacturer's instructions. Protein concentration was measured using the BCA Protein Quantification kit (Yeasen, 20201ES76), and equal quantities of total protein in each sample were resolved by SDS-PAGE and transferred to PVDF (Millipore) membranes. The membranes were blocked with 5% (w/v) skim milk (Becton Dickinson, 232100) at 4°C for 8 h and then probed with primary antibodies at 4°C for 16 h, followed by second antibodies. The following primary antibodies (1:1000) were used: β-Actin (Cell Signaling Technology, 4970s, RRID: AB_2223172), β-Tubulin (Yeasen, 30102ES40), GAPDH (Yeasen, 30102ES40), MAP2 (Sigma-Aldrich, M3696, RRID: AB_1840999), MAP2 (Sigma-Aldrich, M4403, RRID: AB_477193), anti-CaMKII α antibody (Invitrogen, MA1-048, RRID: AB_325403), βCaMKII (Invitrogen, 13-9800, RRID: AB_2533045), Atg7 (Cell Signaling Technology, 8558s, RRID: AB_10831194), LC3B (Abcam, ab48394, RRID: AB_881433), p62/SQSTM1 (MBL, PM045, RRID: AB_1279301), PSD95 (Cell Signaling Technology, 3450s, RID: AB_2292883), mTOR (Cell Signaling Technology, 2983s, RRID: AB_2105622), p-mTOR (Ser2448; Cell Signaling Technology, 5536s, RRID: AB_10691552), p70 S6 Kinase (Cell Signaling Technology, 9202s, RRID: AB_331676), p-p70 S6 Kinase (Thr389; Cell Signaling Technology, 9205s, RRID: AB_330944), p44/42 MAPK (Erk1/2; Cell Signaling Technology, 9102s, RRID: AB_330744), p-p44/42 MAPK (Erk1/2; Cell Signaling Technology, 4370s, RRID: AB_2315112), AMPK (Cell Signaling Technology, 5831s, RRID: AB_10622186), pAMPKα (Thr172; Cell Signaling Technology, 2535s, RRID: AB_331250), c-fos (Cell Signaling Technology, 2250s, RRID:AB_2247211), Akt (Cell Signaling Technology, 4691s, RRID: AB_915783), pAkt (Ser473; Cell Signaling Technology, 4058s, RRID: AB_331168), TFEB (Proteintech, 13372-1-AP), and H3 (Abcam, Ab176842, AB_2493104). Images were collected using an GE HealthcareImager 600. For each protein of interest, band densitometry was normalized to either tubulin or actin (as a loading control) and quantified using ImageJ (National Institutes of Health).

Immunoprecipitation

For immunoprecipitation (IP), neuronal lysates were mixed with specific antibodies at 4°C overnight while shaking slowly, followed by the addition of protein G magnetic beads (Bio-Rad, 161-4023). Immunocomplexes were washed three time with IP buffer containing 50 mm Tris-HCl (pH 7.9), 150 mm KCl, 5 mm MgCl₂, 0.2 mm EDTA, 20% glycerol, 0.1% NP-40, protease inhibitors, and 3 mm β-mercaptoethanol and then eluted with 2× protein loading buffer for 5 min at 100°C. The immunocomplexes were analyzed via SDS-PAGE and Western blot analysis as described above.

Neuron imaging and analysis

For time-lapse imaging, primary cortical neurons expressing GFP-mCherry-LC3 were transferred to an environmental chamber mounted on the sample stage of a Nikon A1+ confocal laser scanning microscope and maintained for time-lapse imaging at 37°C in 5% CO₂ in Neurobasal medium (without phenol red) supplemented with GlutaMAX, B27, and penicillin-streptomycin. Images of fixed cortical neurons in the same experiment were obtained using the same settings. The acquired images were collapsed to produce 2D projections using Nikon NIS-Elements AR software. ImageJ (https://imagej.nih.gov/ij/) and MetaMorph software v. 7.7.5 (Molecular Devices) were used to analyze the images. For analyzing surface AMPAR GluA1 subunits (sGluA1) and surface AMPAR GluA2 subunits (sGluA2) intensity, dendritic areas containing GluA1/GluA2 staining were traced, and the mean intensity of the selected dendrite was calculated; a total of 6–10 dendritic segments (20–50 μm in length) were averaged for each neuron and used for quantification. For each image, background intensity was subtracted by measuring a region lacking dendrites. For synaptic pCaMKII, the intensity of pCaMKII in PSD95 puncta was analyzed. For analyzing the LC3 puncta and cytosol p62 puncta, the “rolling ball” background subtraction method was used in ImageJ to define the puncta (Wheeler et al., 2012). To measure dendritic LC3 puncta, a region of interest was drawn around the circumference of each dendrite; the ImageJ plugin ComDet v.0.5.0 [approximate particle size: 5.00 pixels; intensity threshold (in SD): 5.00 pixels] was then used to measure the number of LC3 puncta in a given dendrite. For each dataset, the threshold was established based on the corresponding control condition.

Real-time PCR

After 14–18 d in culture (Day in vitro14–DIV18), and following drug treatment, cortical neurons were harvested by directly adding 200 μl TRIzol (Invitrogen, 15596018) to each well in a 12-well plate, and total cellular RNA was isolated in accordance with the manufacturer's instructions. Reverse transcription was then performed to synthesize the corresponding cDNA using the Goldenstar RT6 cDNA Synthesis kit (TsingKe, TSK301M) in accordance with the manufacturer's instructions. Real-time PCR studies were performed using SYBR Premix Ex Taq (Tli RNaseH Plus; Takara) in a Bio-Rad CFX96 cycler (Bio-Rad Laboratories) with 40 cycles (95°C for 5 s, 60°C for 30 s). In each experiment, three independent wells of a 12-well plate were used per condition, and each cDNA sample, equivalent to RNA from one well of cortical neurons cultured in a 12-well plate, was run in duplicate for both the housekeeping genes and target genes.

The primer pair sequences used in this paper are listed below:

β-actin

Forward: AGGCCCCTCTGAACCCTAAG

Reverse: CCAGAGGCATACAGGGACAAC

tubulin

Forward: GAGATCCGAAATGGCCCGTAC

Reverse: GCCAATGGTGTAGTGACCACG

Ulk1

Forward: CATGACCTCCCTTGCATGTAAC

Reverse: ACCAGGTGGTGGGTAAGGAAC

Atg5

Forward: CTGTTCGATCTTCTTGCATCA

Reverse: TCCTTTTCTGGAAAACTCTTGAA

Atg7

Forward: GCTGGGAGAAGAACCAGAAA

Reverse: GAGATTCAGATCCACGGATGAC

LC3B

Forward: TCTTTGTAAGGGCGGTTCTG

Reverse: TCACAAGCATGGCTCTCTTC

Atg12

Forward: AAACGTGAGCCAAGGGGATT

Reverse: GGAAACTTGGTGCTGCTTGG

TFEB

Forward: AATGGGAGCAACCGTACTTAGG

Reverse: GAGGGAAGACAGGTCCATGAA

UVRAG

Forward: ACTCCAGACTTGAGGCAAAC

Reverse: ACAGATACTCACCATCTGACC

ATP6V0E1

Forward: CTCGACTGAGAGAAAAGGTGCT

Reverse: ACATCGCTGACTCTCATGTTGT

Cathepsin F

Forward: GCTCTGCAGTTTTGAAGTCCTG

Reverse: CCAGACTCTGTAACCTTGGCAT

Electrophysiology

Rat cortical neurons transfected with the control/scrambled short-hairpin RNA (shRNA) vector or Atg7 shRNA vectors at DIV8–DIV9 and recorded in the whole-cell configuration at DIV15–DIV16. Where indicated, 2 μm TTX was added 24 h before recording. During the recordings, the neurons were bathed in an external solution containing (in mm): 150 NaCl, 3 KCl, 1 CaCl₂, 1 MgCl₂, 6 glucose, and 10 HEPES (pH 7.4), as well as 0.1 μm TTX, 100 μm picrotoxin, and 70 μm D-AP5. The recording pipettes were filled with an intracellular solution containing (in mm): 130 CsMeSO₄, 6 CsCl, 1 MgCl₂, 10 HEPES, 0.3 EGTA, 4 Mg₂ATP, and 0.3 Na₃GTP (pH 7.25–7.3). All recordings were performed at RT in the voltage-clamp mode at a holding potential of −65 mV. Data were collected using a MultiClamp 700B amplifier (Molecular Devices), filtered at 2 kHz, and digitized at 10 kHz using a Digidata 1550B data acquisition system (Molecular Devices). Series resistance was monitored throughout the recording, and recordings in which series resistance increased to >30 MΩ or changed by >20% during an experiment were discarded.

Electron microscopy

Cortical neurons at DIV14 were treated with control media or TTX for 24 h and then fixed in 2.5% glutaraldehyde in 0.1 m cacodylate buffer (pH 7.2) for 2 h at RT. The fixed neurons were then washed three times in cacodylate buffer, postfixed in 1% osmium tetroxide in 0.1 m phosphate buffer (pH 7.4), progressively dehydrated in a graded ethanol series (30–100%), and flat embedded in Araldite epoxy. Ultrathin sections (70–80 nm thick) were placed on grids (200 mesh), double-stained with uranyl acetate and lead citrate, and imaged using a Philips Tecnai 20 transmission electron microscope at the Center for Cryo-Electron Microscopy (Zhejiang University).

Immunogold electron microscopy

Adult rats were deeply anesthetized with diethyl ether and perfused with ice-cold high-sucrose artificial CSF (ACSF) consisting of the following (in mm): 206 sucrose, 26 NaHCO₃,11 glucose, 10 MgCl₂, 2.5 KCl, 1 NaH₂PO₄, and 0.5 CaCl₂ (pH 7.2–7.4). Brain was removed and the PFC was dissected in ice-cold high-sucrose ACSF using Leica vibratome at 300-μm thickness. Slices were transferred to the incubation chamber with normal ACSF consisting of the following (in mm): 122 NaCl, 26 NaHCO₃, 3 KCl, 11 glucose, 2 CaCl₂, 1.3 MgCl₂, and 1.25 NaH₂PO₄, and were allowed to recover for 30 min at 32°C. Slices were then treated with either 300 nm Torin 1 for 2 h at room temperature. ACSF solutions were saturated with 95% O₂ and 5% CO₂ continuously. Slices were then postfixed with 4% PFA and 0.5% glutaraldehyde dissolved in 0.1 m PB (pH 7.4) at 4°C overnight. After washing for three times with 0.1 m PB (15 min each), sections were incubated with 50 mm glycine in 0.1 m PB (pH 7.4) for 30 min, followed by washing in 0.1 m PB for 15 min. Slices were permeabilized with 0.1% Triton X-100 in 0.1 m PB for 15 min. After washing in 0.1 m PB for 15 min, sections were blocked with blocking buffer containing 0.1% BSA-C (Aurion; Immuno Gold Reagents & Accessories) in 0.1 m PBS (pH 7.4) for 30 min, and incubated with the rabbit anti-CaMKII α antibody (Abcam, ab92332, RRID: AB_2049246, 1:50) at 4°C overnight. Slices were then washed with blocking buffer six times (10 min each), followed by incubation with Nano gold-labeled Fab' anti-rabbit secondary antibody in blocking buffer (1:50) overnight at 4°C. After washing with blocking buffer six times and with 0.1 m PB twice (10 min each), slices were postfixed with 2.5% glutaraldehyde in 0.1 m PBS for 4 h at room temperature, followed by washing successively with 0.1 m PB three times (10 min each), with ddH₂O six times (5 min each), and with 0.02 m sodium citrate buffer in ddH₂O (pH 7.0) three times (5 min each). Sections were treated with HQ Silver Enhancement kit (Nanoprobes Inc.) for 5 min in a dark room, and the reaction was stop by washing in ddH₂O six times (10 min each), followed by washing in 0.1 m PBS (pH 7.4) three times (10 min each). Then brain sections were treated with 1% OsO₄ in 0.1 m PBS for 40 min and washed in ddH₂O three times (10 min each). Slices were treated with 2% uranyl acetate in ddH₂O for 30 min, followed by washing in ddH₂O three times (5 min each). Finally, sections were successively dehydrated in 50%, 70% and 90% ethanol (15 min each), then 100% ethanol twice (20 min each), followed by 100% acetone twice (20 min each). Sections were embedded in EPON resin and sliced into sections at 70 nm thickness. Autophagic vesicles were observed using FEI Tecnai G2 spirit 120 kV TWIN TEM (Thermo FEI) at the Center for Cryo-Electron Microscopy (Zhejiang University).

Quantification and statistical analysis

Sample sizes were chosen based on preliminary experiments or similar studies performed previously. For quantification of immunoblots, electrophysiology and real-time PCR experiments, a minimum of three biologically independent experiments were performed. For quantification of immunofluorescence microscopy images (from at least two cultures), images were recorded using the same microscope settings (objective lens and illumination intensity) to ensure reliable quantification across samples and images. Confocal images in the figures represent maximum intensity projections of 3-μm-thick confocal stacks. In addition, images of each biological replicate were captured blindly. All summary data are expressed as the mean ± SEM and the individual data points are shown in bar graphs if n < 10. Statistical analyses were performed using Prism (GraphPad Software; https://www.graphpad-prism.cn), in which the Student's t test, one-way ANOVA, Kolmogorov–Smirnov test, χ² test or one-way repeated measures (RM) ANOVA test were used. Differences were considered significant at p < 0.05.

Resource availability

Lead contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the corresponding author.

Materials availability

All unique resources generated in this study are available on request.

Data and code availability

The data generated during this study are available from the corresponding author on request.

Results

Autophagy flux is increased by chronic neuronal inactivity during synaptic up-scaling

In mammalian central synapses, an increase in AMPARs in the plasma membrane is one end-point effector mediating synaptic up-scaling (O'Brien et al., 1998; Turrigiano et al., 1998; Thiagarajan et al., 2005; Sutton et al., 2006; Wierenga et al., 2006; Cingolani et al., 2008; Ibata et al., 2008). Experimentally, homeostatic changes in AMPARs can be induced in cultured neurons by chronically suppressing neuronal activity using the sodium channel blocker TTX, as well as in brain slices and in vivo (Turrigiano and Nelson, 2004; Pozo and Goda, 2010). To examine autophagy flux together with synaptic scaling, we treated cultured rat cortical neurons with TTX for 24 h to induce synaptic up-scaling and then measured intracellular LC3-II (the lipid-modified form of activated LC3, a biomarker of autophagosomes) in a parallel batch of cultured neurons (Pan et al., 2021). Consistent with previous reports (Turrigiano et al., 1998; Diering et al., 2014), we found that TTX-treated neurons had increased surface AMPAR GluA1 subunits (sGluA1) and an increased amplitude of miniature AMPAR-mediated EPSCs (mEPSCs), reflecting synaptic up-scaling (Fig. 1A). Interestingly, although TTX did not affect the canonical AMP-activated protein kinase (AMPK) pathway (Fig. 1B,C), which induces autophagy by sensing metabolic changes under conditions such as starvation (J. Kim et al., 2011; Lipton and Sahin, 2014; Saxton and Sabatini, 2017), we found that TTX increased LC3-II levels (Fig. 1D,E).

Figure 1.

Chronic neuronal inactivity activates autophagy in primary cortical neurons. A, Schematic illustration of homeostatic synaptic up-scaling in primary cortical neurons (left). At DIV14–DIV18, cortical neurons were treated with TTX (2 μm) for 24 h and stained for sGluA1 subunits (pictures are shown in glow scale; the side bar represents sGluA1 intensity in arbitrary units); MAP2 was also stained to identify neurons (top right). Also shown are mEPSC recordings confirming synaptic up-scaling in TTX-treated cultures (bottom right). B, Representative Western blottings of AMPKα, AMPKα phosphorylated at Thr-172 (p-AMPKα), and tubulin in cortical neurons treated with control media (Ctrl) or TTX for the indicated times. C, Quantification of p-AMPKα levels measured in the groups shown in B; n = 4 experiments/group, Ctrl versus TTX-1 h p = 0.9913, Ctrl versus TTX-3 h p = 0.8726, Ctrl versus TTX-8 h p = 0.4834, Ctrl versus TTX-24 h p > 0.9999, one-way ANOVA followed by Tukey's post hoc test. D, Representative Western blottings of LC3-I, LC3-II, and actin (as a loading control) in cortical neurons treated with Ctrl or TTX for 24 h. E, Quantification of LC3-II levels measured as shown in D; n = 6 experiments/group, p = 0.0006, unpaired t test. F, G, Representative electron micrographs of AVs (F) and quantification of AVs (G) in cortical neurons treated with control media or TTX for 24 h; n = 12–16 neurons/group, p = 0.0006, unpaired t test. AVs in F are indicated by red arrowheads. H, Representative Western blottings of p62 and actin in cortical neurons treated with control media or TTX for 24 h. I, Quantification of p62 levels measured as shown in H; n = 6 experiments/group, p = 0.0041, unpaired t test. J, Immunostaining of the autophagy marker LC3 (green) and the neuronal marker MAP2 (red) in cortical neurons treated with control media or TTX for 24 h. LC3 puncta are indicated by white arrowheads. K, Quantification of LC3 puncta measured in the groups shown in J; n = 49–54 neurons/group, p < 0.0001, unpaired t test. L, Immunostaining of the autophagy adaptor protein p62 (green) and MAP2 (red) in cortical neurons treated with control media or TTX for 24 h. p62 puncta are indicated by white arrowheads and the DAPI channel (blue) is present in merged images to show the nucleus of neurons. M, N, Quantification of p62 puncta and intensity measured in the groups shown in L; n = 45–64 neurons/group for M, p < 0.0001, unpaired t test; n = 45–64 neurons/group for N, p < 0.0001, t = 6.025, unpaired t test. O, P, Representative Western blottings of LC3 (O) and quantification of LC3-II levels (P) measured in DIV14–DIV18 rat cortical neurons treated with control media, TTX, CQ, or both TTX and CQ for 24 h; n = 5 experiments/group, Ctrl versus CQ p = 0.0004, unpaired t test; CQ versus TTX+CQ p = 0.0034, unpaired t test. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001, and ns, not significant. Scale bars: 20 pA/5 s and 10 µm (A), 2 µm (F), and 10 µm (J, L). In this and subsequent figures, summary data are normalized to the unstimulated control and presented as the mean ± SEM.

This increase in LC3-II levels during synaptic up-scaling may reflect an increase in autophagy flux; however, it may also indicate that autophagic degradation at lysosomes is blocked. To test this possibility, we first examined the presence of autophagic vacuoles (AVs), including autophagosomes and autolysosomes, using electron microscopy (Hernandez et al., 2012) and found that TTX-treated neurons had increased AVs (Fig. 1F,G). We then measured p62 (an autophagosome cargo protein in neurons that recruits autophagic substrates to autophagosomes for degradation) using Western blot analysis and found that p62 levels were significantly lower in TTX-treated neurons (Fig. 1H,I), supporting the notion that p62 itself is degraded because of inactivity-induced autophagy flux. Further, we used immunofluorescence with antibodies to stain endogenous LC3 and p62 in cultured neurons treated either with or without TTX; consistent with the results shown above, we found that TTX significantly increased the number of LC3 (Fig. 1J,K) and p62 (Fig. 1L,N) puncta, indicating the recruitment of autophagic substrates to autophagosomes; note that p62 intensity decreased (Itakura and Mizushima, 2011; Fig. 1M). In addition, we assessed the changes in LC3-II accumulation in the presence or absence of chloroquine (CQ), an autophagy inhibitor that disrupts the autophagosome and the lysosome fusion process. We found that treating neurons with CQ alone led to an upregulation of LC3-II, and treating neurons with both CQ and TTX induced a further increase in LC3-II levels (Fig. 1O,P), supporting the notion that TTX treatment increases autophagy flux.

Next, we cultured cortical neurons obtained from transgenic GFP-LC3 mice and directly quantified autophagy flux by measuring GFP-LC3 puncta in neurons identified using the neuron-specific marker microtubule-associated protein 2 (MAP2). We found that TTX-treated neurons had significantly more GFP-LC3 puncta compared with control-treated neurons (Fig. 2A,B), indicating that LC3 is recruited to the autophagosome during autophagy flux. Finally, to determine the destination of these activity-dependent autophagosomes, we expressed a tandem protein containing both acid-stable mCherry-LC3 and acid-sensitive eGFP-LC3 (referred to hereafter as mCherry-eGFP-LC3) in cortical neurons and measured the effect of TTX-induced up-scaling. In the acidic lysosome compartment, eGFP-LC3 fluorescence was quenched, while mCherry-LC3 fluorescence was unaffected. We therefore measured red puncta (reflecting mCherry-LC3 fluorescence alone) to detect autophagosomes and autolysosomes (Kulkarni et al., 2021); yellow puncta (because of the presence of both mCherry-LC3 and eGFP-LC3 fluorescence) indicate the process during which autophagosomes are recruited to autolysosomes for final lysosomal degradation (Pan et al., 2021). We found that TTX-treated neurons had significantly more red puncta both on the somata (Fig. 2C,D) and on dendrites (may include moving AVs; Maday and Holzbaur, 2014, 2016; Fig. 2F,G); in addition, TTX-treated neurons contained more yellow puncta compared with control-treated neurons (Fig. 2C,E). To test this further, we performed time-lapse imaging to monitor puncta formation. As shown previously (Kulkarni et al., 2021), the eGFP-LC3 signal was relatively weak, particularly in the dendrites, which may have been caused by its high background signal (Kulkarni et al., 2021) or because autophagosomes mature and fuse with lysosomes extremely rapidly (Boland et al., 2008). We therefore monitored mCherry-LC3 puncta in neurons and found that these puncta increase significantly following a 24-h treatment with TTX (Fig. 2H–J). Taken together, these results show that autophagy flux in neurons increases during synaptic up-scaling induced by chronic neuronal inactivity.

Figure 2.

Monitoring TTX-induced autophagy flux with fluorescence-tagged LC3 and time-lapse imaging. A, Representative confocal images of GFP-LC3 puncta measured in cortical neurons obtained from GFP-LC3 mice and treated with control media or TTX for 24 h. MAP2 (red) was immunostained to identify neurons, and GFP-LC3 puncta are indicated by white arrowheads. B, Quantification of GFP-LC3 puncta measured in the groups shown in A; n = 49–75 neurons/group, p < 0.0001, unpaired t test. C, Representative confocal images of cortical neurons transfected with mCherry-GFP-LC3 and treated with control media or TTX for 24 h; mCherry-LC3 puncta are indicated by white arrowheads. D, E, Quantification of mCherry puncta (indicating autophagosomes and autolysosomes; D) and yellow puncta (GFP and mCherry; indicating autophagosomes; E) measured in the groups shown in C; n = 53 neurons/group, p = 0.0183 for D, p = 0.0111 for E, unpaired t test. F, Representative confocal images of dendrites in cortical neurons overexpressing the mCherry-GFP-LC3 construct, treated with control media or TTX for 24 h; mCherry puncta are indicated by white arrowheads. G, Quantification of dendritic mCherry puncta measured in the groups shown in F; n = 41 neurons/group, p = 0.0001, unpaired t test. H, I, Representative confocal images of time-lapse images of the same neuron overexpressing the mCherry-GFP-LC3 construct. Images were acquired just before TTX treatment (Pre) and 24 h after TTX treatment. mCherry puncta are indicated by white arrowheads. J, Quantification of mCherry puncta (indicating autophagosomes and autolysosomes) measured in neurons shown in H, I; for Ctrl group, n = 13 neurons, Pre versus 24 h p = 0.9628, Pre versus 24 h + 10 min p = 0.5605, Pre versus 24 h + 20 min p = 0.7848, one-way RM ANOVA test; for TTX group, n = 12 neurons, Pre versus 24 h p < 0.0001, Pre versus 24 h + 10 min p < 0.0001, Pre versus 24 h + 20 min p < 0.0001, one-way RM ANOVA test. *p < 0.05, ***p < 0.001, and ****p < 0.0001. Scale bars: 5 µm (F), 10 µm (A, C, H, and I).

Autophagy is required for TTX-induced synaptic up-scaling

Our finding that autophagy flux is induced by chronic neuronal inactivity prompted us to ask whether autophagy is required for synaptic up-scaling. To address this question, we first treated neurons with CQ to block the autophagy-lysosomal degradation pathway. We found that the TTX-induced increase in sGluA1 was prevented by co-treating neurons with CQ (Fig. 3A,B), indicating that protein turnover via the autophagy-lysosomal pathway is essential for synaptic up-scaling. To clarify the role of autophagy in this process, we treated neurons with either wortmannin (Wort), which blocks autophagosome formation by potently inhibiting phosphatidylinositol 3-kinase (Hou et al., 2008), or SBI-0206965 (SBI), which prevents the initial steps of autophagosome formation by inhibiting the autophagy kinase Unc-51 like autophagy activating kinase (ULK1/2) (Nikoletopoulou et al., 2017; Pandey et al., 2020; Pan et al., 2021). We found that both Wort (Fig. 3C,D) and SBI (Fig. 3E,F) prevented the TTX-induced increase in sGluA1. In addition to sGluA1, surface AMPAR GluA2 subunits (sGluA2) also increase coordinately for synaptic up-scaling induced by chronic TTX treatment (Ibata et al., 2008; Jakawich et al., 2010). In line with our sGluA1 results, this TTX-induced increase in sGluA2 (Fig. 3G,H) was inhibited when autophagy triggered by inactivity was blocked using SBI (Fig. 3I,J). To rule out possible off-target effects caused by these pharmacological inhibitors, we inhibited autophagy using three separate shRNA constructs to knock down Atg7 (Fig. 3K,L), which encodes an enzyme that converts LC3-I to LC3-II; control cells were transfected with a scrambled nonsilencing shRNA. Consistent with our pharmacological data, we found that knocking down Atg7 prevented the TTX-induced increase in LC3-II (Fig. 3M,N) and sGluA1 (Fig. 4A,B).

Figure 3.

Pharmacological inhibition of autophagy prevents TTX-induced synaptic up-scaling. A, B, Representative confocal images of surface GluA1 subunits (A) and quantification of sGluA1 immunofluorescence intensity (B) measured in cortical neurons treated with control media or TTX in the presence or absence of CQ (50 μm). Pictures in A are shown using glow scale (the side bar represents GluA1 staining intensity in arbitrary units). Representative dendritic segments with sGluA1 immunostaining are shown in the insets. n = 44–56 neurons/group, Ctrl versus TTX p < 0.0001, CQ versus TTX+CQ p = 0.7858, one-way ANOVA followed by Tukey's post hoc test. C, D, Representative confocal images of sGluA1 (C) and quantification of sGluA1 immunofluorescence intensity (D) measured in cortical neurons treated with control media or TTX in the presence or absence of Wort (1 μm). Pictures in C are shown using glow scale (the side bar represents GluA1 staining intensity in arbitrary units). Representative dendritic segments with sGluA1 immunostaining are shown in the insets. n = 59–71 neurons/group, Ctrl versus TTX p < 0.0001, Wort versus TTX+Wort p = 0.9223, one-way ANOVA followed by Tukey's post hoc test. E, Immunostaining of sGluA1 in cortical neurons treated for 24 h with control media, TTX (2 μm), SBI (1 μm), or both TTX and SBI. Pictures are shown using glow scale (the side bar represents GluA1 staining intensity in arbitrary units). Representative dendritic segments with sGluA1 immunostaining are shown in the insets. F, Quantification of sGluA1 immunofluorescence intensity measured in the groups shown in E; n = 50–70 neurons/group, Ctrl versus TTX p = 0.0008, SBI versus TTX+SBI p = 0.9974, one-way ANOVA followed by Tukey's post hoc test. G, H, Representative confocal images of sGluA2 (G) and quantification of sGluA2 immunofluorescence intensity (H) measured in cortical neurons treated with control media or TTX in the presence or absence of SBI (1 μm). Pictures in G are shown using glow scale (the side bar represents GluA2 staining intensity in arbitrary units). Representative dendritic segments with sGluA2 immunostaining are shown in the insets. n = 50 neurons/group, Ctrl versus TTX p = 0.0044, SBI versus TTX+SBI p = 0.9502, one-way ANOVA followed by Tukey's post hoc test. I, J, Representative Western blottings of LC3 (J) and quantification of LC3-II levels (J) measured in cortical neurons treated with control media, TTX, SBI, or both TTX+SBI for 24 h; n = 3 experiments/group, Ctrl versus TTX p = 0.0305, SBI versus TTX+SBI p = 0.9998, one-way ANOVA followed by Tukey's post hoc test. K, Representative Western blottings of ATG7 and tubulin in DIV14 cortical neurons infected with a lentivirus expressing a scrambled shRNA (Scr) or Atg7 shRNA constructs (shATG7). L, Quantification of ATG7 levels measured in K. Unless noted otherwise, the shATG7-1 construct was used in subsequent experiments. n = 3 experiments/group, Scr versus shATG7-0 p = 0.0003, Scr versus shATG7-1 p < 0.0001, Scr versus shATG7-2 p < 0.0001, one-way ANOVA followed by Tukey's post hoc test. M, N, Representative Western blottings of LC3 (M) and quantification of LC3-II levels (N) measured in cortical neurons infected with lentivirus expressing Scr or shATG7 and treated with control media or TTX for 24 h; n = 3 experiments/group, Scr versus Scr+TTX p = 0.0470, Scr versus shATG7+TTX p = 0.7843, one-way ANOVA followed by Tukey's post hoc test. Note that SBI or shATG7 in I, J, M, N prevented activity-dependent LC3-II changes but did not appear to affect basal LC3-II levels in neurons. *p < 0.05, ***p < 0.001, ****p < 0.0001, and ns, not significant. Scale bars: 50 µm (A, C, E, and G; insets, 10 µm).

Figure 4.

Knocking down the essential autophagy gene Atg7 prevents synaptic up-scaling. A, Immunostaining of sGluA1 in cortical neurons infected with a lentivirus expressing either the Scr shRNA or shRNA against Atg7 (shATG7-0 through shATG7-2) and treated with control media or TTX for 24 h. Pictures are shown using glow scale (the side bar represents GluA1 staining intensity in arbitrary units). Representative dendritic segments with sGluA1 immunostaining are shown in the insets. B, Quantification of sGluA1 immunofluorescence intensity measured in the groups shown in C; n = 32–74 neurons/group, Scr versus Scr+TTX p < 0.0001, shATG7-0 versus shATG7-0+TTX p > 0.9999, shATG7-1 versus shATG7-1+TTX p = 0.3028, shATG7-2 versus shATG7-2+TTX p = 0.9954, one-way ANOVA followed by Tukey's post hoc test. C, Whole-cell voltage-clamp recordings of cortical neurons infected with a lentivirus expressing either the Scr shRNA or shTAG7; where indicated, the neurons were treated for 24 h with TTX. Shown are representative traces of AMPAR-mediated mEPSCs. D, E, Quantification of mEPSC amplitude (D) and the cumulative distribution of mEPSC amplitude (E) measured from the groups in C; n = 10–16 neurons/group for D, Scr versus Scr+TTX p = 0.0458, Scr versus Scr+TTX p > 0.9999, one-way ANOVA followed by Tukey's post hoc test; n = 10–16 neurons/group for E, Scr versus Scr+TTX p = 0.0002, shATG7 versus shATG7+TTX p = 0.9998, Kolmogorov–Smirnov test. F–I, mEPSC frequency (F), half-width (G), rise time (H), and decay time (I) were measured in cultured neurons infected with lentivirus expressing Scr shRNA or shATG7 and treated with control media or TTX for 24 h; F, Scr versus Scr+TTX p = 0.9961, shATG7 versus shATG7+TTX p = 0.9998; G, Scr versus Scr+TTX p = 0.6774, shATG7 versus shATG7+TTX p = 0.3834; H, Scr versus Scr+TTX p = 0.4238, shATG7 versus shATG7+TTX p = 0.7558; I, Scr versus Scr+TTX p = 0.3006, shATG7 versus shATG7+TTX p = 0.6435; F–I, one-way ANOVA followed by Tukey's post hoc test. *p < 0.05, ****p < 0.0001 and ns, not significant. Scale bars: 50 µm (A; insets, 10 µm), and 20 pA/500 ms (C).

Next, to examine whether autophagy-dependent changes in surface AMPARs are functionally relevant, we recorded mEPSCs in control-treated and TTX-treated neurons expressing either the scrambled shRNA or the Atg7 shRNA. Consistent with previous reports (Turrigiano et al., 1998), TTX increased mEPSC amplitude, but had no effect on mEPSC frequency, half width, rise time, or decay time, in cortical neurons expressing the scrambled shRNA (Fig. 4C–I); in contrast, this TTX-induced increase in mEPSC amplitude was absent in cells expressing the Atg7 shRNA (Fig. 4C–I), suggesting that autophagy plays a functional role in synaptic up-scaling.

Autophagy is required for the TTX-induced reduction in αCaMKII and PSD95

Given that the autophagy pathway is activated by chronic neuronal inactivity and plays a critical role in synaptic up-scaling, we asked whether autophagy is required for synaptic turnover during synaptic up-scaling. To address this question, we first measured αCaMKII, one of the most abundant proteins expressed in excitatory neurons (He et al., 2021; Yasuda et al., 2022), particularly at the postsynaptic density (Kennedy, 2000; Hudmon and Schulman, 2002; Hell, 2014). Previous studies found that following neuronal inactivity, a net decrease in αCaMKII was required for synaptic up-scaling (Thiagarajan et al., 2002; Groth et al., 2011); however, the molecular mechanism underlying this inactivity-dependent loss of αCaMKII remains unclear. Using the iLIR server for the in-silico identification of functional LIR (LC3-interacting region) motifs (http://repeat.biol.ucy.ac.cy/iLIR/), we determined that αCaMKII contains multiple LIR motifs, suggesting that this protein may be targeted by autophagy. To test this, we first performed co-immunoprecipitation experiments using lysates prepared from the cortex, with an antibody against LC3 to pull down endogenous LC3 followed by immunoblotting with an antibody against αCaMKII to detect endogenous αCaMKII. We found that αCaMKII is selectively pulled down by LC3 (Fig. 5A), indicating that αCaMKII may be targeted by autophagy. To confirm this finding, we used the mTOR inhibitor Torin 1 to induce mTOR-dependent autophagy in the cortex (Pan et al., 2021) and then performed immunogold electron microscopy. We found that AVs with the presence of αCaMKII increased significantly following Torin 1 treatment (Fig. 5B,C), suggesting that αCaMKII is an autophagy substrate. Given that CaMKII isoforms in excitatory cortical neurons form either αCaMKII homomultimers or αCaMKII and βCaMKII heteromultimers (Bennett et al., 1983; Brocke et al., 1999), and given that αCaMKII and βCaMKII have similar LIR motifs (http://repeat.biol.ucy.ac.cy/iLIR/), our data indicate that αCaMKII homomultimers are selectively targeted by autophagy, reminiscent of the binding between αCaMKII homomultimers and the proteasome (Bingol et al., 2010).

Figure 5.

TTX-induced loss of αCaMKII is autophagy dependent. A, Representative Western blottings showing LC3 co-immunoprecipitated with αCaMKII in mouse cortical lysates (n = 3 mice). Note that βCaMKII was not pulled down by LC3. B, C, Representative electron micrographs of immunogold-labeled αCaMKII in AVs measured in rat cortical slices treated with control media (Ctrl) or Torin 1 (300 nm) for 2 h (B), and quantification of the percentage of αCaMKII⁺ AVs (indicated by the presence of αCaMKII immunogold labeling) in neurons (C); n = 39–43 neurons/group, p < 0.0001, χ² test. αCaMKII immunogold in AVs is indicated by red arrowheads. D, E, Representative confocal images of αCaMKII (D) and quantification of dendritic αCaMKII immunofluorescence intensity (E) measured in DIV14 rat cortical neurons treated with control media, TTX, SBI, or both TTX and SBI; n = 50–54 neurons/group, Ctrl versus TTX p < 0.0001, SBI versus TTX+SBI p = 0.9285, one-way ANOVA followed by Tukey's post hoc test. Representative dendritic segments with αCaMKII immunostaining are shown in the insets in D. F, G, Representative Western blottings of αCaMKII (F) and quantification of αCaMKII levels (G) measured in cortical neurons treated with control media, TTX, SBI, or both TTX and SBI for 24 h; n = 4 experiments/group, Ctrl versus TTX p = 0.0177, SBI versus TTX+SBI p = 0.7235, one-way ANOVA followed by Tukey's post hoc test. H, I Representative confocal images of dendritic αCaMKII (H) and quantification of dendritic αCaMKII immunofluorescence intensity (I) measured in cortical neurons infected with lentivirus expressing Scr shRNA or shATG7 and treated with control media or TTX for 24 h; n = 66–76 neurons/group, Scr versus Scr+TTX p < 0.0001, shATG7 versus shATG7+TTX p = 0.7160, Scr versus shATG7 p < 0.0001, one-way ANOVA followed by Tukey's post hoc test. Representative dendritic segments with αCaMKII immunostaining are shown in the insets in H. J, K, Representative Western blottings of αCaMKII (J) and quantification of αCaMKII levels (K) measured in cortical neurons infected with lentivirus expressing Scr or shATG7 and treated with control media or TTX for 24 h; n = 5 experiments/group, Scr versus Scr+TTX p = 0.0002, shATG7 versus shATG7+TTXp = 0.9996, Scr versus shATG7 p < 0.0001, one-way ANOVA followed by Tukey's post hoc test. L, Normalized αCaMKII mRNA levels in cortical neurons treated with control media or SBI for 24 h, expressed relative to control levels; n = 4 experiments/group, p = 0.5739, unpaired t test. M, At DIV9, cortical neurons were infected with a lentivirus expressing Scr or shATG7, and αCaMKII mRNA levels were measured and are expressed relative to Scr; n = 4 experiments/group, p = 0.1127, unpaired t test. *p < 0.05, ***p < 0.001, ****p < 0.0001 and ns, not significant. Scale bars: 1 µm (B; insets, 500 nm) and 50 μm (D, H; insets, 10 µm).

To test this at the functional level, we treated cultured neurons with both SBI and TTX to simultaneously block autophagy and drive neuronal inactivity, respectively. In line with previous reports (Thiagarajan et al., 2002), αCaMKII was reduced by TTX (Fig. 5D–K). Interestingly, we also found that αCaMKII protein levels were decreased when autophagy was blocked by knocking down Atg7 under basal conditions (Fig. 5H–K). In contrast, αCaMKII mRNA were unaffected when autophagy was inhibited (Fig. 5L,M), supporting the notion that autophagy is required to generate amino acids (J. Kim et al., 2011; Saxton and Sabatini, 2017) needed for protein synthesis (Mayford et al., 1996; Miller et al., 2002). Nevertheless, we found that the inactivity-induced αCaMKII loss was prevented by SBI and the Atg7 shRNA (Fig. 5D–K). In addition, inhibiting autophagy with SBI or an shRNA targeted to Atg7 mRNA prevented the inactivity-induced phosphorylation of synaptic CaMKII at Thr-286/287 (pCaMKII; Fig. 6A–D). Given that pCaMKII is regulated by the ratio of βCaMKII/αCaMKII at synapses (Thiagarajan et al., 2002; Groth et al., 2011), and given that pCaMKII is upstream of sGluA1 upregulation (Hudmon and Schulman, 2002; Hell, 2014), these findings suggest that αCaMKII is a key autophagy substrate for synaptic up-scaling.

Figure 6.

The role of autophagy in regulating pCaMKII and PSD95. A, B, Representative confocal images of pCaMKII (A) and quantification of synaptic pCaMKII immunofluorescence intensity (B) measured in DIV14 rat cortical neurons treated with control media, TTX, SBI, or both TTX and SBI; n = 51–62 neurons/group, Ctrl versus TTX p = 0.0301, SBI versus TTX+SBI p = 0.8446, one-way ANOVA followed by Tukey's post hoc test. C, D, Representative confocal images of pCaMKII (C) and quantification of synaptic pCaMKII immunofluorescence intensity (D) measured in DIV14 rat cortical neurons infected with lentivirus expressing Scr or shATG7 and treated with control media or TTX for 24 h; n = 51–52 neurons/group, Scr versus Scr+TTX p = 0.0015, shATG7 versus shATG7+TTX p = 0.9988, one-way ANOVA followed by Tukey's post hoc test. E, F, Representative confocal images of PSD95 (E) and quantification of PSD95 puncta (F) measured in cortical neurons infected with lentivirus expressing Scr or shATG7 and treated with control media or TTX for 24 h; n = 66–76 neurons/group, Scr versus Scr+TTX p = 0.0269, shATG7 versus shATG7+TTX p = 0.6836, Scr versus shATG7 p < 0.0001, one-way ANOVA followed by Tukey's post hoc test. Representative dendritic segments with PSD95 immunostaining are shown in the insets in E. G, H, Representative Western blottings of PSD95 (G) and quantification of PSD95 levels (H) measured in cortical neurons treated with control media, TTX, SBI, or both TTX and SBI for 24 h; n = 6 experiments/group, Ctrl versus TTX p = 0.0002, SBI versus TTX+SBI p = 0.3553, one-way ANOVA followed by Tukey's post hoc test. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001, and ns, not significant. Scale bars: 20 µm (A), 10 µm (C) and 50 µm (E; insets, 10 µm).

Next, we measured the postsynaptic protein PSD95, a known target of autophagy that also contains LIR motifs (Nikoletopoulou et al., 2017; Compans et al., 2021; Pan et al., 2021). Consistent with our findings with αCaMKII, we found that the TTX-induced decrease in PSD95 (Ehlers, 2003) was prevented by either expressing the Atg7 shRNA (Fig. 6E,F) or treating the neurons with SBI (Fig. 6G,H). Taken together, these data suggest that synaptic turnover during up-scaling, exemplified by αCaMKII and PSD95, requires autophagy.

Autophagy is not required for bicuculline-induced synaptic down-scaling

Homeostatic synaptic scaling either increases or decreases synaptic strength in response to changes in input. Our finding that autophagy plays a key role in synaptic up-scaling prompted us to ask whether autophagy also plays a role in synaptic down-scaling, a process in which global increases in activity lead to significant down-scaling of the synaptic response (Turrigiano, 2008). To induce synaptic down-scaling, we treated cultured cortical neurons with Bic, a selective antagonist of type A GABA receptors, thus increasing excitatory neuronal activity. Immunostaining of sGluA1 confirmed that treating neurons with Bic for 24 h induced synaptic down-scaling (Fig. 7A–D); however, in contrast to TTX-induced synaptic up-scaling, Bic-induced down-scaling had no effect on LC3-II levels, even after 72 h (Fig. 7E,F). Consistent with this finding, Bic-induced synaptic down-scaling had no effect on p62 puncta (Fig. 7G,H).

Figure 7.

Lack of Effects of bicuculline and all-trans retinoic acid on autophagy. A, Immunostaining of sGluA1 in cortical neurons treated with control media, Bic, Wort (1 μm), or both Bic and Wort for 24 h. Pictures are shown using glow scale (the side bar represents GluA1 staining intensity in arbitrary units). Representative dendritic segments with sGluA1 immunostaining are shown in the insets. B, Quantification of sGluA1 immunofluorescence intensity measured in the groups shown in A; n = 29–46 neurons/group, Ctrl versus Bic p < 0.0001, Wort versus Bic+Wort p < 0.0001, one-way ANOVA followed by Tukey's post hoc test. C, Immunostaining of sGluA1 in cortical neurons treated with control media, Bic, SBI, or both Bic and SBI for 24 h. Pictures are shown using glow scale (the side bar represents GluA1 staining intensity in arbitrary units). Representative dendritic segments with sGluA1 immunostaining are shown in the insets. D, Quantification of sGluA1 immunofluorescence intensity measured in the groups shown in C; n = 29–46 neurons/group, Ctrl versus Bic p < 0.0001, SBI versus Bic+SBI p < 0.0001, one-way ANOVA followed by Tukey's post hoc test. E, Representative Western blottings of LC3 and tubulin in cortical neurons treated with control media or Bic for the indicated times. F, Quantification of LC3-II levels measured in the groups shown in E; n = 6–10 experiments/group, Ctrl versus Bic-24 h p > 0.9999, Ctrl versus Bic-48 h p = 0.9312, Ctrl versus Bic-72 h p = 0.9929, one-way ANOVA followed by Tukey's post hoc test. G, Immunostaining of the autophagy adaptor protein p62 (green) and the neuronal marker MAP2 (red) in cortical neurons treated with control media or Bic for 24 h. H, Quantification of p62 puncta measured in the groups shown in G; n = 97–114 neurons/group, p = 0.5141, unpaired t test. I, Representative Western blottings of LC3 and tubulin in cortical neurons treated with control media or RA (1 μm) for 1.5 h. J, Quantification of LC3-II levels measured in the groups shown in I; n = 6 experiments/group, p = 0.6324, unpaired t test. K, Representative Western blottings of p62 and tubulin in cortical neurons treated with control media or RA for 1.5 h. L, Quantification of p62 levels measured in the groups shown in K; n = 6 experiments/group, p = 0.2517, unpaired t test. M, Immunostaining of sGluA1 in cortical neurons treated with control media, RA, SBI, or both RA and SBI for 1.5 h. Pictures are shown using glow scale (the side bar represents GluA1 staining intensity in arbitrary units). Representative dendritic segments with sGluA1 immunostaining are shown in the insets. N, Quantification of sGluA1 immunofluorescence intensity measured in the groups shown in M; n = 79–115 neurons/group, Ctrl versus RA p < 0.0001, SBI versus RA+SBI p < 0.0001, one-way ANOVA followed by Tukey's post hoc test. ****p < 0.0001, and ns, not significant. Scale bar: 50 µm (insets, 10 µm).

Although autophagy does not occur during synaptic down-scaling, it may still be necessary for this process. To test this possibility, we blocked autophagy using either Wort or SBI while inducing synaptic down-scaling with Bic and found that neither Wort nor SBI prevented the Bic-induced decrease in sGluA1 (Fig. 7A–D), suggesting that autophagy is not required for synaptic down-scaling.

Local synaptic up-scaling is autophagy independent

In addition to classic synaptic homeostasis driven by changes in neuronal firing rate, another form of synaptic scaling can be induced by changes in local synaptic drive (a phenomenon known as local synaptic scaling; Sutton and Schuman, 2006; Aoto et al., 2008; Branco et al., 2008; Lindskog et al., 2010). A characteristic feature of local synaptic scaling is its relatively rapid time course (on the order of ∼2–4 h; Turrigiano and Nelson, 2004); moreover, this form of synaptic scaling engages distinct signaling processes that require protein synthesis (Sutton and Schuman, 2006; Aoto et al., 2008). To test whether autophagy plays a role in this rapid form of up-scaling, we treated neurons with all-trans RA, which has been shown to induce synaptic up-scaling in only 2 h (Aoto et al., 2008). We found that RA had no effect on either LC3-II or p62 (Fig. 7I–L), but significantly increased sGluA1 (Fig. 7M,N). In addition, inhibiting autophagy with SBI had no effect on the RA-induced increase in sGluA1 (Fig. 7M,N), suggesting that autophagy is not required for local synaptic up-scaling.

Autophagy mediates chronic UPS inhibition-induced sGluA1 upregulation

The UPS can be recruited in cultured neurons by chronic Bic treatment and plays an essential role in synaptic down-scaling (Ehlers, 2003; Djakovic et al., 2009; Jakawich et al., 2010; Shin et al., 2012; Dorrbaum et al., 2020). In contrast, simply blocking the UPS for 24 h using proteasome inhibitors increases synaptic strength and occludes synaptic up-scaling induced by blocking neuronal spiking (Jakawich et al., 2010). Thus, these previous findings support the notion that chronic neuronal inactivity and UPS inhibition may use a common signaling pathway for driving synaptic up-scaling. Given the recent finding of crosstalk between the UPS and autophagy (Pohl and Dikic, 2019), we asked whether inhibiting the UPS can trigger autophagy for synaptic up-scaling. We therefore treated cultured neurons with MG132, a potent cell-permeable proteasome inhibitor. Interestingly, we found that neurons treated for 24 h with MG132 had significantly increased levels of LC3-II (Fig. 8A,B). Similar results were obtained using the proteasome inhibitor bortezomib (BTZ) (Fig. 8C,D), suggesting that chronic UPS inhibition indeed activates autophagy in neurons. Consistent with this finding, and consistent with the notion that autophagy can degrade αCaMKII and PSD95, we found that BTZ decreased the expression of these two synaptic proteins and prevented their further decrease during TTX treatment (Fig. 8E–G), indicating that BTZ-induced autophagy and TTX-induced autophagy may preclude each other.

Figure 8.

Autophagy induced by proteasome malfunction occludes TTX-induced changes of synaptic proteins. A, Representative Western blottings of LC3 and tubulin (as a loading control) in cortical neurons treated with control media or MG132 (2 μm) for 24 h. B, Quantification of LC3-II levels measure in the groups shown in A; n = 6 experiments/group, p = 0.0173, unpaired t test. C, Representative Western blottings of LC3 and tubulin in cortical neurons treated with control media or BTZ (10 nm) for 24 h. D, Quantification of LC3-II levels measured in the groups shown in C; n = 7 experiments/group, p = 0.0151, unpaired t test. E, Representative Western blottings of PSD95, αCaMKII, and tubulin measured in cortical neurons treated with control media, TTX, BTZ, or both TTX and BTZ for 24 h. F, G, Quantification of PSD95 levels (F) and αCaMKII levels (G) measured in the groups shown in E; n = 6 experiments/group for F, Ctrl versus TTX p = 0.0002, Ctrl versus BTZ p < 0.0001, BTZ versus TTX+BTZ p = 0.5959, one-way ANOVA followed by Tukey's post hoc test; n = 6 experiments/group for G, Ctrl versus TTX p = 0.0456, Ctrl versus BTZ p = 0.0005, BTZ versus TTX+BTZ p = 0.9868, one-way ANOVA followed by Tukey's post hoc test. H, Immunostaining of c-fos in cortical neurons treated with control media, MG132, or BTZ for 2 h. I, Quantification of c-fos immunofluorescence intensity measured in the groups shown in H; n = 80–87 neurons/group, Ctrl versus MG132 p = 0.1575, Ctrl versus BTZ p = 0.6987, one-way ANOVA followed by Tukey's post hoc test. J, Representative Western blottings of c-fos and tubulin in cortical neurons treated with control media, MG132, or BTZ for 2 h. K, Quantification of c-fos levels measured in the groups shown in J; n = 4 experiments/group, Ctrl versus MG132 p = 0.8700, Ctrl versus BTZ p = 0.8356, one-way ANOVA followed by Tukey's post hoc test. *p < 0.05, ***p < 0.001, ****p < 0.0001, and ns, not significant. Scale bar: 50 µm.

Neuronal activity does not appear to increase when the UPS is blocked (Ehlers, 2003), as shown by measuring the expression of immediate-early gene c-fos following the MG132 or BTZ treatment (Fig. 8H–K). However, we found that chronically inhibiting the UPS (Jakawich et al., 2010; Srinivasan et al., 2021) using either MG132 or BTZ increased sGluA1 levels (Fig. 9), similar to our findings with TTX-treated neurons. Interestingly, the increased sGluA1 levels induced by either MG132 or BTZ was prevented by the autophagy inhibitors SBI and Wort (Fig. 9), suggesting that autophagy mediates the upregulation of sGluA1 mediated by chronic UPS inhibition.

Figure 9.

Increased sGluA1 following proteasome malfunction requires autophagy. A, Immunostaining of sGluA1 in cortical neurons treated with control media, MG132, SBI, or both MG132 and SBI for 24 h. Pictures are shown using glow scale (the side bar represents GluA1 staining intensity in arbitrary units). Representative dendritic segments with sGluA1 immunostaining are shown in the insets. B, Quantification of sGluA1 immunofluorescence intensity measured in the groups shown in A; n = 51–61 neurons/group, Ctrl versus MG132 p < 0.0001, SBI versus MG132+SBI p = 0.7371, one-way ANOVA followed by Tukey's post hoc test. C, Immunostaining of sGluA1 in cortical neurons treated with control media, BTZ, SBI, or both BTZ and SBI for 24 h. Pictures are shown using glow scale (the side bar represents GluA1 staining intensity in arbitrary units). Representative dendritic segments with sGluA1 immunostaining are shown in the insets. D, Quantification of sGluA1 immunofluorescence intensity measured in the groups shown in C; n = 32–44 neurons/group, Ctrl versus BTZ p < 0.0001, SBI versus BTZ+SBI p = 0.8254, one-way ANOVA followed by Tukey's post hoc test. E, Immunostaining of sGluA1 in cortical neurons treated with control media, MG132, Wort, or both MG132 and Wort. Pictures are shown using glow scale (the side bar represents GluA1 staining intensity in arbitrary units). Representative dendritic segments with sGluA1 immunostaining are shown in the insets. F, Quantification of sGluA1 immunofluorescence intensity measured in the groups shown in E; n = 50–62 neurons/group, Ctrl versus MG132 p < 0.0001, Wort versus MG132+Wort p > 0.9999, one-way ANOVA followed by Tukey's post hoc test. G, Immunostaining of sGluA1 in cultured cortical neurons treated with control media, BTZ, Wort, or both BTZ and Wort for 24 h. Pictures are shown using glow scale (the side bar represents GluA1 staining intensity in arbitrary units). Representative dendritic segments with sGluA1 immunostaining are shown in the insets. H, Quantification of sGluA1 immunofluorescence intensity measured in the groups shown in G; n = 28–43 neurons/group, Ctrl versus BTZ p < 0.0001, Wort versus BTZ+Wort p = 0.9900, one-way ANOVA followed by Tukey's post hoc test. ****p < 0.0001 and ns, not significant. Scale bar: 50 µm (insets, 10 µm).

mTOR is dephosphorylated during synaptic up-scaling

Our finding that autophagy is engaged both by chronically inhibiting the UPS and by chronic neuronal inactivity led us to investigate the molecular mechanism underlying these homeostatic processes. The mTOR signaling pathway, particularly complex 1 (mTORC1), has been shown to participate in the crosstalk between the UPS and autophagy by bridging proteasome malfunction with autophagy activation (Pohl and Dikic, 2019). Given that dephosphorylation of mTOR is a key step in initiating classic mTOR-dependent autophagy (J. Kim et al., 2011; Lipton and Sahin, 2014; Saxton and Sabatini, 2017), we hypothesized that both UPS inhibition and neuronal inactivity lead to mTOR dephosphorylation to engage autophagy and increase sGluA1 levels. To test this hypothesis, we measured the level of phosphorylated mTOR (p-mTOR) at Ser-2448 (Chiang and Abraham, 2005) by inhibiting UPS with either MG132 or BTZ. Interestingly, we found that inhibiting UPS decreased not only p-mTOR levels (Fig. 10A–D) but also mTORC1's immediate downstream effector, phosphorylated p70S6K (p-p70S6K; Fig. 10E–H). Moreover, we measured the effect of chronic neuronal inactivity on both p-mTOR and p-p70S6K levels (Figueiredo et al., 2017) and found that they were reduced in TTX-treated neurons compared with control neurons (Fig. 10I–P). Next, we examined the process by which neuronal inactivity leads to mTOR dephosphorylation. Given that p-mTOR is known to be regulated by the MEK-ERK and PI3K pathways in neurons (Hoeffer and Klann, 2010), we measured phosphorylated ERK1/2 at Thr-202/Tyr-204 (p-ERK1/2) and phosphorylated Akt at Ser-473 (p-Akt). Importantly, we found that TTX decreased p-ERK1/2 (Fig. 10Q–S), but not p-Akt (Fig. 10T,U), suggesting that neuronal inactivity may lead to mTOR dephosphorylation via the MEK-ERK pathway.

Figure 10.

mTORC1 signaling is downregulated during synaptic up-scaling. A, Representative Western blottings of mTOR, mTOR phosphorylated at Ser-2448 (p-mTOR), and tubulin in cortical neurons treated with control media or MG132 for 24 h. B, Quantification of p-mTOR levels measured in the groups shown in A; n = 3 experiments/group, p = 0.0303, unpaired t test. C, Representative Western blottings of mTOR, p-mTOR, and tubulin in cortical neurons treated with control media or BTZ for 24 h. D, Quantification of p-mTOR levels measured in the groups shown in C; n = 3 experiments/group, p = 0.0015, unpaired t test. E, Representative Western blottings of p70S6K, p70S6K phosphorylated at Thr-389 (p-p70S6K), and tubulin in cortical neurons treated with control media or MG132 for 24 h. F, Quantification of p-p70S6K levels measured in the groups shown in E; n = 3 experiments/group, p = 0.0415, unpaired t test. G, Representative Western blottings of p70S6K, p-p70S6K, and tubulin in cortical neurons treated with control media or BTZ for 24 h. H, Quantification of p-p70S6K levels measured in the groups shown in G; n = 3 experiments/group, p = 0.0370, unpaired t test. I, Representative Western blottings of mTOR, p-mTOR, and tubulin in cortical neurons treated with control media or TTX for 24 h. J, Quantification of p-mTOR levels measured in the groups shown in I; n = 4 experiments/group, p = 0.0050, unpaired t test. K, Representative Western blottings of p70S6K, p-p70S6K, and tubulin in cortical neurons treated with control media or TTX for 24 h. L, Quantification of p-p70S6K levels measured in the groups shown in K; n = 3 experiments/group, p = 0.0208, unpaired t test. M, Immunostaining of p-mTOR (green) and MAP2 (red) in cortical neurons treated with control media or TTX for 24 h. N, Quantification of p-mTOR immunofluorescence intensity measured in the groups shown in M; n = 94–112 neurons/group, p < 0.0001, unpaired t test. O, Immunostaining of p-p70S6K (green) and MAP2 (red) in cortical neurons treated with control media or TTX for 24 h. P, Quantification of p-p70S6K immunofluorescence intensity measured in the groups shown in O; n = 93–110 neurons/group, p < 0.0001, unpaired t test. Q, Representative Western blottings of Erk1/2 phosphorylated at p-Erk, Erk1/2, and tubulin in cultured cortical neurons treated with control media or TTX for 24 h. R, S, Quantification of p-Erk1 levels (R) and p-Erk2 levels (S) measured in the groups shown in Q; n = 6 experiments/group for R, p = 0.0002, unpaired t test; n = 6 experiments/group for S, p = 0.0006, unpaired t test. T, Representative Western blottings of AKT, Akt phosphorylated at Ser-473 (p-Akt), and tubulin in cortical neurons treated with control media or TTX for 24 h. U, Quantification of p-Akt levels measured in the groups shown in T; n = 4 experiments/group, p = 0.3994, unpaired t test. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001, and ns, not significant. Scale bar: 50 µm (insets, 10 µm). Scale bar: 20 µm.

Autophagy induced by chronic inactivity is transcription dependent

Next, we investigated the mechanism by which autophagy is maintained during synaptic up-scaling. Previous studies found that de novo gene expression and translation are required for maintaining protein synthesis during long-term synaptic plasticity and homeostasis (Alberini and Kandel, 2015); however, mTOR is inactivated during synaptic up-scaling, which in turn prevents translation (Hoeffer and Klann, 2010). Given the previous finding that TTX induces synaptic up-scaling in a transcription-dependent manner (Ibata et al., 2008), we asked whether inactivity-induced autophagy is transcription dependent. To test this possibility, we treated cultured cortical neurons with Anis to block translation and found that TTX-induced increase in LC3-II was prevented by Anis (Fig. 11A,B). Next, we used actinomycin D (ActD) to inhibit transcription and found that this also prevented the TTX-induced increase in LC3-II (Fig. 11C,D).

Figure 11.

Neuronal inactivity-induced autophagy is transcription dependent. A, Representative Western blottings of LC3 and tubulin in cortical neurons treated with control media, TTX, Anis (50 μm), or both TTX and Anis for 24 h. B, Quantification of LC3-II levels measured in the groups shown in A; n = 5 experiments/group, Ctrl versus TTX p = 0.0076, Anis versus TTX+Anis p = 0.7727, one-way ANOVA followed by Tukey's post hoc test. C, Representative Western blottings of LC3 and tubulin in cortical neurons treated with control media, TTX, ActD (1 μm), or both TTX and ActD for 24 h. D, Quantification of LC3-II levels measured in the groups shown in C; n = 6–7 experiments/group, Ctrl versus TTX p = 0.0016, ActD versus TTX+ActD p = 0.6014, one-way ANOVA followed by Tukey's post hoc test. E–I, Normalized mRNA levels of the indicated genes measured in cortical neurons treated with control media, TTX, ActD, or both TTX and ActD for 24 h, expressed relative to control levels; n = 5 experiments/group for E–I; E, Ctrl versus TTX p = 0.0413, ActD versus TTX+ActD p = 0.9610; F, Ctrl versus TTX p < 0.0001, ActD versus TTX+ActD p = 0.9892; G, Ctrl versus TTX p = 0.0020, ActD versus TTX+ActD p = 0.9877; H, Ctrl versus TTX p = 0.0358, ActD versus TTX+ActD p = 0.4839; I, Ctrl versus TTX p = 0.0395, ActD versus TTX+ActD p = 0.8054; one-way ANOVA followed by Tukey's post hoc test. J, Normalized mRNA levels of the indicated autophagy-related genes in cortical neurons treated with control media or BTZ for 24 h, expressed relative to control levels; ULK1, n = 6 experiments/group, p = 0.0011, unpaired t test; ATG5, n = 6 experiments/group, p = 0.0001, unpaired t test; ATG7, n = 3–6 experiments/group, p = 0.0177, unpaired t test; LC3B, n = 4–6 experiments/group, p < 0.0001, unpaired t test; ATG12, n = 3–6 experiments/group, p < 0.0001 unpaired t test. K, Normalized mRNA levels of the indicated autophagy-related genes in cortical neurons treated with control media or Bic for 24 h, expressed relative to control levels; ULK1, n = 5 experiments/group, p = 0.5731, unpaired t test; ATG5, n = 5 experiments/group, p = 0.9555, unpaired t test; ATG7, n = 3–5 experiments/group, p = 0.0468, unpaired t test; LC3B, n = 5 experiments/group, p = 0.5508, unpaired t test; ATG12, n = 5 experiments/group, p = 0.0725, unpaired t test. L, Normalized mRNA levels of the indicated autophagy-related genes in cortical neurons 24 h after TTX washout, expressed relative to control levels; ULK1, n = 4 experiments/group, p = 0.2624, unpaired t test; ATG5, n = 4 experiments/group, p = 0.8866, unpaired t test; ATG7, n = 4 experiments/group, p = 0.8866, unpaired t test; LC3B, n = 4 experiments/group, p = 0.7728, unpaired t test; ATG12, n = 4 experiments/group, p = 0.4567, unpaired t test. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001, and ns, not significant.

One possible explanation for this finding is that transcriptional regulation may be involved in TTX-induced autophagy, as shown previously in several cell types (Fullgrabe et al., 2016; Sakamaki et al., 2018; Di Malta et al., 2019), including neurons (Xu et al., 2011; Nikoletopoulou et al., 2017; Pan et al., 2021). To test this possibility, we measured the expression of several autophagy-related genes, including Ulk1 (involved in autophagosome induction), Atg5 and Atg12 (involved in phagophore expansion), and Atg7 and LC3B (involved in autophagosome assembly), and found that all of these genes were upregulated after a 24-h treatment with TTX or BTZ, whereas Bic treatment had no effect on most of them (Fig. 11E–K); moreover, the TTX-induced gene upregulation is dependent on both neuronal inactivity and transcription, as it returned to baseline within 24 h after removing TTX (Fig. 11L) and was prevented by ActD (Fig. 11E–I), respectively. Taken together, these findings suggest that chronic neuronal inactivity drives the transcription of autophagy genes.

TFEB is required for TTX-induced transcription-dependent autophagy and synaptic up-scaling

Next, we examined the molecular mechanism underlying transcription-dependent autophagy during synaptic homeostasis. We recently showed that the transcription factors CRTC1 and CREB are selectively recruited by LTD-inducing stimuli, thus driving the expression of autophagy-related genes (Pan et al., 2021). To test whether synaptic up-scaling uses the same mechanism, we measured CRTC1/CREB signaling in our cultured neurons and found that neither the distribution of CRTC1 nor CREB phosphorylation at Ser-133 is affected by chronic neuronal inactivity (Fig. 12A–D). Moreover, we found that either knocking down CRTC1 expression using shRNA (Pan et al., 2021) or inhibiting CREB using a dominant negative inhibitor (A-CREB; Ahn et al., 1998) did not prevent TTX-induced autophagy (Fig. 12E–H). These results suggest a key difference in transcription-dependent autophagy between late-phase LTD and synaptic up-scaling, raising the question of which mechanism mediates the inactivity-induced expression of autophagy-related genes.

Figure 12.

CRTC1-CREB signaling is not required for TTX-induced autophagy. A, B, Representative confocal images of CRTC1 (green) and MAP2 (red) immunofluorescence (A) and quantification of the nuclear-to-cytoplasm ratio of CRTC1 (B) measured in cortical neurons treated with control media, TTX, or Bic; n = 49–70 neurons/group, Ctrl versus TTX p = 0.9968, Ctrl versus Bic p < 0.0001, one-way ANOVA followed by Tukey's post hoc test. The nuclei were counterstained with DAPI (blue). C, D, Representative confocal images of phosphorylated CREB (p-CREB; green) and MAP2 (red) immunofluorescence (C) and quantification of p-CREB immunofluorescence intensity (D) measured in cortical neurons treated with control media, TTX, or Bic; n = 48 neurons/group, Ctrl versus TTX p = 0.8107, Ctrl versus Bic p < 0.0001, one-way ANOVA followed by Tukey's post hoc test. The nuclei were counterstained with DAPI (blue). E, F, Representative confocal images of LC3 (green) and MAP2 (red) immunofluorescence (E) and quantification of LC3 puncta (F) measured in cortical neurons treated with control media, TTX, A-CREB (a dominant-negative form of CREB), or both TTX and A-CREB; n = 19–30 neurons/group, Ctrl versus TTX p < 0.0001, A-CREB versus TTX+A-CREB p < 0.0001, one-way ANOVA followed by Tukey's post hoc test. LC3 puncta are indicated by white arrowheads in E. G, H, Representative confocal images of LC3 (green) and MAP2 immunofluorescence (G) and quantification of LC3 puncta (H) measured in cortical neurons expressing the scrambled shRNA (Scr) or CRTC1 shRNA (shCRTC1) and treated with control media or TTX for 24 h; n = 24–38 neurons/group, Ctrl versus TTX p < 0.0001, shCRTC1 versus shCRTC1+TTX p < 0.0001, one-way ANOVA followed by Tukey's post hoc test. LC3 puncta are indicated by white arrowheads in G. I, J, Representative Western blottings of LC3 (I) and quantification of LC3-II levels (J) measured in cortical neurons treated with control media or TTX in the presence or absence of the CaMKK inhibitor STO-609 (3.5 μm); n = 6 experiments/group, Ctrl versus TTX p = 0.0462, STO-609 versus TTX+STO-609 p = 0.0345, one-way ANOVA followed by Tukey's post hoc test. *p < 0.05, ****p < 0.0001, and ns, not significant. Scale bar: 20 µm.

To address this question, we first focused on CaMKIV, as this kinase is present in the nucleus and has been implicated in regulating gene expression during synaptic homeostasis (Ibata et al., 2008). We found that STO-609, which is supposed to inhibit CaMKK and in turn block the activation of CaMKIV (Ibata et al., 2008; Ma et al., 2014; Tyssowski et al., 2018; Li et al., 2020), had no effect on either baseline LC3-II levels or the TTX-induced increase in LC3-II (Fig. 12I,J). Thus, although CaMKIV is likely to be important for synaptic up-scaling in different phases of enduring neuronal inactivation (Ibata et al., 2008; S. Kim and Ziff, 2014; Tyssowski et al., 2018; Li et al., 2020), it may not be key for triggering transcription-dependent autophagy.

We next looked at TFEB, a master regulator of the autophagy–lysosomal pathway that controls the gene network and is involved in various processes such as autophagy and lysosomal biogenesis (Settembre et al., 2011; Martini-Stoica et al., 2016). Given that starvation-induced dephosphorylation of mTOR drives the cytoplasm-to-nucleus translocation of TFEB (Peña-Llopis et al., 2011; Martina et al., 2012), and given that mTOR is dephosphorylated during synaptic up-scaling, we asked whether chronic neuronal inactivity induces TFEB-mediated cytoplasm-to-nucleus signaling to regulate transcription-dependent autophagy. To test this possibility, we measured the subcellular distribution of TFEB in cultured cortical neurons and found that TTX-treated cells had significantly higher levels of nuclear TFEB (Fig. 13A–E). Moreover, this inactivity-driven nuclear translocation requires inactivated (i.e., dephosphorylated) mTOR, as it was prevented by MHY1485 (Fig. 13A,B), a potent cell-permeable mTOR activator that causes mTOR phosphorylation at Ser-2448. Together, these results suggest that, similar to starvation-induced metabolic changes, chronic neuronal inactivity can also inactivate mTOR and recruit TFEB signaling.

Figure 13.

Cytonuclear signaling of TFEB and its important role in inactivity-induced autophagy gene expression. A, Immunostaining of TFEB in cortical neurons treated with control media or TTX in the presence or absence of MHY-1485 (MHY; 2 μm) for 24 h. The nuclei were counterstained with DAPI, and the dashed circles indicate the nucleus. Original pixel intensities from 0 to 255 are represented as a gradient lookup table. B, Quantification of the nuclear-to-cytosolic ratio of TFEB immunofluorescence intensity measured in the groups shown in A; n = 55–90 neurons/group, Ctrl versus TTX p = 0.0005, MHY versus TTX+MHY p = 0.5043, one-way ANOVA followed by Tukey's post hoc test. C–E, Representative Western blottings of TFEB (C) in nuclear and cytoplasmic fraction in cortical neurons treated with control media or TTX for 24 h and quantification (D, E); n = 3 experiments/group, p = 0.0105 for D, p = 0.0079 for E, unpaired t test. Nuclear and cytoplasmic fractions were normalized to H3 and GAPDH, respectively. F, At DIV9, cortical neurons were infected with a lentivirus expressing a scrambled shRNA (Scr) or TFEB shRNA (shTFEB), and TFEB mRNA levels were measured and are expressed relative to Scr; n = 4 experiments/group, p = 0.0015, unpaired t test. G, At DIV9, primary cortical neurons were infected with lentivirus expressing Scr shRNA or shTFEB. The normalized mRNA levels of the indicated genes were then measured after treatment with control media or TTX for 24 h, and are expressed relative to control levels; ULK1, n = 5 experiments/group, Scr versus Scr+TTX p = 0.0171, shTFEB versus shTFEB+TTX p = 0.8583; ATG5, n = 6 experiments/group, Scr versus Scr+TTX p = 0.0181, shTFEB versus shTFEB+TTX p = 0.7469; ATG7, n = 5–6 experiments/group, Scr versus Scr+TTX p = 0.0190, shTFEB versus shTFEB+TTX p = 0.6643; LC3B, n = 6 experiments/group, Scr versus Scr+TTX p = 0.0067, shTFEB versus shTFEB+TTX p = 0.9609; ATG12, n = 6 experiments/group, Scr versus Scr+TTX p = 0.0014, shTFEB versus shTFEB+TTX p = 0.4067; one-way ANOVA followed by Tukey's post hoc test. H–J, Normalized mRNA levels of the indicated genes measured in cortical neurons expressing Scr or shTFEB and treated with control media or TTX for 24 h, expressed relative to Scr; n = 6 experiments/group; H, Scr versus Scr+TTX p = 0.0031, shTFEB versus shTFEB+TTX p = 0.7212; I, Scr versus Scr+TTX p = 0.0462, shTFEB versus shTFEB+TTX p = 0.9967; J, Scr versus Scr+TTX p = 0.0226, shTFEB versus shTFEB+TTX p = 0.4046; one-way ANOVA followed by Tukey's post hoc test. Scale bar: 20 µm.

Finally, to examine the putative role of TFEB in regulating the expression of autophagy-related genes during synaptic up-scaling, we used shRNA to knock down TFEB expression in cultured neurons (Fig. 13F) and found that knocking down TFEB prevented the inactivity-induced upregulation of genes involved in autophagy flux, including genes implicated in regulating the autophagy–lysosomal pathway (Fig. 13G–J). To further show that TFEB plays a key role in autophagy flux, we measured the formation of LC3 puncta in cultured neurons and found that knocking down TFEB prevented the TTX-induced increase in LC3 puncta (Fig. 14A,B). With respect to synaptic up-scaling, we also found that knocking down TFEB prevented the TTX-induced increase in sGluA1 (Fig. 14C,D), suggesting that TFEB, serving as an intermediary between chronic neuronal inactivity and transcription-dependent autophagy, is critical for synaptic up-scaling (Fig. 14E).

Figure 14.

TFEB is required for transcription-dependent autophagy and synaptic up-scaling. A, Immunostaining of the autophagy marker LC3 (green) and MAP2 (red) in cortical neurons infected with lentivirus expressing Scr or shTFEB and treated with control media or TTX for 24 h. LC3 puncta are indicated by arrowheads. B, Quantification of LC3 puncta measured in the groups shown in A; n = 40–50 neurons/group, Scr versus Scr+TTX p = 0.0016, shTFEB versus shTFEB+TTX p = 0.9705, one-way ANOVA followed by Tukey's post hoc test. C, Immunostaining of sGluA1 in cortical neurons infected with lentivirus expressing Scr or shTFEB and treated with control media or TTX for 24 h. Pictures are shown using glow scale (the side bar represents GluA1 staining intensity in arbitrary units). Representative dendritic segments with sGluA1 immunostaining are shown in the insets. D, Quantification of sGluA1 immunofluorescence intensity measured in the groups shown in C; n = 41–48 neurons/group, Scr versus Scr+TTX p < 0.0001, shTFEB versus shTFEB+TTX p = 0.9659, one-way ANOVA followed by Tukey's post hoc test. E, Proposed model of inactivity-induced transcription-dependent autophagy during synaptic up-scaling. Chronic inhibition of neuronal activity (24-h treatment with TTX) leads to mTOR dephosphorylation and recruits cytonuclear TFEB signaling to induce transcription-dependent autophagy, which controls the expression of synaptic proteins such as αCaMKII and therefore plays a key role in synaptic up-scaling. **p < 0.01, ****p < 0.0001, and ns, not significant. Scale bars: 20 µm (A) and 50 µm (B; insets, 10 µm).

Discussion

Temporal and spatial dependence of mTOR signaling during different forms of synaptic scaling

The role of mTOR signaling in various forms of synaptic plasticity may depend on how neuronal activity is manipulated both temporally and spatially, as shown previously for functional plasticity (Hoeffer and Klann, 2010) and structural plasticity (Tavazoie et al., 2005; Tang et al., 2014). For example, mTOR is the key kinase in initiating translation and is involved in translation-dependent synaptic scaling in response to briefly blocking AMPARs, but not NMDARs (Henry et al., 2012). Briefly blocking AMPARs activates synaptic mTOR via the intracellular synthesis of phosphatidic acid through phospholipase-D signaling (Henry et al., 2018), which triggers translation to support rapid synaptic up-scaling. Chronic inhibition of neuronal firing inhibits global calcium signaling (Ibata et al., 2008), which inactivates the MAPK pathway and downstream mTOR signaling (Bateup et al., 2013), which in turn engages transcription-dependent autophagy, thus driving relatively slow synaptic up-scaling.

Cytonuclear signaling, gene regulation and protein turnover in synaptic homeostasis

Following chronic neuronal inactivity, βCaMKK in cortical neurons undergoes nuclear translocation for CaMKIV activation, which drives the nuclear export of the splicing factor Nova-2 and in turn homeostatically increases action potential duration by causing a specific change in alternative splicing (Li et al., 2020). It has also been suggested that up-scaling caused by a relatively brief period of inactivity (4 h TTX) could be mimicked by CaMKIV inhibition (Ibata et al., 2008). Thus, although CaMKIV is likely to be important for regulating activity-dependent events in the nucleus, its functional role may be complicated and even distinct in different phases of scaling following enduring changes of neuronal activity (Ibata et al., 2008; S. Kim and Ziff, 2014; Tyssowski et al., 2018; Li et al., 2020). Our finding that chronic inactivity induces TFEB nuclear translocation and transcription-dependent autophagy provides new insights into the mechanism underlying inactivity-driven nuclear signaling during synaptic homeostasis. Alternative splicing and protein degradation allow chronic neuronal inactivity to simultaneously initiate both protein synthesis and protein turnover, with the flexibility needed to control each process separately using specific forms of cytonuclear signaling machinery, thus providing specific forms of neuronal adaption ranging from changes in action potential duration to changes in synaptic strength.

Studies have shown that the balance between protein synthesis and protein degradation is critical for maintaining synaptic plasticity (Fonseca et al., 2006; Dong et al., 2008, 2014; Srinivasan et al., 2021). Given that protein synthesis changes significantly after synaptic scaling is established (Majdan and Shatz, 2006; Tropea et al., 2006; Aoto et al., 2008; Ibata et al., 2008; Goold and Nicoll, 2010; Penney et al., 2012; Meadows et al., 2015; Schanzenbächer et al., 2016; Steinmetz et al., 2016; Schaukowitch et al., 2017; Garay et al., 2020), we speculate that protein degradation pathways such as autophagy also must be upregulated to counterbalance protein synthesis and maintain synaptic homeostasis, in addition to its role in inducing synaptic scaling. In this regard, the substrates of autophagy may be the newly synthesized plasticity-related proteins (e.g., αCaMKII and PSD95); however, the list of these substrates during neuronal homeostatic maintenance could be expanded to include other autophagy substrates, including plasticity-related proteins targeted by autophagy such as PICK1 and SHANK3 (Shehata et al., 2012; Nikoletopoulou et al., 2017; Glatigny et al., 2019; Pandey et al., 2020; Shen et al., 2020; Compans et al., 2021; Pan et al., 2021; Kallergi et al., 2022) or even organelles such as mitochondria via a process known as mitophagy (Palikaras and Tavernarakis, 2020).

Clinical implications with respect to ASD and other neuropsychiatric conditions

It is generally well-established that autophagy dysregulation plays a key role in diseases involving neuronal degeneration, including Alzheimer's disease (Yue et al., 2009; Rubinsztein et al., 2011; Nixon, 2013). Interestingly, recent studies indicate that autophagy may also play an important role in neuropsychiatric conditions, particularly ASD (Tang et al., 2014). For example, mTOR signaling is overactivated in tuberous sclerosis complex, a relatively rare genetic disorder with a high prevalence of ASD. Importantly, the defects induced by overactivated mTOR, including ASD-like social behaviors and postsynaptic spine pruning, can be rescued in vivo by inducing autophagy (Tang et al., 2014). Moreover, many well-known mechanisms that underlie ASD appear to converge on the inactivity-induced autophagy pathway, including: (1) an autoregulatory feedback loop mediated by dysregulated synaptic scaling (Ramocki and Zoghbi, 2008; Toro et al., 2010; Mullins et al., 2016); (2) changes in activity-dependent gene expression (Ebert and Greenberg, 2013); and (3) excessive mTOR signaling (Hoeffer and Klann, 2010; Tang et al., 2014). In this respect, our findings suggest a new mechanistic link between normal brain physiology and the pathophysiology present in ASD. In addition, lithium, a widely used mood stabilizer, that has also been proposed to exert its antimanic effect via synaptic scaling (Kavalali and Monteggia, 2020). Interestingly, lithium is also a well-known inducer of autophagy (Sarkar et al., 2005), indicating that regulating autophagy during synaptic homeostasis may have important in vivo implications in these brain disorders. Furthermore, given that transcription-dependent autophagy is particularly vulnerable to activity-dependent gene expression and mTOR signaling, both of which are believed to play a causal role in ASD (Hoeffer and Klann, 2010; Ebert and Greenberg, 2013; Tang et al., 2014), it is reasonable to speculate that autophagy is important for homeostatic regulation and may also underlie various neuropsychiatric conditions.

Footnotes

This work was supported by the National Key R&D Program of China Grant 2019YFA0508603 (to H.M.); the Science and Technology Innovation 2030-Major Project Grant 2021ZD0203501 (to H.M.); National Natural Science Foundation of China Grant Numbers 81930030, 82230036, 31771109, and 31722023 (to H.M.), 32200788 (to X.H.), and 31900696 (to W.L.); the Project for Hangzhou Medical Disciplines of Excellence; the Key Project for Hangzhou Medical Disciplines; and the CAMS Innovation Fund for Medical Sciences Grant 2019-I2M-5-057 (to H.M.). We would like to thank Dr. Zhen-Ge Luo for providing the GFP-LC3 mice, Dr. Wei Liu for providing the pEGFP-LC3 plasmid, and the Core Facilities of Zhejiang University School of Medicine for technical assistance. We also thank Dr. Lu Chen for insightful suggestions regarding this study. We are also grateful to Min Zhao, Ruiyun Zhang, and all the other members of the Ma laboratory for technical assistance, helpful discussion, and meaningful suggestions.
The authors declare no competing financial interests.
Correspondence should be addressed to Huan Ma at mah{at}zju.edu.cn

SfN exclusive license.

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