Abstract
Synaptic vesicle (SV) recycling is essential for the maintenance of neurotransmission, with a number of neurodevelopmental disorders linked to defects in this process. Fragile X syndrome (FXS) results from a loss of fragile X mental retardation protein (FMRP) encoded by the FMR1 gene. Hyperexcitability of neuronal circuits is a key feature of FXS, therefore we investigated whether SV recycling was affected by the absence of FMRP during increased neuronal activity. We revealed that primary neuronal cultures from male Fmr1 knock-out (KO) rats display a specific defect in activity-dependent bulk endocytosis (ADBE). ADBE is dominant during intense neuronal activity, and this defect resulted in an inability of Fmr1 KO neurons to sustain SV recycling during trains of high-frequency stimulation. Using a molecular replacement strategy, we also revealed that a human FMRP mutant that cannot bind BK channels failed to correct ADBE dysfunction in KO neurons, however this dysfunction was corrected by BK channel agonists. Therefore, FMRP performs a key role in sustaining neurotransmitter release via selective control of ADBE, suggesting intervention via this endocytosis mode may correct the hyperexcitability observed in FXS.
SIGNIFICANCE STATEMENT Loss of fragile X mental retardation protein (FMRP) results in fragile X syndrome (FXS), however whether its loss has a direct role in neurotransmitter release remains a matter of debate. We demonstrate that neurons lacking FMRP display a specific defect in a mechanism that sustains neurotransmitter release during intense neuronal firing, called activity-dependent bulk endocytosis (ADBE). This discovery provides key insights into mechanisms of brain communication that occur because of loss of FMRP function. Importantly it also reveals ADBE as a potential therapeutic target to correct the circuit hyperexcitability observed in FXS.
Introduction
Information communication between neurons requires the synchronous fusion of neurotransmitter-containing synaptic vesicles (SVs) during neuronal activity. SVs that are mobilized by action potentials (APs) are termed the recycling pool, which can be subdivided into the readily releasable pool (RRP; comprising primed fusion-ready SVs) and the reserve pool (SVs only mobilized during high neuronal activity; Alabi and Tsien, 2012). Several SV endocytosis modes are sequentially recruited by increasing stimulus intensity to replenish these SV pools (Chanaday et al., 2019). Ultrafast endocytosis is dominant during sparse neuronal activity (Watanabe et al., 2013), whereas clathrin-mediated endocytosis becomes prevalent during AP trains (Granseth et al., 2006). Both ultrafast and clathrin-mediated endocytosis saturate during heightened neuronal activity (López-Murcia et al., 2014; Soykan et al., 2017), and under these conditions, activity-dependent bulk endocytosis (ADBE) is the dominant endocytosis mode (Clayton et al., 2008). ADBE generates large endosomes directly from the plasma membrane, from which SVs are formed to replenish the reserve pool (Richards et al., 2000; Cheung et al., 2010).
The translation repressor fragile X mental retardation protein (FMRP) is located in several cellular compartments including central nerve terminals, where it controls the expression of a cohort of presynaptic proteins (Darnell and Klann, 2013). It also has protein synthesis-independent presynaptic functions, such as mediating ion channel gating, activation and density (reviewed in Ferron, 2016). For example, FMRP interacts with Slack potassium channels (Brown et al., 2010), N-type calcium channels (Ferron et al., 2014) and large conductance voltage and calcium-gated big potassium (BK) channels (Deng et al., 2013; Deng and Klyachko, 2016; Kshatri et al., 2020). The absence of FMRP at CA3-CA1 hippocampal synapses leads to reduced BK channel activity and excessive AP broadening (Deng et al., 2013; Wang et al., 2014; Deng and Klyachko, 2016), resulting in increased presynaptic calcium influx during high-frequency stimulation (Deng et al., 2011, 2013). These synapses also display altered short-term plasticity, but only during periods of heightened activity (Deng et al., 2011; Klemmer et al., 2011). Therefore, presynaptic phenotypes that occur in the absence of FMRP are only revealed during intense neuronal activity (Deng et al., 2011, 2013; Ferron et al., 2014).
Hyperexcitability of neuronal circuits is a key feature of fragile X syndrome (FXS; Booker et al., 2019; Das Sharma et al., 2020), one of the most common monogenic causes of intellectual disability (ID) and autism spectrum disorder (Mefford et al., 2012). Most FXS cases are caused by a CGG trinucleotide expansion in the 5′ untranslated region of the FMR1 gene which encodes FMRP, leading to hypermethylation of the promoter and silencing of the FMR1 gene. Additionally, a point mutation which disrupts polyribosome binding (I304N), is sufficient to cause FXS (Feng et al., 1997), whereas a different mutation in FMRP that is unable to bind to and regulate BK channels (R138Q) has been identified in different individuals that display ID and FXS phenotypes (Collins et al., 2010; Myrick et al., 2015; Diaz et al., 2018; Sitzmann et al., 2018).
Since FMRP appears to be required for optimal presynaptic function specifically during intense neuronal activity, we determined whether SV recycling was disproportionately impacted under these conditions in primary neuronal cultures derived from an Fmr1 knock-out (KO) rat model (Asiminas et al., 2019). No significant defect in either SV exocytosis or endocytosis was observed, however Fmr1 KO neurons displayed a robust defect in ADBE. This dysfunction reduced presynaptic performance during periods of intense neuronal activity. Finally, molecular replacement studies with human FMRP mutants revealed that the ADBE defect was because of loss of BK channel interactions and that BK channel activators could restore normal function in Fmr1 KO neurons.
Materials and Methods
Materials
Unless otherwise specified, all cell culture reagents were obtained from Invitrogen. Fetal bovine serum was from BioSera. Papain was obtained from Worthington Biochemical. All other reagents were obtained from Sigma-Aldrich unless specified. Rabbit anti-SV2A was obtained from Abcam (ab32942 RRID: AB_778192). Guinea-pig anti-vGLUT (135304 RID:AB_887878) and rabbit anti-vGAT (274102 RRID:AB_2620000) were obtained from SYSY. Anti-rabbit Alexa Fluor 488 (A11008 RRID: AB_143165), anti-guinea pig Alexa Fluor 568 (A11075 RRID:AB_2534119), and donkey anti-rabbit Alexa Fluor 647 (A32795 RRID:AB_2866496) were obtained from Invitrogen. The mCerN1 (empty vector) was constructed as previously described (Cheung et al., 2010). Synaptophysin-pHluorin (sypHy) was provided by Prof. L. Lagnado (University of Sussex, UK). pGP-CMV-GCamP6f was a gift from Douglas Kim and the GENIE project (Addgene plasmid #40755; RRID:Addgene_40755; Chen et al., 2013). Synaptophysin-mOrange2 was from Prof. Shigeo Takamori (Doshisha University, Koyto, Japan). hEGFP-FMRP isoform 1, was obtained from Dr. B. Bardoni (Institut National de la Santé et de la Recherche Médicale, Institute Pharmacology Moléculaire Et Cellulaire Centre National de la Recherche Scientifique; Khayachi et al., 2018). hEGFP-FMRPR138Q and hEGFP-FMRPI304N were generated using site-targeted mutagenesis (R138Q forward primer GTGCCAGAAGACTTACAGCAAATGTGTGCCAAA; reverse primer TTTGGCACACATTTGCTGTAAGTCTTCTGGCAC: I304N forward primer AAAAATGGAAAGCTGAATCAGGAGATTGTGGAC; reverse GTCCACAATCTCCTGATTCAGCTTTCCATTTTT (mutated bases underlined). The base change was confirmed by Source Bioscience Sanger Sequencing.
Primary neuronal culture
Animal work was performed in accordance with the UK Animal (Scientific Procedures) Act 1986, under Project and Personal License authority and was approved by the Animal Welfare and Ethical Review Body at the University of Edinburgh (Home Office project license 7008878). Specifically, all animals were killed by schedule 1 procedures in accordance with United Kingdom Home Office Guidelines; adults were killed by exposure to CO2 followed by decapitation, whereas embryos were killed by decapitation followed by destruction of the brain.
Heterozygous LE-Fmr1em1/PWC female rats (Asiminas et al., 2019) were mated with wild-type (WT) males to produce either Fmr1 KO or WT males. Male embryos were taken at e18.5–e19.5 for hippocampal dissection. Hippocampi from each embryo were processed separately to avoid contamination across genotypes. Dissociated primary hippocampal cultures were prepared from embryos as previously described (Zhang et al., 2015). Briefly, isolated hippocampi were digested in a 10 U/ml papain solution (Worthington Biochemical, LK003178) at 37°C for 20 min. The papain was then neutralized using DMEM F12 (ThermoFisher Scientific, 21331-020) supplemented with 10% fetal bovine serum (BioSera, S1810-500) and 1% penicillin/streptomycin (ThermoFisher Scientific, 15140-122). Cells were triturated to form a single cell suspension and plated at 5 × 104 cells (with the exception of single cell tetramethylrhodamine (TMR)-dextran uptake experiments, 2.5 × 104 cells) per coverslip on laminin (10 µg/ml; Sigma-Aldrich, L2020) and poly-D-lysine (Sigma-Aldrich, P7886)-coated 25-mm glass coverslips (VWR International Ltd). Cultures were maintained in Neurobasal media (ThermoFisher Scientific, 21103-049) supplemented with 2% B-27 (ThermoFisher Scientific, 17504-044), 0.5 mm L-glutamine (ThermoFisher Scientific, 25030-024), and 1% penicillin/streptomycin. After 2–3 d in vitro (DIV), 1 μm cytosine arabinofuranoside (Sigma-Aldrich, C1768) was added to each well to inhibit glial proliferation. Hippocampal neurons were transfected with sypHy or co-transfected with both GCamP6f and synaptophysin-mOrange2 at DIV7 using Lipofectamine 2000 (ThermoFisher Scientific, 11668027) as per manufacturer's instructions and imaged at DIV13–DIV15. For single-cell TMR-dextran experiments, neurons were transfected with mCerN1 or hEGFP-FMRP constructs using Lipofectamine 2000 3 d before imaging at DIV13–DIV15.
SypHy and GCaMP6f imaging
SypHy-transfected neurons were visualized at 500-nm band pass excitation with a 515-nm dichroic filter and a long-pass >520-nm emission filter on a Zeiss Axio Observer D1 inverted epifluorescence microscope. Images were captured using an AxioCam 506 mono camera (Zeiss) with a Zeiss EC Plan Neofluar 40×/1.30 oil immersion objective. Image acquisition was performed using Zen Pro software (Zeiss). Hippocampal cultures were mounted in a Warner Instruments imaging chamber with embedded parallel platinum wires (RC-21BRFS) and challenged with field stimulation using a Digitimer LTD MultiStim system-D330 stimulator (current output 100 mA, current width 1 ms) either at 10 Hz for 30 s, 40 Hz for 10 s, or 4 × 40 Hz for 10 s with 5-min intervals between trains. Imaging time courses were acquired at 4-s intervals while undergoing constant perfusion with imaging buffer [119 mm NaCl, 2.5 mm KCl, 2 mm CaCl2, 2 mm MgCl2, 25 mm HEPES, and 30 mm glucose at pH 7.4, supplemented with 10 μm 6-cyano-7-nitroquinoxaline-2,3-dione (Abcam, ab120271) and 50 μm DL-2-amino-5-phosphonopentanoic acid (Abcam, ab120044)]. Alkaline buffer (50 mm NH4Cl substituted for 50 mm NaCl) was used to reveal the maximal pHluorin response.
The size of the different functional SV pools in hippocampal neurons was measured by stimulating sypHy-transfected neurons for increasing durations in the presence of 1 μm bafilomycin A1 (Cayman Chemical Company, 11038). The RRP was mobilized by 40 APs (20 Hz), and 10 s later, the remainder of the recycling pool mobilized with a second challenge of 1600 APs (20 Hz). The resting pool was revealed by application of NH4Cl buffer.
Hippocampal neurons co-transfected with GCamP6f and synaptophysin-mOrange2 were mounted on a Zeiss Axio Observer D1 microscope as described above and were visualized at 500-nm band pass excitation with a 515 nm dichroic filter and a long-pass >520 nm. An image of transfected neurons was acquired before 40 Hz (10 s) stimulation using 556/25 nm excitation and 630/98 nm emission bandpass filters (Zeiss) to view synaptophysin-mOrange2 marking of nerve terminals. Imaging time courses were acquired at 2-s intervals while undergoing constant perfusion with imaging buffer.
Time traces were analyzed using the FIJI distribution of ImageJ (National Institutes of Health; NIH). Images were aligned using the Rigid body model of the StackReg plugin (https://imagej.net/StackReg). Nerve terminal fluorescence was measured using the Time Series Analyser plugin (https://imagej.nih.gov/ij/plugins/time-series.html). Regions of interest (ROIs) five pixels in diameter were placed over nerve terminals that responded to the electrical stimulus. The proportion of ROIs that were responsive to stimulation was not different between genotypes (for sypHy experiments: WT 28.6 ± 5.4; KO 26.1 ± 6.2% responsive synapses; for GCamP6f experiments: WT 92.0 ± 5.2%; KO 85.9 ± 7.3% of synaptophysin-mOrange2 puncta respond to the field potential stimulus). A response trace was calculated for each cell by averaging the individual traces from each selected ROI ΔF/F0. Baseline fluorescence was subtracted from GCamP6f time traces to correct for high background fluorescence. For sypHy time traces, fluorescence decay time constants (tau, τ, s) were calculated by fitting a monoexponential decay curve to data from the time point after the end of electrical stimulation.
TMR-dextran uptake
TMR-dextran (ThermoFisher Scientific, D1842) uptake was performed as described previously (Nicholson-Fish et al., 2015). Neurons were mounted on a Zeiss Axio Observer D1 microscope as described above before challenging with 400 APs (40 Hz) in the presence of 50 μm of TMR-dextran (40,000 MW) in imaging buffer. Where the experiment was performed with BK channel modulators, neurons were incubated in 50 μm of TMR-dextran and either DMSO (Sigma-Aldrich, D8418), 10 μm NS 11021 (Bio-Techne Ltd, 4788/10), or 10 μm Paxilline (Bio-Techne Ltd, 2006/10) for 120 s before stimulation. The TMR-dextran solution was immediately washed away after stimulation terminated, and images were acquired using 556/25-nm excitation and 630/98-nm emission bandpass filters (Zeiss) while undergoing constant perfusion. Per coverslip of cells, three to six different fields of view were imaged. The TMR-dextran puncta in each image were quantified using the Analyze Particles plugin of ImageJ (NIH; https://imagej.nih.gov/ij/developer/api/ij/ij/plugin/filter/ParticleAnalyzer.html) to select and count particles of 0.23–0.91 µm2.
Where TMR-dextran uptake was performed on transfected cultures, WT and KO neurons were transfected 3 d before imaging with mCer-N1 plus either hEGFP-FMRPWT, hEGFP-FMRPI304N or hEGFP-FMRPR138Q or mCer-N1 alone. Transfected axons were visualized on DIV13–DIV15 at both 430- and 500-nm excitation (long-pass emission filter >520 nm) to ensure co-transfection. Images of transfected neurons and of TMR-dextran were acquired using 470/27- and 556/25-nm double band pass filters and emission filters 512/30 and 630/98 nm, respectively (Zeiss). Per coverslip of cells, two to eight neurons were imaged. Axon length was calculated using the Simple Neurite Tracer plugin of ImageJ (NIH; https://imagej.net/SNT). TMR-dextran puncta (0.23–0.91 µm2) were counted along transfected axons with the final value normalized for axon length. For all experiments, for each condition, at least one unstimulated coverslip was imaged to correct for the background level of TMR-dextran uptake.
Immunofluorescence staining
Immunofluorescence staining was performed as previously described (Nicholson-Fish et al., 2015). Briefly, hippocampal neurons were fixed with 4% paraformaldehyde (Sigma-Aldrich, 47608) in PBS for 15 min. Excess paraformaldehyde was quenched with 50 mm NH4Cl in PBS. Cells were then permeabilized in 1% bovine serum albumin (BSA; Roche Diagnostics GmbH, 10735078001) in PBS-Triton X-100 0.1% solution for 5 min and blocked in 1% BSA in PBS at room temperature for 1 h. After blocking, cells were incubated in rabbit anti-SV2A (1:200 dilution) for 1 h, after which the cultures were washed with PBS and incubated with fluorescently conjugated secondary antibodies (anti-rabbit Alexa Fluor 488; 1:1000 dilution) for 1 h. The coverslips were mounted on slides for imaging with FluorSave (Millipore, 345789). SV2A puncta were visualized at 500-nm band pass excitation with a 515-nm dichroic filter and a long-pass >520-nm emission filter on a Zeiss Axio Observer D1 inverted epifluorescence microscope. Images were captured using an AxioCam 506 mono camera (Zeiss) with a Zeiss EC Plan Neofluar 40×/1.30 oil immersion objective. SV2A puncta in each image were quantified using the Analyze Particles plugin of ImageJ to select and count particles of 0.23–3.18 µm2.
Fluorescein (FITC)-dextran uptake
Hippocampal neurons were exposed to FITC-dextran (40,000 MW, 50 μm) in imaging buffer during challenge with 400 APs (40 Hz) in an identical manner to TMR-dextran uptake. Neurons underwent an identical immunofluorescence staining protocol to that described above, with the exception that neurons were incubated with guinea pig anti-vGLUT and rabbit anti-vGAT primary antibodies (both at 1:1000 dilution) and goat anti-guinea pig Alexa Fluor 568 and donkey anti-rabbit Alexa Fluor 647 (1:1000 dilution). Fluorescent neurons were visualized using an AxioObserver Z1 inverted epifluorescence microscope and an AxioCam 506 mono camera. Excitation was provided by a Colibri 7 Solid-State Light Source using 475-, 555-, and 630-nm wavelengths, respectively. Emission was monitored using the following bandpass filters 525/50, 605/70, and 690/50 nm.
Per coverslip of cells, three to nine different fields of view were imaged and FIJI was used for image processing. The background was subtracted from all images using the Rolling Ball algorithm (https://imagej.net/plugins/rolling-ball-background-subtraction). Images with vGLUT and vGAT staining were auto-thresholded using Otsu's algorithm (Otsu, 1979). Masks were then generated of particles sized 5–150 px, representing antibody staining at nerve terminals. For FITC images, the threshold was set at 5, 255 and masks with particles 3–19 px were generated, representing internalized FITC-dextran. To determine the overlap between masks, a combined image was created using image calculator (NIH) and overlapping particles were counted.
Horseradish peroxidase (HRP) uptake
Hippocampal cultures were mounted in the RC-21BRFS stimulation chamber and challenged with 400 APs (40 Hz) in the presence of 10 mg/ml HRP (Sigma-Aldrich, P8250) supplemented imaging buffer. Immediately following the end of stimulation, cultures were washed in imaging buffer to remove noninternalized HRP and fixed with a solution of 2% glutaraldehyde (Electron Microscopy Sciences, 16019) in PBS. After washing in 0.1 m Tris buffer, HRP was developed with 0.1% 3,3'-diaminobenzidine (Fluka Chemica, 22204001) and 0.2% v/v hydrogen peroxide (Honeywell, 216763) in Tris buffer. After further washing in Tris buffer, cultures were then stained with 1% osmium tetroxide (TAAB laboratory and microscopy, O015/1) for 30 min. Samples were then dehydrated using an ethanol series and polypropylene oxide (Electron Microscopy Sciences, 20411) and embedded using Durcupan resin (Sigma-Aldrich, 44610). Samples were sectioned, mounted on grids, and viewed using an FEI Tecnai 12 transmission electron microscope. Intracellular structures that were <61 nm in diameter were arbitrarily designated to be SVs, whereas larger structures were considered endosomes. The area of individual endosomes was obtained by tracing the circumference using the freehand selections tool in ImageJ and measuring the resulting area. Typically, 30 fields of view were acquired for one coverslip of cells. In nerve terminals that contained HRP, the average number of HRP-labeled endosomes and SVs per nerve terminal was calculated for each coverslip and represents the experimental n.
Experimental design and statistical analysis
Microsoft Excel (Microsoft) and Prism 6 software (GraphPad Software Inc.) were used for data processing and analysis. The experimenter was blinded to genotype during the acquisition and to both the genotype and drug treatment or transfection condition during data analysis. For all figures, results are presented with error bars as ±SEM, and the n for each condition represents the number of coverslips imaged. For all assays, cells were obtained from at least two cultures, each containing at least three independent preparations from individual embryos. In sypHy assays, at least 10 ROIs were collected from each coverslip. The number of ROIs examined was comparable for all experiments. Normality was determined using a D'Agostino and Pearson omnibus normality test. For comparison between genotypes, an unpaired Student's t test was performed where data followed a normal distribution, with a Mann–Whitney test performed for those that did not. For comparison between more than two conditions, a one-way ANOVA was performed, whereas a two-way ANOVA was performed for comparison between genotypes across multiple stimulation trains. For both one-way and two-way ANOVA tests, results were corrected for multiple comparisons.
Results
Fmr1 KO hippocampal neurons show no obvious defect in SV recycling
FMRP is proposed to control SV fusion (Ferron et al., 2014), presynaptic AP duration (Deng et al., 2011), and short-term synaptic plasticity (Deng et al., 2011; Klemmer et al., 2011) in different murine models. To determine whether SV exocytosis or endocytosis were altered in a recently generated Fmr1 KO rat model (Asiminas et al., 2019), we examined SV recycling using sypHy in primary hippocampal cultures from KO or WT littermate controls. SypHy consists of the SV protein synaptophysin with a pH-sensitive EGFP (pHluorin) fused to an intraluminal loop (Granseth et al., 2006). Since sypHy reports the pH of its immediate environment, an exocytosis-dependent increase in its fluorescence signal is observed during SV fusion (Fig. 1A,B). After endocytosis, sypHy fluorescence is quenched by SV acidification. This loss of fluorescence is an estimate of the kinetics of SV endocytosis, since this is rate-limiting (Atluri and Ryan, 2006; Granseth et al., 2006). When WT neurons were challenged with a train of 300 APs delivered at 10 Hz, they displayed a characteristic sypHy response, with an evoked increase in fluorescence followed by an exponential poststimulation decrease to baseline (Fig. 1B). To determine the amount of exocytosis as a proportion of the total SV pool, fluorescence traces were normalized to the sypHy response in the presence of an alkaline solution (to reveal the maximal unquenched signal; Fig. 1B). No significant difference in this parameter was observed when WT and KO were compared (Fig. 1C). The lack of effect was confirmed using the vacuolar-type ATPase inhibitor bafilomycin A1 which allows SV exocytosis to be reported without confounding input from SV endocytosis (Sankaranarayanan and Ryan, 2001; Fig. 1E,F). The time constant, τ, of SV endocytosis was also not significantly different between WT and KO neurons (Fig. 1D). Therefore, loss of FMRP does not impair SV exocytosis or endocytosis during low-frequency stimulation.
Fmr1 KO hippocampal neurons show no obvious defect in SV recycling during low-frequency stimulation. A–D, Hippocampal neurons derived from either Fmr1 KO or WT littermate controls were transfected with sypHy on DIV7 and imaged DIV13–DIV15. Transfected neurons were challenged with a train of 300 APs (10 Hz) and left to recover for 3 min before exposure to an alkaline buffer (NH4Cl). A, Representative images of nerve terminals (indicated by arrows) transfected with sypHy at different points during this experiment (Rest, 10 Hz, Recovery, and NH4Cl). Scale bar: 5 µm. B, Mean trace displaying the average sypHy fluorescent response of WT (black) and KO (red) hippocampal neurons in response to stimulation ± SEM. Traces are ΔF/F0 and are presented as a fraction of the total SV pool, revealed by NH4Cl application. Bar indicates the period of stimulation. C, Mean sypHy peak height ± SEM during stimulation. D, Mean sypHy retrieval time constants (τ ± SEM) following stimulation. n = 10 for both WT and KO, unpaired Student's t test, C, p = 0.65; D, p = 0.21. E, F, Hippocampal neurons were transfected, imaged, and stimulated in an identical manner to above except neurons where exposed to 1 μm bafilomycin A1. E, Mean trace displays the average sypHy response to 300 APs delivered at 10 Hz (ΔF/F0 as a fraction of the total SV pool, revealed by NH4Cl). Bar indicates stimulation period. F, Mean sypHy peak height during 10-Hz stimulation ± SEM, n = 7 for both WT and KO, unpaired Student's t test, p = 0.31.
Previous studies examining loss of FMRP function revealed presynaptic defects during high-frequency stimulation (Deng et al., 2011; Ferron et al., 2014; Wang et al., 2014). Therefore, we next examined sypHy responses evoked by a train of 400 APs delivered at 40 Hz (Fig. 2). Under these stimulation conditions, there was no difference in either the extent of SV exocytosis, measured with or without bafilomycin A1 (Fig. 2B,D,E) or the kinetics of SV endocytosis (Fig. 2C). Taken together, this indicates that deletion of Fmr1 does not play a significant role in SV recycling during either low or high-frequency activity.
Fmr1 KO hippocampal neurons show no obvious defect in SV recycling during high-frequency stimulation. A–C, Hippocampal neurons derived from either Fmr1 KO or WT littermate controls were transfected with sypHy on DIV7 and imaged DIV13–DIV15. Transfected neurons were challenged with a train of 400 APs (40 Hz) and left to recover for 3 min before exposure to an alkaline buffer (NH4Cl). A, Mean trace displaying the average sypHy fluorescent response of WT (black) and KO (red) hippocampal neurons in response to stimulation ± SEM. Traces are ΔF/F0 and are presented as a fraction of the total SV pool, revealed by NH4Cl application. Bar indicates the period of stimulation. B, Mean sypHy peak height ± SEM during stimulation. C, Mean sypHy retrieval time constants (τ ± SEM) following stimulation. n = 13 for both WT and KO, Mann–Whitney test B, p = 0.26; unpaired Student's t test C, p = 0.84. D, E, Hippocampal neurons were transfected, imaged, and stimulated in an identical manner to above except neurons where exposed to 1 μm bafilomycin A1. D, Mean trace displays the average sypHy response to 400 APs delivered at 40 Hz (ΔF/F0 as a fraction of the total SV pool, revealed by NH4Cl). Bar indicates stimulation period. E, Mean sypHy peak height during 40-Hz stimulation ± SEM (n = 8 for both WT and KO, unpaired Student's t test, p = 0.38).
Previous studies that examined presynaptic morphology in Fmr1 KO cultures have produced contradictory observations on the number and density of SV in nerve terminals (Deng et al., 2011; Klemmer et al., 2011). To determine whether there was any alteration in the size of functional SV pools rather than SV numbers themselves, we inhibited SV acidification with bafilomycin A1 in sypHy-transfected neurons (Sankaranarayanan and Ryan, 2001; Fig. 3). In these experiments, the RRP was mobilized by 40 APs (20 Hz), and the remainder of the recycling pool by 1600 APs (20 Hz). The resting pool (which cannot be mobilized by APs) was revealed by application of an alkaline solution (Fig. 3A). In these experiments, we observed no difference in either the size of the total SV pool (Fig. 3B) or the RRP between genotypes (Fig. 3C). However, a small, but significant, increase in the SV recycling pool was observed in Fmr1 KO cultures (Fig. 3D).
Fmr1 KO hippocampal neurons have a larger recycling pool. Hippocampal neurons derived from either Fmr1 KO or WT littermate controls were transfected with sypHy on DIV7 and imaged DIV13–DIV15 in the presence of 1 μm bafilomycin A1. Transfected neurons were challenged with two trains of APs (20 Hz, 2 s and 20 Hz 80 s) before exposure to alkaline buffer (NH4Cl). A, Mean traces displaying the average sypHy fluorescent response of WT (black) and KO (red) hippocampal neurons is displayed ± SEM. Traces are ΔF/F0 and normalized to the total SV pool, revealed by NH4Cl application. Bars indicate periods of stimulation. B, Mean sypHy fluorescence during exposure to NH4Cl ± SEM. C, Mean sypHy peak height ± SEM during 20-Hz 2-s stimulation (RRP, % of total pool). D, Mean sypHy peak height ± SEM during 20-Hz 80-s stimulation (Recycling pool, % of total pool). n = 14 for WT and KO, unpaired Student's t test; B, p = 0.91; C, p = 0.79; D, *p = 0.0498.
Fmr1 KO hippocampal neurons display defective ADBE
Our experiments thus far have revealed that the absence of FMRP has little effect on either SV recycling or functional SV pool size. However, since presynaptic deficiencies in the absence of FMRP are only revealed during high-frequency stimulation (Deng et al., 2011; Ferron et al., 2014; Wang et al., 2014), we next assessed whether ADBE was affected in Fmr1 KO neurons. ADBE is the dominant mode of SV endocytosis during physiological patterns of activity in dissociated neuronal culture (Clayton et al., 2008) and intact neuronal circuits (Imig et al., 2020), and in response to environmental stimuli in vivo (Körber et al., 2012). Therefore altered ADBE may contribute to the activity-specific changes in neurotransmission previously observed in Fmr1 KO synapses.
ADBE was monitored optically using uptake of 40-kDa TMR-dextran, which is a fluid phase marker that is selectively accumulated via this endocytosis mode (Clayton et al., 2008). WT and Fmr1 KO neurons were challenged with a train of 400 APs (40 Hz) to maximally trigger ADBE in the presence of TMR-dextran (Clayton et al., 2008; Nicholson-Fish et al., 2015). The number of nerve terminals that performed ADBE were revealed as discrete TMR-dextran fluorescent puncta (Fig. 4A,B). There was a significant decrease in TMR-dextran puncta in Fmr1 KO neurons relative to WT littermate controls (Fig. 4C). Importantly, this phenotype was not because of synapse loss in the Fmr1 KO cultures, since there was no difference in nerve terminal numbers as measured by staining for the SV protein SV2A (Fig. 4D). The absence of synapse loss was confirmed by quantifying the number of excitatory and inhibitory nerve terminals in WT and Fmr1 KO cultures using antibodies against the glutamate and GABA transporters vGLUT and vGAT, respectively. When this analysis was performed, no significant difference in nerve terminal number was observed (Fig. 4E,F). Co-labeling with a fixable form of 40-kDa dextran (FITC-dextran) during a 400 AP train (40 Hz) confirmed the inhibition of ADBE (Fig. 4G), while the number of vGLUT and vGAT puncta positive for FITC-dextran was consistent with previous reports (Wenzel et al., 2012; vGLUT puncta with FITC-dextran, 4.62 ± 0.45%: vGAT puncta, 5.43 ± 0.46%, n = 17 WT, n = 13 KO). Taken together, these results reveal a small but robust decrease in the number of nerve terminals undergoing ADBE, suggesting that FMRP may be required for the optimal functioning of this mode of endocytosis.
Fmr1 KO hippocampal neurons show a defect in ADBE. A–C, Hippocampal neurons derived from either Fmr1 KO or WT littermate controls were challenged with a 40-Hz 10-s AP train in the presence of 50 μm TMR-dextran at DIV13–DIV15. A, B, Representative images of TMR-dextran-loaded nerve terminals of WT (A) and KO (B) hippocampal neurons. Scale bar: 30 µm. C, Mean TMR-dextran uptake is presented as a proportion of total WT uptake ± SEM, n = 14 for both WT and KO, unpaired Student's t test **p = 0.0054. D, Fmr1 KO neurons or WT littermate controls were fixed and stained with anti-SV2A as a marker of nerve terminals. Bar graph represents the mean number of SV2A-stained puncta for both KO and WT normalized to WT ± SEM, n = 14 for both WT and KO, unpaired Student's t test p = 0.54. E–G, Hippocampal neurons derived from either Fmr1 KO or WT littermate controls were challenged with a 40-Hz 10-s AP train in the presence of 50 μm FITC-dextran at DIV13–DIV15, fixed and then stained for either vGLUT or vGAT. Bar graph represents the mean number of vGLUT (E) or vGAT (F) puncta for both KO and WT (normalized to WT ± SEM, n = 17 for WT, n = 13 for KO, unpaired Student's t test vGLUT p = 0.739; vGAT p = 0.212). G, Mean FITC-dextran uptake as a proportion of total WT uptake ± SEM, n = 17 for WT, n = 13 for KO, unpaired Student's t test *p = 0.012. H–K, Hippocampal neurons derived from either Fmr1 KO or WT littermate controls were challenged with a 40-Hz 10-s AP train in the presence of 10 mg/ml HRP at DIV13–DIV15. H, I, Representative images of HRP-labeled endosomes and SVs in WT (H) and KO (I) nerve terminals. Black arrows in indicate HRP endosomes. Scale bar: 200 nm. J, Mean HRP-labeled endosome number per nerve terminal in WT and KO cultures ± SEM in nerve terminals that contain HRP-labeled endosomes. K, Mean HRP-labeled endosome areas in WT and KO ± SEM. WT n = 8; KO n = 9, unpaired Student's t test G, ***p = 0.0004; H, p = 0.67.
TMR-dextran uptake reports the number of nerve terminals undergoing ADBE, not the extent to which ADBE occurs in each nerve terminal (Clayton et al., 2008). Therefore, we applied the fluid phase marker HRP during AP stimulation (40 Hz 10 s) to confirm the ADBE defect in Fmr1 KO neurons (Fig. 4H,I). We observed a significant reduction in the number of HRP-labeled endosomes in Fmr1 KO neurons compared with WT in nerve terminals that contained HRP (Fig. 4J), whereas the endosome area across both genotypes was not significantly different (Fig. 4K). This reduction in the generation of bulk endosomes reveals that Fmr1 KO neurons display a specific defect in both the extent and prevalence of ADBE during intense neuronal activity.
ADBE is reduced at stimulus frequencies above 40 Hz (Clayton et al., 2008). This reduction may result from excess activity-dependent calcium influx, since this inhibits various forms of SV endocytosis in both large atypical and small central nerve terminals (von Gersdorff and Matthews, 1994; Cousin and Robinson, 2000; Wu and Wu, 2014; Leitz and Kavalali, 2016). Furthermore, it has been reported that Fmr1 KO neurons display enhanced calcium influx during AP stimulation because of either dysfunctional BK channel function or altered trafficking of voltage-gated calcium channels (Deng et al., 2013; Ferron et al., 2014, 2020). Therefore, to determine whether activity-dependent calcium influx may underlie the depression of ADBE in Fmr1 KO neurons, we monitored changes in presynaptic intracellular free calcium during trains of high-frequency AP stimulation. Primary cultures of both WT and Fmr1 KO neurons were co-transfected with both the genetically-encoded reporter GCamP6f and synaptophysin-mOrange2 (to mark nerve terminals) and were then challenged with a train of 400 APs (40 Hz). The extent of the evoked GCamP6f response at synaptophysin-mOrange2 puncta was not significantly different when WT and Fmr1 KO neurons were compared (Fig. 5). Therefore, the depression of ADBE observed in Fmr1 KO neurons was not a result of increased presynaptic calcium influx during intense neuronal activity.
Fmr1 KO hippocampal neurons do not display altered intracellular free calcium responses during high-frequency stimulation. Hippocampal neurons derived from either Fmr1 KO or WT littermate controls were co-transfected with GCamP6f and synaptophysin-mOrange2 (Syp-mOr2) on DIV7 and imaged DIV15. Transfected neurons were challenged with a train of 400 APs (40 Hz). A–C, Representative image of transfected neurons before (A, Rest), during (B, 40 Hz) and after (C, Recovery) stimulation. Scale bar: 1 µm. D, Mean trace displaying the average GCamP6f fluorescent response of WT (black) and KO (red) at Syp-mOr2 positive puncta in response to stimulation (F/F0 ± SEM). Bar indicates the period of stimulation. E, Mean peak GCamP6f fluorescence response ± SEM during stimulation, n = 6 for both WT and KO, unpaired Student's t test, p = 0.223.
Defective ADBE in Fmr1 KO neurons results in decreased presynaptic performance
Since ADBE is the dominant endocytosis mode during intense neuronal activity (Clayton et al., 2008), the impact of defective ADBE on SV exocytosis may only be revealed during patterns of high-frequency stimulation. To determine this, we monitored the amount of sypHy that visited the plasma membrane as a surrogate of neurotransmitter release in response to multiple trains of high-frequency stimulation (Fig. 6A). The prediction was that Fmr1 KO neurons would not be able to sustain performance to the same extent as WT, because of a lack of new SVs being provided via ADBE (Nicholson-Fish et al., 2015). Cultures were stimulated with four trains of high-frequency APs (40 Hz 10 s) separated by 5-min intervals. WT neurons displayed a sequential decrease in the extent of the evoked sypHy response, consistent with a depletion of SVs available for exocytosis. When the same protocol was performed for Fmr1 KO neurons, the sypHy response during the final stimulation was significantly lower when compared with WT (Fig. 6B). This suggests FMRP helps to sustain neurotransmitter release during increased neuronal activity via its control of ADBE.
The ADBE defect in Fmr1 KO neurons results in decreased presynaptic performance. Hippocampal neurons derived from either Fmr1 KO or WT littermate controls were transfected with sypHy on DIV7 and imaged DIV13–DIV15. Transfected neurons were stimulated four times with 40 Hz 10 s at 5-min intervals, before exposure to an alkaline buffer (NH4Cl). A, Mean traces displaying the average sypHy fluorescent response traces of WT (black) and KO (red) hippocampal neurons is displayed ± SEM. Traces are ΔF/F0 and normalized to the sypHy peak response to the first train of APs. Bars indicate the periods of stimulation. B, Mean sypHy peak heights for each 40-Hz 10-s stimulation, normalized to the first ± SEM WT n = 11 and KO n = 14, two-way ANOVA *p = 0.028, Bonferroni's multiple comparison test, Stim1 p > 0.99, Stim2 p > 0.99, Stim3 p = 0.15, Stim4 *p = 0.02.
BK channel activation corrects ADBE defects in Fmr1 KO neurons
To determine the mechanism underlying the control of ADBE by FMRP, we examined the ability of specific loss of function mutants to rescue TMR-dextran uptake in Fmr1 KO cultures. First, cultures from Fmr1 KO rats or WT littermate controls were transfected with either an empty fluorescent mCer vector or mCer plus EGFP-FMRP (FMRP) 3 d before imaging. Overexpression of FMRPWT did not affect evoked TMR-dextran uptake in WT neurons (Fig. 7A,B), indicating that excess FMRP has no detrimental effect on ADBE. It also signifies that additional FMRP does not enhance ADBE, as observed with constitutively active Rab11 mutants (Kokotos et al., 2018). Importantly, FMRPWT expression was sufficient to fully rescue the impairment in TMR-dextran uptake in Fmr1 KO neurons (Fig. 7B).
The R138Q FMRP mutant does not correct the ADBE defect in Fmr1 KO neurons. A, B, Hippocampal neurons derived from either Fmr1 KO or WT littermate controls were transfected with mCerulean (Emp) or mCerulean and GFP-FMRP (FMRP) 3 d before imaging. On DIV13–DIV15, neurons were challenged with a 40-Hz 10-s AP train in the presence of 50 μm TMR-dextran. A, Representative images of TMR-dextran uptake on axons transfected with empty or FMRP vectors, arrows indicate TMR-dextran puncta. Scale bar: 10 µm. B, Mean TMR-dextran uptake per 100 µm of transfected axon is presented normalized to WT control ± SEM WT n = 11; WT FMRP n = 7; KO n = 11; KO FMRP n = 10, one-way ANOVA with Bonferroni multiple comparison test comparing WT empty and KO empty *p = 0.043; and KO empty and KO FMRP *p = 0.048. C, D, Hippocampal neurons derived from Fmr1 KO embryos were transfected with mCerulean (Emp) or mCerulean and GFP-FMRP (WT), GFP-FMRPR138Q (R138Q), GFP-FMRPI304N (I304N) 3 d before imaging. On DIV13–DIV15, neurons were challenged with a 40-Hz 10-s AP train in the presence of 50 μm TMR-dextran. C, Representative images of TMR-dextran uptake on axons transfected with empty or FMRP vectors, arrows indicate TMR-dextran puncta. Scale bar: 10 µm. D, Mean TMR-dextran uptake per 100 µm of transfected axon is presented normalized to FMRPWT control ± SEM. Emp n = 8; WT n = 10; R138Q n =12; I304N n = 8, one-way ANOVA with Dunnett's multirole comparison test comparing all values to WT, empty *p = 0.043, R138Q *p = 0.038, I304N p = 0.79.
We exploited the ability of WT FMRP to fully rescue TMR-dextran uptake to perform molecular replacement studies using two loss of function mutants. These were: FMRPI304N, which does not bind ribosomes (De Boulle et al., 1993), and FMRPR138Q, which does not bind BK channels (Myrick et al., 2015; Kshatri et al., 2020; Fig. 7C). When these experiments were performed, FMRPWT restored TMR-dextran uptake in Fmr1 KO cultures as previously observed (Fig. 7D). TMR-dextran uptake in FMRPI304N expressing neurons was not significantly different to neurons expressing FMRPWT (Fig. 7D). This indicates that FMRP-dependent control of protein translation is dispensable for ADBE. Interestingly, expression of FMRPR138Q failed to restore TMR-dextran uptake in Fmr1 KO neurons, with levels equivalent to those observed with an empty vector (Fig. 7D). These results strongly suggest that FMRP controls ADBE through interactions with BK channels.
The absence of ADBE rescue by FMRPR138Q suggests that BK channel modulation by FMRP may be important for this endocytosis mode. Therefore, we investigated whether altering BK channel activity impacted ADBE. First, we determined whether enhancing channel activity with the activator NS 11021 could restore ADBE in Fmr1 KO neurons. BK channel activation with NS 11021 did not affect TMR-dextran uptake in WT neurons when compared with a vehicle control (Fig. 8A). It did however restore TMR-dextran uptake in Fmr1 KO neurons to WT levels (Fig. 8A). Therefore, activation of BK channels is sufficient to correct dysfunctional ADBE in Fmr1 KO neurons.
BK channel activation corrects ADBE defects in Fmr1 KO neurons. A, Hippocampal neurons from either Fmr1 KO or WT littermate controls were incubated with DMSO or 10 μm NS 11021 (NS), and 50 μm TMR-dextran for 2 min before a 40-Hz 10-s stimulus on DIV13–DIV15. Mean TMR-dextran uptake as a proportion of total WT uptake ± SEM is displayed. WT DMSO n = 11; WT NS n = 11; KO DMSO n = 10; KO NS n = 11, one-way ANOVA with Bonferroni's multiple comparison test comparing WT DMSO and KO DMSO **p = 0.0039; KO DMSO and KO NS *p = 0.032; WT DMSO and KO NS p > 0.9999; WT DMSO and WT NS p > 0.9999. B, WT hippocampal neurons were incubated with DMSO or 10 μm Paxilline (Pax) and 50 μm TMR-dextran for 2 min before a 40-Hz 10-s stimulus on DIV13–DIV15. Mean TMR-dextran uptake as a proportion of total WT uptake ± SEM is displayed. n = 13 for both DMSO and Pax, unpaired Student's t test p = 0.62.
To determine whether loss of BK channel function was responsible for defective ADBE in Fmr1 KO neurons, we attempted to mimic the deficit using the BK channel antagonist Paxilline in WT hippocampal cultures. Interestingly, this maneuver did not impact activity-dependent TMR-dextran uptake (Fig. 8B). Taken together, these results suggest that while BK channel dysfunction is not responsible for ADBE defects in Fmr1 KO neurons, BK channel activation can correct this fault.
Discussion
Loss of FMRP results in dysregulated translation of a specific cohort of mRNAs culminating in altered postsynaptic function (Darnell and Klann, 2013). However, FMRP also has noncanonical roles at the presynapse, particularly in the control of ion channel trafficking and activity. Disruption of these roles results in presynaptic dysfunction during intense neuronal activity (Deng et al., 2013; Wang et al., 2014; Ferron, 2016). We reveal that the absence of FMRP translates into a specific defect in ADBE, an endocytosis mode that is dominant during heightened neuronal activity. This defect results in reduced presynaptic performance during repetitive neuronal activity in Fmr1 KO neurons. Loss of BK channel interactions, but not channel activity, may be responsible for reduced ADBE, since FMRPR138Q failed to rescue function in KO neurons and BK channel inhibitors did not recapitulate the defect in WT neurons. Finally, the ability of BK channel activation to restore ADBE indicated that the defect may be an adaptation to hyperactivity in Fmr1 KO neurons.
Previous studies have produced contradictory observations on SV numbers within CA1 nerve terminals in Fmr1 KO mice in situ (Deng et al., 2011; Klemmer et al., 2011). Here, we provide a functional confirmation that the SV recycling pool is increased in Fmr1 KO neurons. From first principles, ADBE inhibition should limit recycling SV pool size, however a complex relationship exists between ADBE and SV pools. For example, ADBE inhibition upregulates single SV pathways (Clayton et al., 2009, 2010; Smillie et al., 2013). Furthermore, VAMP4 is required for ADBE, but its selective trafficking independently impacts release probability, providing additional functional regulation than just SV reformation (Ivanova et al., 2021). Therefore, the increase in SV recycling pool size may be an adaptation to reduced ADBE, which occurs in ∼40% of nerve terminals [vGLUT/vGAT labeling reveals 10% perform ADBE, whereas 25% of nerve terminals respond to a 40-Hz AP train: WT 28.9 ± 3.6; KO 24.6 ± 2.6 responsive synapses (n = 13 for both); Nicholson-Fish et al., 2016].
The increased recycling pool in Fmr1 KO neurons did not result in increased number of SVs fusing during an AP train during either low or high-frequency stimulation, confirming the lack of a fundamental role for FMRP in SV exocytosis. This agrees with previous studies, where neurotransmitter release in intact hippocampal circuits from Fmr1 KO mice was not different to WT after stimulation frequencies below 20 Hz (Deng et al., 2011). In contrast, studies in dorsal root ganglion neurons suggested that shRNA-induced depletion of FMRP enhanced SV fusion during high-frequency stimulation at 30°C (Ferron et al., 2014). Therefore, to confirm a lack of role for FMRP, we performed experiments at either physiological temperatures or by depleting FMRP with shRNA in hippocampal cultures. In both instances the absence of FMRP had no effect on SV exocytosis (K. Bonnycastle, unpublished observation). It is unclear why this disparity exists, however in primary cultures of rat hippocampal Fmr1 KO neurons we observe no obvious SV fusion phenotype.
We observed specific and selective effects on ADBE in Fmr1 KO neurons, which is the dominant form of endocytosis during high activity in both in vitro and in vivo (Clayton et al., 2008; Körber et al., 2012). TMR-dextran specifically reports ADBE because of size exclusion from single SVs (Clayton et al., 2008). Furthermore, ∼90% of TMR-dextran puncta co-localize with the vital dye FM1-43 after high-frequency AP trains, indicating that this fluid phase marker it is only accumulated by active central nerve terminals (Clayton and Cousin, 2009). This uptake accurately reflects ADBE, since interventions that selectively block this endocytosis mode abolish TMR-dextran uptake (Clayton et al., 2009, 2010; Smillie et al., 2013; Morton et al., 2015). These same interventions also arrest the activity-dependent formation of HRP-labeled endosomes (Clayton et al., 2009, 2010; Morton et al., 2015). HRP-labeled endosomes reflect newly formed bulk endosomes, since the number of unlabeled presynaptic endosomes does not change after stimulation, whereas additional HRP endosomes appear (Clayton et al., 2009, 2010; Smillie et al., 2013).
In this study, we observed that FMRP containing the I304N mutation fully restored TMR-dextran uptake in Fmr1 KO neurons, while the R138Q mutation (which fully supports protein translation; Myrick et al., 2015; Kshatri et al., 2020) did not. Therefore, the inability of R138Q FMRP to rescue defective ADBE in KO neurons supports the hypothesis that presynaptic defects are translation-independent (Deng and Klyachko, 2016). The failure of R138Q FMRP to restore ADBE in Fmr1 KO neurons could be because of the loss of BK channel regulation. BK channels control AP duration and shape, and in the absence of FMRP, Fmr1 KO synapses display enhanced calcium influx because of AP broadening (Deng et al., 2013). However, this does not appear to be the case, since inhibition of BK channels in WT neurons does not recapitulate the defect in ADBE. An intriguing alternative is that the FMRP-dependent control of ADBE is independent of the modulation of BK channel activity. In this scenario, BK channels act as a docking station to place FMRP in the correct location to facilitate ADBE. Without this interaction, FMRP is mislocalized and unable to perform its role. In support, FMRP and its interaction partners, CYFIP2 and NUFIP2 are enriched on bulk endosomes formed via ADBE (Kokotos et al., 2018).
Another alternative is that the R138Q mutation results in a loss of FMRP function that is independent of BK channels. The FMRP N terminus interacts with at least 76 different proteins in non-neuronal cells (Taha et al., 2021), including CYFIP1/2 and NUFIP1/2 (Bardoni et al., 1999, 2003; Schenck et al., 2001). While the cellular role of most of these proteins are still unclear, CYFIP1/2 are key components of the Wave Regulatory Complex required for actin remodeling (Chen et al., 2010; Cioni et al., 2018). Actin dynamics are essential for optimal ADBE (Soykan et al., 2017), suggesting an additional noncanonical action of FMRP, similar to the actin-organizing role in SV endocytosis recently reported for Kv3.3 channels (Wu et al., 2021) Interestingly, Cyfip1+/− neurons display an increased recycling SV pool in primary culture (Hsiao et al., 2016), similar to our observations with Fmr1 KO neurons in this study. Therefore, loss of FMRP function in ADBE may be because of disruption of parallel interactions within its N terminus.
The R138Q mutation is found in individuals presenting with ID and in some cases FXS phenotypes, seizures and autism (Collins et al., 2010; Myrick et al., 2015; Diaz et al., 2018; Sitzmann et al., 2018), although a direct link between the R138Q mutation and FXS has been questioned, since a cohort of individuals that display no discernable form of ID also carry this mutation (https://gnomad.broadinstitute.org). There may be explanations for this, however. For example, the cohort that carry the R138Q mutation may display a mild form of ID that was not detected in clinical assessments. Alternatively, other genetic modifiers may compensate for this mutation in these individuals, reducing its impact. Following this logic, this would mean that the R138Q FMRP mutation would have highly variable penetrance, dependent on the genetic composition of each individual subject. In support of a causal role, a recently generated knock-in mouse expressing the R138Q mutation displays cellular phenotypes and socio-cognitive defects similar to those observed in Fmr1 KO mice (Prieto et al., 2021).
The correction of ADBE by BK channel activation suggests that the most likely mechanism for the observed reduction in ADBE is a compensatory or homeostatic mechanism to counteract the increased excitability in FXS and Fmr1 KO models (Zhang et al., 2014; Booker et al., 2019; Das Sharma et al., 2020). In support, BK channel openers decrease hyperexcitability in Fmr1 KO models both in vitro and in vivo by increasing hyperpolarization, resulting in decreased calcium influx in nerve terminals (Hébert et al., 2014; Zhang et al., 2014). However, the correction of ADBE via BK channel opening is not mediated at the level of calcium influx, since we demonstrate no change in evoked intracellular free calcium responses in Fmr1 KO neurons.
A homeostatic reduction in ADBE in response to increased hyperexcitability would not be a homogenous adaptation across the brain, since neurons that display high firing rates would be disproportionately impacted. Therefore, even if reduced ADBE is not a causal mechanism in FXS, it may still be a valuable therapeutic intervention point to correct hyperexcitability in specific circuits (Booker et al., 2019; Das Sharma et al., 2020). Future studies are now required to determine whether modulation of ADBE can sculpt circuit activity in FXS (and other autism models that display hyperexcitability such as SynGAP haploinsufficiency disorder; Gamache et al., 2020). However, regardless of whether ADBE dysfunction contributes to FXS, the role of FMRP in the control of this key event provides valuable new information into the mechanisms of SV recycling that regulate presynaptic function.
Footnotes
This work was supported by grants from the Simons Foundation (529508), The RS McDonald fund, and a College of Medicine and Veterinary Medicine studentship. We thank Jennifer Darnell for expert advice and Steven Mitchell for excellent technical assistance.
The authors declare no competing financial interests.
- Correspondence should be addressed to Michael A. Cousin at m.cousin{at}ed.ac.uk
References
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