Baker–Gordon Syndrome-Associated Synaptotagmin-1 Mutations Reduce Synaptic Strength in Mouse Primary and Human-Induced Neuronal Culture Models

Abstract

Baker–Gordon syndrome (BAGOS) is a neurodevelopmental disorder (NDD) linked to a series of de novo mutations in the synaptic vesicle protein, Synaptotagmin-1 (SYT1). SYT1 is the major calcium sensor for synaptic transmission, and therefore a key molecule in neuronal communication. Several approaches have been used to reveal the underlying molecular mechanisms that lead to BAGOS pathology. While the murine genetic deletion, loss-of-function approach has proven valuable for modeling human diseases, human-induced pluripotent stem cells (hiPSCs) offer a powerful new strategy. In this study, we compare the phenotypes of BAGOS-associated SYT1 mutant variants in murine and human neuron models of either sex. In the well-established murine SYT1 knock-out (KO) model, we found that although all SYT1 mutant variants were correctly localized to the synaptic compartment, none could effectively rescue synaptic transmission. To examine the phenotype of BAGOS-associated SYT1 mutations in the context of human neurons, we generated a SYT1 KO hiPSC line via CRISPR/Cas9 gene editing and used this to derive neurons. As in mouse neurons, SYT1 KO in hiPSCs-derived human neurons strongly impairs synchronous release. Surprisingly, fast synaptic transmission could be rescued to varying extents in the human SYT1 KO model using BAGOS SYT1 mutants. However, overexpression of BAGOS SYT1 mutants in either WT mouse neurons or hiPSC-derived human neurons, a condition closer to the heterozygotic genotype of patients, revealed a dominant-negative effect of the mutant proteins. Our findings suggest that impaired neurotransmitter release efficacy caused by mutations in synaptic proteins may contribute to NDD pathophysiology.

Significance Statement

Synaptotagmin-1 (SYT1) plays crucial roles in synaptic transmission. Reports have linked mutations in the critical C2B Ca²⁺-binding loops of SYT1 to the neurodevelopmental disorder Baker–Gordon syndrome (BAGOS). We compare the physiological phenotypes of BAGOS-associated SYT1 mutant variants in SYT1 KO models using both mouse and human neurons. While we observe species-dependent variation in the functional impairments caused by SYT1 mutants in the KO models, we find that overexpression of BAGOS-associated variants in either WT mouse or human-induced stem cell-derived neurons consistently produces a dominant-negative effect on synaptic transmission. Our work strongly supports the notion that structure-function experiments in species-specific contexts to uncover how patient-associated mutations in synaptic proteins contribute to the underlying pathophysiology of neurological disorders.

Introduction

Synaptotagmin-1 (SYT1) plays a critical role in fast neurotransmission across species (DiAntonio and Schwarz, 1994; Jorgensen et al., 1995; Fernandez-Chacon et al., 2001). At nerve terminals, SYT1 is involved in synaptic vesicle (SV) docking (Jorgensen et al., 1995; Reist et al., 1998; de Wit et al., 2009; Chang et al., 2018; Chen et al., 2021), priming (Liu et al., 2009; Huson et al., 2020; Bouazza-Arostegui et al., 2022), Ca²⁺-triggered (Geppert et al., 1994; Fernandez-Chacon et al., 2001; Brunger et al., 2018) and spontaneous (Chicka et al., 2008; Xu et al., 2009; Bouazza-Arostegui et al., 2022) neurotransmitter (NT) release, as well as the retrieval of the SVs after fusion (Poskanzer et al., 2003; Yao et al., 2011). Extensive structure-function analysis of SYT1 has revealed that the highly evolutionarily conserved C2 cytosolic domains, namely, C2A and C2B, are indispensable for SYT1's synaptic functions (Fernandez et al., 2001; Fernandez-Chacon et al., 2001; Bacaj et al., 2013, 2015; Chang et al., 2018).

Numerous disorders are associated with mutations in genes encoding synaptic proteins (Claes et al., 2001; McTague et al., 2016; Alten et al., 2021; Bonnycastle et al., 2021). A series of heterozygous de novo mutations in the human SYT1 gene are linked to a neurodevelopmental disorder known as Baker–Gordon syndrome (BAGOS; Baker et al., 2015, 2018; Melland et al., 2022; van Boven et al., 2024). These disease-associated mutations are clustered within the Ca²⁺-binding loops of the C2B domain, a region that is critical for initiating Ca²⁺-triggered NT release (Bacaj et al., 2013). Individuals affected by this syndrome exhibit a range of symptoms including infantile hypotonia, profound intellectual disability, and movement disorders, with symptom severity varying widely between patients (Baker et al., 2018). The underlying reasons for this variability remain unclear.

Until now, most studies of SYT1 mutations have used mouse models, and no clear correlation between synaptic phenotypes of SYT1 BAGOS variants and patient symptoms has been identified (Baker et al., 2015, 2018; Bradberry et al., 2020; van Boven et al., 2024). Notably, work involving a subset of SYT1 BAGOS-associated variants overexpressed (OE) in cultured mouse neurons revealed the dominant-negative nature of the mutations on Ca²⁺-evoked synaptic transmission (Bradberry et al., 2020), thus shedding light on the autosomal dominance of the BAGOS mutations. Do BAGOS-associated SYT1 mutations impair neurotransmission through similar mechanisms in human synapses? Neurons differentiated from human-induced pluripotent stem cells (iPSCs) have proven an invaluable tool for examining disease mechanisms (Lee et al., 2009; Marchetto et al., 2010; Brennand et al., 2011; Pasca et al., 2011) and have been shown to replicate disease-associated phenotypes observed in primary mouse neurons (Chanda et al., 2013).

In our study, we compared the phenotype of BAGOS-associated SYT1 mutations in primary mouse neurons and iNs generated from human iPSCs. In the SYT1 KO mouse model, all mutant variants exhibited loss-of-function in synaptic transmission. To test SYT1 mutant variants in human synapses, we generated a SYT1 KO iPSC line using CRISPR/Cas9 gene editing. Genetic deletion of SYT1 in induced human neurons resulted in a drastic reduction in synchronous, calcium-triggered NT release, confirming that SYT1 is a critical calcium sensor for synaptic transmission in human synapses (Houcek et al., 2024). Expression of BAGOS-related SYT1 mutants in SYT1 KO human iNs (hiNs) resulted in less efficient synaptic transmission than rescue with WT human SYT1, but the phenotype was much milder than the full loss-of-function phenotype observed in the mouse SYT1 KO model. The severity of the SYT1 mutant phenotypes in the human KO model was graded and reflected the graded severity of patient phenotypes. To better evaluate the autosomal dominant nature of BAGOS-related SYT1 mutants, we also performed overexpression experiments in WT mouse neurons and human iNs. In both models, patient-specific variants showed a reduction in the efficacy of synaptic transmission, suggesting a dominant-negative phenotype of the calcium-binding site SYT1 mutants, as previously reported (Bradberry et al., 2020). Our research highlights the significance of species and genetic contexts in elucidating SYT1's impact on synaptic transmission.

Materials and Methods

All animal procedures were approved by the Animal Welfare Committee of Charité - Universitätsmedizin and the Berlin state government agency for Health and Social Services (license no. T 0220/09).

Induced pluripotent stem cells

Human-induced pluripotent stem cells (iPSCs) were provided by the Berlin Institute of Health Core Facility Stem Cells. The iPSC line BIHi005-A was derived from fibroblasts from human skin biopsy using Sendai virus vectors (Miltenyi mRNA, Miltenyi Biotec). The BIHi005-A iPSC line was maintained in feeder-free conditions using StemFlex medium (Thermo Fisher Scientific) in a standard incubator at 37°C and 5% CO₂.

Preparation of microdot patterned glass coverslips

Glass coverslips with a diameter of 30 mm (Glaswarenfabrik Karl Hecht) were first cleaned by shaking them overnight with 1 M HCl (Carl Roth), followed by rinsing with deionized water (ddH₂O) three times, and storage in 95% ethanol (Carl Roth). Prior to use, the coverslips were flamed to remove residual ethanol and then placed into 6-well plates (Corning). A thin layer of liquefied 0.15% agarose type IIa (Sigma-Aldrich) was then coated onto the coverslips, which creates a surface on which cells do not adhere well. Using a custom-built stamping machine, circular microislands of a growth solution containing collagen and poly-d-lysine (Gibco Life Technologies and Sigma-Aldrich) were applied onto the agarose-coated glass, resulting in uniform microislands with a diameter of ∼200 µm and a spacing of 500 µm between them. The plates with micropatterned glasses were then sterilized with ultraviolet light and stored at room temperature until use.

Preparation of astrocytes

Astrocytes were isolated from the cortices of newborn C57/BL6-N mice. The mice were killed, and their brains were placed in cold Hanks balanced salt solution (HBSS). The cortices were dissected, meninges removed, and then digested in 0.05% trypsin-EDTA for 20 min at 37°C. The tissue was triturated in DMEM medium (10% FCS and penicillin/streptomycin) to obtain a cell suspension. This suspension was divided into two T75 flasks with prewarmed DMEM medium and incubated at 37°C with 5% CO₂ for 1–2 weeks. Once 80% confluency was reached, astrocytes were detached using 0.05% trypsin-EDTA, stopped with DMEM medium, and centrifuged. The cells were then plated at 5 × 10³ cells per cm² on microdot-coated coverslips 1 week before adding human-induced neurons (hiNs) or primary murine neurons.

Primary murine neuronal culture preparation

Synaptotagmin-1 WT (SYT1+/+) and KO (SYT1−/−) mice in a C57BL/6 background were generated by interbreeding heterozygous mice as previously described (Xue et al., 2008). Embryonic (E18.5) SYT1-KO or C57BL/6 mice were decapitated, and hippocampi were dissected, placed in 500 µl NBA medium, and maintained at 37°C and 750 rpm until genotype confirmation. After confirming the genotypes, hippocampi were treated with 25 U/ml papain, bubbled with carbogen gas to adjust pH, for 45 min at 37°C. The digestion was stopped using a prewarmed inactivation solution. Neurons were then triturated in NBA medium and plated onto astrocyte feeder microislands. For WT only cultures, postnatal day 0–2 (P0–2) C57BL/6N mice were decapitated and hippocampi were dissected, treated, and triturated as described for SYT1-KO embryos.

Lentivirus

Lentiviruses were produced using HEK293T cells cotransfected with 3.9 µg of pRSV-REV (Addgene Plasmid #12253), 8.1 µg pMDLg/pRRE (Addgene Plasmid #12251) packaging plasmids, 6 µg of envelope plasmid VSVg (Addgene Plasmid #12259), and 12 µg of complementary DNA (cDNA) transfer plasmids containing the construct of interest (e.g., Tet-O-FUW or FUGW) in a T75 flask with calcium phosphate-mediated transfection. Prior to transfection, 6.8 × 10⁶ HEK cells were seeded in the T75 bottle and cultured in DMEM supplemented with 10% fetal calf serum (FCS). The plasmids were mixed with 111.6 µl of 2 M CaCl₂ and 768 µl of H₂O, added to 900 µl 2× HBS, and incubated for 20 min at room temperature. The transfection mix was added to the HEK cells and incubated at 37°C for 5 h. The media was then replaced with DMEM containing 10% FCS, and the cells were incubated at 37°C for 48 h. Lentivirus particles were collected from the supernatant and purified using Amicon Ultra-15 Centrifugal Filter units Ultracel-100K. The purified virus was stored at −80°C until use. Only virus batches with >90% infection efficiency, as assessed by green fluorescent reporter expression (FUW-eGFP) or by 90% survival cells by bright-field puromycin resistance (FUW-TetO-Ngn2-T2A-Puro), were used for experiments. cDNA constructs for rtTA (FUW-rtTA), NGN2 (FUW-TetO-Ngn2-T2A-Puro), and eGFP (FUW-eGFP) were provided by Thomas C. Südhof (Stanford University School of Medicine). Human SYT1 cDNA (NCBI accession number NM_005639) was synthesized at Eurofins Genomics and provided within a plasmid (pEX-A128 from Eurofins). Subsequently the studied human mutations (M303K, D304G, D366E, I368T, N371K) were introduced separately in the cDNA plasmid using the site-directed mutagenesis kit Quick change II (Agilent) and subcloned in lentiviral shuttle vectors in frame after a red fluorescent protein (RFP) fused to a P2A self-cleavage site. These RFP-P2A-SYT1 expression cassettes (WT and mutants) were controlled by the human Synapsin-1 promoter [f(syn)NLS-RFP-P2A-hsSYT1-WPRE].

Generation of iN cells from human iPSCs

Excitatory human iNs (hINs) were produced as described previously (Fenske et al., 2019). On Day −1, iPSCs were dissociated with Accutase cell dissociation reagent (Thermo Fisher Scientific) and reseeded on 6-well plates coated with Matrigel (Corning; 300,000 per well) in StemFlex Medium containing 2 µM Thiazovivin (Tocris) and lentiviral particles for transduction of Neurogenin-2 (NGN2), rtTA, and eGFP. On Day 0, the culture medium was changed to DMEM/F12 (DMEM+; Thermo Fisher Scientific), supplemented with N-2 (Thermo Fisher Scientific; 1:100), nonessential amino acids (Thermo Fisher Scientific; 1:100), human brain-derived neurotrophic factor (BDNF; 10 ng/ml, PeproTech), human NT-3 (10 ng/ml, PeproTech), mouse laminin (0.2 mg/L, Thermo Fisher Scientific), as well as doxycycline (2 µg/ml, Sigma-Aldrich) to induce TetO-dependent gene expression. Doxycycline was included in all media until the end of the experiments. On Day 1, iNs containing the NGN2 plasmid with puromycin resistance cassette were selected for via the addition of 0.5 µg/ml puromycin. On Day 3, the DMEM+ medium was replaced with NBA+ medium (Thermo Fisher Scientific), supplemented with B-27 (Thermo Fisher Scientific; 1:50), GlutaMAX (Thermo Fisher Scientific; 1:100), human BDNF (10 ng/ml), human NT-3 (10 ng/ml), mouse laminin (0.2 mg/L), cytosine β-d-arabinofuranoside (Ara-C; 2 µg/ml, Sigma-Aldrich), and doxycycline (2 µg/ml). Additionally, mouse astrocytes were added at the density of 3 × 10⁵ cells/cm². From Day 7 on, half of the medium in each well was replaced every 5 d by NBA+ medium containing B-27 supplement (1:50), GlutaMAX (1:100), doxycycline (2 µg/ml), and 2.5% fetal bovine serum (PAN-Biotech). After 50–70 d, cells were redissociated for plating on astrocyte microdots. Cells were washed twice with 0.5 mM EDTA/PBS (Thermo Fisher Scientific) and dissociated with Accutase Cell Dissociation Reagent (twice for 7 min at 37°C). GFP-positive cells were counted in a hemocytometer under a fluorescence microscope and seeded at a density of 4,000 per well on astrocyte feeder microislands to obtain induced human neuronal autaptic cultures. Cells on microislands were seeded in media containing NBA for human iN mass cultures supplemented with human BDNF (10 ng/ml), human NT-3 (10 ng/ml), and mouse laminin (0.2 mg/L). Autaptic hiN cultures were maintained in a humidified incubator (37°C and 5% CO₂) for 13–21 d.

Immunocytochemistry

Autaptic primary murine neurons were fixed in 4% paraformaldehyde (PFA, Sigma-Aldrich)/4% sucrose (Sigma-Aldrich) in PBS (Merck Millipore) for 10 min at room temperature (RT) between days in vitro (DIV) 14–16. The cells were washed three times with PBS and permeabilized in PBS-T (PBS with 0.1% Tween-20, Carl Roth GmbH + Co. KG) for 15 min. The cells were then incubated in a quenching solution (100 mM glycine in PBS) for 30 min and blocked with 5% normal donkey serum (Jackson ImmunoResearch) in PBS-T for 1 h at RT.

Primary antibodies were diluted in blocking solution and added to the autaptic neurons for overnight incubation at 4°C. The following primary antibodies were used: chicken anti-microtubule-associated protein 2 (MAP2) at a dilution of 1:2,000 (Merck Millipore, AB5543), mouse anti-synaptotagmin-1 at a dilution of 1:200 (Synaptic Systems, 105 011), and rabbit anti-vglut1 at a dilution of 1:4,000 (Synaptic Systems, 135 302).

After primary antibody incubation, neurons were washed three times with PBS-T, and secondary Alexa-Fluor 405, 555, or 647 antibodies at a dilution of 1:500 (Jackson ImmunoResearch) were added for 1 h at RT in PBS-T. Cells were washed twice with PBS-T and once with PBS before they were mounted on glass slides with Mowiol (Thermo Fisher Scientific). For quantitative assessment, all groups compared in one experiment were processed in parallel using identical antibodies solutions and other reagents.

Imaging and morphological analysis

For morphological analysis of mouse autaptic neuronal cultures, stacks of 16 bit fluorescence images were acquired on an Olympus IX81 inverted fluorescence microscope equipped with a 60× water immersion objective. Images were acquired with a CCD camera (Princeton MicroMax; Roper Scientific) using MetaMorph software (Molecular Devices). All images were analyzed using Fiji software—2.0.0-rc-65/1.52b National Institutes of Health. Fifty synapses per neuron were selected as regions of interest (ROIs), and hSYT1 fluorescence intensity signals were measured within the defined ROIs. The relative expression of hSYT1 was calculated by normalizing the measured intensities of hSYT1 to that of Syph. The data were normalized per culture to the hSYT1 WT rescue.

For each image stack, maximum intensity projections were generated. Uniform background subtraction and optimal threshold adjustment was performed on all images. Two independent cultures were imaged and analyzed per group for every experiment.

Western blot

For quantification of SYT1 protein levels (Figs. 1, 3, 4), mass cultures of primary mouse neurons or hiNs were lysed 14 d post SYT1 transduction by lentivirus infection at 4°C with lysis buffer [50 mM Tris, pH 8.0, 150 mM NaCl, 0.2% NP-40, protease inhibitor cocktail complete mini (Roche Diagnostics)]. Equal amounts of total protein from the lysates of cells transduced with different SYT1 constructs or empty vector control were separated on SDS polyacrylamide gel and transferred to a nitrocellulose membrane. Membranes were blocked for 1 h with 5% skim milk in PBS-T and incubated at 4°C overnight with primary antibodies in blocking solution: anti-SYT1 (105 011 Synaptic Systems) and anti-Synaptophysin-1 (101 004 Synaptic Systems, Sigma-Aldrich). Following three 10 min washes with PBS-T, the membranes were incubated with horseradish peroxidase-conjugated secondary antibodies (Jackson ImmunoResearch) in PSB-T for 1 h at RT. Detection was performed by using ECL Plus Western Blotting Detection Reagents (GE Healthcare Biosciences) in a Fusion FX7 detection system (Vilber Lourmat). Data were analyzed offline using ImageJ. The expression levels of the different SYT1 constructs were normalized over the levels of Synaptophysin-1.

Figure 1.

BAGOS-associated human SYT1 mutant variants are localized to synapses in mouse SYT1 KO neurons. A, An example Western blot against Synaptotagmin 1 (SYT1) and Synaptophysin 1 (SyPhy) from protein lysate of cultured SYT1 KO mouse neurons without rescue or transduced to express WT full-length mouse SYT1 (msSYT1-FL), WT full-length human SYT1 (hSYT1-FL), or human SYT1 variants carrying mutations M303K, D304G, D366E, I368T, or N371K. B, Quantification of SYT1 variant expression level over multiple Western blots. SYT1 band intensity was first normalized to SyPhy band intensity per blot. The SYT1/SyPhy signals were then normalized to that of the hSYT1-FL group. C, Example images of cultured SYT1 KO mouse neurons immunolabeled with antibodies against vesicular glutamate transporter 1 (VGLUT1), SYT1, and MAP2, with no rescue or expressing human SYT1 variants as indicated. D, Quantification of SYT1 intensity over VGLUT1 intensity over multiple fields of view in cultured SYT1 KO mouse neurons without rescue (red) or expressing human SYT1 variants as indicated by lentiviral transduction.

Figure 3.

SYT1 drives synchronous neurotransmitter release in human synapses. A, Scheme for targeting of guide RNAs to generate stop codons in exon 5 of the human SYT1 gene using CRISPR/Cas9 gene editing. B, Representative Western blot against SYT1 in human neurons derived from induced pluripotent stem cells (iPSCs; iNs) in isogenic control (WT) or clones with stop codons inserted into exon 5 of the gene. C, Example traces of EPSCs from glutamatergic neurons derived from iPSCs (hiNs) in isogenic control (WT, black) or two edited SYT1 KO clone lines (red, blue). Neurons were derived and split into autapses as previously described (Fenske et al., 2019). Stimulation artifacts have been blanked for illustrative purposes. D, E, Quantification of the EPSC amplitude (D) and charge (E; 1 s charge integration) in SYT1 KO hiNs and isogenic controls. Bars represent mean ± SEM.

Figure 4.

BAGOS-associated SYT1 mutant variants show graded rescue in SYT1 KO hiN correlated to disease severity. A, Representative western blot of protein lysate from SYT1 hiNs without rescue or expressing human SYT1 variants as indicated and immunolabeled with SYT1 and synaptophysin 1 (SyPhy) antibodies. B, Quantification of SYT1 band intensity divided by SyPhy band intensity in Western blots from cultured SYT1 KO hiNs without rescue or expressing human SYT1 variants as indicated. C, Examples traces of EPSCs recorded from SYT1 KO hiNs without rescue or expressing human SYT1 variants as indicated. D, E, Quantification of EPSC amplitude (D) and charge (E; 1 s charge integration) in glutamatergic SYT1 KO hiN autapses without rescue or expressing human SYT1 variants as indicated. F, Average cumulative charge illustrating the time course from hiN EPSCs represented in plots D and E in groups as indicated. G, Example traces of hypertonic sucrose-evoked currents in human hiN SYT1 KO autapses without rescue or expressing human SYT1 variants as indicated. H, Quantification of charge during transient component of sucrose response representing the readily releasable pool (RRP) of synaptic vesicles over many hiN SYT1 KO autapses in indicated rescue groups. I, Quantification of vesicular release probability in hiN SYT1 KO autapses with indicated SYT1 variant rescues by dividing the charge of the EPSC by the charge of the RRP per cell. Bar graphs represent the mean ± SEM.

Electrophysiology

Whole-cell patch-clamp recordings were performed in single mouse neurons between DIV 14 and 21 or single hiNs between day 63 and 91 postinduction at RT using a MultiClamp 700B amplifier (Molecular Devices) controlled by Clampex 10 software (Molecular Devices). Data were digitally sampled at 10 kHz and were filtered using a low-pass Bessel filter at 3 kHz. The series resistance was compensated by 70% and only cells with series resistances of <12 MΩ were analyzed. The recording chamber was constantly perfused with extracellular solution containing the following (in mM): 140 NaCl, 2.4 KCl, 10 HEPES, 2 CaCl₂, 4 MgCl₂, and 10 glucose (pH adjusted to 7.3 with NaOH, 300 mOsm, all Carl Roth). Borosilicate glass patch pipettes were pulled using a multistep puller (model P-1000, Sutter Instrument). Pipettes with resistance 3–5 MΩ were filled with KCl-based intracellular solution containing the following (in mM): 136 KCl, 17.8 HEPES, 1 EGTA, 4.6 MgCl₂, 4 ATP-Na₂, 0.3 GTP-Na₂, 12 creatine phosphate, and 50 U/ml phosphocreatine kinase (300 mOsm, pH 7.4, all Carl Roth). For whole-cell voltage-clamp recordings, neurons were maintained at −70 mV holding potential. Excitatory postsynaptic currents (EPSCs) were evoked by a 2 ms somatic depolarization from −70 to 0 mV producing an unclamped axonal action potential (AP). EPSC charge was measured by integrating the inward current for the 1 s period following stimulation. The paired-pulse ratio (PPR) was computed by dividing the second EPSC (EPSC2) amplitude by the first (EPSC1). To determine the size of the readily releasable pool (RRP) of synaptic vesicles, 500 mM sucrose (Sigma-Aldrich) solution was applied for 5 s, and the resulting transient inward current was integrated with the steady-state current set as the baseline (Rosenmund and Stevens, 1996). Vesicular release probability (Pvr) was calculated as the ratio between the charge of the EPSC and the evoked-sucrose charge.

Genetic modification by CRISPR/Cas9

A constitutive iPSC knock-out (KO) line for SYT1 was generated using a CRISPR/Cas9 ribonucleoprotein (RNP) complex. The complex was formed by mixing 2 µl of sgRNA (designed and validated for on-target specificity and minimal off-target effects) with 2 µl of HiFi Cas9 nuclease (IDT) and incubating the mixture for 20 min at RT. The sgRNAs used in the initial trials were ATTGCAATTAAGGCCCACGG, GCTATTGCAATTAAGGCCCA, GTTTCTTTCAGTGCCACCGT, and TGTTTCTTTCAGTGCCACCG. After assessing efficiency, the sgRNA ATTGCAATTAAGGCCCACGG was selected for final clone generation. Prior to nucleofection, 2 × 500,000 iPSCs were seeded per nucleofection. The sgRNA/Cas9 complex was then mixed with 82 µl P3 buffer, 18 µl Supplement 1 (Amaxa Primary Cell Kit P3), and 1 µl electroporation enhancer. iPSCs were detached using Accutase, centrifuged at 300 × g for 5 min, and resuspended in the nucleofection mix. Nucleofection was performed using the Amaxa 4D Nucleofector system with the CM-150 program. Immediately post-nucleofection, cells were seeded into a 6-well plate containing StemFlex medium supplemented with 2 µM Thiazovivin to enhance survival. The medium was replaced after 4 h.

Clonal selection, verification, and quality control

Single-cell clones were isolated through limited dilution and expanded. DNA was extracted from these clones to confirm the desired genetic modification. Clonality was verified using Amplicon NGS, while potential off-target effects were assessed via Sanger sequencing and Cas-OFFinder analysis.

Comprehensive quality control testing was conducted in collaboration with the BIH core facility for stem cells. This included assessments for viral pathogens, mycoplasma, bacteria, yeast, fungi, morphology, PluriTest, G-banding, virtual karyotyping using Illumina Infinium Global Screening Array-24 BeadChip, and Promega GenePrint 10 System. All clones successfully passed these quality controls.

Statistical analysis

We collected the same number of neurons (n) for each experimental group each day to reduce data variability. Data were collected from 3–5 independent human iN or primary mouse hippocampal autaptic cultures (N = 3–5) to account for potential culture-to-culture variability. For each replicate, independent hiNs cultures were generated via an individual induction process from each human iPSC line. Data from each experimental group were pooled except for those that were normalized to the mean value of the control group. All data were first subjected to Pearson’s omnibus K2 normality testing. Two-tailed unpaired t test or one-way ANOVA test for normally distributed data followed by Dunnett's multiple comparisons and Mann–Whitney test or Kruskal–Wallis ANOVA test for non-normally distributed data followed by Dunnett's post hoc multiple-comparison were performed using GraphPad Prism version 10.0 for Mac OS, GraphPad Software, www.graphpad.com. Significance and p values (*p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001) were calculated and are shown in the corresponding figures.

Results

Human SYT1 variants are properly expressed and localized in murine hippocampal neurons

Proper localization of the protein mutants to the synapse is required for structure-function experiments investigating the role of specific residues of synaptic proteins in the process of synaptic transmission. Previous assessment of human SYT1 mutant variants associated with the BAGOS suggested some isoforms (e.g., M303K) display abnormal trafficking and expression levels at synapses (Baker et al., 2015). Therefore, before exploring the physiological implications of these mutants, we investigated the expression and trafficking of human SYT1 mutant variants (M303K, D304G, D366E, I368T, and N371K) in a SYT1 knock-out (KO) mouse model (Fig. 1). Using Western blotting, we found that all constructs exhibited expression patterns comparable with full-length wild-type human SYT1 (hSYT1 FL) when reintroduced using lentiviral infection (Fig. 1A,B).

To further examine the expression levels and subcellular localization of the human SYT1 mutant variants, we performed immunocytochemistry on cultured mouse SYT1 KO hippocampal neurons. All SYT1 variants exhibited presynaptic localization (Fig. 1C) and were expressed at levels similar to hSYT1 FL in synaptic compartments (Fig. 1D). These results indicate that the expression and trafficking properties of all tested hSYT1 variants are comparable with the wild-type isoform.

BAGOS-related human SYT1 mutant do not rescue transmission in SYT1 KO murine synapses

The pathogenic mutations investigated here are located within the C2B top loops, which are required by SYT1 to perform its Ca²⁺-triggering function (Hui et al., 2009; Evans et al., 2015; Chang et al., 2018). Using mouse neurons lacking SYT1, we characterized the impact of the full panel of BAGOS-associated SYT1 mutants on synaptic transmission. We performed whole-cell voltage-clamp recordings of autaptic neurons cultured from the hippocampus of SYT1 KO mice expressing human WT full-length SYT1 (hSYT1 FL) or patient-associated mutant hSYT1 variants. The amplitudes of the glutamatergic EPSCs in neurons expressing mutant SYT1 were dramatically reduced compared with those expressing WT hSYT1 (Fig. 2A–C), suggesting a loss of Ca²⁺ triggering. This is consistent with previous study of a subset of BAGOS-associated mutants in inhibitory mouse SYT1 KO synapses (Bradberry et al., 2020) as well as mutations specifically targeting the Ca²⁺ binding region of SYT1 (Geppert et al., 1994; Fernandez et al., 2001; Fernandez-Chacon et al., 2001; Mackler et al., 2002; Xu et al., 2009). Additionally, further analysis revealed that more than half of the BAGOS-associated mutants demonstrated on average a smaller EPSC amplitude and charge than the SYT1 KO synapses (Fig. 2B,C), hinting at a dominant-negative phenotype in the remaining asynchronous release in SYT1 KO synapses, likely through an inhibition of the Ca²⁺-triggered change in SYT1 conformation, leaving the machinery in a primed state while preventing asynchronous release.

Figure 2.

BAGOS-associated human SYT1 variants show loss-of-function phenotype in cultured mouse SYT1 KO autapses. A, Example traces of excitatory postsynaptic currents (EPSCs) in mouse SYT1 KO glutamatergic autapses. SYT1 KO EPSC amplitude (red; no rescue) was compared with SYT1 KO neurons expressing human variants of Syt 1, including the full-length WT human SYT1 protein (black; hSYT1 FL) or BAGOS-associated human SYT1 mutant variants (as indicated). Stimulation artifacts were blanked for illustrative purposes. B, C, Quantification of EPSC amplitude (B) or charge (C; integrated over 1 s) in mouse SYT1 KO autapses without rescue or expressing human SYT1 variants as indicated. D, E, Average cumulative charge over the first second (D) or 100 ms (E) from EPSC onset for mouse SYT1 KO autapses without rescue or expressing human SYT1 protein variants as indicated. Sold line represents the mean with transparent extensions representing standard error of the mean. F, Example traces of current evoked by 5 s application of hypertonic sucrose solution (500 mM) by fast-flow perfusion system in mouse SYT1 KO autapses without rescue or expressing human SYT1 variants as indicated. G, Quantification of charge calculated by integrating the area of the transient component of the sucrose response (base-lined at the steady-state). H, Quantification of vesicular release probability (Pvr) calculated by dividing the charge of the EPSC by the charge of the sucrose response per cell in mouse SYT1 KO autapses without rescue or expressing human SYT1 variants as indicated. I, Example traces of paired-pulse stimuli in glutamatergic mouse SYT1 KO autapses without rescue or expressing human SYT1 variants as indicated. Stimulation artifacts have been blanked for illustrative purposes. J, Quantification of the paired-pulse ratio (PPR) calculated by dividing the second EPSC amplitude over the first EPSC amplitude. In all bar graphs, bars represent mean ± SEM.

Furthermore, we evaluated the time course of synaptic responses by integrating the EPSC charge for 1 s following stimulation (Fig. 2D,E). This analysis revealed that certain mutations showed a partial rescue of the fast synchronous component (M303K and D366E), albeit to a dramatically reduced extent compared with WT SYT1. Interestingly, these two variants show the mildest clinical phenotype within the patients (Baker et al., 2015, 2018; Bradberry et al., 2020). The remaining three mutants (D304G, I368T, and N371K) displayed no synchronous component and showed slower asynchronous component than even SYT1 KO synapses when compared in the first 100 ms (Fig. 2E). In addition to acting as the calcium sensor for synaptic transmission, SYT1 also plays a role in synaptic vesicle priming prior to Ca²⁺ triggering (Toulme et al., 2024). We tested the priming activity of the BAGOS-associated SYT1 mutants by evoking release of the readily releasable pool (RRP) of synaptic vesicles with a 5 s application of hypertonic sucrose solution (Rosenmund and Stevens, 1996; Fig. 2F). As expected, SYT1 KO neurons displayed a reduced RRP size (∼40%) when compared with neurons expressing the human SYT1 FL wild-type isoform (p = 0.0502; Kruskal–Wallis, multiple-comparison test vs KO; Fig. 2F,G). SYT1 mutant variants showed RRP sizes ranging from a significant rescue compared with SYT1 KO (∼40% increase from SYT1 KO; I368T) to a trend toward increase from SYT1 KO (∼30–40% increase, D366E, D304G) to no increase from SYT1 KO RRP at all (N371K; Fig. 2G). While the rescue of the RRP size from SYT1 KO with most of the SYT1-BAGOS variants is not strongly significant, it suggests that the SYT1-BAGOS associated mutants can support the precalcium, synaptic vesicle priming function of SYT1.

To investigate how this series of mutations might impact release probability, we calculated vesicular release probability (Pvr) for each group by dividing the charge of the EPSC by the sucrose-evoked charge (Rosenmund and Stevens, 1996). We found that none of the groups rescued the deficiency in the Pvr value displayed by the SYT1-lacking neurons (Fig. 2H). In fact, we saw a decrease in Pvr in all the SYT1 mutants tested (Fig. 2H), possibly suggesting a repression of the asynchronous component of release observed in SYT1 KO synapses. Furthermore, we performed paired-pulse experiments with two consecutive pulses given at 40 Hz (Fig. 2I). We found that while the human SYT1 FLWT isoform shifted the paired-pulse ratio (PPR; EPSC2/EPSC1) from facilitatory (EPSC2 > EPSC1) as displayed by the SYT1 KO neurons toward a depressive phenotype (EPSC2 < EPSC1), the SYT1 mutants all tended toward paired-pulse facilitation (Fig. 2I,J), a short-term plasticity behavior typically associated with lower release probability synapses.

Generation of an iPSC SYT1 KO line to study SYT1 function in human synapses

Expression of BAGOS-associated SYT1 mutant variants in the well-established murine SYT1 KO model system revealed proper expression and synaptic localization of all proteins but impaired calcium-triggered exocytosis. To investigate whether the function of human SYT1 mutant variants is species specific, we generated a constitutive SYT1 KO human iPSC line using CRISPR/Cas9 (Fig. 3). We targeted exon 5 of the SYT1 gene, adding stop codons to result in a severely truncated protein (Fig. 3A). We validated the resulting cell lines according to standard procedures to ensure clonality, absence of off-target effects, and lack of detectable abnormalities (see Materials and Methods). Two independently generated SYT1 KO iPSC lines were induced into glutamatergic neurons using the NGN2 protocol (Zhang et al., 2013) and later plated as autapses for electrophysiological characterization (Fenske et al., 2019). As observed in the murine SYT1 KO model, we saw a strong effect of eliminating SYT1 in neurons derived from the two independently generated iPSC lines compared with their respective isogenic controls (Fig. 3C–E). SYT1KO human-induced neurons (hiNs) displayed a significantly reduced EPSC amplitude (Fig. 3C,D) and displayed asynchronous release (Fig. 3C), as reported previously in the mouse model (Broadie et al., 1994; Geppert et al., 1994). The EPSC charge as integrated over 1 s was not significantly decreased (Fig. 3E). These results suggest that SYT1 is indeed a major synaptotagmin isoform in human synapses mediating regulated release and that our CRISPR/Cas9 generated SYT1 KO hiNs provide a valid platform to test the structure-function effects of BAGOS-associated SYT1 mutants in a human, species-relevant context. This result is consistent with a recent study in a human embryonic stem cell based SYT1 KO model (Houcek et al., 2024).

BAGOS-associated SYT1 mutants show milder deficits in a human SYT1 KO model

We next sought to characterize the synaptic properties of SYT1 KO human iN (hiNs) cells overexpressing three different SYT1 patient variants associated with three different levels of severity: mild (M303K), intermediate (D366E), and severe (N371K; Fig. 4). The expression levels of these three mutant variants in SYT1 KO hiNs were not significantly different from the group overexpressing the human SYT1 FL wild-type isoform when assessed by Western blot (Fig. 4A,B). Next, we performed electrophysiological recordings on autaptic SYT1 KO hiNs expressing human WT SYT1 (hSYT1 FL), BAGOS-associated mutants (M303K, D366E, N371K), or no exogenous SYT1 protein (null). Surprisingly, all three mutant variants showed a partial rescue of EPSC amplitude (Fig. 4C,D) and charge (Fig. 4E) and restored synchronous release (Fig. 4F). RRP size, as evoked by hypertonic sucrose, was not significantly different across the SYT1 KO and SYT1 variant rescue groups (Fig. 4G,H), suggesting that the hiN SYT1 KO model may not have the sensitivity to reveal the precalcium, priming function of SYT1 (Chang et al., 2018; Bouazza-Arostegui et al., 2022; Toulme et al., 2024). However, it should be noted that the RRP size measured in hiNs is ∼10-fold smaller than that in mouse neurons (Fig. 3). While this decreases the sensitivity of the measurement, as the steady-state component of the pool is more difficult to define, we are confident that major alterations of the pool would be detectable. Further analysis revealed that the mutation associated with the mildest disease phenotype, M303K, was able to restore the Pvr to similar values as the wild-type SYT1 variant (Fig. 4I). These results suggest that (1) species-specific context may play a role in the function of proteins at the synapse and (2) the hiN SYT1 KO model, allowed for the detection of a graded phenotype of BAGOS-related SYT1 mutant function that correlates with the phenotypic severity of patient disease (Baker et al., 2018).

The expression of the SYT1 variants impairs SYT1’s endogenous function during neurotransmitter release

Having characterized BAGOS-associated mutant protein function in SYT1 KO mouse and human synapses, we next aim to examine their effects in a more physiologically relevant context—in the presence of endogenous WT SYT1, mimicking the heterozygous patient genotype (Fig. 5). We first turned back to the well-established mouse system, by overexpressing human SYT1 FL and mutant variants in WT primary hippocampal autaptic neurons and performed electrophysiology (Fig. 5). WT synapses overexpressing human SYT1 FL exhibited a significant increase in both EPSC amplitude (Fig. 5A,B) and charge (Fig. 5C), likely due to the increased functional presence of SYT1 (Bouazza-Arostegui et al., 2022). In contrast, the neurons overexpressing each patient variant presented either a significant reduction or a trend toward a decreased EPSC amplitude or charge compared with WT mouse hippocampal synapses (Fig. 5A–C). Hypertonic sucrose-evoked RRP measurements revealed that overexpressing human full-length or BAGOS-associated SYT1 mutants in WT mouse hippocampal neurons has a mild effect on SV priming (Fig. 5D,E). The RRP size was significantly increased with overexpression of D304G and D366E and trended toward an increase with overexpression of FL SYT1 or M303K (p = 0.067, 0.061, respectively) compared with the RRP size in WT hippocampal neurons.

Figure 5.

BAGOS-associated SYT1 mutant variants show dominant-negative phenotypes with overexpression in WT mouse neurons. A, Representative traces of EPSCs recorded in glutamatergic autapses cultured from WT mice as control or with overexpression of indicated human SYT1 variants. Stimulation artifacts have been blanked for illustrative purposes. B, C, Quantification of the EPSC amplitude (B) and charge (C) when human SYT1 variants are overexpressed in WT mouse autapses. D, E, Representative traces of sucrose-evoked current (D) and quantification of the charge of the readily releasable synaptic vesicle pool (RRP; E) recorded from WT mouse autapses overexpressing indicated variants of human SYT1. F, Average vesicular release probability (Pvr) for glutamatergic WT mouse autaptic neurons overexpressing human SYT1 variants. Pvr is calculated by dividing the EPSC charge by the sucrose-evoked charge per autapse. G, H, Example traces (G) and quantification (H) of the paired-pulse ratio for WT mouse glutamatergic autpases expressing indicated human SYT1 variants. Stimulation artifacts have been blanked for illustrative purposes. All bar graphs represent the mean ± SEM.

Exploring the impact of human SYT1 mutants on release efficiency, we observed that the overexpression of any of the SYT1 disease variants led to a reduced Pvr (Fig. 5F) and more strongly facilitating responses during paired-pulsed stimulation (Fig. 5G,H). Taken together, these data show that BAGOS SYT1 variants, even when expressed alongside endogenous SYT1, impair synaptic strength—likely by competing with WT SYT1 or disrupting its function, consistent with previous reports of dominant-negative effects in mouse neurons (Bradberry et al., 2020).

Overexpression of BAGOS-associated SYT1 mutations decreases synaptic transmission in induced human neurons

Overexpression of SYT1 BAGOS-associated mutant variants in WT hippocampal mouse neurons resulted in a dominant-negative effect on synaptic transmission (Fig. 5). However, since we saw distinct phenotypes when expressing SYT1 mutant variants in mouse versus human SYT1 KO models (Figs. 2, 4, respectively), we wanted to investigate whether a species-specific effect of STY1 mutant overexpression was also present. Therefore, we overexpressed SYT1 patient variants in control hiNs and examined the properties of synaptic transmission (Fig. 6). This experimental paradigm is closer to the patient situation, in which synapses contain copies of both WT and mutant human SYT1. We saw that overexpression of most variants caused a significant decrease in EPSC amplitude (D304G, D366E, I368T). Interestingly, overexpression of M303K and N371K showed no significant decrease in EPSC amplitude compared with WT hiNs (p = 0.20, p = 0.96, respectively), and these mutants displayed the strongest and weakest rescue phenotypes, respectively, in the human SYT1 KO iN model (Fig. 4). In contrast to the mouse model (Fig. 5), the overexpression of the human SYT1 wild-type isoform did not lead to any enhancement of EPSC amplitude (Fig. 6B). A significant reduction in EPSC charge was only observed for I368T (Fig. 6C), although M303K, D304G, and D366E showed strong trends toward decreased charge (∼40–60%) without reaching significance (p = 0.11, 0.18, 0.07, respectively). These results suggest that the SYT1 BAGOS variants may act as dominant-negative suppressors of evoked Ca²⁺-triggered release even in human synapses that express WT SYT1.

Figure 6.

BAGOS-associated STY1 variants show dominant-negative suppression of release probability in wild-type human iNs. A–C, Examples traces (A) and quantification of average EPSC amplitudes (B) and charge (C, 1 s integration) for WT hiN autaptic glutamatergic neurons in control condition (gray), overexpressing (OE) SYT1 full-length WT (FL; black) or indicated mutant variants. Stimulation artifacts in A were blanked for illustrative purposes. D, E, Example traces of currents evoked by hypertonic sucrose solution (500 mM) in hiN autaptic neurons expressing human SYT1 variants. The average charge of sucrose-evoked currents were quantified across cells from groups illustrated in D and represented in E. F, Quantification of vesicular release probability (Pvr) per autapse in hiN neurons in control condition (gray) and expressing human SYT1 variants as indicated. Pvr was calculated as the EPSC charge divided by the charge generated by sucrose application. All bars graphs represent the mean ± SEM.

To further examine this, we quantified RRP size and Pvr in WT hiNs overexpressing SYT1 mutant variants. We found that overexpression of the mutants did not significantly affect the priming of SVs in hiNs, as indicated by the RRP size evoked by hypertonic sucrose (Fig. 6D,E). However, comparing the EPSC charge and RRP measurements, we calculated Pvr and found that overexpression of four of five mutants displayed a Pvr that was significantly decreased from WT hiNs (M303K, D304G, I368T, and N371K; Fig. 6F). D366 mutant showed a strong trend toward decreased Pvr (p = 0.06), reflecting a lower release probability. Taken together, these results indicate that in the context of a human synapse, with WT SYT1 present, the BAGOS-associated SYT1 mutants can decrease the efficacy of Ca²⁺-evoked neurotransmitter release, resulting in a reduction of synaptic strength.

Discussion

The SYT1 mutations causing Baker–Gordon Syndrome are autosomal dominant and distinct mutations are associated with a graded severity of disease in patients (Baker et al., 2015, 2018). Given that in the heterozygous mouse SYT1 KO model, synaptic transmission is largely preserved (Bouazza-Arostegui et al., 2022), it seems unlikely that simple loss-of-function of a single SYT1 allele would cause severe neurodevelopmental deficits. Therefore, to better understand the pathophysiology of BAGOS, we systematically studied the physiological phenotypes of BAGOS-associated SYT1 mutants in the absence (KO) and presence of WT SYT1 protein. In addition, we took species-specific context into account and compared how these human SYT1 protein mutant variants function in the long-established mouse neuron model and in human synapses. We found that in the mouse SYT1 KO model, the BAGOS-associated SYT1 mutants failed to rescue synaptic transmission, though they were all appropriately targeted to the synaptic compartment. In fact, all the mutants exhibited a slight inhibition of the remaining, asynchronous synaptic transmission in the mouse SYT1 KO synapses, hinting at a dominant-negative role for the mutants, consistent with work done with synonymous mutations in mouse SYT1 (Bradberry et al., 2020). Interestingly, in our human SYT1 KO model, generated by CRISPR/Cas9 in a human iPSC line, we observed a more potent rescue of synaptic transmission, but in a manner where rescue efficacy correlated to disease severity in patients (Baker et al., 2015, 2018). Nevertheless, overexpression of human SYT1 mutants in WT mouse or human neurons impaired synaptic transmission, indicating that these variants act via a dominant-negative mechanism.

SYT1's major role is to act as the Ca²⁺ sensor triggering SV exocytosis. As all the mutants associated with BAGOS tested in our study occur in the putative Ca²⁺ binding domain of SYT1, it follows that the physiological impact would involve a loss-of-function in calcium triggering. Indeed, in our rescue experiments in the mouse SYT1 KO model we saw that no BAGOS-associated human SYT1 mutant variant could rescue synaptic transmission (Fig. 2). One major caveat to this finding is that a mislocalization of mutant SYT1 could lead to a similar phenotype. For instance, previous research suggested that M303 variant has a trafficking defect (Baker et al., 2015). However, we found that in the mouse SYT1 KO model all SYT1 mutant variants were expressed at synapses at levels similar to WT SYT1 protein rescue. Another phenotype of the BAGOS-associated SYT1 mutants revealed by our rescue experiments in the mouse SYT1 KO neurons was the ability of specific mutants (M303K, D366E) to restore a component of synchronous neurotransmitter release, albeit at a small fraction of the magnitude of WT human SYT1 rescue. This was a similar result as reported previously for the D365E variant in inhibitory synaptic transmission (Bradberry et al., 2020). The phenotype of the remaining mutants (D304G, I368T, N371K) revealed not only a lack of synchronous release component, but in fact a further inhibition of the asynchronous release sustained in the SYT1 KO mouse neuron synapses in the first 100 ms following stimulation (Fig. 1A–E). Taken together, we speculate that while all these mutants affect the ability of Ca²⁺ to bind SYT1's C2B domain to trigger synchronous neurotransmitter release, they may do so with different efficacy—from a significantly decreased affinity to no binding capacity at all. The suppression of asynchronous release in the SYT1 KO mouse synapses is also consistent with previous studies revealing a dominant-negative phenotype of SYT1 Ca²⁺-binding mutants (Mackler et al., 2002; Zhou et al., 2015; Toulme et al., 2024). Overall, these experiments highlight the utility of fundamental structure-function experiments performed in established models, such as the mouse SYT1 KO, to better understand the mechanisms of pathophysiology in human diseases.

While performing structure-function experiments with BAGOS-associated human SYT1 mutant variants in mouse SYT1 KO synapses revealed important insight about the function of mutated residues, we could not be sure that the mutant proteins would function in the same manner in their native context of the human synapse. Therefore, it was crucial for us to extend our research to human synapses to investigate interspecies differences in SYT1 mutant function. To do so, we generated a SYT1 homozygous knock-out in human iPSCs using the CRISPR/Cas9 system and subsequently induced cells from this line into human SYT1 KO glutamatergic neurons. First, we show that knocking out SYT1 protein itself results in phenotypes consistent with the SYT1 KO mouse models, such as a reduction in phasic EPSCs and an increase in tonic release (Fig. 3C–E). Importantly, these data confirm that SYT1 is indeed the dominant isoform mediating synchronous synaptic transmission in human synapses. In the human SYT1 KO neuron model, we performed structure-function experiments with three representative mutants, M303K, D366E, and N371K—variants that show a graded severity in patients from least to most, respectively (Baker et al., 2015, 2018). We saw that these mutants could rescue the SYT1 KO phenotype in hiNs to a much greater extent than observed in the SYT1 KO mouse model (Fig. 4 vs Fig. 2). This could reflect a robustness in the synaptic vesicle fusion machinery in human synapses, guaranteeing at least some neurotransmitter release regardless of mutations in crucial proteins. However, it may also be due to a narrower window for successful neurotransmission in the immature hiN synapses, wherein the maximum amount of neurotransmitter release is reached even in substandard conditions and quickly plateaus. Additionally, it is important to note that a second synaptotagmin isoform, SYT7, is expressed at a higher level early in development and declines with age (Bouazza-Arostegui et al., 2022). Is a high level of SYT7 aiding the mutant SYT1 variants to rescue synaptic transmission in hiN synapses? While we have shown previously that SYT7 supports synaptic vesicle priming (Bouazza-Arostegui et al., 2022), we have more recently discovered that the presence of SYT7 inhibits calcium-triggered neurotransmitter release (Zobel et al., unpublished). Therefore, we argue that the presence of SYT7 at hiN synapses is unlikely to account for the rescue capability we observed in SYT1 KO neurons.

In addition to a general rescue of synaptic transmission by the select BAGOS mutants in the hiN SYT1 KO neurons, we also observed a graded rescue of synaptic function that correlated with the severity of disease observed in the patients with the different mutations (Baker et al., 2015, 2018; Fig. 4). Specifically, the M303K mutation, which is associated with a milder disease phenotype, rescued synaptic function with a greater efficacy than two mutations associated with more severe disease phenotypes. This graded rescue phenotype was not observed in our own work or other previous studies using the mouse synapse to investigate SYT1 mutant variant function (Bradberry et al., 2020). Therefore, our findings emphasize the need to consider species and genetic context in understanding the impact of mutations in synaptic proteins associated with human diseases.

BAGOS arises from heterozygous SYT1 mutations, meaning patient synapses also contain WT SYT1 protein. To understand how human SYT1 mutants function in an environment analogous to the heterozygous condition seen in patients (Baker et al., 2015, 2018), we overexpressed patient-specific variants alongside endogenous SYT1 in both mouse and human neurons. Surprisingly, even though rescue experiments with human SYT1 mutant variants differed in the mouse and human SYT1 KO models (Figs. 2, 4, respectively), overexpression of the SYT1 patient variants in the two WT models showed similar results (Figs. 5, 6). While in both models overexpression of SYT1 mutants trended toward a decrease in the amplitude of synaptic transmission (Figs. 5A,B, 6A,B), the most striking result was a decrease in the efficiency of synaptic transmission. In both mouse and human synapses, release probability was significantly decreased with overexpression of all mutants (Figs. 5F, 6F, respectively). This significant decrease in release probability was accompanied by a significant increase in paired-pulse facilitation (Figs. 5G,H, 6G,H). These results shed light on how BAGOS-associated SYT1 mutations may act in a dominant-negative manner even in the presence of WT SYT1 proteins in the case of patient synapses. We can also speculate that deviations in the plasticity characteristics and neurotransmitter release efficacy from the WT setpoint can have drastic consequences on neural network function and information processing. Therefore, it is still of high importance to understand the detailed mechanisms of how proteins function in the synapse. Grasping the intricate interplay between mutations and synaptic functions is essential for designing targeted therapeutic strategies for SYT1-related disorders.