Synaptotagmin 9 modulates spontaneous neurotransmitter release in striatal neurons by regulating substance P secretion

Synaptotagmin 9 (SYT9) is a tandem C2-domain Ca2+ sensor for exocytosis in neuroendocrine cells; its function in neurons remains unclear. Here, we show that, in mixed-sex cultures, SYT9 does not trigger rapid synaptic vesicle exocytosis in mouse cortical, hippocampal, or striatal neurons, unless it is massively overexpressed. In striatal neurons, loss of SYT9 reduced the frequency of spontaneous neurotransmitter release events (minis). We delved into the underlying mechanism and discovered that SYT9 was localized to dense core vesicles that contain substance P (SP). Loss of SYT9 impaired SP release, causing the observed decrease in mini frequency. This model is further supported by loss-of-function mutants. Namely, Ca2+ binding to the C2A domain of SYT9 triggered membrane fusion in vitro, and mutations that disrupted this activity abolished the ability of SYT9 to regulate both SP release and mini frequency. We conclude that SYT9 indirectly regulates synaptic transmission in striatal neurons by controlling SP release.


INTRODUCTION
The synaptotagmins (SYTs) are a large family of membrane trafficking proteins; seventeen isoforms have been identified in mammals, each encoded by a distinct gene (Craxton, 2010;Wolfes and Dean, 2020). In addition, most isoforms are thought to undergo alternative splicing, generating further potential diversity within this family (Craxton, 2010). Most This work is licensed under a Creative Commons Attribution 4.0 International License, which allows reusers to distribute, remix, adapt, and build upon the material in any medium or format, so long as attribution is given to the creator. The license allows for commercial use.
isoforms have a single N-terminal transmembrane domain (TMD) with a short intra-luminal domain. In all isoforms, the cytoplasmic domain largely comprises tandem C2-domains that are connected by a short linker. In at least eight of the seventeen isoforms, the C2-domains serve as Ca 2+ -dependent anionic lipid-binding motifs (Rickman et al., 2004;Bhalla et al., 2008). Ca 2+ binding is thought to be mediated by acidic side chains in two flexible loops that protrude from each C2-domain (Sutton et al., 1995). In the case of SYT1, Ca 2+ triggers the partial penetration of these loops into lipid bilayers and facilitates interactions with SNARE proteins (Chapman, 2008).
A current goal is to understand the functional diversity within the SYT family of proteins. SYT1 and SYT2 have been heavily studied and shown to serve as Ca 2+ sensors that trigger rapid, synchronous synaptic vesicle (SV) exocytosis in neurons (Geppert et al., 1994;Pang et al., 2006a). Importantly, SYT1 and SYT2 form a closely related clade with one more family member, SYT9 (Rickman et al., 2004). This homology prompted the idea that SYT9 might also function as a fast Ca 2+ sensor for SV exocytosis, which is a point we address in detail in the current study. For clarity, we note that SYT9 is sometimes referred to as SYT5, and vice versa; the SYT9 studied here comprises 386 residues (NCBI Gene accession # 53420) whereas SYT5 is 491 residues in length (NCBI Gene accession #60510).
Along with SYT1, SYT9 co-regulates catecholamine secretion from dense core vesicles (DCV) in PC12 cells (Fukuda et al., 2002;Lynch and Martin, 2007). SYT9 also regulates the release of follicle-stimulating hormone (FSH) from DCVs in the gonadotrophs of female mice in a sex-specific manner (Roper et al., 2015). However, while SYT9 is widely expressed in the brain (Mittelsteadt et al., 2009) (note: these authors referred to SYT9 as SYT5), its function in neurons remains controversial. One report concluded that SYT9 serves as a Ca 2+ sensor for rapid SV exocytosis in cultured striatal neurons (Xu et al., 2007). In contrast, SYT9 failed to support SV exocytosis in Syt1 KO hippocampal neurons, and pHluorin-tagged SYT9 underwent regulated exocytosis in both axons and dendrites of hippocampal neurons, with kinetics suggestive of DCV fusion (Dean et al., 2012).
To address and clarify the function of SYT9 in neurons, we studied synaptic transmission in cultured hippocampal, cortical, and striatal neurons obtained from Syt9 KO mice; for completeness Syt1 KO neurons were examined in parallel. We show that all three classes of neurons express both SYT9 and SYT1, but loss of SYT9 had no discernable effect on evoked neurotransmitter release, while loss of SYT1 disrupted rapid transmission in all three neuronal preparations. SYT9 was able to rescue the loss of synchronous release that is characteristic of Syt1 KO neurons, but only when over-expressed by ~25-fold, which matched the expression levels of endogenous SYT1. Interestingly, loss of SYT9 in cultured striatal neurons resulted in a specific and unexpected phenotype: a decrease in the spontaneous SV fusion rate (i.e., mini frequency). We then discovered that SYT9 was co-localized with substance P (SP), a DCV peptide hormone known to be enriched in striatal neurons. Moreover, loss of SYT9 impaired action potential-triggered SP release. Importantly, in patch-clamp experiments, supplementing the bath with this neuropeptide rescued the mini phenotype. Moreover, the structural elements of Syt9 that underlie Ca 2+ triggered membrane fusion in vitro also support evoked SP release and spontaneous SV fusion in living striatal neurons. These findings reveal a mechanism in which SYT9 indirectly regulates spontaneous transmission in striatal neurons by directly regulating secretion of the neuromodulator, SP.

Animals
All neuronal cultures were prepared from early postnatal (P0-P1) mice of either sex in accordance with all relevant regulations and the approval of the University of Wisconsin Institutional Animal Care and Use Committee. Conditional Syt9 KO mice were obtained from Jackson Labs (Xu et al., 2007). These conditional Syt9 KO mice were terminally crossed with an E2a-CRE mouse (Jackson Labs, Bar Harbor, ME, USA) to yield a population of constitutive Syt9 KO mice; the mice used in this study are all products of this constitutive Syt9 KO mouse. Syt1 cKO (conditional KO) mice were a gift from W.

Genotyping
The Syt9 KO mouse line was maintained as heterozygous breeders so that offspring WT and KO littermates could be studied. To genotype the neonatal pups, tissue samples were taken by collecting ~2 mm of the end of the tail and extracting DNA from the tissue sample using the Kapa Express Extract Kit (Kapa Biosystems, Wilmington, MA, USA). The extracted DNA was then subjected to PCR using in-house designed primers to identify WT and KO alleles. The primers used to identify the WT allele yield a 209 bp amplicon: F-primer 5'-GCGTCAATGGGAGGAGAGGTCA-3'; R-primer 5'-GTCCACTAGGGCTAGCCCAGG-3'. The primers used to identify the KO allele yield a 301 bp amplicon: F-primer 5'-CAGTAGGGAGCGAGCAGAAATTAAAAT-3'; R-primer 5'-AACTTCGTCGATCGACCTCGAATAA-3'. The PCR product was subjected to agarose gel electrophoresis and imaged on a ChemiDoc MP imaging system (BioRad, Hercules, CA, USA).
SYT9 Ca 2+ ligand mutant (CLM) constructs correspond to D-to-N substitutions of two Ca 2+ ligands in each C2-domain as follows: C2A m B; D197N and D199N; C2AB m ; D330N and D332N. All four mutations are present in the C2A m B m construct. These constructs were made by performing site-directed mutagenesis on the Syt9 cDNA with the QuikChange Lightning Site-Directed Mutagenesis Kit (Agilent, Santa Clara, CA, USA). Full-length CLM constructs were subcloned into pFSynW and used for CLM rescue experiments, and isolated C2-domain CLM constructs were subcloned into pGEX and used for biochemistry experiments. cDNA for SNAP-25B was provided by M. C. Wilson (University of New Mexico School of Medicine, Albuquerque, NM, USA), and full-length synaptobrevin 2 (SYB2) and syntaxin 1A (SYX) were provided by J. Rothman (Yale University, New Haven, CT); these SNARE proteins were subcloned into pTrcHis vectors to yield His 6 -tagged proteins .

Cell culture, transient transfection, and lentiviral transduction
Cortical, hippocampal, and striatal regions were obtained by microdissection and kept in ice-cold Hibernate-A (BrainBits, Springfield, IL, USA) with 2% B-27 until completion of dissection. Neural tissue was then incubated with 0.25% trypsin-EDTA (Corning Inc., Corning, NY, USA) for 25 minutes at 37°C. The tissue was washed with DMEM (Gibco, Carlsbad, CA, USA) supplemented with FBS (Atlanta Biologicals, Flowery Branch, GA, USA) and penicillin-streptomycin (Thermo Scientific, Waltham, MA, USA) and triturated 10-15 times with a P1000 pipette tip until complete dissociation of the tissue. Cells were plated at a density of 100,000 cells/cm 2 onto poly D-lysine-coated glass coverslips. Supplemented DMEM was replaced after 1 hour with Neurobasal-A (Gibco) supplemented with B-27 (Thermo Scientific) and GlutaMAX (Thermo Scientific). For SP-pH release experiments, sparse transfection of SP-pH and a cytosolic mRuby marker was required for identifying and imaging single neurons. This was achieved using Lipofectamine LTX with PLUS reagent (Invitrogen, Carlsbad, CA, USA) on DIV 8-9. For localization studies, rescue experiments, and Ca 2+ imaging, full coverage was desirable and was achieved via lentiviral transduction. Lentivirus encoding CRE-recombinase was added to Syt1 cKO neurons at DIV 0-2, and lentiviruses that expressed SYT9 rescue constructs, the Ca 2+ imaging construct synaptophysin-HaloTag (SYP-Halo) (Bradberry and Chapman, 2022), or SP-pH for localization studies, were added to neurons at DIV 5-7.

Electrophysiological studies
Whole-cell voltage-clamp recordings of cultured neurons (DIV 14-18) were performed at room temperature in artificial cerebrospinal fluid (ACSF) containing (in mM): 128 NaCl, 5 KCl, 1.5 CaCl 2 , 1 MgCl 2 , 30 glucose, 25 HEPES, pH 7.4 with an internal solution containing (in mM): 130 KCl, 10 HEPES, pH 7.4, 1 EGTA, 2 ATP, 0.3 GTP, 5 phosphocreatine. Recordings were performed using a MultiClamp 700B amplifier and Digidata 1550B digitizer (Molecular Devices, San Jose, CA, USA) under the control of Clampex 10 software (Molecular Devices). GABA A -mediated currents were pharmacologically isolated using 20 μM CNQX (Tocris Bioscience, Bristol, UK) and 50 μM D-AP5 (Tocris) in the bath solution; for mIPSC recordings, 1 μM tetrodotoxin (Tocris) was included in the bath. 5 mM QX 314 chloride (Tocris) was included in the pipette solutions for all recordings. Neurons were held at −70 mV in all experiments without correction for liquid junction potentials. Recordings were discarded if series resistance rose above 15 MΩ. For evoked recordings, a concentric bipolar electrode (FHC, Bowdoin, ME, USA) was placed 100-200 μm away from the patched soma and stimulation currents were adjusted to evoke maximal responses. For mIPSC recordings, 300 s of data were recorded for each striatal neuron, and 180 s of data were acquired for cortical and hippocampal neurons. mIPSCs were quantified for each recording using a template-matching algorithm in Clampfit (Molecular Devices). For pharmacology rescue experiments, SP (Tocris), GR73632 (Tocris), Leu-enkephalin (Tocris), dynorphin A (Tocris), and cholecystokinin (Tocris) were added to the ACSF at 100 nM, and SR140333 (Tocris) was added at 10 nM. Neurons were treated with peptides or drugs for ten minutes before patch-clamp experiments were conducted.

Immunocytochemistry and confocal microscopy
Immunocytochemistry was conducted as previously described (Courtney et al., 2019). For the pH-SYT9 rescue experiments, immediately before fixation and permeabilization, cortical neurons were incubated in primary antibody directed against the pHluorin tag of pH-SYT9 (1:250) (α-GFP; Abcam Cat# ab290, RRID: AB_303395) in culture medium to label only the surface-resident pH-SYT9, as the antibody is unable to enter intact, live neurons. Unbound antibody was removed by washing, and then cells were fixed and permeabilized and incubated a second time with primary antibodies to label synaptophysin (1:1000) (α-SYP; Synaptic Systems Cat# 101 004; RRID:AB_1210382) and pHluorin (1:1000) (α-GFP; Abcam Cat# ab13970, RRID:AB_300798). This second α-GFP antibody labeled all pHluorin tags that were not labeled by the first α-GFP 'blocking' antibody, to reveal the internal fraction of pH-SYT9. This strategy allowed for differential secondary labeling of the different pH-SYT9 populations with species-specific secondary antibodies.
Images were acquired on a Zeiss LSM 880 laser scanning confocal microscope with a 63x Plan-Apochromat 1.4NA DIC oil immersion objective. Identical laser and gain settings were used for all samples within each experimental paradigm. Primary antibodies were:

Live cell imaging
DIV 14-18 striatal neurons were washed twice in ASCF before being mounted in an imaging chamber with parallel platinum electrodes (Warner Instruments, Hamden, CT, USA) with 500 μL of ACSF. Imaging was carried out with a 40x, 1.4 NA oil-immersion objective on an inverted epifluorescence microscope (IX81, Olympus, Tokyo, Japan) equipped with a CMOS camera (Orca Flash 4.0 V2, Hamamatsu Photonics, Hamamatsu, Japan), motorized stage (Mad City Labs, Madison, WI, USA), and a custom illumination source containing three LEDs (470, 530, 625 nm) (Thorlabs, Newton, NJ, USA). The system was controlled using Micro-Manager (Edelsteinet al. 2010). Stimulation was delivered with a stimulation box (SD9, Grass Instrument Co., West Warwick, RI, USA) controlled via a HEKA EPC10 DAQ-amplifier and PatchMaster software (HEKA, Holliston, MA, USA), which was also used to synchronize the start of image sequence acquisition. Images were collected using 2 x 2 binning (0.325 μm x 0.325 μm pixel size).
For SP-pH release imaging, an exposure time of 100 ms was used. Neurons were sparsely co-transfected with the SP-pH construct and a cytosolic mRuby construct. Individual transfected neurons were identified for imaging by first detecting the cytosolic mRuby signal with the 530 nm LED and standard RFP filter set (49004, Chroma, Bellows Falls, VT, USA); this cytosolic mRuby construct also allowed us to ensure that no overlap with other cells was present in the imaging field, ensuring single-cell analysis. SP-pH release was imaged with the 470 nm LED, a standard green fluorescent protein (GFP) filter set (49002, Chroma). Ten seconds of baseline were obtained before stimulation was given-16 trains of 50 APs at 50 Hz with 0.5 sec between each train. Seventy seconds after the start of stimulation, 500 μL of 2x NH 4 Cl ACSF, containing (in mM): 100 NH 4 Cl, 28 NaCl, 5 KCl, 1.5 CaCl 2 , 1 MgCl 2 , 30 glucose, 25 HEPES, pH 7.4, was added to dequench pHluorin fluorescence to visualize the total number of SP-pH-bearing vesicles.
For Ca 2+ -imaging, the 625 nm LED, custom three-band pass dichroic mirror (Chroma) and far-red emission filter, and a 10 ms exposure time were used. DIV 14-18 striatal neurons that were transduced with a virus that induced expression of SYP-Halo were first washed twice in ACSF before being incubated in ACSF containing 100 nM of the acetoxymethyl ester of JF 646 -BAPTA bearing a HaloTag ligand (JF 646 -BAPTA-AM-HTL) (Deo et al., 2019), a gift from L. Lavis (Janelia Research Campus, Ashburn, VA, USA). After a 20-minute incubation at 37°C, the cells were again washed with dye-free ACSF and incubated for 20 minutes at 37°C before being mounted in the imaging chamber. Five fields of view (FOV) were selected per coverslip. Three seconds of baseline were recorded before a saturating stimulus (50 AP @ 50Hz) was delivered to establish a maximum fluorescence (F max ) for each field of view. At least five minutes elapsed between imaging each FOV to give the neurons time to recover and return to basal conditions.

Image analysis
For colocalization analysis, the Coloc2 plugin in Fiji (Schindelin et al., 2012)was used to determine the Mander's overlap coefficients and Pearson's correlation coefficients; Costes' automatic threshold regression was used for both determinations.
For synapse counting analysis, images were exported to SynD (Schmitz et al., 2011), where the image of MAP2 staining was used to create a neurite mask, then synaptic density was quantified by counting the number of overlapping VGAT and Gephyrin puncta within the neurite mask and dividing the total number by the length of dendrite measured to yield number of synapses per micron.
For SP-pH release analysis, Fiji with the DCV_pHluorin toolset (https://github.com/ alemoro/DCVpHluorin_toolseet) (Moro et al., 2021) was used; 3 pixels x 3 pixels regions of interest (ROIs) were placed over each observed increase in fluorescence, and somatic events were excluded. Fusion events were defined as a sudden increase of 2 standard deviations (SD) above the average fluorescence of the first 100 frames in the ROI (F 0 ). Time lapses were also manually scored along with the DCV_pHluorin analysis algorithm to ensure a thorough and complete characterization of each release event. Greater than 95% of ROIs contained only a single stepwise change in fluorescence after stimulation, so single ROIs were counted as single fusion events. The start of a fusion event was defined as the first frame with a signal that was at least twice the background noise (i.e., 2 x the SD of F 0 ), and end of a fusion event was defined by the first frame below this threshold. Most ROIs that contained a release event also showed increases in fluorescence after NH 4 Cl perfusion, indicating incomplete release of SP-pH. Pool size estimations were calculated using the same rules used for fusion event detection, after perfusion with NH 4 Cl. Analysis was blinded, and the number of fusion events, SP-pH vesicle pool size estimation, event duration, and fusion onset timing were all determined using DCV_pHluorin. Output data were processed in Excel and plotted using GraphPad Prism.
Ca 2+ -imaging analysis was performed as previously described (Bradberry and Chapman, 2022) with a modified ImageJ script taken from (Vevea and Chapman, 2020). Briefly, SYP-HaloTag-JF 646 -BAPTA traces were background subtracted, and ROIs were created based on a custom workflow to identify changes in fluorescence of the selected image series. An average intensity projection of pre-stimulus baseline frames was subtracted from a maximum intensity projection of the entire timelapse. This result was duplicated and mean filtered using a rolling ball radius of 10 pixels. The mean filtered image was subtracted from the original result, and this result was used to threshold changes in fluorescence. After thresholding, images were made binary, and a watershed function was run to separate overlapping objects. Using this procedure, objects (>10 pixels) were created and defined as ROIs. These ROIs were used to measure fluorescence changes over time from the original image series. The fluorescence traces of each ROI were then imported into AxoGraph where the basal, pre-stimulation fluorescence of each ROI (F r ) was subtracted from the max fluorescence at indicator saturation (F max ) to give a ΔF. The following equation was used to calculate presynaptic basal [Ca 2+ ] i : where K D is the dissociation constant of the indicator, F r is the baseline fluorescence of an ROI pre-stimulation, F max is the fluorescence achieved with the delivery of a saturating stimulus, R f is the dynamic range of the indicator, and n is the Hill coefficient. For K D , R f and n, we used in vitro measurements (K D = 140 nM, R f = 5.5, n = 1) from (Deo et al., 2019).
Trafficking analysis was conducted using the Multi Kymograph plugin in Fiji; 30-second time-lapses, taken after NH 4 Cl perfusion in the SP-pH release experiments, were used. Ninety μm segments that began within 20 μm of the soma were chosen. The kymograph was created from this time-lapse and every path was traced and quantified.

In vitro Fusion Assays
Fusion assays between SYB2 and SYX1A vesicles were conducted using a scaled-down version of the referenced method (Bhalla et al., 2006). For split t-SNARE fusion assays, 1 μM of the cytoplasmic domain (C2AB) or 2 μM of isolated C2-domains, SYB2 vesicles, SYX1A vesicles, 0.2 mM EGTA, and 7 μM soluble SNAP-25B were incubated for 20 min at 37°C. Ca 2+ (1 mM final free concentration) was added and the reaction was monitored for an additional 60 min. N-dodecyl-β-d-maltoside was added at the end of the experiment, to yield the maximum fluorescence signal. NBD donor fluorescence was monitored using a Synergy HT multidetection microplate reader (Bio-Tek, Santa Clara, CA, USA). Traces were normalized to the first time point, and the maximum fluorescence signal, to determine the %F max .

Co-sedimentation Assays
WT SYT9 C2A, C2B, or C2AB (4 μM) were combined with protein-free liposomes (1 mM total lipid; %PS as indicated in the Results) in 0.2 mM EGTA or 1 mM Ca 2+ and incubated for 15 min at RT. Samples were then centrifuged in an Optima MAX-E tabletop ultracentrifuge (Beckman Coulter, Brea, CA, USA) at 184,000 x g (65,000 rpm) for 45 min at 4°C. After centrifugation, the supernatant (S), and the pellet (P) were collected, and equal fractions were subjected to SDS-PAGE and Coomassie blue staining. Bands were quantified by densitometry and plotted.

Isothermal Titration Calorimetry (ITC)
WT and CLM forms of the isolated C2-domains of SYT9 were dialyzed overnight in ITC buffer (50 mM HEPES-NaOH, pH 7.4, 200 mM NaCl, and 10% glycerol) in the presence of Chelex-100 resin (BioRad), to remove residual divalent cations. ITC buffer was subsequently filtered and used to make Ca 2+ and protein dilutions. Heat of binding was measured, using a MicroCal ® iTC200 (Malvern Panalytical, Malvern, UK), at 25 °C, in response to twenty injections of Ca 2+ into the sample cell containing 100 μM protein. Heat of dilution was corrected by subtracting the signal that arises when Ca 2+ is diluted into buffer. Data were analyzed using the "sequential binding site" model in the data software package.]

Experimental design and statistical analysis
Graphically, bars represent the mean and error bars indicate the SEM. For electrophysiological experiments and live-cell imaging experiments, each n represents a single neuron and n's were obtained from at least three separate animal preparations. All data were tested for normality with the D'Agostino & Pearson test; the appropriate statistical test was applied based on the reported normality of the data. All statistical tests were twosided. Expected sample sizes were not estimated or predetermined. Statistical significance was assessed with two-tailed t-tests, ANOVAs with post-hoc pairwise comparisons, Mann-Whitney tests, or Kruskal-Wallis tests with post-hoc pairwise comparisons as indicated in the text or legends. Specific values t, df, Mann-Whitney U, Kruskal-Wallis test statistic, p, and r 2 values are reported in the Figure legends. All statistical analysis was conducted using GraphPad Prism 9.4 (GraphPad Software).

Synaptotagmin 9 is not required for fast, synchronous neurotransmitter release in cultured neurons
Immunoblot analysis of neuronal culture lysates revealed that SYT1 and SYT9 are expressed in cortical, hippocampal, and striatal neurons (Fig. 1A,E). By including known amounts of purified recombinant SYT1 cytoplasmic domain (C2AB) or SYT9 (full-length) as standards on these blots we were able to determine the absolute expression levels via quantitative densitometry (Fig. 1B,F). Briefly, recombinant protein standards were included in all immunoblots to generate standard curves. Ten micrograms of neuronal lysate protein were also immunoblotted, and the absolute levels of either SYT1 or SYT9 in the lysate were calculated from the standard curve in the same blot. This analysis revealed that SYT1 and SYT9 were ~0.1% and ~0.004% of the total protein in each of the neuronal lysates, respectively. Hence, SYT1 expression is ~25 times greater than SYT9 in all three classes of neurons.
A prior study suggested that SYT1 and SYT9 serve as partially redundant Ca 2+ sensors in striatal neurons and that SYT9 can fully rescue fast release in Syt1 KO cortical neurons (Xu et al., 2007). However, this is difficult to reconcile with the well-established findings that loss of SYT1 in hippocampal and cortical neurons disrupts virtually all synchronous release (Geppert et al., 1994;Maximov and Südhof, 2005). If SYT9 is a sensor for synchronous release, some fast release should have persisted in the Syt1 KO neurons. To address this conundrum, whole-cell patch-clamp experiments were conducted using cultured neurons from Syt1 conditional KO (cKO) and Syt9 KO mice. In striatal Syt1 cKO neurons, ablation of SYT1 expression with application of CRE lentivirus eliminated fast, synchronous neurotransmitter release with an 8-fold reduction in evoked inhibitory postsynaptic current (eIPSC) amplitude (Fig. 1C,D). Disruption of SYT9 expression, however, did not result in any change in eIPSC amplitude or kinetics in striatal neurons (Fig. 1G,H). Analogous results were obtained using cultured cortical and hippocampal neurons (Fig. 2). Evoked release during a stimulus train was also unaffected in Syt9 KO cortical, hippocampal, and striatal neurons, further demonstrating that this isoform does not impact evoked transmission (Fig.  3). We conclude that endogenous SYT9 is unable to support synchronous release in absence of SYT1 in any of the neuronal preparations studied here, perhaps because of the relatively low expression levels of this protein. Xu et al. (2007) reported that SYT9 over-expression can functionally rescue synchronous neurotransmitter release in cultured Syt1 KO cortical neurons (Xu et al., 2007). Similar observations were made upon over-expression of SYT9 in a calyx of Held slice preparation lacking the primary Ca 2+ sensor at this synapse, SYT2 (Kochubey et al., 2016). Given the observation that endogenous SYT9 is expressed at levels ~25-fold lower than SYT1 (Fig. 1B,F), a quantitative approach was taken to re-evaluate this previously reported rescue activity. Cultured Syt1 cKO cortical neurons were treated at DIV1 with a CRE lentivirus to disrupt SYT1 expression and then transduced at DIV5 with lentivirus to express SYT9 with an N-terminal pHluorin tag (pH-SYT9). The pH-SYT9 lentivirus was titered such that expression of pH-SYT9 in the + CRE neurons would be either 1x or 25x the endogenous SYT9 expression level; the latter condition matching the levels of endogenous SYT1 ( Fig.  4A; Fig. 1). Ablation of SYT1 expression again resulted in a loss of fast, synchronous neurotransmission, and 1x over-expression of pH-SYT9 had no significant effect on eIPSC amplitude, and while there was a decrease in the time-to-peak this did not reach significance ( Fig. 4B-E). However, 25x over-expression of pH-SYT9 significantly rescued the eIPSC amplitude (60% compared to − CRE) and completely rescued the eIPSC time-to-peak ( Fig.  4B-E). The 25x over-expression condition also rescued the decay time of the + CRE condition (Fig. 4F). So, only supraphysiological SYT9 expression levels rescue evoked responses in neurons lacking SYT1 (Xu et al., 2007;Kochubey et al., 2016).

SYT9 rescues SYT1 KO phenotypes only when over-expressed
SYT1 is a multifunctional protein that also serves to clamp spontaneous neurotransmitter release under resting conditions (Liu et al., 2014;Bai et al., 2016;Courtney et al., 2019;Vevea and Chapman, 2020). To investigate whether over-expressed pH-SYT9 can clamp spontaneous fusion, miniature inhibitory postsynaptic currents (mIPSCs, also called minis) were recorded in the presence of tetrodotoxin, to block action potentials (Fig. 4G). Disruption of SYT1 resulted in a ~3-fold increase in mIPSC frequency with no rescue observed in the 1x pH-SYT9 over-expression condition. In contrast, 25x over-expression of SYT9 resulted in a complete rescue of clamping activity (Fig. 4H). mIPSC amplitude was unaffected across conditions (Fig. 4I). Taken together, these findings indicate that SYT9 can rescue evoked release and clamp spontaneous release, in Syt1 KO neurons, only when massively over-expressed.
To investigate how the localization of over-expressed SYT9 functions to support SV release, we utilized the N-terminal pHluorin tag to differentiate between pools in the plasma membrane versus internal compartments (see Methods and (Courtney et al., 2019). Briefly, prior to fixation and permeabilization, cultured neurons were incubated with a rabbit anti-GFP antibody to label all surface-resident pH-SYT9. Then, after fixation and permeabilization, the cultures were incubated in a chicken anti-GFP antibody to label all pH-SYT9 on internal compartments. Using different species of anti-GFP antibodies allowed for differential labeling by fluorescent secondary antibodies. We co-stained with antibodies against synaptophysin (SYP) to label SVs. The internal fraction in the 1x over-expression condition showed some degree of colocalization with SYP, and the internal fraction of 25x pH-SYT9 had a higher degree of colocalization. We note that endogenous SYT9 colocalized with SYP to a similar degree as 1x over-expressed pH-SYT9, while 25x over-expressed pH-SYT9 colocalized with SYP to a similar degree as endogenous SYT1 (Fig. 5). Overexpression of SYT9 results in a greater degree of colocalization with SYP, suggesting that some fraction of the over-expressed protein is targeted to SVs. This increased SV localization likely underlies the apparent rescue of both fast, synchronous neurotransmitter release and clamping of spontaneous release in Syt1 KO neurons. So, while SYT9 has the intrinsic ability to regulate these aspects of SV release, it does so only after over-expression and mislocalization. These results are congruent with the idea that some SYT9 may be present on SVs, as indicated in proteomic studies (Takamori et al., 2006;Taoufiq et al., 2020). However, these levels are too low to support evoked release or to clamp spontaneous release, at least in the types of neurons studied here.

Decreased mIPSC frequency in SYT9 KO striatal neurons
To further characterize neurotransmission in Syt9 KO mice, spontaneous release from cultured striatal neurons was measured and compared to WT controls. We reiterate that all aspects of release evoked by single action potentials were normal (Fig. 1G); however, the mIPSC frequency in Syt9 KO striatal neurons was reduced 2-fold ( Fig. 6A,C), with no effect on the amplitude or kinetics of these spontaneous quantal events (Fig. 6B). Interestingly, when the same experiment was performed using cortical and hippocampal neurons, loss of SYT9 did not affect spontaneous release rates (Fig. 7), suggesting that this phenotype was specific for striatal neurons, most of which are GABAergic medium spiny neurons (MSNs) (Kemp and Powell, 1971). Importantly, the density of synapses was unchanged between WT and Syt9 KO striatal neurons (Fig. 8).
We note that most mIPSCs are Ca 2+ -dependent and are thought to use the same Ca 2+ sensors as evoked synchronous and asynchronous release, including SYT1 and Doc2 (Groffen et al., 2010;Courtney et al., 2018). The insufficiency of endogenous SYT9 to drive evoked release argues against the idea that it is a Ca 2+ sensor for mIPSCs. Furthermore, the specificity of the mIPSC phenotype for striatal versus hippocampal or cortical neurons suggests a cell-specific mechanism for the action of SYT9 in spontaneous release. Notably, SYT9 is targeted to DCVs in a variety of tissues (Fukuda et al., 2002;Iezzi et al., 2005;Lynch and Martin, 2007;Roper et al., 2015), and some DCV neuropeptides have been shown to regulate aspects of synaptic transmission (Dean et al., 2009;Cao et al., 2011;He et al., 2019). This prompts the idea that SYT9 might regulate minis indirectly, potentially by controlling the release of neuromodulators from DCVs.

SYT9 regulates the secretion of substance P-pHluorin from striatal neurons
To investigate the mechanism by which SYT9 regulates mIPSC frequency, we conducted further localization studies. Since the Syt9 KO phenotype was specific for striatal neurons, we focused on striatum-specific DCV neuropeptides. Among these, substance P (SP) was of particular interest because it has been shown to regulate mIPSC frequency in striatal neurons through its action on the neurokinin 1 receptor (NK1R) (He et al., 2019). Interestingly, SP is not enriched in hippocampal or cortical neurons (Shults et al., 1984), where the mIPSC phenotype was not observed.
Specific antibodies for SP are not available, so we transduced striatal neurons with a lentivirus that expresses an SP-pHluorin (SP-pH) fusion protein and performed immunocytochemistry (ICC) to assess co-localization of the pHluorin tag with endogenous SYT9. Strikingly, SYT9 was colocalized with SP-pH (Fig. 9A,B). The correct localization of the SP-pH construct was confirmed by staining for chromogranin B (CHGB), a canonical DCV marker in neurons (Fig. 9C,D). In summary, these findings demonstrate the presence of SYT9 on DCVs that bear a neuropeptide known to regulate mIPSC frequency (He et al., 2019).
Next, we sought to determine whether SYT9 regulates SP release from DCVs in striatal neurons. To optically measure SP secretion events at single-event resolution, WT and Syt9 KO neurons were sparsely transfected with a plasmid encoding SP-pH along with a cytosolic mRuby construct to enable identification of transfected neurons and imaging of individual neurons within a network culture. SP-pH fluorescence is quenched in the low pH environment of the DCV lumen; upon exocytosis, the pHluorin is rapidly dequenched, yielding a robust increase in fluorescence intensity (Fig. 9E). In cultured neurons, repetitive high-frequency electrical stimulation (i.e., 16 trains of 50 APs at 50Hz) has been shown to effectively trigger these release events (Persoon et al., 2018); this paradigm was used in our experiments and yielded robust exocytosis ( Fig. 9F; Video 1). DCV fusion was quantified in WT and Syt9 KO striatal neurons, where the released fraction was calculated as the number of fusion events divided by the total number of SP-pH vesicles in a single neuron. The total SP-pH DCV pool was estimated by perfusing neurons with NH 4 Cl to alkalinize the lumen of DCVs and dequench the pHluorin reporter; no significant differences were observed between the WT and Syt9 KO (Fig. 9G). In WT neurons the released fraction was ~10%. In Syt9 KO neurons this was reduced by ~61% (Fig. 9H,I), with only ~3.9% of the vesicles being released. Fusion event duration was also measured in WT and Syt9 KO neurons, and there appeared to be a small population of long-duration release events that was absent in the WT (Fig. 9J), hinting at a potential role of SYT9 in regulating the fate of fusion pores, as shown for other SYT isoforms (Wang et al., 2001;Rao et al., 2014). Finally, loss of SYT9 had no apparent effect on the transport of SP-pH vesicles, suggesting that the observed reductions in SP secretion were not secondary to trafficking defects (Fig. 10).

Application of exogenous substance P rescues the SYT9 KO mIPSC phenotype
Since our data suggest that SYT9 regulates SP release (Fig. 9), and SP is known to modulate mIPSC frequency in striatal neurons (He et al., 2019), we sought to determine whether the Syt9 KO phenotype could be rescued via acute application of this neuropeptide.
In addition to SP, enkephalins, dynorphins, and cholecystokinin are also enriched in the striatum (Krause et al., 1987;Angulo and McEwen, 1994), so we also assayed the effects of each of these neuropeptides (Fig. 11A). Acute addition of SP to the bath fully rescued the decreased mIPSC frequency in Syt9 KO neurons. Rescue was also observed using GR73632, a synthetic neurokinin 1 receptor (NK1R) agonist. In contrast, dynorphin A (DynA), Leuenkephalin (L-Enk), and cholecystokinin (CCK8) had no effect, demonstrating that the mIPSC phenotype is SP-specific (Fig. 11B). Also, we found that SP supplementation did not significantly change the mIPSC frequency of WT neurons, indicating a high degree of SP-receptor activation under basal conditions.
This finding is consistent with a previous study utilizing a different NK1R antagonist (He et al., 2019), and confirms that the SP-NK1R pathway serves to regulate mini frequency in striatal neurons. Consistent with this model, SR140333 had no effect on mini frequency in Syt9 KO neurons, presumably due to the large reduction in SP release from these cells and the concomitant decrease in NK1R activation. No differences in mIPSC amplitude were observed across all conditions (Fig. 11C). In summary, SYT9 regulates SP release, and activation of the NK1R by SP is pivotal in the regulation of mIPSCs in striatal neurons.
The binding of SP to the NK1 receptor presumably activates G q/11 , resulting in the generation of IP 3 and the mobilization of Ca 2+ from internal stores. While basal presynaptic [Ca 2+ ] i did not appear to be altered in the Syt9 KO striatal neurons (Fig. 12), it has been shown that local, transient increases in [Ca 2+ ] i , mediated by IP 3 receptors in the presynaptic ER, drive spontaneous neurotransmitter release in pyramidal neurons (Emptage et al., 2001;Simkus and Stricker, 2002). We therefore speculate that decreased SP-mediated IP 3 signaling in the Syt9 KO underlies the decreased mIPSC phenotype observed in our studies.

Ca 2+ -binding to the C2A domain of SYT9 is crucial for its function in vitro and in neurons
The findings presented thus far suggest a model in which SYT9 regulates SP release which, in turn, modulates mini frequency. To further evaluate this model and directly determine whether SYT9 serves as a Ca 2+ sensor for SP release, we sought to identify the structural elements in SYT9 that regulate membrane fusion in vitro, and then correlate these findings with the function of SYT9 in regulating both SP release and mini frequency. For these experiments, we used soluble, cytoplasmic fragments of SYT9, and for comparison, SYT1, in a 'split t-SNARE' in vitro fusion assay (Bhalla et al., 2006) that mimics fusion of SVs with the plasma membrane in cells. In this assay, SYT fragments must first fold soluble SNAP-25B onto membrane-embedded syntaxin 1A (SYX1A) for fusion with synaptobrevin 2 (SYB2)-bearing liposomes to occur. Consistent with a previous study (Roper et al., 2015), the cytoplasmic domains (C2AB) of both SYT1 and SYT9 stimulated fusion to similar extents in the presence of Ca 2+ (omitted for clarity in Fig. 13A left panel but quantified in Fig. 13A right panel). We extended these experiments by evaluating the isolated C2-domains of each isoform and found that the C2A domain of SYT9 strongly stimulated fusion in response to Ca 2+ , while C2B had little effect (Fig. 13A). This is in sharp contrast to SYT1, where C2B, but not C2A, facilitated fusion ( Fig. 13A) (Gaffaney et al., 2008). Additionally, we utilized Ca 2+ ligand mutants (CLMs), in which native Asp residues (D197, D199 in C2A and D330, D332 in C2B) that serve as conserved Ca 2+ ligands in numerous isoforms were substituted with Asn, in either C2A or C2B of SYT9. These mutations disrupted the Ca 2+ binding activity (Fig. 13C) of each C2-domain and abolished the ability of the C2A domain of SYT9 to stimulate membrane fusion in response to Ca 2+ (Fig. 13A). Hence, these two related isoforms have diverged unexpectedly, with C2A serving as the primary Ca 2+ sensing domain in SYT9, while C2B is more crucial in SYT1.
Since SYT1 regulates fusion via interactions with membranes that harbor anionic phospholipids (Bhalla et al., 2005;Liu et al., 2014;Bai et al., 2016), we assayed the ability of SYT9 to bind phosphatidylserine (PS)-harboring membranes in the presence and absence of Ca 2+ . A previous study reported that for SYT9 C2A, but not C2B, bound to PS-bearing vesicles (Shin et al., 2004). We found that each of the isolated C2-domains efficiently bound to liposomes, in response to Ca 2+ , when the mole fraction of PS was increased from 15% to 25%; this interaction is physiologically relevant since PS is ~22% of the inner leaflet of the plasma membrane (Hamilton et al., 2000) (Fig. 13B).
We also directly measured Ca 2+ binding to the isolated C2-domains of SYT9, using isothermal titration calorimetry (ITC); calorimetry data for SYT1 has been previously reported (Radhakrishnan et al., 2009;Evans et al., 2016). Ca 2+ binding to SYT9 C2A was endothermic and binding to C2B was exothermic (Fig. 13C). When fitted with a sequential binding site model, C2A exhibited three binding sites with K D values of 53 μM, 182 μM, and 337 μM; C2B had two binding sites, with K D values of 43 μM and 500 μM (see Table  1 for all thermodynamic values). Interestingly, the K D value for the third binding site of C2A (337 μM) was significantly lower than the value reported for SYT1 (K D > 1 mM) (Radhakrishnan et al., 2009;Evans et al., 2016). Titrations were also performed on the CLM forms of isolated C2A or C2B, described above, and these constructs yielded little to no signal, confirming they fail to bind Ca 2+ (Fig. 13C). These experiments demonstrate that while both isolated C2-domains bind Ca 2+ , only C2A was able to effectively stimulate membrane fusion.
Next, if our model is correct, the biochemical results above suggest that Ca 2+ binding to C2A, but not C2B, should underlie the ability of SYT9 to promote SP release. To test this, full-length variants of the SYT9 CLMs were generated and expressed in SYT9 KO neurons via lentiviral transduction; WT SYT9 served as a positive control. SP release was monitored using the optical assay described in Fig. 9. Importantly, full length SYT9 constructs with mutated Ca 2+ ligands in the C2A domain or both C2 domains, referred to as C2A m B and C2A m B m , respectively, failed to rescue release, while the full-length construct with mutated Ca 2+ ligands in the C2B domain, referred to as C2AB m , exhibited full rescue ( Fig.  14A) with no change in DCV pool size (Fig. 14B). Hence, the ability of these mutants to regulate reconstituted membrane fusion reactions in vitro, and to drive SP release from living neurons, were well correlated; in both cases, the C2A domain functioned as the crucial Ca 2+ -sensing motif. These findings further support the idea that SYT9 functions as a Ca 2+ sensor for DCV exocytosis.
Finally, we conducted patch-clamp recordings of Syt9 KO striatal neurons that expressed each full-length CLM construct. Again, complete rescue of mIPSC frequency was observed using WT SYT9. In sharp contrast, SYT9 C2A m B m , which cannot bind Ca 2+ via either C2domain, failed to rescue the mini phenotype. Consistent with our model, C2A m B also failed to rescue (Fig. 14C), while C2AB m was as effective as the WT protein ( Fig. 14D; note: no changes in mini amplitude were observed across conditions [ Fig. 14E]). In summary, Ca 2+ binding to the C2A domain, but not the C2B domain, underlies the ability of SYT9 to regulate membrane fusion to, in turn, indirectly control the rate of spontaneous release in striatal neurons by controlling SP secretion.

DISCUSSION
SYT9 is widely expressed in the brain (Mittelsteadt et al., 2009) (note: these authors referred to SYT9 as SYT5), but KOs lacking this protein have only been characterized using the calyx of Held slice preparation (Kochubey et al., 2016) and cultured striatal neurons (Xu et al., 2007). SYT9 appears to be expressed in the calyx of Held, but loss of this protein did not affect AP-triggered neurotransmitter release (Kochubey et al., 2016). In contrast, in cultured striatal neurons, one study reported a 57% reduction in eIPSC amplitude in Syt9 KO striatal neurons (Xu et al., 2007). However, this finding has not been reproduced, and it is difficult to reconcile with the expression patterns of SYT9 and SYT1. Namely, SYT9 is expressed at similar levels in cortical, hippocampal, and striatal neurons, and it is well-established that loss of SYT1 abolishes synchronous release in both cortical and hippocampal neurons (Geppert et al., 1994;Maximov and Südhof, 2005) (Fig. 1). So, it is apparent that endogenous SYT9 does not function as a Ca 2+ sensor for synchronous SV exocytosis in either of these two classes of neurons ( Fig. 1; Fig. 2). Here, we extended these observations to cultured striatal neurons and found that SYT1, but not endogenous SYT9, drives rapid SV exocytosis. We went on to show that SYT9 is largely localized to SP-bearing DCVs in striatal neurons, where it regulates the release of this neuropeptide to indirectly control mIPSC frequency. Evidently, at least some isoforms of SYT have become specialized to control the release of neuropeptides that impact synaptic function (Dean et al., 2009;Cao et al., 2011). It is tempting to speculate the divergence of numerous SYT isoforms might enable the differential release of myriad distinct neuromodulators and hormones, in response to different patterns of activity (Moghadam and Jackson, 2013).
We found that SYT9 is expressed at ~25-fold lower levels than SYT1 (Fig. 1). Given the estimate of ~15 copies of SYT1 per SV (Takamori et al., 2006), the abundance of SYT9 on SVs would, on average, be expected to be low, particularly since its primary localization appears to be on DCVs (Fig. 8). Some fraction of SYT9 does colocalize with canonical SV markers ( Fig. 5; (Xu et al., 2007), and SYT9 was detected in proteomic studies of purified SVs (Takamori et al., 2006;Taoufiq et al., 2020) (note: these authors referred to SYT9 as SYT5). Thus, it remains possible that there could be limited populations of SVs, in discrete brain regions, which harbor a sufficient copy number of SYT9 molecules to trigger SV exocytosis. However, in our hands, this was not the case with cortical, hippocampal, or striatal neurons ( Fig. 1; Fig. 2). It should also be emphasized that DCVs are often present in or near presynaptic nerve terminals (Persoon et al., 2018;Radhakrishnan et al., 2021), and this can potentially exaggerate the presence of SYT9 on SVs as determined via immunocytochemistry.
While our experiments indicate that SYT9 functions as a Ca 2+ sensor for SP release from striatal neurons, some degree of regulated release persisted in the KO (Fig. 9), suggesting that additional Ca 2+ sensors are associated with SP-bearing DCVs. Indeed, SYT1 was present on SP-bearing DCVs purified from rabbit optic nerve (Berg et al., 2000). Additionally, in hippocampal neurons, SYT1 and SYT7 have both been implicated in DCV exocytosis (van Westen et al., 2021), suggesting multiple roles for these Ca 2+ sensors. We also note that SYT4, which does not bind Ca 2+ , negatively regulates the release of brain-derived neurotrophic factor (BDNF) (Dean et al., 2009), and SYT10, which binds Ca 2+ , positively regulates the secretion of IGF-1 from DCVs in neurons (Cao et al., 2011). Collectively, these findings, combined with the observations in the current study, support the idea that the SYTs have diverged, at least in part, to control different forms of DCV exocytosis (Dean et al., 2012). We also reiterate that the Syt9 KO mIPSC phenotype appears to be specific for striatal neurons, which robustly express SP. The function of SYT9 in cortical and hippocampal neurons remains unknown. We speculate that it regulates the release of neuropeptides; a future goal is to address this issue via co-localization studies and by isolating and characterizing SYT9-bearing organelles from these classes of neurons.
The loss of SYT9 in striatal neurons results in a decrease in spontaneous neurotransmitter release (Fig. 6). Spontaneous release is thought to play an important role in synapse function and structure. For example, minis modulate excitability by regulating the tonic activity of post-synaptic receptors (Sutton et al., 2004 and maintaining postsynaptic receptor density (Ehlers et al., 2007). Spontaneous release also regulates dendritic protein synthesis to control aspects of synaptic plasticity . Further work is now needed to address how the change in mini frequency, in SYT9 KO striatal neurons, affects their function and plasticity in native circuits.
Regarding the diversity of SYT isoforms, our finding that the C2A domain of SYT9 serves as the crucial Ca 2+ -sensing motif for fusion was unexpected. SYT9 forms a clade with SYT1/2, and Ca 2+ ligand mutations in the C2B-domain of SYT1 clearly disrupt function (Nishiki and Augustine, 2004), while similar mutations in C2A have been reported to either have no effect (Fernández-Chacón et al., 2002), to be gain-of-function (Stevens and Sullivan, 2003;Pang et al., 2006b), or to cause partial loss-of-function (Shin et al., 2009). Moreover, the isolated C2B domain of SYT1, but not the C2A domain, regulates fusion in vitro ( Fig.   13; [Gaffaney et al., 2008]). In contrast, Ca 2+ -binding to the C2A domain of SYT9, but not the C2B domain, was required to both stimulate fusion in vitro (Fig. 13) and promote SP release from striatal neurons to regulate minis (Fig. 14). Apparently, this shuffling of C2-domain function is tolerated, as SYT9 can rescue evoked neurotransmitter release and clamp spontaneous release in Syt1 KO neurons, when sufficiently over-expressed (Fig. 4).
Returning to the physiology, we emphasize that the striatum is a specialized structure with a dense local network largely composed of medium spiny neurons (MSNs) and their axon collaterals. While a substantial fraction of these neurons projects to other brain regions, direct synaptic communication between MSNs also occurs (Tunstall et al., 2002); this is the condition recapitulated in our experiments using cultured striatal neurons. Subpopulations of MSNs release SP within the striatum (Gerfen, 1992) and harbor NK1 receptors (Haneda et al., 2007). We observed modulation of MSN-MSN transmission by SP in culture, and the next step will be to measure such modulation in acute striatal slice preparations. In addition, MSNs project to targets in the substantia nigra and globus pallidus, and recordings from brain slices that encompass the striatum along with these other structures will address the role of SYT9-regulated SP release at terminals that innervate these target cells. Moreover, SP enhances center-surround contrast of striosomal dopamine signals by modulating dopamine release from neurons that project from the substantial nigra onto striatal targets (Brimblecombe and Cragg, 2015). Hence, slice recordings, to measure dopamine release, are an important future direction of this work.
We propose that SP modulates spontaneous release rates in striatal neurons by binding the NK1 receptor and activating G q/11 , resulting in the generation of IP 3 and the mobilization of Ca 2+ from internal stores (Fig. 15), thus increasing mini frequency. Indeed, localized increases in [Ca 2+ ] i , mediated by IP 3 receptors in the presynaptic ER, have been reported to drive spontaneous release in pyramidal neurons (Emptage et al., 2001;Simkus and Stricker, 2002). A future goal is to compare these brief Ca 2+ transients, akin to Ca 2+ sparks in muscle cells, in WT and Syt9 KO striatal neurons. Finally, we emphasize that the highest expression levels of SYT9 in mouse brain occur in the pituitary gland and hypothalamus, both of which are particularly rich sources of numerous classes of DCVs (Roper et al., 2015). It will be interesting to investigate whether SYT9 regulates the release of hypothalamic hormones.

SIGNIFICANCE STATEMENT
Synaptotagmin 9 (SYT9) has been described as a Ca 2+ sensor for dense core vesicle (DCV) exocytosis in neuroendocrine cells, but its role in neurons remains unclear, despite widespread expression in the brain. This article examines the role of SYT9 in synaptic transmission across cultured cortical, hippocampal, and striatal neuronal preparations. We found that SYT9 regulates spontaneous neurotransmitter release in striatal neurons by serving as a Ca 2+ sensor for the release of the neuromodulator substance P from DCVs. This demonstrates a novel role for SYT9 in neurons and uncovers a new field of study into neuromodulation by SYT9, a protein that is widely expressed in the brain.    A, Immunoblots of DIV14 cortical neuronal lysates from Syt1 cKO mice, ± CRE lentivirus.

Video 1. SYT9 regulates the secretion of SP-pH from striatal neurons
Representative time-lapse of SP-pH release events in a WT striatal neuron during electrical stimulation (16 x [50 AP at 50 Hz]). Stimulation starts at t = 10 s and ends at t = 34 s; NH 4 perfusion starts at t = 80 s. Scale bar = 40 μm. Thermodynamic properties of Ca 2+ binding to isolated C2 domains of SYT9 using ITC.