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. Author manuscript; available in PMC: 2025 Dec 10.
Published in final edited form as: J Neurosci. 2025 Oct 22;45(43):e1052252025. doi: 10.1523/JNEUROSCI.1052-25.2025

Presenilin loss impairs synaptic transmission and causes axonal degeneration through ryanodine receptor dysfunction, independent of γ-secretase activity

Xinran Du 1,1, Longgang Niu 1,1, Michal Ragan 1, Zuzanna Fracz 1, Zhao-Wen Wang 1,*
PMCID: PMC12548705  NIHMSID: NIHMS2122234  PMID: 40925676

Abstract

Presenilin mutations are the most common cause of familial Alzheimer’s disease (FAD), but the mechanisms by which they disrupt neuronal function remain unresolved, particularly in relation to γ-secretase activity. Using C. elegans, we show that the presenilin ortholog SEL-12 supports synaptic transmission and axonal integrity through a pathway involving the ryanodine receptor RYR-1. Loss-of-function mutations in either sel-12 or ryr-1 reduce neurotransmitter release and cause neuronal structural defects, with no additional impairment in double mutants, suggesting a shared pathway. Transgenic expression of a γ-secretase–inactive SEL-12 variant or human presenilin 1 restores normal synaptic transmission in sel-12 mutants. Notably, sel-12 loss does not alter ryr-1 transcript or protein levels. These findings define a novel γ-secretase–independent role for presenilin in maintaining neuronal function via ryanodine receptor signaling, providing new mechanistic insight into presenilin-linked neurodegeneration and pointing to potential therapeutic strategies for FAD.

Introduction

Alzheimer’s disease (AD), a leading cause of age-related cognitive decline, imposes a devastating societal burden, with familial AD (FAD) offering critical insights into pathogenic mechanisms. Approximately 90% of FAD cases stem from mutations in PSEN1 and PSEN2, encoding presenilin 1 (PS1) and presenilin 2 (PS2) (Ho and Shen, 2011; De Strooper et al., 2012), the catalytic core of the γ-secretase complex responsible for amyloid β (Aβ) peptide generation (Oikawa and Walter, 2019). Prevailing pathogenic models posit that most FAD-linked presenilin mutations are gain-of-function in nature, driving neurodegeneration by enhancing the production of pathogenic Aβ peptides (Burdick et al., 1992; Jarrett and Lansbury, 1993; Suzuki et al., 1994; Fernandez et al., 2014; Sun et al., 2017). However, clinical trials targeting Aβ production or clearance have repeatedly failed to halt cognitive decline (Kametani and Hasegawa, 2018; Makin, 2018; Huang and Liu, 2020; Imbimbo et al., 2020), underscoring the urgent need to explore Aβ-independent pathways.

At least a subset of FAD cases results from loss-of-function (lf) mutations in presenilins (De Strooper et al., 2012). Similarly, mice with conditional double knockout (cDKO) of Psen1 and Psen2 exhibit progressive neurodegeneration and brain atrophy with age (Feng et al., 2004; Saura et al., 2004). While these findings indicate that presenilin lf mutations can cause neurodegeneration, the underlying molecular mechanisms remain largely unknown.

Dysregulated Ca²⁺ mobilization from the endoplasmic reticulum (ER) has emerged as a key factor in presenilin-linked neurodegeneration. Presenilins modulate ER Ca2+ flux via ryanodine receptors (RyRs), inositol 1,4,5-triphosphate receptors (IP3Rs), and sarco/endoplasmic reticulum Ca²⁺-ATPase (SERCA) (Honarnejad and Herms, 2012). Some studies suggest that presenilins may function as Ca²⁺ channels in the ER membrane, with pathogenic mutations impacting Ca²⁺ release (Tu et al., 2006; Nelson et al., 2007; Nelson et al., 2011; Supnet and Bezprozvanny, 2011; Zampese et al., 2011). Notably, cDKO of Psen1 and Psen2 in presynaptic hippocampal neurons reduces neurotransmitter release through RyR dysfunction (Zhang et al., 2009; Wu et al., 2013). However, it remains unclear whether RyR dysfunction contributes to neurodegeneration and whether the synaptic role of presenilins depends on their γ-secretase activity.

The nematode C. elegans provides a uniquely powerful model to dissect Aβ-independent mechanisms of presenilin-mediated neurodegeneration. Although it lacks the amyloid β precursor protein (Daigle and Li, 1993), C. elegans exhibits neurodegeneration upon loss of sel-12 (Alexander et al., 2014; Alvarez et al., 2022), a gene homologous to mammalian presenilins. It also offers a convenient system to investigate whether presenilin deficiency-associated RyR dysfunction contributes to neurodegeneration, as it possesses a single RyR gene, ryr-1 (also known as unc-68), whose deletion phenocopies the presynaptic defects observed in presenilin-deficient mice (Liu et al., 2005; Chen et al., 2017; Mueller et al., 2023). However, whether sel-12 deficiency impairs synaptic transmission in C. elegans, as it does in mice (Zhang et al., 2009; Wu et al., 2013), remains unknown.

Here, leveraging C. elegans genetics, electrophysiology, and axonal morphology analyses, we uncover a conserved, γ-secretase-independent pathway through which presenilin safeguards synaptic function and neuronal structure via RyRs. We demonstrate that sel-12(lf) and ryr-1(lf) mutations non-additively impair neurotransmitter release and induce axonal degeneration. Remarkably, synaptic defects in sel-12 mutants are rescued by neuronal expression of either human PSEN1 or a SEL-12 variant lacking γ-secretase activity. These findings redefine presenilin’s role in aging-associated neurodegeneration, highlighting RyR-related molecular pathway as a potential therapeutic target for FAD and related neurodegenerative disorders.

Materials and Methods

C. elegans culture and strains.

All C. elegans strains were maintained on nematode growth medium (NGM) plates seeded with OP50 Escherichia coli at 22°C in an environmental chamber. The strains used in this study are listed in Table S1.

Rescue experiments.

sel-12 mutant rescue experiments were performed by microinjection of plasmids (20 ng/µl) containing a cell-specific promoter driving wild-type sel-12 cDNA (F35H12.3). Plasmids used included Prab-3::sel-12 (wp2133) for neuronal rescue, Pmyo-3::sel-12 (wp2132) for body-wall muscle rescue, Pglr-1::sel-12 (wp2271) for command interneuron rescue (AVA, AVD, AVE, PVC), and Punc-17;unc-47::sel-12 (wp2204) for rescue in cholinergic and GABAergic motor neurons. The sel-12 cDNA was amplified from first-strand cDNA synthesized from wild-type animals using primers listed in Table S2. In addition, the fosmid WRM0633C (80 ng/µl) was used to rescue sel-12(zw101). Pmyo2::mStrawberry (wp1613) served as the transformation marker.

Knockdown experiments.

sel-12 and ryr-1 knockdowns were achieved by expressing ~500 bp fragments of complementary sense and antisense cDNA sequences under cell-specific promoters. The following plasmids were used to induce knockdown (sense/antisense plasmid pairs are indicated in parentheses): pan-neuronal knockdown, Prab-3::sel-12 RNAi (wp2232/2233); command interneuron-specific knockdown (AVA, AVD, AVE, PVC), Pglr-1::sel-12 RNAi (wp2238/2239); and A-type cholinergic motor neuron-specific knockdown, Punc-4::sel-12 RNAi (wp1609/wp1610). The latter plasmid pair was used in our previous study (Chen et al., 2017). AVA-specific ryr-1 knockdown was performed by co-injecting three plasmids: Pflp-18::loxP::LacZ::STOP::loxP::ryr-1 RNAi (sense/antisense: wp2310/2311), and pNP269::Pgpa-14::Cre (Schmitt et al., 2012). Primer sequences used to amplify the sel-12 and ryr-1 cDNA fragments are listed in Table S2. Pmyo2::mStrawberry (wp1613) was used as the transformation marker.

Generation of the sel-12(zw101) allele.

A CRISPR/Cas9 strategy was used to generate the sel-12 knockout strain. A single guide RNA (sgRNA) targeting the first exon of sel-12 (5′-TCCACAAGGAGACAACAGG-3′) was cloned into the Pu6::unc-119 sgRNA plasmid (Addgene plasmid #46169), replacing the original unc-119 sgRNA to create the Pu6::sel-12 sgRNA construct (wp2186). This plasmid was co-injected into wild-type worms along with the Peft-3::Cas9-SV40_NLS::tbb-2 (Addgene plasmid #46168) and a transformation marker, Pmyo2::mStrawberry (wp1613). Transgenic animals were isolated, and genomic DNA from their progeny was PCR-amplified for sequencing to verify editing. The primers used for amplification and sequencing were 5′-ATTGCCGACCATTGCAA-3′ (forward) and 5′-CGAATGAAAATTGCAGGGGGTAT-3′ (reverse).

Tagging RYR-1 with GFPnovo2.

To tag RYR-1 with GFPnovo2, a DNA sequence encoding GFPnovo2 (including artificial introns and flanking glycine residues, 8 on each side) was inserted between the codons for amino acids V1392 and R1393 in ryr-1b (K11C4.5b) using CRISPR/Cas9 (Suny Biotech, Fuzhou, China), resulting in the knock-in strain ryr-1(syb7697[GFPnovo2]).

sel-12 expression analysis.

To analyze sel-12 expression pattern, a 518 bp Psel-12 fragment immediately upstream of the start codon was fused to mScarlet cDNA to generate a Psel-12::mScarlet plasmid (wp2246). This plasmid was linearized with PstI and co-injected with the fosmid WRM0633C, which was linearized with NotI. Fluorescence and DIC images were captured using a Leica Thunder Imager equipped with 20x and 63x objectives and a DFC9000 GT sCMOS camera (Leica, Wetzlar, Germany).

Analysis of ryr-1 transcription and RYR-1 protein levels.

To assess whether sel-12(lf) affects ryr-1 transcription in neurons, the Pryr-1neuronal::GFP transgene (Chen et al., 2017) in a wild-type background was crossed into sel-12(zw101). To examine RYR-1 protein levels, the knock-in strain ryr-1(syb7697[GFPnovo2]) was also crossed into sel-12(zw101). Fluorescence and DIC images were acquired as described above. Young adult hermaphrodites were imaged under identical exposure conditions, and maximal z-projection fluorescence intensities were compared between wild-type and sel-12(zw101) animals.

Axonal morphology analysis.

Axonal morphologies of PLM and AVA neurons were analyzed in integrated transgenic strains expressing GFP in mechanosensory neurons (zdIs5[Pmec-4::GFP]) (Wu et al., 2007) or mStrawberry in AVA interneurons (Liu et al., 2020). These transgenes were crossed from wild-type backgrounds into mutant strains including sel-12(zw101), ryr-1(syb216), and sel-12(zw101);ryr-1(syb216) for comparative analysis. Images of the AVA posterior axon (relative to the vulva) and the distal PLM axon were obtained using the Leica imaging system. Axonal defects were scored as the percentages of neurons exhibiting breaks (clear discontinuities in axonal fluorescence) or beads (focal swellings), using a approach similar to that described previously (Sarasija et al., 2018).

Electrophysiology.

C. elegans hermaphrodites were used for all electrophysiological experiments, following methods similar to those previously described (Liu et al., 2005; Liu et al., 2007; Niu et al., 2020). Briefly, a young adult animal was immobilized on a Sylgard-coated glass coverslip by applying a cyanoacrylate adhesive (3M Vetbond Tissue Adhesive) to the dorsal midsection in a solution identical to the bath solution used during recording. A longitudinal incision was made along the glued region, and internal organs were removed. The resulting cuticle flap was folded back and secured to the coverslip. The exposed preparation was treated with 0.5 mg/ml collagenase A (10103578001, Roche Applied Science) for approximately 10 seconds, then rinsed thoroughly by perfusing the bath solution. Borosilicate glass pipettes (3–5 MΩ tip resistance) were used as electrodes. Whole-cell configuration was achieved by applying negative pressure through the recording pipette. Voltage-clamp recordings were performed using a Nikon FN1 microscope equipped with a digital patch clamp amplifier (Sutter Instrument, Novato, CA) and SutterPatch software. Signals were filtered at 2 kHz and sampled at 10 kHz. Cells were held at −60 mV to record ePSCs, minis, and muscle responses to exogenous acetylcholine and GABA. ePSCs were elicited by delivering a 20-V pulse (0.5 ms) through a stimulation electrode positioned near the ventral nerve cord. Exogenous acetylcholine or GABA was diluted to the desired final concentration (in the bath solution) and applied via pressure ejection (3 psi, 1 s) through a glass pipette (tip diameter ~2 μm) using a Picospritzer III microinjector (Parker Hannifin). Histamine was added to the recording chamber bypipetting. The bath solution consisted of (in mM): 140 NaCl, 5 KCl, 5 CaCl₂, 5 MgCl₂, 11 dextrose, and 5 HEPES, with the pH set to 7.2 using NaOH. The pipette solution contained (in mM): 120 KCl, 20 KOH, 5 Tris, 0.25 CaCl₂, 4 MgCl₂, 36 sucrose, 5 EGTA, and 4 Na₂ATP, with the pH adjusted to 7.2 using HCl.

Optogenetic stimulation.

Worm expressing ChR2 were grown to L4 stage on regular culture plates and then transferred to new plates either with or without all-trans retinal 1 day before the experiment. The retinal plates were prepared by spotting each plate (60-mm diameter with 10 ml agar) with 200 µl OP50 E. coli containing 2 mM retinal. Photostimulation was applied through the 40x water-immersion objective using a light source (Lambda XL with SmartShutter, Sutter Instrument) equipped with a 470±20 nm excitation filter (59222, Chroma Technology Corp.). The measured light intensity at the specimen position was 6.7 mW/mm2, as described in our previous study (Liu et al., 2013).

Chemicals.

Acetylcholine (AC159170050, Thermo Scientific), GABA (AC103280250, Thermo Scientific), and histamine (H7250, Sigma-Aldrich) were dissolved in water to prepare 1.0 M stock solutions, which were stored at −20°C. All-trans retinal (R2500, Sigma-Aldrich) was dissolved in DMSO to prepare a 100-mM stock solution.

Data analysis.

ePSC amplitudes, mean mini amplitudes, and mini frequencies were quantified using Clampfit (version 11, Molecular Devices, Sunnyvale, CA, USA). The threshold for mini detection was set at 5 pA. Graphs and statistical analyses were generated using OriginPro 2024 (OriginLab Corporation, Northampton, MA, USA). Data are presented as mean ± SEM. Statistical comparisons were performed using either one-way ANOVA with Fisher’s LSD post hoc test or t-tests, as indicated in figure legends. For comparisons of mini and ePSC properties, all groups were analyzed together before being presented in separate figures. A p-value of less than 0.05 was considered statistically significant. The sample size (n) refers to the number of animals analyzed.

Results

Loss of sel-12 impairs neurotransmitter release

We investigated the role of SEL-12 in synaptic transmission by analyzing evoked postsynaptic currents (ePSCs) and miniature postsynaptic currents (minis) at the C. elegans neuromuscular junction (NMJ). We began with two available sel-12 mutants: sel-12(ok2078), which carries a large deletion encompassing most exons and introns, and sel-12(ty11), which contains a premature stop codon (Cinar et al., 2001). Compared to wild type, both mutants exhibited significantly reduced ePSC amplitudes and mini frequencies, with no change in the mean amplitude of minis (Figure 1A, B). These findings indicate that SEL-12 is essential for normal synaptic transmission.

Figure 1. Presynaptic deficiency underlies neuromuscular synaptic transmission defects in sel-12 mutants.

Figure 1.

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A. Representative ePSC and mini traces recorded from body-wall muscle cells. B. Quantification of ePSC amplitudes, mini frequencies, and mini amplitudes across the indicated genotypes. Prab-3 and Pmyo-3 were used for neuron- and muscle-specific expression of wild-type SEL-12, respectively. C. Chromatograms of the sel-12(zw101) allele showing 5 bp insertion in exon 1 and the resulting premature stop codon (*). D. Representative traces of muscle whole-cell currents elicited by pressure-ejected acetylcholine (ACh, 100 µM)) or GABA (100 µM) in wild type and sel-12(zw101) animals, along with statistical comparisons of current amplitudes. Asterisks indicate statistically significant differences compared to wild type (*p < 0.05; **p < 0.01, one-way ANOVA, Fisher’s LSD post hoc test). Sample sizes are shown within bars.

Because these mutants were generated by chemical mutagenesis and may carry additional background mutations not easily removed by backcrossing, we generated a new sel-12 allele, sel-12(zw101), using CRISPR-Cas9. This allele has a 5-base pair insertion in the first exon, leading to a frameshift and premature stop codon (Figure 1C), and displayed synaptic phenotypes similar to those of the original mutants (Figure 1A, B). Furthermore, expression of a fosmid (WRM0633C) containing the full sel-12 gene, including its 5′- and 3′-regulatory regions, rescued the synaptic defects in sel-12 mutants (Figure 1A, B), providing additional evidence that SEL-12 regulates synaptic transmission.

To identify the site of SEL-12 function, we performed cell-specific rescue experiments in sel-12(zw101) mutants by expressing wild-type sel-12 under either the pan-neuronal Prab-3 (where “P” indicates promoter) and the muscle specific Pmyo-3. Pan-neuronal expression fully rescued the synaptic defects, while muscles-specific expression showed no effect (Figure 1A, B). In addition, neuron-specific knockdown of sel-12 using Prab-3 recapitulated the mutant phenotype (Figure 1A, B), while postsynaptic (muscle) responses to exogenous acetylcholine and GABA were comparable between wild-type and sel-12(zw101) animals (Figure 1D). Together, these findings indicate that SEL-12 acts presynaptically to regulate neurotransmitter release at the NMJ, consistent with the presynaptic roles of presenilins in the mouse hippocampus (Zhang et al., 2009; Wu et al., 2013).

C. elegans has three presenilin-like genes: sel-12, spe-4, and hop-1. Among these, SEL-12 shares 61-63% identity and 79% similarity with human PS1 and PS2, whereas HOP-1 and SPE-4 show lower identity (~34% and ~25%, respectively) and similarity (~56% and ~50%, respectively). SEL-12 is therefore considered the ortholog of human presenilins based on sequence homology. To test whether the presynaptic function of presenilin is specific to SEL-12, we examined hop-1(ar179), a lf mutant caused by a missense mutation (R179H) (Agarwal et al., 2018). This mutant was indistinguishable from wild type in synaptic function (Figure 1A, B). spe-4 was excluded from our electrophysiological analysis because it is exclusively expressed in the germline (Gosney et al., 2008). The lack of synaptic defects in hop-1 mutants indicates that SEL-12 plays a unique role in regulating neurotransmitter release at the C. elegans NMJ.

SEL-12 functions in AVA command interneurons to regulate neuromuscular transmission

To test whether SEL-12 acts cell-autonomously in ventral cord motor neurons to regulate neurotransmitter release, we performed rescue and knockdown experiments targeting these neurons. For rescue, we used a chimeric promoter composed of the distal region of the unc-17 (vesicular acetylcholine transporter) promoter and the unc-47 (vesicular GABA transporter) promoter, which drives expression in both cholinergic and GABAergic motor neurons (Chen et al., 2017). For knockout, we used Punc-4, which is specific to A-type cholinergic motor neurons (Miller and Niemeyer, 1995) that are directly controlled by AVA command interneurons through chemical and electrical synapses (White et al., 1986; Liu et al., 2017). Surprisingly, motor neuron-targeted expression of wild-type SEL-12 in sel-12(zw101) mutants, or motor neuron-targeted knockdown of endogenous sel-12 in wild-type animals, did not alter ePSC or mini properties (Figure 2A, B), indicating that the synaptic phenotypes in sel-12 mutants are not due to loss of SEL-12 in motor neurons.

Figure 2. SEL-12 regulates neuromuscular transmission by acting in AVA interneurons.

Figure 2.

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A. Representative ePSC and mini traces from body-wall muscle cells of indicated genotypes. Cell-specific sel-12 rescue or knockdown used promoters targeting cholinergic and GABAergic motor neurons (Punc-17;unc-47), command interneurons (Pglr-1: AVA, AVD, AVE, PVC), or AVA alone (Cre-LoxP with Pflp-18 and Pgpa-14). B. Quantification of ePSC amplitudes, mini frequencies, and mini amplitudes.. C. Representative ePSC and mini traces from AVA-specific HisCl1 strain: untreated (Control) vs. histamine (10 µM). D. Comparison of mini and ePSC properties between Control and Histamine. E. Sample traces of minis recorded from body-wall muscle cells in wild-type and sel-12(zw101) mutant animals expressing channelrhodopsin-2 in AVA, with or without all-trans retinal pretreatment. The blue horizontal line above each trace indicates the period of blue light stimulation. F. Quantification of mini frequency and total charge transfer during the 20-s blue light stimulation (“Light on”) and the preceding 20-sec control period (“Light off”). *p < 0.05, **p < 0.01 compared with wild type (panel B, one-way ANOVA, Fisher’s LSD post hoc test) or with the control group (panels D and F, unpaired t-test). Sample sizes are shown within bars.

We next asked whether SEL-12 functions in upstream interneurons to regulate neuromuscular transmission. In C. elegans, five pairs of command interneurons, including AVA, AVB, AVD, AVE, and PVC, control cholinergic motor neurons through chemical and electrical synapses (de Bono and Maricq, 2005). Among them, AVB communicates exclusively through electrical synapses, while AVA uses both chemical and electrical synapses, and AVD, AVE, and PVC use only chemical synapses (de Bono and Maricq, 2005).

We found that expressing wild-type sel-12 under Pglr-1, which is active in all command interneurons except AVB (Pereira et al., 2015), fully rescued the transmission defects in sel-12(zw101) mutants. Conversely, Pglr-1-driven knockdown of endogenous sel-12 in wild-type animals recapitulated the mutant synaptic phenotype at the NMJ (Figure 2A, B). These results indicate that SEL-12 functions in command interneurons to regulate neuromuscular transmission.

Among these interneurons, AVA plays a central role in controlling ventral cord A-type cholinergic motor neurons (A-MNs) (Liu et al., 2017). Although AVD and AVE also form chemical synapses with A-MNs (de Bono and Maricq, 2005), only AVA generates the spontaneous postsynaptic currents observed in these motor neurons (Liu et al., 2017). To test whether SEL-12 acts specifically in AVA, we silenced sel-12 specifically in AVA of wild-type animals using a Cre-loxP system driven by two promoters with overlapping expression in AVA (Schmitt et al., 2012). These animals showed mini and ePSC phenotypes indistinguishable from those of sel-12 mutants (Figure 2A, B). Together, these findings indicate that SEL-12 functions in command interneurons, specifically in AVA, to regulate neurotransmitter release from motor neurons.

It was unexpected that SEL-12 functions in AVA rather than in motor neurons, given that minis and ePSCs at the NMJ are typically thought to result from spontaneous exocytosis and direct stimulation of motor neurons, respectively. To further confirm the role of AVA in neuromuscular transmission, we acutely inhibited AVA using histamine in a strain expressing the histamine-gated chloride channel HisCl1 specifically in AVA (Pokala et al., 2014). Histamine application significantly reduced both ePSC amplitudes and mini frequencies without affecting mini amplitudes at the NMJ (Figure 2C, D), confirming that AVA activity is critical for normal neuromuscular transmission. These results suggest that tonic input from AVA elevates motor neuron excitability, thereby enhancing mini frequency, and that electrical stimulation of the ventral nerve cord evokes ePSCs through stimulation of the neuropils of both motor neurons and AVA, with the latter influencing ePSCs indirectly via cholinergic motor neurons.

To determine whether optogenetic activation of AVA could compensate for sel-12(lf) effects, we compared optogenetically evoked postsynaptic currents at the NMJ between wild-type and sel-12(zw101) strains expressing channelrhodopsin-2 in AVA neurons. Following pre-incubation with all-trans retinal (2.0 mM, 24-30 hours), wild-type animals showed significant increases in both mini frequency and total charge transfer upon blue light stimulation compared to their vehicle control group (Figure 2E,F). In contrast, sel-12 mutants exhibited no such increases, indicating that AVA hyperactivation cannot overcome the synaptic transmission deficits caused by sel-12(lf).

sel-12 is expressed in command interneurons but not in motor neurons

The findings described above prompted us to examine the expression pattern of sel-12 using two complementary approaches. First, we generated a transcriptional reporter by expressing mScarlet under the control of Psel-12 using an in vivo homologous recombination strategy. This approach involved co-injecting a plasmid containing 518 bp of Psel-12 fused to mScarlet with a fosmid (WRM0633C) encompassing the full sel-12 gene (including the promoter and 3′ UTR) and neighboring genes. Homologous recombination between the plasmid and fosmid was expected to produce a Psel-12::mScarlet fusion that preserved all regulatory sequences upstream of the sel-12 start codon. In transgenic animals, we observed mScarlet fluorescence in a small number of head neurons and one pair of tail neurons, but not in ventral cord motor neurons (Figure 3A).

Figure 3. Expression of sel-12 in command interneurons.

Figure 3.

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sel-12-expressing neurons were visualized by driving mScarlet expression under Psel-12. GFP was coexpressed to label specific neuronal subtypes: command interneurons (AVA, AVD, AVE, PVC) using Pglr-1; AVA specifically via Cre-LoxP recombination with Pflp-18 and Pgpa-14; or mechanosensory neurons using Pmec-4. A. Fluorescence and corresponding DIC images of anterior and posterior regions of a transgenic animal expressing Psel-12::mScarlet. B. Fluorescence and DIC images of an animal coexpressing Psel-12::mScarlet and Pglr-1::GFP. Only AVA and PVC interneurons are labeled. C, D. Fluorescence and DIC images of animals coexpressing Psel-12::mScarlet and GFP, with GFP expression restricted to AVA (C) or to mechanosensory neurons, including PLM (D). Scale bar: 60 µm in a, 30 µm in b-d (all panels).

To determine whether the mScarlet-positive neurons included command interneurons, we introduced a Pglr-1::GFP transgene into the strain carrying the Psel-12::mScarlet reporter. In this double-transgenic strain, we observed overlapping expression of mScarlet and GFP in several head neurons, including AVA, and in one pair of tail neurons corresponding to PVC (Figure 3B). This co-expression pattern indicates that sel-12 is expressed in command interneurons.

To confirm sel-12 expression in AVA specifically, we crossed the Psel-12::mScarlet reporter into a strain that expresses GFP exclusively in AVA (strain ID: ZX1023) (Schmitt et al., 2012). In the resulting animals, mScarlet and GFP signals colocalized in a pair of head neurons, confirming that sel-12 is expressed in AVA (Figure 3C). This expression pattern is consistent with our electrophysiological findings and supports the role of SEL-12 in command interneurons.

We also determined whether sel-12 is expressed in PLM mechanosensory neurons by introducing a Pmec-4::GFP transcriptional reporter into the Psel-12::mScarlet strain. Co-labeling of PLM by GFP and mScarlet (Figure 3D) confirmed that sel-12 is expressed in PLM neurons.

γ-Secretase activity is not required for SEL-12’s synaptic function

To determine whether SLE-12’s proteolytic activity is required for its role in synaptic transmission, we tested whether substitution of a key aspartate residue in SEL-12’s active site (D226) by alanine impairs its synaptic function. D226 corresponds to D257 in human PS1, which is required for PS1’s γ-secretase activity (Wolfe et al., 1999; Xia et al., 2002; Kimberly et al., 2003; Kim et al., 2005). The critical role of D226 in SEL-12’s γ-secretase activity was demonstrated by a knock-in D226A mutation that produced phenotypes indicative of impaired γ-secretase-dependent Notch signaling, including severe egg-laying defects and protruding vulva (Ashkavand et al., 2020). This mutation serves as a useful tool to evaluate whether specific SEL-12 functions depend on its γ-secretase activity (Sarasija et al., 2018; Ashkavand et al., 2020; Ryan et al., 2024). We found that neuronal expression of sel-12(D226A) fully rescued the defective minis and ePSCs in sel-12(zw101) worms (Figure 2A, B), indicating that γ-secretase activity is not required for SEL-12’s role in regulating neurotransmitter release.

Human PS1 can substitute for SEL-12 in synaptic function

The high degree of amino acid sequence homology between SEL-12 and human PS1 led us to test whether human PSEN1 (the gene encoding PS1) could rescue synaptic phenotypes in C. elegans sel-12 mutants. Pan-neuronal expression of human PSEN1 driven by Prab-3 fully restored minis and ePSCs in sel-12(zw101) mutants (Figure 2A, B). This functional rescue demonstrates conserved roles for SEL-12 and human PS1 in synaptic regulation.

SEL-12 and RYR-1 function together to regulate synaptic transmission

The defects in minis and ePSCs observed in sel-12 mutants resemble those we previously reported in ryr-1 mutants (Liu et al., 2005; Chen et al., 2017; Mueller et al., 2023). These mutant phenotypes arise from presynaptic rather than postsynaptic deficiencies, as they are rescued by neuronal expression of wild-type ryr-1 using Prab-3, but not by expression in body-wall muscle cells using Pmyo-3 (Liu et al., 2005). The similarity in phenotypes, together with the findings that cDKO of Psen1 and Psen2 in mouse impairs neurotransmitter release via RyRs (Zhang et al., 2009; Wu et al., 2013), promoted us to investigate whether SEL-12 and RYR-1 functionally interact to regulate synaptic transmission. To this end, we analyzed two different ryr-1 alleles, e540 and syb216, as well as their double mutants with sel-12(ok2078) or sel-12(zw101). The ryr-1(e540) allele carries a premature stop codon (Sakube et al., 1997; Mueller et al., 2023), while the ryr-1(syb216) allele selectively abolishes neuronal ryr-1 expression due to a deletion of the first exon and the preceding promoter of neuron-specific isoforms of ryr-1 (Marques et al., 2020). Both ryr-1 mutants exhibited similar reductions in ePSC amplitude and minis frequency as sel-12 mutants, and these effects were non-additive in sel-12;ryr-1 double mutants (Figure 4A, B and Figure 5A, B). Intriguingly, the mean amplitude of minis was significantly decreased in the ryr-1 single mutants as well as their double mutants with sel-12, but not in sel-12 single mutants (Figure 4A, Figure 5A, B). The non-additive mutant effects on ePSC amplitude and mini frequency indicate that SEL-12 and RYR-1 function in a common molecular pathway to regulate neurotransmitter release. However, the additional effect of ryr-1(lf) on mini amplitudes suggests a mechanistic difference between sel-12 and ryr-1 mutants with respect to neuromuscular synaptic transmission.

Figure 4.

Figure 4.

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SEL-12 and RYR-1 function together to regulate neuromuscular synaptic transmission. Knockout of ryr-1 in AVA, AVD, AVE, and PVC interneurons was driven by Pglr-1; AVA-specific knockout was achieved using a Cre-LoxP approach with Pflp-18 and Pgpa-14; and motor neuron (MN)-specific knockout was driven by Punc-4. A. Representative ePSC and mini traces recorded from body-wall muscle cells. B. Quantification of ePSC amplitudes, mini frequencies, and mini amplitudes. Asterisks indicate statistically significant differences compared to wild type (**p < 0.01; one-way ANOVA, Fisher’s LSD post hoc test). Sample sizes are shown within bars.

Figure 5.

Figure 5.

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SEL-12 and RYR-1 function together to regulate synaptic transmission at the neuromuscular junction. A. Representative traces of ePSCs and minis recorded from body-wall muscle cells. B. Quantification of ePSC amplitudes, mini frequencies, and mini amplitudes. * p < 0.05, ** p < 0.01 compared to wild type and #p < 0.05 compared between the indicated groups (one-way ANOVA, Fisher’s LSD post hoc test). Sample sizes are shown within bars.

Cellular expression differences underlie differential mutant effects on mini amplitudes

Since sel-12 is expressed primarily in command interneurons while ryr-1 is broadly expressed in the nervous system (Chen et al., 2017), we hypothesized that the distinct effects of sel-12(lf) and ryr-1(lf) mutations on mini amplitudes arise from differences in their motor neuron expression. To test this, we analyzed synaptic transmission at the NMJ in strains with ryr-1 knockdown targeted to: (1) multiple command interneurons (including AVA), (2) AVA alone, (3) A-MNs specifically, or (4) both AVA and A-MNs. All strains exhibited reduced mini frequencies and ePSC amplitudes compared to wild type; however, only strains with ryr-1 knockdown in A-MNs (either alone or with AVA) showed a significant decrease in mean mini amplitude (Figure 4A, B). These results demonstrate that RYR-1 regulates mini amplitude cell-autonomously in motor neurons, and that the differential effects of sel-12 and ryr-1 mutants reflect their distinct expression patterns.

SEL-12 and RYR-1 function together to maintain neuronal health

The similar synaptic defects observed in ryr-1(lf) and sel-12(lf) mutants promoted us to examine whether ryr-1(lf) also leads to a neurodegenerative phenotype, as previously shown for sel-12(lf) (Sarasija et al., 2018; Parvand et al., 2024), and whether these effects are additive in sel-12;ryr-1 double mutants. To answer these questions, we analyzed the morphology of the AVA posterior (relative to the vulva) axon and the PLM distal axon (Figure 6A). AVA was chosen because it expresses sel-12 and is presynaptic to A-MNs, while PLM is known to exhibit neurodegenerative phenotypes in sel-12(lf) mutants (Sarasija et al., 2018). AVA neurons were specifically labeled using an integrated mStrawberry transgene (Liu et al., 2020), and PLM neurons were labeled with GFP driven by Pmec-4 (Wu et al., 2007). Compared to wild type, both sel-12(zw101) and ryr-1(syb216) mutants displayed defective axonal morphologies characterized by breaks and/or bead-like swellings. These defects were not further exacerbated in the sel-12;ryr-1 double mutant (Figure 6B, C). These findings indicate that neuron-specific ryr-1(lf) is sufficient to induce neurodegenerative phenotypes and support the idea that the neurodegenerative effects of sel-12(lf) may result, as least in part, from impaired RyR-1-mediated Ca2+ release from the ER.

Figure 6.

Figure 6.

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Loss of sel-12 or ryr-1 leads to axonal degeneration with non-additive effects in double mutants. A. Diagram showing the AVA and PLML neuron positions and the analyzed axonal segments (enclosed by dashed-line squares). B. Representative images of AVA and PLM axons across genotypes. PLM neurons were labeled with GFP under Pmec-4, and AVA interneurons with mStrawberry via Pflp-18 and Ppga-14 in a Cre-LoxP strategy. Scale bar = 20 µm. C. Quantification of axonal defects (breaks or beads/swellings) in AVA and PLM neurons. Asterisks denote statistically significant differences compared to wild type (** p < 0.01, *** p < 0.001; one-way ANOVA, Fisher’s LSD post hoc test). Sample sizes in bars.

ryr-1 transcription and translation are not altered by sel-12 mutation

To determine whether sel-12(lf) affects ryr-1 transcription or RYR-1 protein levels, we used two strategies: (1) expression of a GFP reporter driven by the neuron-specific Pryr-1neuronal (Chen et al., 2017), (2) tagging of endogenous RYR-1 with GFPnovo2. The GFPnovo2 knock-in strain, ryr-1(syb7697[GFPnovo2]), was generated by inserting GFPnovo2 between amino acids V1392 and R1393 (as numbered in the RYR-1b isoform) into all RYR-1 isoforms in a wild-type background. This site was chosen based on a prior study showing that GFP insertion at the corresponding position in mouse RyR2 does not impair channel function (Hiess et al., 2015). Consistent with this, we observed no differences in ePSC and mini properties at the NMJ between the knock-in and wild-type strains (Figure 7A), indicating that RYR-1 function is preserved in the knock-in strain.

Figure 7.

Figure 7.

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Loss of sel-12 does not alter ryr-1 transcription or RYR-1 protein levels. A. Representative traces of ePSCs and minis, and quantification of their amplitudes and frequencies in wild-type and RYR-1::GFPnovo2 animals. “ns” indicates no statistically significant difference (unpaired t-test). B. Fluorescence and DIC images showing fluorescence signals in head neurons of wild-type and sel-12(zw101) animals expressing GFP using the neuron-specific Pryr-1, and quantification of fluorescence intensity. C. Fluorescence and DIC images showing fluorescence signals in the head region of wild-type and sel-12(zw101) animals with endogenous RYR-1 tagged by GFPnovo2. Sample sizes are shown within bars. Scale bar = 20 µm in B and C.

We next crossed both Pryr-1neuronal::GFP and ryr-1(syb7697[GFPnovo2]) from their original wild-type backgrounds into the sel-12(zw101) mutant to generate two independent strains for fluorescence comparison. In both cases, GFP intensity was comparable between wild-type and sel-12(zw101) animals (Figure 7B, C), suggesting that sel-12(lf) does not significantly affect ryr-1 transcription or RYR-1 protein expression. However, it is important to note that the GFPnovo2 signal was predominantly observed in body-wall muscle cells rather than neurons.

Discussion

Our findings establish that SEL-12 and RYR-1 act synergistically to maintain synaptic transmission and neuronal integrity in C. elegans, with loss of either gene inducing non-additive synaptic deficits and axonal degeneration. These results illuminate a conserved, γ-secretase-independent pathway linking presenilin dysfunction to RyR-mediated Ca2+ dysregulation, a mechanism with direct implications for age-related neurodegenerative processes. While our morphological analyses focused on axons, the observed degeneration may reflect broader neuronal vulnerability that escalates with aging. The premature lethality of sel-12(lf) mutants under standard conditions, driven by internal hatching within the parental body and subsequent rupture, precluded direct analysis of age-dependent progression. Nonetheless, the convergence of synaptic and structural deficits in young adults highlights the essential role of presenilin-RyR interplay in sustaining neuronal resilience.

Notably, sel-12(lf) did not alter RYR-1::GFP expression in muscle cells, contrasting with murine studies where presenilin loss reduced RyR protein levels in neurons (Wu et al., 2013). Our data suggest SEL-12 likely regulates RyR function rather than abundance—consistent with evidence that PS1 fragments directly modulate RyR gating (Payne et al., 2013). However, this conclusion was based on RYR-1::GFP fluorescence in muscle cells, as neuronal RYR-1::GFP fluorescence was not assayed due to low expression. Crucially, synaptic defects in sel-12(lf) mutants were rescued by human PSEN1, demonstrating evolutionary conservation of presenilin’s synaptic role. They were also rescued by a γ-secretase-inactive SEL-12 variant, indicating that this function is independent of amyloidogenic processing. These findings challenge Aβ-centric models of FAD.

Although intracellular Ca2+ accumulation is often linked to presenilin mutation-associated neurodegeneration (Bezprozvanny and Mattson, 2008), our study shows that ryr-1(lf) alone induces neurodegeneration. This finding is consistent with previous reports that FAD-linked mutations in PSEN1 and PSEN2 decrease, rather than increase, cytosolic Ca2+ levels (Zatti et al., 2004; Giacomello et al., 2005; Zatti et al., 2006), which prompted calls to revise the Ca2+ overload hypothesis of AD pathogenesis (Giacomello et al., 2005). How might presenilin mutations cause neurodegeneration through reduced cytosolic Ca2+? Evidence from this and previous studies (Zhang et al., 2009; Wu et al., 2013) points to impaired synaptic transmission as one plausible mechanism. Synaptic dysfunction is considered a major determinant of AD (Lepeta et al., 2016) and disease onset is thought to involve early changes in synaptic efficacy followed by neuronal degeneration (Selkoe, 2002). Conceivably, defective synaptic function could trigger neurodegeneration through several possible pathways, including reduced neurotrophic support, impaired synaptic plasticity and maintenance, and excitotoxicity from an imbalance between excitatory and inhibitory synaptic inputs.

Mechanistically, SEL-12 functions in AVA command interneurons to trans-synaptically regulate neuromuscular transmission. Our previous study demonstrated the crucial role of AVA command interneurons in controlling postsynaptic currents at the NMJ via cholinergic motor neurons (Liu et al., 2017). The trans-synaptic effects observed here suggest that neurodegenerative changes can influence downstream neurons that are not directly connected to the degenerating neurons. Conceivably, such trans-synaptic effects could disrupt network homeostasis and thereby accelerate cognitive decline.

In summary, we identify an evolutionarily conserved mechanism in which presenilin maintains synaptic fidelity and axonal integrity through RyR-dependent Ca2+ signaling, independent of its γ-secretase activity. This work reframes presenilin-linked FAD as a disorder of Ca2+ dysregulation and highlights RyR modulators and presenilin interactors as potential therapeutic targets for AD..

Supplementary Material

Tables S1 and S2

Significance Statement.

Mutations in presenilins are the major cause of familial Alzheimer’s disease and are commonly associated with impaired synaptic transmission and neurodegeneration. However, the molecular mechanisms underlying these effects remain poorly understood. This study shows that loss of presenilin in C. elegans impairs neurotransmitter release and causes axonal degeneration through dysfunction of ryanodine receptors (RyRs), independent of presenilin’s γ-secretase activity. Notably, RyR expression remains unchanged, suggesting that presenilins likely regulate RyR function. These findings uncover a γ-secretase-independent pathway linking presenilin dysfunction to synaptic and neuronal deficits. The findings of this study offer new insight into the pathogenesis of Alzheimer’s disease.

Acknowledgements

This work was supported by NIH grant R01MH085927. We thank Dominique A. Glauser for the ryr-1(syb216) strain, Cori Bargmann for the CX14865 strain, Alexander Gottschalk for the ZX1023 strain, Max Planck Institute for the fosmid WRM0633C, DNASU Plasmid Repository for the human presenilin 1 cDNA, and Caenorhabditis Genetics Center for the sel-12(ok2078), sel-12(ty11), hop-1(ar179), ryr-1(e540), and zdIs5[Pmec-4::GFP] strains.

Footnotes

Conflict of Interest Statement

The authors declare no competing financial interests.

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Supplementary Materials

Tables S1 and S2
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