A brain-gut excitatory peptide/CCHamide homolog regulates satiation and motivational state transitions in the Aplysia feeding circuit

TL;DR

This study shows how a brain-gut peptide in Aplysia controls satiation and shifts between feeding behaviors.

Contribution

The paper identifies a novel EP/CCHa signaling pathway in Aplysia and its role in regulating feeding circuits.

Findings

apEP/CCHa inhibits food intake and shifts motor programs from ingestion to egestion.

apEP/CCHa modulates the excitability of key feeding interneurons in the central pattern generator.

Two apEP/CCHa receptors were identified, with distinct phylogenetic origins and expression patterns.

Abstract

Excitatory peptide (EP) and CCHamide (CCHa) are protostome neuropeptides originally identified in lophotrochozoans (including annelids and mollusks) and arthropods, respectively, and are homologous to the deuterostome endothelin (ET) and gastrin-releasing peptide (GRP)/neuromedin-B (NMB) systems. These peptides are brain-gut peptides: in vertebrates, GRP/NMB function as satiety peptides, whereas arthropod CCHa displays species-specific actions, either inhibiting or promoting feeding. However, the mechanisms by which these peptides modulate feeding circuits remain unknown. Here, we investigated the EP/CCHa signaling pathway in Aplysia, a mollusk with a well-defined feeding circuit. We identified a single precursor encoding Aplysia EP/CCHa (apEP/CCHa). Mass spectrometry demonstrated that an apEP/CCHa peptide is present in the central ganglia. In situ hybridization and immunohistochemistry revealed apEP/CCHa-positive neurons in the CNS, immunopositive cells in the gut, and immunopositive fibers in the gut-innervating esophageal nerve. To identify potential targets, we cloned two novel apEP/CCHa receptors. Phylogenetically, one receptor clusters with lophotrochozoan EP/CCHa receptors, whereas the other unexpectedly clusters with arthropod receptors, suggesting independent lineages for the two receptors. Single-cell RNA sequencing showed that both receptors are expressed in the key feeding central pattern generator (CPG) interneurons B20 and B34. Functionally, apEP/CCHa inhibited food intake in vivo and converted ingestive motor programs to egestive ones in vitro. At the circuit level, apEP/CCHa modulated excitability of B20 and B34, and two additional interneurons (B40, B64). In summary, we demonstrate that apEP/CCHa is a brain–gut peptide that functions as a satiation signal, and identify specific feeding CPG elements through which apEP/CCHa regulates motivational state transitions.

Figures

Full text

Neuropeptides, by acting on their G-protein coupled receptors (GPCR), modulate neural networks to regulate the expression of multiple behaviors (1, 2, 3, 4, 5, 6, 7, 8). A prominent subset of peptides is referred to as “brain-gut” because they are expressed in both the central nervous system (CNS) and the gut. Some of these peptides function as satiation or satiety signals. They are released following food intake and subsequently inhibit feeding. In mammals, Peptide YY (PYY) and Cholecystokinin (CCK) are well-established examples (9, 10, 11, 12). Data indicate that CCK (13) and neuropeptide Y (NPY) (14) have a similar function in the mollusk Aplysia californica. Despite this progress, additional satiation/satiety peptides are likely to exist. Here, we present evidence that a newly identified excitatory peptide (EP)/CCHamide (CCHa) in Aplysia constitutes one such candidate.

EP was initially discovered in earthworms Eisenia foetida and Pheretima vittate (15). Subsequently, it was identified in other annelids (16, 17, 18, 19) and several mollusks (20, 21). It was named for its myo-excitatory effects (15, 16, 20). CCHa was independently identified in the arthropod Bombyx mori (22), and later in other arthropods, including Drosophila melanogaster (23). EP and CCHa share key structural features—most notably a disulfide bond and a C-terminal amidation—and are widely regarded as homologous (24, 25, 26, 27, 28, 29). Accumulating evidence indicates that arthropod EP/CCHa peptides function as brain-gut peptides that regulate feeding, diuresis, reproduction, metabolic homeostasis, and circadian rhythms (23, 30, 31, 32, 33, 34). Interestingly, their effects on feeding are species-specific: in Drosophila, they promote food intake (31, 35), whereas in Aedes aegypti (36) and Gryllus bimaculatus (37), they suppress it. Within lophotrochozoans—including annelids and mollusks—the widespread expression of EP/CCHa in both CNS and digestive tissues (16, 17, 19, 20, 38, 39) strongly suggests a comparable regulatory role in feeding, although this has not been functionally demonstrated.

Subsequent bioinformatic analyses showed that protostome EP/CCHa signaling systems are homologous to mammalian neuromedin-B (NMB)/gastrin-releasing peptide (GRP) and endothelin (ET)-related signaling families, based primarily on receptor homology rather than ligand similarity (26, 27, 28, 29, 40, 41, 42). Among deuterostomes, the peptide bombesin (BN; pQQRLGNQWAVGHLM-NH_2_) was first isolated from the amphibian skin (43, 44), and an orthologous peptide was recently identified in an echinoderm starfish, Asterias rubens (42). Because of similarities in peptide structure and receptor homology, BN signaling in non-mammalian species is considered an ancestral counterpart of mammalian GRP/NMB systems. Consequently, GRP/NMB are also referred to as BN-type peptides (45, 46, 47, 48, 49). Current evidence suggests BN-type peptides inhibit feeding and operate as satiation/satiety peptides (50, 51, 52). Similarly, A. rubens bombesin (ArBN) inhibits feeding (42). Taken together, these evolutionary and functional considerations led us to hypothesize that molluscan EP/CCHa peptides function as satiation/satiety signals.

To test this hypothesis, we sought to study the EP/CCHa signaling system in the experimentally advantageous molluscan model Aplysia. This species has been the focus of extensive neurobiological research, and both its feeding behaviors and the underlying neural circuit are well characterized (53, 54, 55, 56, 57, 58, 59, 60, 61). The accessibility of identified neurons and well-defined central pattern generator (CPG) circuits enables mechanistic analyses that are not feasible in some other organisms, including direct assessments of how peptides modulate CPG activity.

Here, we characterized one apEP/CCHa precursor and two cognate receptors in Aplysia. We characterized the primary structure of the bioactive neuropeptide by tandem mass spectrometry, mapped the distribution of apEP/CCHa-expressing neurons in the CNS and gut using in situ hybridization and immunohistochemistry, and showed that apEP/CCHa is a bona fide brain-gut peptide. We then demonstrated that it inhibits food intake in vivo and promotes egestive motor activity in vitro. Using single-cell RNA sequencing, we localize the two receptors to the CPG interneurons, B20 and B34 and electrophysiological recordings show that apEP/CCHa modifies the excitability of these and other interneurons to implement its network effects. This work provides a comprehensive functional characterization of an EP/CCHa neuropeptide system in mollusks and supports the conclusion that these peptides act as a satiation/satiety signal, illuminating the neural mechanisms through which they influence feeding behavior.

Results

Identification of the EP/CCHa precursor in Aplysia

To identify a precursor for Aplysia EP/CCHa (apEP/CCHa), we used two EP precursors from the mollusk Thais clavigera (BAQ25802.1 and BAQ25803.1) (38) to perform BLAST searches. An initial BLAST search in NCBI yielded no significant matches. However, a BLAST in the AplysiaTools database did return an mRNA sequence (TRINITY_DN9791_c1_g1_i6) (Fig. 1A and Table S1).Figure 1Gene expression mapping and cloning of the Aplysia EP/CCHa precursor. A, a genome sequence from AplysiaTools (HiC_scaffold_6:45471172–46843127) transcribes an mRNA (TRINITY_DN9791_c1_g1_i6). It encodes a previously uncharacterized protein, which is similar to EP/CCHa from other lophotrochozoans. B, a PCR product for apEP/CCHa precursor (apEP/CCHa pre) gene with a length of 426 bp. Lane 1: DNA marker (M); Lane 2 and lane 3: the target gene. C, the complete protein sequence of apEP/CCHa precursor illustrating the signal peptide (green) and a predicted peptide (red). The predicted mature peptide undergoes two post-translational modifications: amidation of the terminal glycine residue at the C-terminus (purple) and the formation of a disulfide bond between the two cysteine residues (blue). The dibasic cleavage site (KR) is highlight in yellow.

To verify the presence of this transcript in Aplysia, we designed primers (Table S2) based on the putative sequence and performed PCR on complementary DNA (cDNA) from the Aplysia CNS. This produced a 426 bp product (Fig. 1B) identical to the coding sequence (CDS) of TRINITY_DN9791_c1_g1_i6.

SignalP 5.0 predicted that the putative apEP/CCHa precursor contains a 29-amino acid signal peptide with a cleavage site between Ala^29^ and Lys^30^ (Fig. 1C). We used NeuroPred (62) to predict peptides encoded by this precursor and found that it encodes one EP/CCHa-like peptide: apEP/CCHa (KCHGRWAIHACFGGN-NH_2_) (Fig. 1C). Similar to EP/CCHa peptides in other lophotrochozoans, apEP/CCHa contains two cysteine residues with eight amino acids in between. We also compared Aplysia EP/CCHa to EP/CCHa in other protostome species (Fig. 2 and Table S3). These results showed that the two cysteines are present in all protostome species, which could potentially form disulfide bonds. In addition, there is a conserved C-terminal GGNamide motif in most lophotrochozoans, whereas it is a GGHamide or GAHamide motif in arthropods.Figure 2Comparison of EP/CCHa peptides from protostomes. A, comparison of selected EP/CCHa peptides from protostomes using BioEdit (ClustalW Multiple Alignments-Graphic View). “#” indicates that the peptide has been verified. B, a frequency plot for these sequences using Weblogo v2.8.2 (http://weblogo.berkeley.edu/logo.cgi). C-terminal amidation and the formation of a disulfide bond between two cysteine residues are highly conserved across protostome species. See Table S3 for the information about the sequences. -S-S-: disulfide bridge; -NH_2_: the C-terminal amidation. Note that EP/CCHa peptides purified and chemically identified from Eisenia foetida and Pheretima vittata (15) are the only ones characterized by non-amidated C-termini.

Distribution of apEP/CCHa precursor mRNA and peptides in the Aplysia CNS

To map the distribution of the apEP/CCHa precursor mRNA in the Aplysia CNS, we performed in situ hybridization experiments. We observed apEP/CCHa mRNA-positive neurons in all central ganglia, i.e., in the buccal (Fig. 3, A and B), cerebral (Fig. 3, C and D), pleural-pedal (Fig. S1, A–D), and abdominal ganglia (Fig. S1, E and F).Figure 3Detection of apEP/CCHa peptide in the CNS: positive neurons in buccal/cerebral ganglia (whole mounts) and mass spectrometry validation. In situ hybridization: A and B, the rostral (A) and caudal (B) surface of a buccal ganglion. C and D, the dorsal (C) and ventral (D) surface of a cerebral ganglion. Immunohistochemistry: E and F, the rostral (E) and caudal (F) surface of a buccal ganglion. The ganglion was photographed with a coverslip. Arrows in each surface point to the cells that are more in focus on that surface. G and H, the dorsal (G) and ventral (H) surface of a cerebral ganglion. Scale bar: 200 μm. All panels have the same magnification; Each experiment was performed 3 times and representative results are shown. Buccal abbreviations: N1, nerve 1; N2, nerve 2; N3, nerve 3; EN, esophageal nerve; RN, radula nerve; CBC, cerebral-buccal connective. Cerebral abbreviations: UL, upper labial nerve; PT, posterior tentacular nerve; AT, anterior tentacular nerve; CBC, cerebral-buccal connective; CPe, cerebral-pedal connective; CPl, cerebral-pleural connective. I, the tandem MS spectrum for apEP/CCHa neuropeptide in central ganglia. The tandem MS spectrum collected from a CNS extract is shown on top, and the predicted theoretical spectrum based on the amino acid sequence of apEP/CCHa peptide is shown on the bottom, both illustrating matching b- and y-ion series in blue and red, respectively. Key identification metrics, including the confidence identification score and accuracy of detection, are summarized in Table S5 to confirm apEP/CCHa peptide detection and identification via mass spectrometry.

Subsequently, we used immunohistochemistry experiments to further determine whether the apEP/CCHa peptide was expressed in the CNS of Aplysia, and identified apEP/CCHa-positive neurons (Fig. 3, E–H and Fig. S1, G–L). In the buccal ganglion, there are apEP/CCHa-positive neurons, some of which are large, on both rostral and caudal surfaces (Fig. 3, E and F). In the cerebral ganglion, we observed several apEP/CCHa-positive neurons on both dorsal and ventral surfaces (Fig. 3, G and H). Based on their locations, some neurons in the cerebral ganglion may be cerebral-buccal interneurons (CBIs), which are higher-order neurons essential for feeding initiation and modulation (58, 63, 64, 65, 66, 67, 68, 69, 70). The buccal ganglion contains feeding motoneurons and pattern-generating interneurons.

In addition to the staining observed in the somata of neurons, immunostaining also allowed us to observe apEP/CCHa-positive axons extending through the anterior and posterior tentacular nerves (AT and PT) (Fig. 3, G and H) in the cerebral ganglion. In the buccal ganglion, the strongest staining was present in the esophageal nerve (EN), which innervates the gut (esophagus). Immunopositive fibers were also present in buccal nerves 1, 2, and 3, but the staining was less intense (Fig. 3, E and F). Collectively, these observations suggest that apEP/CCHa might play an important role in feeding circuit regulation.

In other ganglia, including the pleural-pedal and abdominal ganglia, there are some apEP/CCHa-positive neurons (Fig. S1, G–L). Notably, one cluster of strongly immunopositive neurons appeared near the P5 and P6 nerves on the ventral surface of the pedal ganglia (Fig. S1, I and J), suggesting a possible role of apEP/CCHa in locomotion (71, 72).

Identification of mature apEP/CCHa in Aplysia by mass spectrometry

In a previous work (13), we utilized liquid chromatography-tandem MS (LC-MS/MS) to detect peptides in the EN extracts, particularly CCK. Given that our new immunohistochemistry data showed significant apEP/CCHa-positive staining in the EN (Fig. 3, E and F), we reanalyzed the previous data (13) to determine whether apEP/CCHa was present in these extracts. Indeed, these EN extracts contained apEP/CCHa precursor-derived peptides (Fig. S2A and Table S4). However, we did not detect the mature apEP/CCHa peptide in the EN data, likely because its structural features—a large circular domain, a short terminal linear sequence, and continuous C-terminal glycine residues—reduce fragmentation of the peptide molecular ions during collision-induced activation in the mass spectrometer.

To characterize the primary structure and PTMs of the mature apEP/CCHa, peptide extracts from the central ganglia were subject to reduction/alkylation of cysteine residues to efficiently disrupt disulfide bridges (73). This treatment linearizes the peptide chains, thereby facilitating peptide analysis by mass spectrometry. We were then able to detect apEP/CCHa (Fig. 3I, Fig. S2B and Table S5). The data provided direct evidence for the presence of apEP/CCHa, most likely with a disulfide bond, in the CNS.

Peripheral sources of Aplysia EP/CCHa

EP/CCHa functions as a brain-gut peptide in arthropods (22, 30, 74, 75), but its role in lophotrochozoans, particularly mollusks, remains unclear. To identify peripheral sources of apEP/CCHa in Aplysia, we conducted immunocytochemistry experiments on peripheral tissues, including the esophagus and crop. We observed a small number of apEP/CCHa-positive cells near the anterior region of the dorsal longitudinal folds (DLFs) (Fig. 4, A–C) of the esophagus. Notably, a recent study (13) has identified apCCK immunopositive cells ventral to the DLFs, which do not appear to overlap with apEP/CCHa positive cells. In addition, we found large apEP/CCHa positive cells in the stomatogastric ring (Fig. 4, D and E), which surrounds the digestive tract between the crop and the gizzard. These large cells may be motoneurons that regulate gut motility (76).Figure 4Distribution of peripheral apEP/CCHa-positive cells. A, the esophagus and posterior buccal cavity (pharynx) lining, with the luminal side facing upward. B and C, EP/CCHa-positive cells (arrows) in the esophagus and pharynx, anterior to DLF. B, bright-field view and C, fluorescent view showing immunostaining. D, caudal crop and stomatogastric ring, shown with the outside surface facing upward. E, two large apEP/CCHa-positive neurons and positive fibers in the stomatogastric ring. DLF: Dorsal longitudinal fold, D: dorsal, V: ventral, A: anterior, P: posterior. Images were taken from a light fluorescent microscope, except panels**B and C, which were taken from a confocal microscope.

These findings support the classification of apEP/CCHa as a brain-gut peptide, with this function appearing to be conserved between lophotrochozoans and arthropods.

Identification of apEP/CCHa receptors

Next, we sought to identify putative apEP/CCHa receptors in Aplysia. Using the Platyneris dumerilii EP receptor sequence (AKQ63032.1) (28), the first EP receptor identified in annelids, we performed a BLAST search in NCBI. This search returned two candidate sequences, named CCHamide-1 receptor (XP_005093521.2) and CCHamide-2 receptor (XP_005098643.1). We then used these two sequences to perform BLAST searches in AplysiaTools, retrieving similar sequences (TRINITY_DN9288_c0_g1_i6 and TRINITY_DN13622_c3_g1_i4) (Table S1, Figs. S3 and S4). We tentatively referred to them as apEP/CCHaR1 and apEP/CCHaR2. In both cases, the NCBI and AplysiaTools sequences only differed by a single amino acid (Fig. S3, A and B).

To determine whether the two sequences are GPCRs, we used NCBI Conserved Domain Search and TMHMM Server 2.0. Both tools predicted that the two sequences we obtained have the seven transmembrane domains expected for GPCRs (Fig. 5A). To confirm the presence of these sequences in Aplysia, we designed primers (Table S2) based on their open reading frames (ORFs), and successfully cloned the mRNAs for both putative receptors (Fig. 5B). Sequencing showed that our cloned sequences are identical to the sequences in AplysiaTools (Fig. S3, A and B).Figure 5Bioinformatics and cloning of two putative EP/CCHaRs in Aplysia. A, prediction of 7 TM domains of putative apEP/CCHaRs: apEP/CCHaR1 and apEP/CCHaR2 using TMHMM 2.0. Conserved motifs in transmembrane domains 3 (TM3: D/ERY) and TM7 (NPXXXY) are shown. The amino acids different from the motifs are shown in red. B, the PCR products of two putative apEP/CCHaRs: apEP/CCHaR1 (1302 bp), apEP/CCHaR2 (1755 bp). C, a phylogenetic tree of the two Aplysia receptors with multiple molluscan class A GPCR sequences from Jiang et al. (77) (see the results and Table S6 for information of these sequences) using MEGA X. A class-B GPCR, Crassostrea gigas parathyroid hormone peptide receptor, was used as an outgroup. The tree suggests that apEP/CCHaR1 and apEP/CCHaR2 are likely Aplysia EP/CCHa receptors. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. Numbers at the nodes are bootstrap values as percentage. Only bootstrap values greater than 50 are shown.

Phylogenetic analysis based on a balanced tree from Jiang et al. 2022 (77) (Table S6) after adding EP/CCHa receptors from molluscan species showed that apEP/CCHaR1 and apEP/CCHaR2 cluster with the putative Crassostrea gigas CCHa-1 receptor (Fig. 5C), supporting the idea that they are Aplysia EP/CCHa receptors.

We next tested whether the apEP/CCHa peptide could activate the two putative receptors using an inositol monophosphate (IP1) accumulation assay. The peptide was applied at two concentrations (10^–12^ M and 10^–5^ M). At 10^–12^ M, receptor activation was minimal, so these data were used as a control. At 10^–5^ M, only apEP/CCHaR1 responded to apEP/CCHa (Fig. 6A). Given the negative results for apEP/CCHaR2, we conducted an additional experiment in which we co-transfected both apEP/CCHaR1 and apEP/CCHaR2 with a promiscuous Gαq-family protein (78). This protein couples with most GPCRs and allows ligand activation regardless of the receptor’s native signaling pathway. With Gαq co-transfection, both apEP/CCHaR1 and apEP/CCHaR2 responded to apEP/CCHa (Fig. 6B), indicating that apEP/CCHaR2 does not associate efficiently with native Gαq proteins in CHO-K1 cells. Consequently, we performed all subsequent IP1 assay experiments on both receptors using Gαq co-transfection.Figure 6Activation of putative receptors by apEP/CCHa as determined by IP1 accumulation assay. A and B, screening for ligands on putative receptors using two concentrations: 10^−12^ M and 10^−5^ M (n = 3), without co-transfection (A) or with co-transfection (B) of the promiscuous Gαq. At 10^-12^ M, a peptide minimally activates a receptor, serving as a control. Paired two-tailed t test: ns, not significant; ∗∗p < 0.01; ∗∗∗p < 0.001; error bars: S.D. C, representative examples of dose-response curves showing the ability of apEP/CCHa to activate apEP/CCHaRs with co-transfection with the promiscuous Gαq in CHO-K1 cells. Each data point is from two wells. D, comparison of pEC_50_ shown in (C) (n = 3). Paired t test, ns, not significant. E, a summary of the average pEC_50_ and EC_50_ values for apEP/CCHaR1 and apEP/CCHaR2. pEC_50_ values are reported as the mean ± SD from three independent experiments. CHO-K1 cells, Chinese hamster ovary-K1 cells; IP1, inositol monophosphate.

We generated dose–response curves for peptide activation using apEP/CCHa concentrations from 10^-12^ M to 10^-4^ M (Fig. 6C). Both receptors exhibited similar half-maximal effective concentrations (EC_50_): 30 nM for apEP/CCHaR1 and 28 nM for apEP/CCHaR2 (Fig. 6, D and E).

Expression profiling (GEO accession number: [GSE79231](GSE79231)) showed that apEP/CCHaR1 is broadly distributed in both the CNS and peripheral tissues, whereas apEP/CCHaR2 expression is primarily in the CNS (Fig. S5). In situ hybridization revealed sparse apEP/CCHaR1 expression in the buccal and pleural-pedal ganglia, while apEP/CCHaR2 was primarily present in the cerebral and pleural-pedal ganglia (Fig. S6). Taken together, these findings suggest that apEP/CCHaR1 and apEP/CCHaR2 might participate in modulating multiple physiological functions, including feeding, as the cerebral and buccal ganglia are involved in the feeding control.

Phylogenetic relationships between the protostome EP/CCHa, and the deuterostome BN-type peptide, and ET signaling systems in bilaterians

The identification of the two apEP/CCHaRs enabled us to further investigate the phylogenetic relationships among EP/CCHa receptors, BN-type peptide receptors (GRPR, NMBR, and BRS-3), and ET receptors (Fig. 7, see Table S7 for information about these sequences). In particular, previous phylogenetic trees (27, 29, 40, 41, 42) included fewer numbers (≤5) of molluscan sequences. Thus, we used the two Aplysia receptor sequences to find 17 additional sequences in mollusks (Table S7). With this set of sequences, we constructed a phylogenetic tree. The tree showed that apEP/CCHaR1 and some other molluscan sequences cluster with annelid and nemertean EP/CCHa receptors, while apEP/CCHaR2 and several other molluscan sequences formed a clade with CCHa receptors from arthropods (Fig. 7). This suggests the presence of two distinct EP/CCHaR types in mollusks. Interestingly, although there are also two receptors in most arthropods, these two arthropod receptors all cluster together, rather than forming two clades. Additionally, all lophotrochozoan receptors formed a large clade with arthropod CCHa receptors. Notably, protostome EP/CCHa receptors were grouped within the same branch as deuterostome ET receptors, whereas BN-type peptide receptors formed a separate branch. These findings support the hypothesis that EP/CCHa and ET signaling systems share a common evolutionary origin, whereas the BN-type peptide system represents a distinct but related lineage. This conclusion is consistent with phylogenetic analyses in other studies (27, 40, 42).Figure 7Phylogeny of bilaterian EP/CCHa, ET and BN-type peptide receptors, including Aplysia californica receptors, apEP/CCHaR1 and apEP/CCHaR2. The tree was constructed using MEGA X with 10,000 replicates using JTT + G model (see the results and Table S7 for information about these sequences). “∗” indicates that the receptor has been verified. “Neuropeptide S receptor Plakobranchus ocellatus” was used as an outgroup. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. Numbers at the nodes are bootstrap values as a percentage. Only bootstrap values greater than 50 are shown. NMBR = BB1 receptor; GRPR = BB2 receptor; BRS3 = BB3 receptor.

apEP/CCHa reduces food intake in intact animals

To determine whether apEP/CCHa might function as a satiation/satiety signal in mollusks, we examined its effects on food intake in intact Aplysia. Animals were injected with either artificial seawater (ASW) or apEP/CCHa at multiple concentrations. Initial experiments showed that injection of apEP/CCHa at 1 × 10^-8^ M produced variable effects on feeding (Fig. 8A). In contrast, injections at higher concentrations (2 x 10^-8^ M to 10^-7^ M) completely suppressed feeding behavior. However, at these higher concentrations, some animals exhibited body curling and failed to perform righting behavior, suggesting possible nonspecific systemic effects.Figure 8Effects of apEP/CCHa on food intake and feeding programs. A, effects of 1 x 10^-8^ M and 1.5 x 10^-8^ M apEP/CCHa on food intake. Whereas 1 x 10^-8^ M apEP/CCHa did not have a significant effect, 1.5 x 10^-8^ M apEP/CCHa did. F (2, 19) = 24.21, p < 0.0001 (ASW, n = 11; 1 x 10^-8^ M, n = 6, and 1.5 x 10^-8^ M, n = 5). B, a representative example; a single cycle of a motor program was elicited by simulating CBI-2 at 10 Hz until the end of the protraction phase, monitored by the onset of the abrupt ending of I2 nerve activity. The protraction phase (open bar) was defined by the activity in the I2 nerve. The retraction phase (orange filled bar) was defined as a period of activity of the radula closing motoneuron B8 that occurs after the protraction phase ends. Upon wash, protraction duration returned to its control value. C and D, group data for the effects of apEP/CCHa on protraction duration (C) and retraction duration (D) (n = 7). Protraction: F (3, 18) = 30.36, p < 0.0001. Retraction: F (3, 18) = 4.683, p > 0.05. E, group data. Plot of average B8 activity during protraction versus retraction of CBI-2-elicited programs (n = 4). One-way ANOVAs are used in (A), (C) and (D) with their Tukey post hoc tests: ns, not significant, ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001, ∗∗∗∗p < 0.0001. Error bars: S.D.

Importantly, injection of apEP/CCHa at 1.5 x 10^-8^ M significantly reduced food intake (Fig. 8A), and at the end of the feeding session, all the animals performed righting behavior and exhibited normal locomotion. The narrow “working” concentration range suggests that higher doses may engage additional physiological systems beyond those regulating feeding. Together, these results suggest that apEP/CCHa potently suppresses feeding, consistent with a role as a satiation/satiety peptide in Aplysia.

apEP/CCHa modulates feeding programs in isolated ganglia

To explore the neural basis of the behavioral actions of apEP/CCHa, we first determined its effects on feeding motor programs in isolated ganglia. Aplysia generates two major types of motor programs, ingestive and egestive. Each cycle of motor programs consists of a protraction-retraction sequence, and different types of programs can be distinguished by activity phasing of radula closer motoneuron B8. During ingestive programs, B8 is primarily active during retraction to pull food into the buccal cavity, whereas during egestive programs, B8 is primarily active during protraction to push inedible food/object out (55, 63, 64, 79, 80, 81).

apEP/CCHa was tested on programs induced by stimulation of CBI-2, a command-like interneuron that is active during food-elicited feeding behavior (82, 83) and provides potent excitatory inputs to buccal CPG interneurons (13, 14, 63, 64, 84), orchestrating ingestive motor programs. Stable single-cycle motor programs were evoked by stimulating CBI-2 at 8 to 10 Hz throughout the protraction phase with a 90-s intertrial interval. Two concentrations, 10^-6^ M and 10^-5^ M, of apEP/CCHa were tested. Under control conditions, CBI-2 stimulation elicited an ingestive motor program (i.e., B8 was predominantly active during retraction). When 10^-6^ M and 10^-5^ M apEP/CCHa were perfused, there was a concentration-dependent shortening of protraction (Fig. 8, B and C, n = 7). The retraction phase was not significantly altered (Fig. 8D, n = 7). To further determine whether apEP/CCHa might alter the type of feeding motor program induced, we analyzed a subset of preparations that showed ingestive characteristics under control conditions (n = 4). Group data showed that B8 activity during protraction was increased in a concentration-dependent manner. In contrast, B8 activity during retraction was decreased in a concentration-dependent manner (Fig. 8E).

These results suggest that apEP/CCHa can transform ingestive feeding programs to egestive programs with a shorter protraction, consistent with its behavioral effects on feeding suppression.

The effects of apEP/CCHa on CPG interneurons in isolated ganglia

Motor neuron activity in the Aplysia feeding circuit is shaped by CPG interneurons, most of which are located in the buccal ganglion. To identify potential CPG neurons that apEP/CCHa might target, we analyzed previously generated single-cell RNA-seq data (GEO accession number: [GSE296741](GSE296741)) (13). This dataset included two identified CPG interneurons B34 and B20, both of which expressed transcripts for apEP/CCHaR1 and apEP/CCHaR2 (Fig. 9A). Functionally, B34 promotes long protraction (83), whereas B20 promotes egestive programs (63, 64) and shorter protraction phases (85).Figure 9The effects of apEP/CCHa on the excitability of CPG interneurons B34, B20, B40 and B64. A, Single cell RNA-seq showing expression of the two receptors, apEP/CCHaR1 and apEP/CCHaR2, in B34 and B20 neurons. Relative counts were obtained by removing batch effects (n = 3). B and C, apEP/CCHa decreased the excitability of B34 in a concentration-dependent manner; representative examples (B); group data (C) (n = 6). F (3, 15) = 41.05, p = 0.0002. D and E, apEP/CCHa enhanced the excitability of B20 at higher concentration (10^-5^ M); representative examples (D); group data (E) (n = 5). F (3, 12) = 22.28, p = 0.0009. F and G, apEP/CCHa decreased the excitability of B40; representative examples (F); group data (G) (n = 3). F (3, 6) = 85.14, p = 0.0042. H and I, apEP/CCHa enhanced the excitability of B64 following the initial perfusion of the concentration at 10^-5^ M; representative examples (H), group data (I) (n = 4). F (3, 12) = 79.71, p = 0.0007. One-way ANOVAs are used in (C), (E), (G) and (I) with their Tukey post hoc tests: ∗p < 0.05, ∗∗p < 0.01. Error bars: S.D.

We additionally examined two other CPG interneurons, i.e., B40 and B64, selected for their well-established roles in feeding pattern generation. B40 promotes ingestive programs and long protraction (64, 83), whereas B64 is active during retraction and terminates protraction when activated (86, 87). We assessed the effects of apEP/CCHa on the excitability of all four interneurons. apEP/CCHa reduced the excitability of B34 (Fig. 9, B and C) while increasing the excitability of B20 (Fig. 9, D and E). Moreover, apEP/CCHa decreased the excitability of B40 in a concentration dependent manner (Fig. 9, F and G) and transiently increased the excitability of B64 during the 10^-5^ M application, an effect that was not maintained during prolonged exposure (Fig. 9, H and I).

All excitability experiments were performed in high-divalent saline to suppress polysynaptic transmission. Therefore, the observed changes in B40 and B64 excitability likely reflect direct actions of apEP/CCHa, suggesting that one or both apEP/CCHa receptors are expressed in these neurons, although single-cell RNA-seq are not yet available for them.

Overall, the effects of apEP/CCHa on these CPG neurons align with the network-level changes expected when apEP/CCHa converted CBI-2-elicited ingestive programs to egestive programs with shortened protraction (Fig. 10).Figure 10Schematic diagram for apEP/CCHa-mediated satiation/satiety mechanism. Food presentation to the mouth activates the cerebral higher-order neuron CBI-2, which preferentially excites B40 relative to B20, thereby promoting ingestive motor programs. Peripheral apEP/CCHa-expressing cells in the gut (fluorescent image reproduced from Fig. 4C) are likely activated by gut distension during and after a meal. These cells are proposed to project via the EN (the esophageal nerve) to the buccal ganglion, where apEP/CCHa is released to act on the pattern-generator elements; additional sensory inputs (e.g., CCK-expressing neurons) may also contribute. Within the buccal ganglion, apEP/CCHa binds to apEP/CCHa receptors (apEP/CCHaRs) in CPG interneurons, including B20 and B34: increasing the excitability of B20 (green arrow) while decreasing the excitability of B34 (red arrow). apEP/CCHa also reduces the excitability of B40 (red arrow) and transiently increases the excitability of the retraction interneuron B64 (green arrow); both B40 and B64 are presumed to express apEP/CCHaRs. Collectively, these neuron-specific changes in excitability bias the feeding network toward a transition from ingestion to egestion. GPCR, apEP/CCHa receptors; PM, protraction motoneurons; RM, retraction motoneurons.

Discussion

In this study, we used the tractable and the well-characterized molluscan model Aplysia to investigate the roles of EP/CCHa signaling in feeding. By integrating molecular, cellular, chemical, anatomical, behavioral, and neurophysiological approaches, we provide a comprehensive characterization of the EP/CCHa neuropeptide signaling system in a lophotrochozoan. The two receptors we characterized represent the first receptors described in mollusks, and phylogenetic analyses suggest that they belong to two separate lineages. We also provide the first functional exploration of EP/CCHa peptide in Aplysia, demonstrating that apEP/CCHa modulates the excitability of multiple feeding CPG interneurons. These findings establish EP/CCHa as a key regulator of feeding, supporting its function as a brain-gut peptide that mediates satiation or satiety.

Protostome EP/CCHa precursors and peptides

EP/CCHa peptides were first identified biochemically in annelid E. foetida and P. vittate (15), with subsequent identification of single precursors in E. foetida, Hirudo nipponia and Pheretima vittata (18, 88). In mollusks, two precursors in each of the two species: T. clavigera (38) and Deroceras reticulatum (21), were identified. Here, we identified a single EP/CCHa precursor in mollusk Aplysia (Fig. 1B) that yields a 15-amino acid peptide containing a disulfide bond and a C-terminal amidation (Fig. 1C). To ensure detection of predicted cyclic apEP/CCHa, we adapted a cysteine derivatization protocol to the neuropeptide extracts to open potential disulfide bonds in peptides, using an approach described in our earlier work (73), which allowed us to successfully confirm the peptide sequence and the presence of disulfide bridge (Figs. 3I, S2B and Table S5) using LC-MS/MS.

Comparative analyses across protostomes reveal broad variation in EP/CCHa precursor number. Most annelids harbor a single precursor, as in Platynereis dumerilii (89) and Dinophilus gyrociliatus (19). Mollusks vary, with some species having two precursors, such as T. clavigera (38), Charonia tritonis (90) and D. reticulatum (21), and others, including Aplysia, Lottia gigantea (24), C. gigas (91), and Patinopecten yessoensis (92), containing only one. Importantly, each precursor encodes only a single mature peptide.

In arthropods, most species possess two precursors, as in D. melanogaster (23), B. mori (22), Spodoptera exigua (75), A. aegypti (36) and Carausius morosus (93). However, several species, such as Marsupenaeus japonicus (94), Daphnia pulex (95), and Plutella xylostella (96), contain one. In nematodes (which, together with arthropods, belong to superphylum ecdysozoa), no EP/CCHa precursors, nor receptors, have been identified so far, suggesting that this peptide signaling system was lost during evolution (27).

This variability underscores the evolutionary plasticity of the EP/CCHa signaling system and raises important questions regarding functional diversification across protostomes.

EP/CCHa receptors in protostomes

Prior to this work, only two lophotrochozoan EP/CCHa receptors had been experimentally validated: one in the annelid P. dumerilii (28) and one in the nemertean Lineus longissimus (29). Here, we provide the first experimental characterization of two functional EP/CCHa receptors in mollusks: apEP/CCHaR1 and apEP/CCHaR2 (Fig. 5). Both respond to apEP/CCHa with comparable EC_50_ values (30 and 28 nM, Fig. 6), similar to those reported in P. dumerilii (7.9–15 nM) (28) and L. longissimus (59–78 nM) (29). Most arthropods, including B. mori (97), D. melanogaster (25), and A. aegypti (36), possess two receptors preferentially activated by distinct peptides derived from their two corresponding precursors. The EC_50_ values for both receptors in B. mori (97) and D. melanogaster ranged from 2 to 100 nM (25), but in A. aegypti, they were 8 to 15 nM for CCHa-2R, but were higher (420 pM –984 nM) for CCHa-1R (36). Overall, these remain broadly similar in magnitude to those in lophotrochozoans and support conservation of receptor pharmacology across protostomes.

Phylogenetic relationships between protostome EP/CCHa signaling systems and deuterostome ET and BN-type peptide signaling systems

Using the two newly identified Aplysia receptors and additional predicted molluscan receptors (two in some, e.g., Crassostrea angulata, Biomphalaria glabrata, and Ostrea edulis, and one in others (Fig. 7 and Table S7)), our phylogenetic analyses revealed distinct receptor clusters. One (cluster 1) includes apEP/CCHaR1 and single-copy receptors from other mollusks, grouping together with annelid and nemertean receptors. The other (cluster 2), containing apEP/CCHaR2, clusters with arthropod receptors (Fig. 7). The low sequence identity (17.32%) and modest similarity (30.39%) between the two Aplysia receptors (Fig. S3C) further support significant divergence or functional specialization.

Thus, although EP/CCHa receptors are broadly conserved across protostomes, the two molluscan receptors have distinct evolutionary affinities: one aligns with other lophotrochozoan receptors, whereas the other may represent an ancestral lineage that gave rise to arthropod receptors.

The broader significance of the protostome EP/CCHa signaling system derives from its deep evolutionary relationship to deuterostome ET and bombesin-type (BN-type) peptide signaling systems. BN-type peptides—including BN, GRP, and NMB—comprise three related subfamilies (Fig. S7). BN was originally isolated from the skin of the frog Bombina (43, 44), followed by ranatensin from the northern leopard frog (Rana pipiens) (98) and phyllolitorin from Phyllomedusa sauvagii (99). These peptides share highly similar C-terminal sequences. GRP (45) and NMB (46) were later identified in mammals, where they regulate physiological processes, such as feeding and digestion (100, 101, 102). Similar to EP/CCHa, BN-type peptides contain a C-terminal amidation (Figs. 2 and S7), although they lack the disulfide bonds characteristic of EP/CCHa.

In mammals, NMB and GRP selectively activate the BB1 (Bombesin 1) receptor (i.e., NMBR in mammals) and BB2 receptor (i.e., GRPR in mammals), respectively, with orthologs present across vertebrates (47, 48, 49, 103, 104). Another homologous receptor, bombesin receptor subtype 3 (BRS-3) (or BB3 receptor), first identified from the human genome (105), is widely conserved among mammals (106, 107, 108, 109). However, unlike NMBR and GRPR, BRS-3 displays extremely low affinity for known BN-type peptides and remains an orphan receptor (110, 111).

ETs, by contrast, were first isolated from porcine aortic endothelial cells (112), and represent the most potent vasoconstrictor known. Two additional ET ligands and their cognate receptors were subsequently identified (113, 114, 115). Like EP/CCHa peptides in protostomes, ETs contain disulfide bonds; however, they possess two such bonds and lack C-terminal amidation (Figs. 2 and S7). Although a BN-type neuropeptide was recently characterized in A. rubens (42), a bona fide ET signaling system has not yet been confirmed in echinoderms.

Despite the low primary sequence similarity among EP/CCHa, ETs and BN-type peptides (Figs. 2 and S7), their evolutionary relationships become evident at the receptor level. Earlier phylogenetic analyses (27) and receptor-based clustering (26, 28) using EP/CCHa and NMB/ET receptors, as well as more recent analyses (28, 29, 40, 41, 42), support their homology. By incorporating molluscan receptors, including the Aplysia receptors and others predicted by us, and echinodermata BN-type peptide receptors (42), our phylogenetic analyses reaffirm these relationships. Protostome EP/CCHa receptors cluster within the same branch as deuterostome ET receptors, whereas BN-type peptide receptors form a separate clade (Fig. 7), indicating that protostome EP/CCHa receptors share a closer evolutionary relationship with ET receptors.

In summary, our results strengthen the view that the protostome EP/CCHa signaling system is homologous to deuterostome ET/BN-type peptide signaling systems. Intriguingly, although EP/CCHa peptides exhibit greater functional similarity to BN-type peptides (see discussion below), their receptors show higher sequence homology to ET receptors. This divergence between ligand function and receptor evolution underscores the complexity of neuropeptide–receptor co-evolution across bilaterians.

EP/CCHa, BN-type peptides, and ETs as brain-gut peptides

Homology between protostome EP/CCHa and deuterostome BN-type peptides and ETs extends beyond their phylogenetic relatedness. Across the species examined to date, these peptide families consistently exhibit dual localization in the CNS and the gastrointestinal system, supporting their designation as brain-gut peptides.

EP/CCHa was originally identified in annelids (E. foetida and P. vittate) through HPLC/MS analysis of gut and whole-body tissues (15), followed by detection in annelid brains via immunohistochemistry (17, 18, 19). In other lophotrochozoans, specifically mollusks, two T. clavigera EP peptides (TEP-1 and TEP-2) show distinct CNS distributions: TEP-1 is prominent in the pleural, subesophageal, pedal, and visceral ganglia, whereas TEP-2 is mainly found in the cerebral ganglia (38), but whether the EP/CCHa peptides are present in the gut has not been established.

Here, we demonstrate that Aplysia EP/CCHa is expressed in both the CNS (Figs. 3 and S1) and the digestive system (Fig. 4). Within the gut, we identified small cells situated anterior to DLF (116) in the posterior oral cavity or the esophagus (Fig. 4, A–C), consistent with previously described sensory neurons that project via the EN to the buccal ganglion (13), where the feeding CPG resides. We also observed large cells near stomatogastric rings (Fig. 4, D and E), which resemble gut motoneurons implicated in gastrointestinal motility (76). These findings establish EP/CCHa as a bona fide brain–gut peptide in mollusks.

In arthropods, EP/CCHa peptides similarly show dual localization. They have been identified in both the gut and CNS of B. mori (22), D. melanogaster (30), Chilo suppressalis (74), and S. exigua (75), reinforcing their conserved role as brain-gut peptides across protostomes.

In deuterostomes, homologous BN-type peptides also occur in both peripheral tissues (including the gut) and CNS in amphibian, mammals (117, 118, 119), and A. rubens (42). ET peptides display a similar pattern: although ET-1 and ET-2 are predominantly peripheral (120) and ET-3 is largely CNS-enriched (121, 122, 123), all three subtypes are present in both the periphery—including the gut—and the CNS.

Collectively, these data indicate that EP/CCHa peptides in protostomes and ET/BN-type peptides in deuterostomes represent evolutionarily conserved brain-gut peptides. Our findings provide the first demonstration that EP/CCHa serves as a brain-gut peptide in mollusks, further linking this lineage with homologous systems in arthropods and deuterostomes.

Actions of EP/CCHa in feeding: Roles as a satiation/satiety signal

The observation that protostome EP/CCHa and deuterostome BN-type peptides and ETs are brain-gut peptides suggests a potential role in feeding regulation. Because relatively few studies have examined the effects of ETs on feeding, our discussion focuses primarily on EP/CCHa and BN-type peptides.

Prior to this study, functional analyses of protostome EP/CCHa signaling were largely confined to arthropods, particularly insects, and revealed species-specific effects on feeding. In D. melanogaster, genetic knockdown of EP/CCHa or its receptor suppresses hunger-driven feeding (31, 32), whereas exogenous EP/CCHa promotes food intake (30). In contrast, in adult female A. aegypti, CCHa-2 knockdown increases sucrose consumption, and peptide injection decreases sucrose intake over 24 h (36); with similar action on food intake observed in G. bimaculatus (37). Thus, EP/CCHa signaling contributes to feeding regulation across arthropods, but its net effect—stimulatory or inhibitory—varies among species.

In lophotrochozoans, EP/CCHa peptides have been reported to exert myotropic effects on tissues including the gut (15, 16, 17, 18, 20). However, whether these actions translate into changes in overall feeding behavior had not been demonstrated before the present study.

In deuterostomes, BN-type peptides serve as anorexigenic signals in mammals (50, 51, 52) and echinodermata (42). For example, earlier studies showed that BN injection in rats suppresses feeding (50, 124), and mice lacking BN-type peptide receptors (NMBR, GRPR, or BRS-3) exhibit hyperphagia and obesity (125).

Our results indicate that apEP/CCHa functions analogously to BN-type peptides and to CCHa in certain insects (e.g., A. aegypti). The role of apEP/CCHa in feeding regulation is supported by convergent evidence from behavioral assays, in vitro motor program analyses, and cellular-level effects on identified elements of the feeding CPG. Specifically, in intact Aplysia, apEP/CCHa decreases food intake (Fig. 8A), while in isolated ganglia it shifts feeding motor programs from ingestion to egestion (Fig. 8E). Coupled with its distribution as a brain-gut peptide, these findings are consistent with EP/CCHa acting as a satiation/satiety signal in mollusks.

In vertebrates, meals consist of a series of feeding bouts regulated by satiation signals—such as CCK—which emerge within minutes of meal initiation to shape meal size (10, 126, 127, 128). In contrast, satiety governs the duration of the intermeal interval and is often mediated by post-absorptive signals, including GLP-1 (128, 129, 130) and PYY (128, 131). Although satiation and satiety are behaviorally distinguishable, their underlying neural circuits and neuromodulatory mechanisms may not be fully segregated and could operate through overlapping pathways (126). Based on our current data, apEP/CCHa appears to act primarily to terminate an ongoing meal, akin to CCK. However, further experiments are required to determine whether apEP/CCHa also influences longer-term satiety.

While Drosophila CCHa-2 promotes feeding by acting on insulin-producing cells in the brain (35), the underlying circuit-level mechanisms remain largely unknown in insects or any other species. Here, we defined such mechanisms in Aplysia. apEP/CCHa shortens the protraction phase of the feeding motor program by reducing the excitability of protraction interneurons B34 and B40 (Fig. 9, B, C, F and G), which normally serve to prolong protraction (64, 83, 132). A transient enhancement of excitability of B64—the protraction terminator (86, 87)—likely contributes further (Fig. 9, H and I). The shift toward egestive programs induced by apEP/CCHa could be attributed to increased excitability of the egestion-promoting interneuron B20 (63) (Fig. 9, D and E), and the concomitant reduced excitability of the ingestion-promoting interneuron B40 (64) (Fig. 9, F and G). To our knowledge, these results provide the first mechanistic demonstration of how EP/CCHa peptides regulate a feeding CPG (Fig. 10).

Previous studies in Aplysia have identified several neuropeptides that promote egestive motor programs and likely act as satiation/satiety signals, including NPY (14), SCP (133), and CCK (13). Our identification of apEP/CCHa adds another peptide to this group. Notably, during the transition from hunger to satiation—before complete cessation of feeding—food rejection has also been observed in mammals, including rodents (130) and monkeys (134). The presence of multiple satiation/satiety-promoting peptides with distinct but complementary roles appears to be a recurring feature in both vertebrates (e.g., CCK, PYY, GLP-1, see Discussion above) (9, 10, 127, 128, 130) and Aplysia. The Aplysia feeding circuit provides a tractable system for comparing the cellular effects of these peptides. All known satiation/satiety-related peptides are present in the EN and commonly enhance B20 activity while suppressing B40 activity. Some actions, however, differ. For example, at high concentrations, CCK suppresses protraction motoneuron activity during CBI-2-elicited programs by reducing their synaptic input (13). Moreover, although both CCK and EP/CCHa exhibit immunoreactivity in esophageal cells near the DLF, they reside in distinct populations. EP/CCHa-immunoreactive cells exhibit a more restricted and anterior distribution (Fig. 4, A–C) relative to CCK-positive cells. These differences suggest that distinct gut-derived peptide populations may be differentially engaged by feeding-related signals, enabling complementary modes of satiation/satiety control.

In conclusion, apEP/CCHa acts as a brain-gut satiation/satiety peptide that inhibits feeding by modulating CPG elements governing protraction duration and program selection, likely through its two receptors. These findings establish an evolutionarily conserved feeding-inhibitory role for EP/CCHa peptides in mollusks and underscore the broader diversity of EP/CCHa signaling across protostomes. More generally, brain-gut peptides and their receptors have emerged as central regulators of metabolic disorders (135, 136, 137), diabetes (138), and mental health and neurological diseases (139, 140). Indeed, peptide mimics now rank among the top five therapeutics worldwide (141, 142, 143). Our findings highlight the ancient origins and enduring significance of these neuromodulatory systems.

Limitations of the study

By leveraging the well-defined feeding behavior and circuit of Aplysia, our results support the role of apEP/CCHa in satiation and/or satiety. However, the lack of genetic tools for neuron-specific labeling and manipulation in Aplysia limits our ability to directly test whether peripheral apEP/CCHa-expressing neurons are activated by food intake and whether they are required for feeding suppression during satiation. Such experiments would also help address concerns regarding the narrow effective concentration range observed in peptide injection experiments, which may reflect nonspecific systemic effects at higher doses.

In addition, the development of compatible protocols for simultaneous fluorescence in situ hybridization and immunofluorescence would enable direct assessment of apEP/CCHa peptide and receptor expression within the same neuronal populations. Finally, characterization of intracellular signaling pathways downstream of apEP/CCHa receptor activation will be necessary to explain how a single peptide produces opposite effects on the excitability of distinct CPG elements.

Experimental procedures

Subjects and reagents

A. californica (70–350 g) were obtained from both Marinus Scientific in Los Angeles, CA, and RSMAS National Resource for Aplysia in Miami, Florida. Animals were maintained in an aquarium with a circulating artificial seawater at 14 °C - 16 °C and the room was equipped with a 12:12 h light-dark cycle, with 7 AM to 7 PM as the daylight. Before dissection, animals were anesthetized by injection of isotonic 333 mM MgCl_2_ (about 50% of body weight) into the body cavity. All reagents were purchased from Sigma-Aldrich unless otherwise stated.

Bioinformatic analysis of peptide precursors and receptors

To identify Aplysia EP/CCHa precursors and receptors, we used NCBI and AplysiaTools databases (Dr Thomas Abrams, University of Maryland) (144) to search specific sequences of interests. These latter databases (http://aplysiatools.org) include databases for Aplysia transcriptome and genome. The open reading frames (ORFs) from the apEP/CCHa precursor, and the putative apEP/CCHa receptors were obtained using ORF Finder (https://www.ncbi.nlm.nih.gov/orffinder/). For the apEP/CCHa precursor, the putative signal peptide was predicted using SignalP 5.0 (http://www.cbs.dtu.dk/services/SignalP/) and the putative peptides encoded by the apEP/CCHa precursor were predicted using NeuroPred (http://stagbeetle.animal.uiuc.edu/cgi-bin/neuropred.py). We also compared the apEP/CCHa with those of other species using BioEdit software and generated a frequency plot of each amino acid (aligned from the C-terminus) using Weblogo software (http://weblogo.berkeley.edu/logo.cgi). For the putative apEP/CCHa receptors, transmembrane domains were predicted using TMHMM Server v.2.0 (http://www.cbs.dtu.dk/services/TMHMM/). For proteins that were difficult to annotate using BLAST, we used the Pfam database (http://pfam.xfam.org/search#tabview=tab1) to determine what type of a protein it is. The phylogenetic trees of sequences from different species were constructed by MEGA X software (https://www.megasoftware.net/) using alignment by Clustal W and the maximum likelihood method with 1000 replicates. For Figure 5C, we used JTT + G + F model to generate the phylogenetic tree; for Figure 7, LG + G model was performed, and the resulting tree was visualized with ChiPlot (https://www.chiplot.online/) (145); The selection of the models was based on the results of MEGA analysis. The other parameters were set as default.

Cloning of mRNA in Aplysia

RNA extraction

RNA was prepared from the Aplysia ganglia using the TRIzol reagent method. After anesthesia, Aplysia ganglia were dissected out. The dissected ganglia were placed into 200 μl TRIzol (Sigma, T9424) and stored at −80 °C until use. The frozen ganglia in TRIzol were thawed and homogenized with a plastic pestle, then TRIzol was added to a total volume of 1 ml, which were incubated at room temperature for 10 min. Then, 200 μl chloroform was added, and the solution was mixed thoroughly by shaking, and let stand on ice for 15 min. The solution was centrifuged (12,000×g, 4 °C, 15 min), and the supernatant was added to an equal volume of isopropanol. The tube was shaken gently by hand and let stand at −20 °C for 2 h. After 2 h, it was centrifuged (12,000×g, 4 °C, 15 min) again, the supernatant was discarded, 1 ml of 75% ethanol/water was added, and the centrifuge tube was shaken gently by hand to suspend the pellet. It was centrifuged (12,000×g, 4 °C, 10 min), the supernatant discarded and the precipitant was dried at room temperature for 5 to 10 min. Finally, 30 μLof nuclease-free water was added to dissolve the RNA pellet, and the RNA concentration was determined with a Nanodrop ND-1000 spectrophotometer (Thermo Fisher Scientific).

Reverse transcription

Using the above extracted RNA as a template, cDNA was synthesized by reverse transcription using PrimeScript RT Master Mix Kit (Takara, RR036 A) according to the manufacturer’s instructions and then stored at - 20 °C until use.

PCR

The synthesized cDNA above was used as a template for PCR. Each pair of specific primers was designed (Table S2) in Primer Premier six and Oligo7, based on protein-coding sequences for the apEP/CCHa precursor and putative receptors. The PCR reaction was performed with 98 °C/2 min pre-denaturing; 98 °C/10 s denaturing; ∼60 °C (depending on the specific primers: see Table S2)/15s annealing; 72 °C/30 s extension and 72 °C/5 min re-extension for 35 cycles. The PCR products were subcloned into vector pcDNA3.1(+) and sequenced to ensure that the sequence was correct.

IP1 accumulation assay

Inositol monophosphate (IP1) accumulation assay measures the concentration of IP1, that is hydrolyzed from the second messenger, inositol triphosphate (IP3). IP3 is generated by Gαq pathway when a GPCR expressed in CHO-K1 cells (Procell, CL-0062) is activated by an appropriate ligand. IP1 accumulation assay was performed as described previously (146, 147, 148, 149, 150, 151). To express the Aplysia putative apEP/CCHa receptors transiently in CHO-K1 cells, the cDNA was cloned into the mammalian expression vector pcDNA3.1(+). CHO-K1 cells were cultured in F-12K medium (Gibco,21,127–022) with 10% fetal bovine serum (Genial, G11–70500) at 37 °C in 5%CO_2_. Transfection experiments were performed when the cells were grown to 70 to 90% confluence. In preliminary experiments, for each dish (6-well plate), 2 μg of the putative receptor plasmids in pcDNA3.1(+) (1.5 μg plasmids, when co-transfecting with 1.5 μg Gα16) were mixed with 200 μl jetPRIME buffer (Polyplus, 101000046) in a 1.5 mL EP tube, followed by the addition of 4 μl (6 μl, when co-transfecting with Gα16) of jetPRIME Transfection reagent (Polyplus, 101000046) mixture. The CHO-K1 cells with the reagents added above were mixed gently, and incubated at room temperature for 15 to 20 min. The mixture dropwise was then added to the dish, and the cells were incubated at 37 °C in 5% CO_2_ overnight.

After 24 h, the cells were trypsinized and reseeded in 384-well tissue culture-treated plates (Corning, 3570) at a density of 20,000 cells/well in F-12K and 10% FBS and incubated at 37 °C in 5%CO_2_ overnight. On the third day, the activation of the putative receptor was detected by monitoring IP1 accumulation using an IP1 detection kit (Cisbio, 62IPAPEB) in Tecan Spark. Except for using 0.5x reagent, all other procedures were performed by following the IP1 detection kit manufacturer’s instructions. Peptides were synthesized by Sangon Biotech Co., Ltd and are aliquoted in 50 nmol/tube and stored at −20 °C until use.

In situ hybridization

pSPT18 vector containing the full sequence of apEP/CCHa precursor or apEP/CCHaRs was constructed for synthesizing sense and antisense RNA probes depending on the sequence insertion direction. PCR was performed to linearize the plasmid and amplify the insert, and then the amplified PCR product was purified using the MiniBEST Agarose Gel DNA Extraction Kit (TaKaRa, 9762). Sense and antisense RNA probes labeled with digoxigenin (DIG) were synthesized using a DIG RNA Labeling Kit (SP6/T7) (Roche, 11175025910).

The ganglia of Aplysia were dissected and fixed overnight at 4 °C with 4% paraformaldehyde (PFA) in PBS. The ganglia were washed and dehydrated in an ascending ethanol series. The ganglia were rehydrated in a descending ethanol series and prehybridized at 55 °C for 8 h and hybridized overnight (12–16 h) at 55 °C in hybridization buffer (50% Deionized formamide, 5 mM EDTA, 5×SSC, 1×Denhardt’s solution, 0.1% Tween-20, and 0.5 mg/ml yeast tRNA) containing 2 ng/μl DIG-labeled antisense RNA probes. After washout of the probes, ganglia were then incubated overnight at 4 °C with a 1:200 dilution of alkaline phosphatase-conjugated anti-digoxigenin antibody (Roche, 11093274910) in PBS containing 0.1% Tween-20, 0.2% bovine serum albumin (BSA), and 1% normal goat serum. After washing with PBST (PBS with 0.1% TritonX-100 and 0.2% BSA) to remove unbound antibodies, ganglia were washed with detection buffer (100 mM NaCl, 50 mM MgCl_2_, 0.1% Tween-20, 1 mM levamisol, and 100 mM Tris-HCl, pH 9.5) and developed with 4.5 μl of nitro blue tetrazolium (NBT) and 3.5 μl of 5-bromo-4-chloro-3-indolyl phosphate (BCIP) (Roche,11681451001) in 1 ml of detection buffer. The staining reaction was monitored visually and stopped by washing with PBS when the level of staining was adequate. The stained ganglia were observed and photographed using a fluorescence microscope (Olympus) with epi-illumination against a white background. Photographs were taken with a Nikon CoolPix 990 digital camera.

Immunohistochemistry

For immunohistochemistry experiments, a polyclonal antibody against apEP/CCHa (KCHGRWAIHACFGGN-NH_2_) was raised by ChinaPeptides Co, Ltd (Shanghai). Specifically, keyhole limpet hemocyanin (KLH) was conjugated with amino group of apEP/CCHa. Immunohistochemistry was performed in whole mount preparations as described previously (13, 150). The tissue was fixed in a buffer (4% PFA, 0.2% picric acid, 25% sucrose, and 0.1 M NaH_2_PO_4_, pH 7.6), for either 3 h at room temperature or overnight at 4 °C. All subsequent incubations were done at room temperature. The tissue was washed with PBSe (154 mM NaCl, 20 mM PB (pH 7.6), 1 mM EDTA-2Na (pH 8), and was permeabilized and blocked by overnight incubation in blocking buffer (10% normal goat serum, 2% Triton X-100, 1% BSA, 154 mM NaCl, 50 mM EDTA-2Na, 0.01% thimerosal, and 10 mM Na_2_HPO_4_, pH 7.4). The primary antibody was diluted 1:500 in blocking buffer and incubated with the tissue for 4 to 7 days. The tissue was then washed twice per day for 2 to 3 days with washing buffer (2% Triton X-100, 1% BSA, 154 mM NaCl, 50 mM EDTA-2Na, 0.01% thimerosal, and 10 mM Na_2_HPO_4_, pH 7.5). After washing, the tissue was incubated with a 1:500 dilution of secondary antibody (Goat Anti-Rabbit IgG Antibody, (H + L) Rhodamine conjugate, Sigma, AP307 R) in blocking buffer for 2 to 3 days and then washed again two times with washing buffer for 1 day and four times with storage buffer (1% BSA, 154 mM NaCl, 50 mM EDTA-2Na, 0.01% thimerosal, and 10 mM Na_2_HPO_4_, pH 7.5) for 1 day. Finally, the tissue was observed and photographed using either a fluorescence microscope (Olympus) or a TCS SP8 MP Multiphoton Confocal Microscope.

Peptide measurements by LC-MS/MS

To characterize the primary structure and PTM of actual apEP/CCHa precursor gene products in the Aplysia CNS, peptide extracts from all central ganglia of the CNS and the EN were sequenced de novo by LC-MS/MS (152, 153). Isolated and desheathed ganglia were homogenized in acidified methanol (80:18:2 MeOH: water: acetic acid) using Precellys bead beater set to the “Soft” method (3 × 15 s, 5800 rpm protocol). Homogenates were collected and centrifuged at 14,000 x g for 15 min to precipitate tissue debris. Resulting supernatants were transferred into another set of vials, dried using a speed-vac. Dry crude extracts were reconstituted in 100 mM ammonium bicarbonate (NH_4_HCO_3_, pH 8.0) at a volume of 200 - 500ul. Samples were mixed thoroughly until fully dissolved, centrifuged, and the resulting supernatant was collected for further processing; the pellet was discarded.

To ensure detection of neuropeptides with disulfide bonds, the ganglion extracts were subjected to reduction and alkylation following reconstitution (73). To reduce disulfide bonds in the CNS extract, dithiothreitol (DTT, 0.5 M stock in water) was added to the samples to the final concentration of 4 mM. The mixtures were aspirated back and forth twice to ensure proper mixing and incubated in a water bath, with agitation, at 60 °C for 30 min. Following incubation, the samples were briefly cooled on ice to reduce temperature.

For alkylation of free thiols, iodoacetamide (IAA) was added to each sample to final concentration of 10 mM. The mixtures were incubated in the dark at 25 °C for 30 min. To quench excess alkylating agent, DTT was added to a final concentration of 10 mM, followed by an additional incubation at 25 °C for 30 min in the dark. Finally, concentrated formic acid was added to each sample at 0.2% to acidify the solution. Samples were then desalted using Pierce C18 spin columns following the manufacturer’s instructions prior to LC–MS/MS analysis. Note that the EN samples were not derivatized in our previous work (13).

To determine the apEP/CCHa precursor gene products in the Aplysia CNS, peptide extracts were characterized using the Bruker nanoElute 2 LC hyphenated to Bruker timsTOF Pro two via CaptiveSpray two source with external column oven (Bruker Daltonics, Billerica, MA). Samples were first injected onto Acclaim PepMap 100 C18 pre-column trap for additional desalting and preconcentration, followed by separation on a Bruker PepSep column (75 μm ID, 1.9 μm particle size, 250 mm length) at a flow rate of 400 nl/min and a column temperature of 35 °C. The mobile phase consisted of solvent A (0.1% formic acid in water) and solvent B (0.1% formic acid in acetonitrile). A gradient of solvent B was applied, starting from 2% and increasing to 10% in 5 min and then to 50% over next 80 min, followed by a rapid ramp to 95% for wash, and finally equilibration with starting conditions. The positive ion mass spectra were acquired in Parallel Accumulation Serial Fragmentation (PASEF) mode with a cycle time of 1.1 s and 1/k0 range 0.6 to 1.6 V s/cm^2^.

Peptide identification by bioinformatics

Raw PASEF data were imported into Peaks Online 12 (Bioinformatics Solutions) and processed for de novo sequencing (154) using the following settings: no enzyme, semi-specific digestion, Precursor accuracy 20 PPM Fragment accuracy 0.05 DA, CCS Error Tolerance was 0, alignment local confidence ≥50% (ALC, %). PTMs included [C] Carbamidomethylation; [A] Amidation; [O] Oxidation (M); [P] Pyro-glu from E; [P] Pyro-glu from Q; [S] Sulfation (TYR). The deduced sequence tags were mapped to the EP/CCHa prohormone via database search protocol in Peaks Online 12. False Discovery Rate for Peptide Spectrum Matches (FDR PSM) was 0.3%, FDR for Peptide Sequences was 1.0%, −10LgP Cutoff for Peptides was 28.44.

Behavioral experiments

Behavioral experiments were performed in three batches, each using animals from separate shipments (average body weight: 107 ± 19.24 g). Animals were food-deprived for 5 days. In each experiment, two weight-matched animals were housed in separate round net cages that were filled with circulating aerated ASW (14–16°C). To ensure that the animals were responsive to food, they were first pretested with a small piece of seaweed. After pretesting, drugs and vehicle were injected into the hemocoel of each animal using a syringe needle. Experimental animals were injected with apEP/CCHa freshly dissolved in 1.5 ml ASW. Control animals were injected with 1.5 ml of ASW. Three minutes later, the feeding session commenced. Animals were hand-fed seaweed (Shirako, Tokyo, Japan) cut in 1 × 10 cm strips, weighing ∼0.1 g, increasing 10-fold when hydrated. This hand-feeding approach has been used in prior study of Aplysia feeding behavior (13, 155, 156).

Consumed seaweed was converted to hydrated weight and normalized to the animal’s body weight. Each strip of seaweed was delivered to the mouth area with forceps. Feeding was terminated after 1 h or when animals stopped ingesting seaweed for 3 min. The experimenter who fed the animals did not know which animal was injected with apEP/CCHa. After feeding sessions, the experimenter gently picked up the animals and released them upside down in the water to test whether they displayed a normal righting reflex, i.e., they turned themselves right side up and began to locomote.

Single-cell RNAseq (SMART-seq)

The sheath covering the ganglia was removed. The neurons were initially identified based on their anatomical location and electrophysiological properties. Following identification, fast green buffer was injected into the target neuron to enhance visibility under microscopic observation. The ganglion was fixed using ethyl alcohol for approximately 1 to 2 min. Subsequently, the cells surrounding the target neuron were carefully removed using forceps. The target neuron was then isolated carefully using scissors and forceps. Once isolated, the single neuron was transferred into a tube containing RNAase inhibitor and lysis buffer (Takara Bio, Cat. No. 635013). The tubes were rapidly placed into liquid nitrogen for flash freezing and then stored at −80 °C. Single-cell RNA sequencing using the switching mechanism at the 5′ end of the RNA template (SMART-seq) technology was performed by Quintara Biosciences Co., Ltd or Annoroad Gene Technology (Beijing) Co., Ltd.

We aligned sequencing data from multiple batches of sequencing using software HISAT2 and the reference genome AplCal3.0 (GCF_000002075.1) and obtained read counts for each gene sample using the featureCounts tool. Subsequently, we employed “sva” package in R version 4.1.1 to eliminate batch effects from the read counts of the multi-batch samples. The “relative counts” in the figures refer to the read counts of each sample after removal of batch effects.

Electrophysiology

Intracellular and extracellular recordings were made as described previously (13, 58, 69, 70, 157, 158). Because apEP/CCHa has myoactive properties similar to other EPs (15, 16, 20), ganglia were pretreated to prevent peptide-induced movement of ganglia during recording sessions. In feeding motor program experiments, we treated ganglia with 0.8% protease IX for 1 to 2 min. For single-cell excitability experiments, we treated ganglia with 0.4% glutaldehyde for 20 to 30 s. Ganglia were desheathed, transferred to a recording chamber containing ∼1.5 mLof ASW (460 mM NaCl, 10 mM KCl, 11 mM CaCl_2_, 55 mM MgCl_2_, and 10 mM HEPES, pH 7.6), continuously perfused at 0.3 mL/min, and maintained at 14 to 17 °C. Intracellular recordings were obtained using 5 to 10 MΩ sharp microelectrodes filled with an electrolyte (0.6 M K_2_SO_4_ plus 60 mM KCl). Extracellular recordings were acquired from polyethylene suction electrodes. Grass S88 and WPI Pulsemaster A300 stimulators were used to provide timing signals for intracellular and extracellular stimulation. Electrophysiological recordings were digitized online using AxoScope (software version 10.7, Molecular Devices, LLC) and were plotted by CorelDraw (version 2018, Corel Corp.).

To determine the effect of apEP/CCHa on Aplysia feeding programs, both the cerebral and buccal ganglia were included. For excitability experiments, the buccal ganglion was included with or without the cerebral ganglion. Freshly made apEP/CCHa solution in ASW was perfused into the recording chamber.

Peptide concentrations for in vitro experiments were set at 10^-6^ M, which is higher than those used in vivo behavioral assays. It is common in Aplysia electrophysiological studies for in vitro concentrations to exceed those used in vivo (13, 14, 67, 159, 160), because peptide access to synaptic and extrasynaptic targets in isolated ganglia relies on passive diffusion rather than active circulation via the hemolymph. Based on extensive prior experience, lower concentrations often produce little or no measurable physiological effect in vitro.

Feeding-motor programs were elicited by stimulation of command-like neuron CBI-2 at 8 to 10 Hz throughout the duration of protraction monitored by activity in the I2 nerve of the buccal ganglion. Excitability experiments were performed in a high divalent saline (368 mM NaCl, 8 mM KCl, 13.8 mM CaCl_2_, 115 mM MgCl_2_, and 10 mM HEPES, pH 7.6), which increases the spiking threshold of neurons and therefore curtails polysynaptic activity.

Statistical analysis

Statistical analysis was performed using Prism software (GraphPad version 8). Data are expressed as the mean ± SD of at least three independent experiments. For comparisons between two groups, data were analyzed by paired two-tailed t test. For comparisons between multiple groups, one-way ANOVA with Tukey post hoc multiple comparisons was used. In all statistical tests, the significance level was set at p < 0.05.

Data availability

All data are included in this article and the Supporting information. The nucleotide sequences reported in this paper have been submitted to the GenBank/EBI Data Bank with accession number(s): PX759564 (apEP/CCHa precursor), PX759565 (apEP/CCHaR1), PX759566 (apEP/CCHaR2).

Supporting information

This article contains supporting information.

Conflict of interest

The authors declare that they have no conflicts of interest with the contents of this article.