The zebrafish genome contains a single sema7a gene that produces two transcripts. One transcript encodes Sema7A-GPI, a full-length, GPI-linked, cell-surface-attached protein. The second transcript yields Sema7A^sec, a protein with a truncated C-terminus, which we conjecture is secreted into the local environment. Each transcript encodes an N-terminal signal sequence and a single copy of the conserved Sema domain (Yazdani and Terman, 2006). To detect the presence of both sema7a transcripts in the hair cells of the lateral-line neuromasts, we used fluorescence-activated cell sorting (FACS) to capture the labeled hair cells (Baek et al., 2022) and isolated total RNA. Using primers that flank the regions encoding the conserved Sema domain and the distinct C-termini of the two transcripts, we performed reverse transcription and polymerase chain reactions (RT-PCR) to identify both the membrane-anchored and the secreted transcript in developing larvae (Figure 1B–D; Figure 1—figure supplement 1A–D). Single-cell RNA sequencing data also have shown that the GPI-linked sema7a transcript is particularly enriched in the immature and mature hair cells of the neuromast during early larval development (Lush et al., 2019), a period when sensory axons of the posterior lateral line contact hair cells and form synapses with them (Nagiel et al., 2008; Dow et al., 2015). Indeed, we have observed Sema7A to be specifically localized in both mature and immature hair cells (Figure 1E; Figure 1—figure supplement 2A–D).

As neuromasts mature, the contacts between sensory axons and hair cells increase in number, become stabilized, and establish synapses (Nagiel et al., 2008; Dow et al., 2015). Because Sema7A can modulate axon guidance (Jeroen Pasterkamp et al., 2003; Carcea et al., 2014) and synapse formation (Inoue et al., 2018), we wondered whether the level of Sema7A also changes during neuromast development. Upon quantifying the average intensity of Sema7A, we found similar expression in rostrally and caudally polarized hair cells of neuromasts at 1.5, 2, 3, and 4 days post fertilization (dpf). The average Sema7A intensity increased significantly over this period in both rostrally and caudally polarized hair cells. At 1.5 dpf, the neuromasts harbor primarily immature hair cells that make minimal contacts with sensory axons and do not form stable association with axonal terminals (Dow et al., 2015). At this stage the average Sema7A intensity in hair cells remained low. By 2–4 dpf the hair cells mature to form stable contacts and well-defined synapses with the sensory axons (Nagiel et al., 2008; Pujol-Martí et al., 2014). The average Sema7A intensity in hair cells at each of these stages rose to levels that significantly exceeded those of 1.5 dpf neuromasts (Figure 1F). This rise in the amount of Sema7A during neuromast maturation supports a role for Sema7A in the guidance of sensory axons and their interaction with hair cells.

We additionally demonstrated an anisotropic distribution of Sema7A along the apicobasal axis of each hair cell. As neuromasts develop through 2, 3, and 4 dpf, Sema7A remained highly enriched in the subapical region of the hair cell, the site of the Golgi network (Figure 1E; Figure 1—figure supplement 2E). This localization suggests that Sema7A—like other proteins of the semaphorin family (De Wit et al., 2005)—is directed to the plasmalemma through Golgi-mediated vesicular trafficking. Using the transgenic line Tg(cldnb:lyn-mScarlet) that expresses membrane-tethered mScarlet in all cells of the neuromast (Dalle Nogare et al., 2020), we detected Sema7A at the plasma membrane and potentially in vesicles (Figure 1—figure supplement 2F–I). While Sema7A maintained a high level in the subapical region, the protein increasingly accumulated at the base of the hair cell: the Sema7A level at the hair cell base was low at 2 dpf, but increased sharply by 3 dpf and 4 dpf (Figure 1G; Figure 1—figure supplement 2J). During this period the hair cells increase in number and mature, whereas the associated sensory axons arborize to contact multiple hair cells and form robust synaptic boutons at their bases (Nagiel et al., 2008; Pujol-Martí et al., 2014). Does progressive basal accumulation of Sema7A facilitate association of sensory axon terminals to the hair cell? To address this question, we utilized doubly labeled transgenic fish Tg(myo6b:actb1-EGFP;neurod1:tdTomato) that marked the hair cells and the sensory axons of the posterior lateral line with distinct fluorophores (Figure 1A; Ji et al., 2018). We quantified the average Sema7A intensity and the accumulation of sensory arbors, measured by td-Tomato intensity, at the bases of the hair cells. Across developmental stages, the basal association of the sensory arbors showed a positive correlation with Sema7A accumulation (Figure 1H, I; Figure 1—figure supplement 3A–E, Video 1). The progressive enrichment of Sema7A at the hair cell base with the sensory axons suggests that Sema7A acts as a mediator of contacts between sensory axons and hair cells.

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If Sema7A guides and restricts the sensory arbors at and around the hair cells clustered in a neuromast, then inactivating Sema7A function should disrupt this process. To test this hypothesis, we obtained a sema7a mutant allele (sema7a^sa24691) with a point mutation that introduces a premature stop codon in the conserved sema domain (Figure 2—figure supplement 1A). The homozygous mutant, hereafter designated sema7a^–/–, should lack both the secreted and the membrane-attached forms of Sema7A. The average intensity of Sema7A immunolabeling in hair cells diminished by 61% in sema7a^–/– larvae (0.34±0.01 arbitrary units) in comparison to controls (0.87±0.01 arbitrary units). In a few cases, we observed minute accumulations of Sema7A at the subapical region, but little or no protein at the base of the mutant hair cells (Figure 2A–E). We quantified the number of hair cells in the neuromasts of the control and the sema7a^–/– larvae. We did not observe significant changes in the hair cell number between the genotypes across developmental stages (Figure 2—figure supplement 1B). Furthermore, we tested the mechanotransduction ability of sema7a^–/– mutant hair cells by labeling with the styryl dye FM 4–64 (Nagiel et al., 2008). The hair cells of the 4 dpf sema7a^–/– larvae incorporated the FM 4–64 as effectively as the age-matched control animals. We quantified the FM 4–64 intensities from 60 hair cells and six neuromasts from three control larvae, and from 60 hair cells and six neuromasts from three sema7a^–/– mutant larvae. The average intensities of FM 4–64 labeling between the genotypes did not show a significant difference, which suggests that the mutation does not impact essential hair cell functions (Figure 2—figure supplement 1C and D; Videos 2 and 3).

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The source data contains comparisn of hair cell numbers, Sema7A intensity, and FM4-64 intensities between the control and the mutant larvae.

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To characterize the impact of the sema7a^–/– mutation on arbor patterning, we utilized doubly labeled transgenic fish Tg(myo6b:actb1-EGFP;neurod1:tdTomato) that distinctly labeled the hair cells and the sensory arbors (Ji et al., 2018). In control neuromasts, the sensory axons approached, arborized, and consolidated around the bases of clustered hair cells (Figure 2F). Although the sensory axons approached the hair cells of sema7a^-/- neuromasts, they displayed aberrant arborization patterns with wayward projections that extended transversely to the organ (Figure 2G). To quantitatively analyze the arborization morphology, we traced the sensory arbors in three dimensions to generate skeletonized network traces that depict–as pseudocolored trajectories–the increase in arbor length from the point of arborization (Figure 2H, I).

To visualize the hair cell clusters and associated arborization networks from multiple neuromasts, we aligned images of hair cell clusters, registered them at their hair bundles, from neuromasts at 2 dpf, 3 dpf, and 4 dpf (Figure 2—figure supplement 1E and F). Throughout development, the arborization networks of control neuromasts largely remained within the contours of the hair cell clusters; the few that reached farther nonetheless lingered nearby (Figure 2J; Figure 2—figure supplement 1G1). This result suggests that an attractive cue retained the axons near the sensory organ. In sema7a^-/- neuromasts, however, the neuronal arbors failed to consolidate within the boundaries of hair cell clusters and extended far beyond them (Figure 2K; Figure 2—figure supplement 1H and J; Video 4).

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We quantified the distribution of the sensory arbors around the center of the combined hair cell clusters. For both the control and the sema7a^-/- neuromasts, the arbor densities peaked proximal to the boundaries of the hair cell clusters. Beyond the cluster boundaries, in control neuromasts at 2 dpf, 3 dpf, and 4 dpf the arbor densities fell sharply at respectively 31.7 µm, 36.6 µm, and 32.7 µm from the center. In sema7a^-/- neuromasts of the same ages, the sensory arbors extended respectively 38.5 µm, 51.4 µm, and 56.6 µm from the center (Figure 2L; Figure 2—figure supplement 1K and L; Figure 2—figure supplement 2A). Furthermore, we quantified the degree of contact of the sensory arbors to their hair cell clusters in individual neuromasts from both control and sema7a^-/- mutants. In control neuromasts, the degree of contact was 83 ± 1% (mean ± SEM) at 2 dpf and 84 ± 1% at 3 dpf, but significantly increased to 90 ± 1% at 4 dpf (Figure 2M; Figure 2—figure supplement 2B). This observation indicates that the sensory arbors reinforce their association to the hair cell clusters as neuromasts develop. However, in sema7a^-/- neuromasts such contact was significantly reduced. Only 69 ± 4% of sensory arbors at 2 dpf were closely associated with the corresponding hair cell clusters, a value that remained at 67 ± 2% at both 3 dpf and 4 dpf (Figure 2M). These findings signify that, with simultaneous disruption of both signaling modalities of Sema7A, the organ fails to restrict the localization of sensory arbors and has significantly diminished contact with them.

Accurate spatial arrangement of the neuronal projections around hair cells is essential for proper retrieval and transmission of sensory information. Mutations that perturb proper assembly of sensory neuronal microcircuits lead to hearing impairment (Diaz-Horta et al., 2016). Quantitative insight into the wiring pattern is therefore essential to identify systematic aberrations in the microcircuit arising from genetic deficits. The three-dimensional skeletonized network traces allowed us to accurately delineate the microcircuit topology across individuals, developmental stages, and genotypes from living animals (Figure 2D and E). To quantitatively analyze the arbor network, we defined five topological attributes: (Lykissas et al., 2007) nodes, denoted as junctions where arbors branch, converge, or cross; (Vaccaro et al., 2022) loops, comprising minimal arbor cycles forming topological holes; (Nguyen-Ba-Charvet et al., 2008) bare terminals, defined as arbors with free ends; (Jeroen Pasterkamp et al., 2003) the node degree, calculated as the number of first-neighbor nodes incident to a node; and (Nagiel et al., 2008) the arbor curvature, measured as the total curvature of a bare terminal (Figure 3A). We quantified these attributes to accurately determine how the circuit topology is established during maturation of the sensory organ in control larvae and how it is impacted by sema7a deficit.

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To determine how internode connectivity is distributed across networks, we quantified the distribution of node degrees along the relative height of the hair cell cluster across individuals, developmental stages, and genotypes. As for most other real-world networks (Newman, 2018), we observed a highly skewed node-degree distribution: the majority of nodes had low node degrees (three and four) and a small number of nodes—often acting as hubs, like the arborization initiation point—had high node degrees (five to seven). The node degree varied from three to seven in control larvae and three to six in sema7a^–/– larvae. Across developmental stages and genotypes, the high node degrees primarily occurred at the base (region from 0 % to 10 % of the cluster height) of the hair cell cluster and acted as hubs for the majority of branching events. Low node degrees primarily occurred along the basolateral region (region from 10 % to 60 % of the cluster height) of the hair cell cluster. The apical region (region from 60 % to 100 % of the cluster height) was sparsely populated with low node degrees. The network’s hierarchical organization along the organ’s axis suggests that arbors receive sensory inputs at multiple levels, allowing the network to carry out robust neural computations. Even though nodes of low degree were most prevalent between genotypes across developmental stages, their distribution along the relative organ axis became significantly different at 4 dpf, with nodes in control larvae localized higher along the axis than in sema7a^–/– larvae (Figure 3A; Figure 3—figure supplement 1A–C; Table 1). These observations indicate that Sema7A regulates both the spatial organization and node-degree complexity of the sensory arbor network.

An average of 9.3 sensory axonal branches approach the hair cell cluster of a neuromast (Dow et al., 2018). Pathfinding of sensory axons around the hair cell cluster results in the formation of axonal fascicles that form numerous loops and bare terminals, which determine the microcircuit topology (Figure 2B and D; Dow et al., 2018). To optimally surround and contact the hair cell cluster, do the sensory arbors prefer to adopt one topological feature over the other? To address this question, we measured the correlation between the number of bare terminals and the number of loops across individuals, developmental stages, and genotypes. In both 2 dpf and 3 dpf larvae, the distributions remained highly scattered across individuals and genotypes. However, by 4 dpf in the control larvae the arbor network contained either a high number of loops with a low number of bare terminals or a high number of bare terminals with a low number of loops. This result suggests that the stereotypical circuit topology observed in the mature organ may emerge through transition of individual arbors from forming bare terminals to forming closed loops encircling topological holes. That we did not observe a similar transition in sema7a^-/- neuromasts of the same age suggests that Sema7A may direct the switch between these two distinct topological features (Figure 3B; Figure 3—figure supplement 1D, E; Table 1).

Measurement of the contour length of both bare terminals and loops additionally revealed that across developmental stages in control larvae the sensory arbors followed a three-step process to choose the appropriate topological feature: arbors shorter than 10 µm always formed bare terminals; arbors between 10 µm and 30 µm began to switch between features, with arbors longer than 20 µm strongly preferring to form loops; and arbors longer than 30 µm always formed loops. These results imply that the switch between bare terminals and loops is partly dependent on the length of the arbors. However, this characteristic topological preference was significantly perturbed in the sema7a^–/– larvae across developmental stages, as many of the sensory arbors longer than 30 µm remained as bare terminals (Figure 3C; Figure 3—figure supplement 1F and G; Table 1). Furthermore, in 4 dpf control neuromasts the majority of the loops contained four nodes and utilized 30–40 µm of arbors to form closed cycles. These conserved topological attributes were significantly perturbed in the sema7a^–/– larvae of same age (Figure 3D and E; Figure 3—figure supplement 2A–D; Table 1). These findings indicate that Sema7A tightly monitors the sensory arbor loop formation, including the preferred loop length and number of nodes per loop, to properly surround and contact the hair cell cluster.

As observed previously, bare terminals in control neuromasts project transverse to the organ and linger nearby, often curving back to form new loops and contact the hair cell cluster (Figure 2D and F). To analyze the propensity of the bare terminals to curl backward, we measured their total curvatures, defined as the sum of the curvature at each point along the length of an arbor. Henceforth, we address the total curvature as the curvature of a bare terminal. We classified the bare terminals in two categories: high-curvature arbors with curled morphology and curvature greater than π/6 (30 degrees); and low-curvature arbors with linear morphology and curvature less than π /6. In control neuromasts 44.8% of arbors had low curvature compared to 56.4% in sema7a^-/- neuromasts at 4 dpf. Concomitantly, 55.2% of arbors had high curvature in control neuromasts compared to 43.6% in sema7a^-/- neuromasts at 4 dpf (Figure 3F; Figure 3—figure supplement 2E, F; Table 1). These observations further confirm our hypothesis that a Sema7A-mediated attractive cue is involved in orienting bare neural arbors toward the hair cell cluster to potentially form loops.

In summary, we propose that Sema7A mediates four key aspects of the topology of the arborization network: (1) the node degree complexity along the organ’s axis; (2) the transition of arbors from bare terminals to loops; (3) the preference for forming loops containing four nodes with 30–40 µm of arbors; and (4) the preference of bare terminals to adopt higher curvature.

Because genetic inactivation of the sema7a gene disrupts both signaling modalities of the cognate protein, we could not assess the specific activity of the Sema7A^sec diffusive cue in guiding lateral-line sensory axons. To independently verify the potential role of that isoform, we therefore expressed the secreted variant ectopically and investigated the resultant arborization patterns.

Analysis of the microcircuit connectivity of the neuromasts has demonstrated bare sensory-axonal terminals in the perisynaptic compartments, where they do not contact the hair cell membrane (Dow et al., 2018). We speculate that these bare terminals are attracted by a Sema7A^sec diffusive cue. To test this hypothesis, we ectopically expressed Sema7A^sec protein tagged with the fluorophore mKate2, then analyzed its effect on the morphology of sensory arbors. Fertilized one-cell embryos were injected with the hsp70:sema7a^sec-mKate2 plasmid and raised to 3 dpf. Larvae expressing the transgenesis marker—the lens-specific red fluorescent protein mCherry—were heat-shocked, incubated to allow expression, and subsequently mounted for live imaging. In these larvae, a random mosaic of cells—often embryonic muscle progenitors in the dermomyotome or mature myofibers—expressed the exogenous Sema7A^sec-mKate2 protein. We selected only those neuromasts in which a single mosaic ectopic integration had occurred near the network of sensory arbors (Figure 4A and B). In all 22 such cases, we observed robust axonal projections from the sensory arbor network toward the ectopically expressing Sema7A^sec cells. When the exogenous sema7a^sec integration occurred in embryonic muscle progenitor cells, which reside in a superficial layer external to the muscle fibers and adjacent to the larval skin (Sharma et al., 2019), the projections were able to form direct contacts with them (Figure 4C; Videos 5 and 6). But more often, when mature myofibers expressed the exogenous Sema7A^sec deep in the myotome, the axonal projections approached those targets but failed to contact them (Figure 4D, Videos 7 and 8). This behavior likely arose because other components in the myotomal niche—including the fibrous epimysium, other myofibers, myoblasts, and immature myotubes whose surroundings are permeable to diverse diffusive cues (Keenan and Currie, 2019)—physically obstructed direct interaction of the extended neurites with the ectopically expressing myofibers. All 18 of the injected but not heat-shocked control larvae did not express ectopic Sema7A^sec, and we did not observe aberrant projections from the sensory arbor network (Figure 4E).

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The source data depicts the accuracy of the 18 extended axonal arbors in finding ectopic Sema7Asec sources.

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To measure the accuracy of the extended axonal projections in finding the ectopic Sema7A^sec source, we defined three parameters: (1) the source path, denoted as the linear distance from the point of arborization to the boundary of an ectopically expressing Sema7A^sec cell; (2) the projection path, representing the linear distance from the point of arborization to the terminus of the extended projection; and (3) the projection proximity, calculated as the linear distance from the terminus of the extended axonal projection to the nearest boundary of the ectopically expressing Sema7A^sec cell (Figure 4B). We quantified these parameters from 18 of the 22 mosaic integration events; in the remainder the extended axonal projections–although following the ectopic source–reentered the posterior lateral-line nerve so that identification of the axon terminals was not possible (Figure 4—figure supplement 1A–A"; Video 9). The projection paths of the extended axons reached from 27.1 µm to 142.5 µm while following the ectopic Sema7A^sec cues. When we plotted the projection path length against the source-path length, we observed a nearly perfect correlation (Figure 4F). This result signifies that irrespective of the location of the ectopic source–whether proximal or distal to the sensory arbor network–the Sema7A^sec cue sufficed to attract axonal terminals from the sensory arbor. Furthermore, the low average projection-proximity length of 3.5±0.7 µm (mean ± SEM) indicated that the extended projection terminals remained either in contact (8 of 18) or in the vicinity (10 of 18) of the ectopic Sema7A^sec sources (Figure 4G). These observations together demonstrate that the Sema7A^sec diffusive cue is sufficient to provide neural guidance in vivo.

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As the posterior lateral-line ganglion matures, newly formed neurons extend their axonal growth cones along the posterior lateral-line nerve to reach the neuromasts (Sato et al., 2010). We wondered whether the axonal projections that had yet to be associated with a neuromast could also respond to an ectopic source of Sema7A^sec. Indeed, on seven occasions in which sema7a^sec integration had occurred between two neuromasts, we detected neurite extensions from the posterior lateral-line nerve to the ectopic Sema7A^sec sources (Figure 4H; Figure 4—figure supplement 1B–B", Videos 10–12). Live time-lapse video microscopy further revealed dynamic association between the extending neurites and the ectopic Sema7A^sec sources. Long and thin along their entire extents, the neurites were highly motile, extending and retracting rapidly near the Sema7A^sec expressing cells. Many neurites maintained their contact with the ectopic sources of Sema7A^sec from one hour to almost eight hours (Figure 4I and J; Figure 4—figure supplement 2A, Videos 13–16). The responsiveness of the posterior lateral-line axons to Sema7A^sec might therefore be an intrinsic property of the neurons that does not require the neuromast microenvironment.

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We speculate that the GPI-anchored transcript variant of the sema7a gene can function only through contact-dependent pathways. To test this hypothesis, we expressed the Sema7A-GPI ectopically and investigated the resultant arborization patterns of the sensory axons. Fertilized one-cell embryos were injected with the hsp70:sema7a-GPI-2A-mCherry plasmid and raised to 3 dpf. Larvae expressing the transgenesis marker—the lens-specific green fluorescent protein (GFP)—were heat-shocked, incubated to allow expression, and subsequently mounted for live imaging (Figure 5A). In these larvae, a few mature myofibers expressed the exogenous Sema7A-GPI protein. We selected only those neuromasts in which an ectopic integration had occurred near the network of sensory arbors. In all 15 such cases, no sensory axon left the arbor network to approach the ectopic Sema7A-GPI sources. In all 17 of the uninjected and heat-shocked control larvae, the sensory arbor network did not show any perturbation (Figure 5B–C, Videos 17–19). These data suggest that the Sema7A-GPI, unlike its secreted counterpart, functions at the surfaces of the expressing cells.

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Contact stabilization between an axon and its target cell is essential for synapse formation (Shen and Cowan, 2010). Neural GPI-anchored proteins can act as regulators of various synaptic-adhesion pathways (Um and Ko, 2017). In the mouse olfactory system, GPI-anchored Sema7A is enriched in olfactory sensory axons and mediates sustained interaction with the dendrites of mitral and tufted cells in the olfactory bulb to establish synapses (Inoue et al., 2018). In the developing mouse brain–particularly in Purkinje cells–GPI-anchored Sema7A instead regulates the elimination of synapses onto climbing fibers (Uesaka et al., 2014). The role of anchored Sema7A in regulating synaptic architecture can thus vary according to the cell type and developmental stage. Our demonstration that Sema7A is necessary to consolidate the contact between sensory axons and hair cells during neuromast development suggests a possible role in regulating synaptic structure.

Proper synaptogenesis requires the correct spatial organization of the presynaptic and postsynaptic apparatus at the apposition of two cells. At the interface between hair cells and sensory axons, the synaptic network involves two scaffolding proteins in particular: ribeye, a major constituent of the presynaptic ribbons (Sheets et al., 2014), and membrane-associated guanylate kinase (MAGUK), a conserved group of proteins that organize postsynaptic densities (Oliva et al., 2012). Utilizing these scaffolding proteins as markers, we investigated the impact of the sema7a^-/- mutation on synapse formation. To analyze the distribution of presynaptic densities–immunolabeled with an antibody against the ribeye-associated presynaptic constituent C-terminal binding protein (CTBP; Sheets et al., 2014)–we counted the number and measured the area of CTBP punctae from multiple neuromasts at several stages of neuromast development (Figure 6A and B). The CTBP punctae in each hair cell ranged from zero to seven across developmental stages. In the zebrafish’s posterior lateral line, the formation of presynaptic ribbons is an intrinsic property of the hair cells that does not require contact with lateral-line sensory axons. However, sustained contact with the sensory axon influences the maintenance and stability of ribbons (Suli et al., 2016). As neuromasts matured from 2 dpf to 4 dpf, we identified similar distribution patterns with a characteristic increase in the number of CTBP punctae in both control and sema7a^-/- larvae (Figure 6C and D). This result implies that the formation of new presynaptic ribbons is not perturbed in the mutants, even though there are on average fewer CTBP punctae (Figure 6—figure supplement 1A–C). Measurement of the areas of CTBP punctae also showed similar distributions in both genotypes (Figure 6E and F) but the average area of presynaptic densities was reduced in the mutants (Figure 6—figure supplement 1D–F, Figure 6—figure supplement 2). Because GPI-anchored Sema7A lacks a cytosolic domain, it is unlikely that Sema7A signaling directly induces the formation of presynaptic ribbons. We propose that the decrease in average number and area of CTBP punctae likely reflects decreased stability of the presynaptic ribbons owing to lack of contact between the sensory axons and the hair cells.

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The source data represents the number and area of pre and post synaptic aggregates across genotypes.

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Our analysis of the skeletonized network traces of the sensory arbor network identified distinct topological features that are resultant of collective assemblies of each sensory axon. Because postsynaptic aggregates are organized at the sensory axons, we wondered whether they are associated with a specific topological feature of the network. To identify the distribution of the postsynaptic aggregates along the arbor network, we utilized the transgenic animal TgBAC(neurod1:EGFP) that labeled the sensory axons of the posterior lateral line in green fluorescence protein (GFP), and marked the postsynaptic aggregates through immunostaining with a pan-MAGUK antibody (Figure 6G, Video 20).

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We generated skeletonized network traces of the arbor network and determined the three-dimensional spatial locations of the MAGUK punctae (Figure 6—figure supplement 2). Comparison of the topological features of the network with the spatial locations of the MAGUK punctae revealed that the postsynaptic densities occur more extensively on the loops than on the bare terminals (Figure 6G, Video 20). We measured 567 MAGUK punctae from 16 sensory arbor networks, where 531 punctae were located on the loops and only 36 punctae on the bare terminals (Figure 6H). Furthermore, the MAGUK punctae predominantly aggregate near the node and their density sharply diminishes moving farther from the node (Figure 6I, Figure 6—figure supplement 2). These observations suggest that the distribution of the postsynaptic aggregates is closely associated with the topological attributes of the sensory circuit. Because in the sema7a^-/- mutants the arbor networks have significantly lower number of loops and nodes, and more bare terminals, we hypothesize that in the mutants the occurrence of postsynaptic aggregates is perturbed. Moreover, Sema7A-GPI-mediated juxtracrine signaling onto the apposed axonal membrane is essential to induce postsynaptic assembly (Inoue et al., 2018).

To characterize the impact of sema7a mutation on the formation of postsynaptic aggregates, we counted the number and measured the area of MAGUK punctae from multiple neuromasts at several stages of neuromast development (Figure 6J and K). At 4 dpf, 68.3% of the hair cells in the control neuromasts were associated with at least one MAGUK punctum. In the sema7a^-/- mutant this value fell to 37.5% (Figure 6L and M). The average number of MAGUK punctae also declined in the mutants (Figure 6—figure supplement 1G–I). As control neuromasts matured from 2 dpf to 4 dpf, the MAGUK punctae showed a shift toward smaller sizes, suggesting that the postsynaptic structure fragmented into smaller entities as observed in other excitatory synapses (Taschenberger et al., 2002). At 4 dpf, 77.5% of the hair cells in the control neuromasts had postsynaptic densities with areas between 0.10 µm² and 0.40 µm². In the sema7a^-/- mutant the corresponding value was only 42.5% (Figure 6N and O). The average area of MAGUK punctae was also reduced in the mutants compared to the controls (Figure 6—figure supplement 1J–L, Figure 6—figure supplement 2). We speculate that the abnormalities in the postsynaptic structure of the sema7a^-/- mutants arose from reduced contact between the hair cells and the sensory axons and perturbed topological attributes of the sensory circuit or from the lack of Sema7A-GPI-mediated juxtracrine signaling onto the apposed neurite terminals (Inoue et al., 2018).

Experiments were performed on 1.5–4 dpf zebrafish larvae in accordance with the standards of Rockefeller University’s Institutional Animal Care and Use Committee (IACUC) with protocol number 22,081 H (PRV 19,076 H). Naturally spawned and fertilized eggs were collected, cleaned, staged, and maintained at 28.5 °C in system-water with 1 μg/mL methylene blue. Wild-type AB and heterozygous mutant semaphorin7a^sa24691 zebrafish were obtained from the Zebrafish International Resource Center. In addition, the following transgenic zebrafish lines were used: Tg(myo6b:actb1-EGFP) (Kindt et al., 2012), Tg(neurod1:tdTomato) (Ji et al., 2018), Tg(cldnb:lyn-mScarlet) (Dalle Nogare et al., 2020), and TgBAC(neurod1:EGFP) (Obholzer et al., 2008).

Immediately before dissection, 4-day-old Tg(myo6b:actb1-EGFP) and wild-type larvae were anesthetized in 600 μM 3-aminobenzoic acid ethyl ester methanesulfonate in system water with 1 μg/mL methylene blue. The anterior half of each larva was removed with a fine surgical blade. We used approximately 400 GFP-positive and 50 GFP-negative larvae, the latter for controls of the gating procedure. Dissociation of the tails were performed using a protocol (Baek et al., 2022) with minor modification; detailed protocol is available upon request. The resuspended cells were incubated for 5 min on ice and sorted with an influx cell sorter (BD Biosciences, San Jose, CA, USA) with an 85 μm nozzle at 20 psi and 1 X DPBS (Sigma-Aldrich, St Louis, USA). Laser excitation was conducted at 561 nm and 488 nm. Distinct populations of GFP-positive hair cells were isolated based on forward scattering, lateral scattering, and the intensity of GFP fluorescence.

Sorted cells were collected in a lysis-buffer solution supplemented with β-mercaptoethanol and RNA extraction was performed immediately using a standard protocol (RNeasy Plus Micro Kit; QIAGEN). The yield and quality of the product were measured with a bioanalyzer (Agilent 2100). Only samples with an RNA integrity score (RIN) greater than 8.0 were selected for the preparation of cDNA libraries using iScript cDNA Synthesis Kit (Bio-Rad). One nanogram of total RNA was amplified to several micrograms of cDNA.

The sema7a gene expresses two transcript variants: variant 1, sema7a-GPI anchored, NM_001328508.1; and variant 2, sema7a^sec secreted, NM_001114885.2. The variant 2 lacks several C-terminal exons, and its 3'-terminal exon extends past a splice site that is used in variant 1. The encoded variant 2 has a shorter and distinct C-terminus than variant 1. To individually identity these transcripts, each of them was PCR amplified from the cDNA library using the following primers: For variant 1, sema7aF = 5´-GGTTTTTCTGAGGCCATTCC-3´, sema7aR1=5´-GGCACTCGTGACAAATGCTA-3´; and for variant 2, sema7aF = 5´-GGTTTTTCTGAGGCCATTCC-3´, sema7aR2=5´-TGTGGAGAAAGTCACAAAGCA-3´. To detect pvalb8 transcript we used the following primers: pvalb8F=5´-ATGTCTCTCACATCTATCCT-3´ and pvalb8R=5´-TCAGGACAGCACCATCGCCT-3´. To detect s100t transcript we used the following primers: s100tF = 5´-ATGCAGCTGATGATCCAGAC-3´ and s100tR = 5´-CTATTTCTTGTCATCCTTCT-3´.

For the immunofluorescence labeling of wholemounted wild-type control and sema7a^-/- larvae, 1.5–4 dpf larvae were fixed overnight at 4 °C in 4% formaldehyde in phosphate-buffered saline solution (PBS) with 1% Tween-20 (1% PBST). Larvae were washed four times with 1% PBST for 15 min each, then placed for 2 hr in blocking solution containing PBS, 2% normal donkey serum (NDS), 0.5% Tween-20, and 1% BSA. Primary antibodies were diluted in fresh blocking solution and incubated with the specimens overnight at 4 °C. The primary antibodies were goat anti-Sema7A (1:200; AF1835, R&D Systems) (Figure 1—figure supplement 4; Carcea et al., 2014), rabbit anti-myosin VI (1:200; Proteus 25–6791), murine anti-GM130 (1:200; 610822, BD Transduction Laboratories), murine anti-CTBP (1:200; B-3, SC-55502, Santa Cruz Biotech), and rabbit anti-mCherry (1:200; GTX128508, GeneTex). Larvae were washed four times with 0.1% PBST for 15 min each. Alexa Fluor-conjugated secondary antibodies (Invitrogen, Molecular Probes) applied overnight at 1:200 dilutions in 0.2% PBST included anti-rabbit (488 nm), anti-goat (555 nm), and anti-mouse (647 nm). Larvae were then washed four times in 0.2% PBST for 15 min each and stored at 4 °C in VectaShield (Vector Laboratories).

For the immunofluorescence labeling of postsynaptic density in wholemounted wild-type control and sema7a^-/- larvae, 2–4 dpf larvae were fixed with 4% formaldehyde in PBS for 4.5–6 hr at 4 °C. Larvae were washed five times with 0.01% PBST for 5 min each and then rinsed in distilled water for 5 min. The larvae were permeabilized with ice-cold acetone at –20 °C for 5 min. They were rinsed in distilled water for 5 min, washed five times with 0.01% PBST for 5 min each, and blocked overnight with PBS buffer containing 2% goat serum and 1% BSA. Murine anti-pan-MAGUK antibody (1:500; IgG1 NeuroMab, K28.86, 75–029 and MABN72, Millipore) was diluted in PBS containing 1% BSA, added to the larvae, and incubated for 3 hr at room temperature. Larvae were washed five times for 5 min each with 0.01% PBST. Alexa Fluor 555-conjugated anti-mouse secondary antibody (1:1000) diluted in PBS containing 1% BSA was added to the larvae and incubated for 2 hr at room temperature. Larvae were washed five times with 0.01% PBST for 5 min each, rinsed in distilled water for 5 min, and stored at 4 °C in VectaShield (Vector Laboratories).

Posterior segments of fixed larvae were mounted on a glass slide and imaged on an Olympus IX81 microscope with a microlens-based, super-resolution confocal microscope (VT-iSIM, VisiTech international) under 60 X and 100 X, silicone-oil objective lenses of numerical apertures 1.30 and 1.35, respectively. Static images at successive focal depths were captured at 200 nm intervals and deconvolved with the Microvolution software in ImageJ.

The average Sema7A fluorescence intensities of the hair cells from 1.5 dpf, 2 dpf, 3 dpf, and 4 dpf neuromasts were measured by determining the mean gray level within each hair cell labeled by actin-GFP expression. The intensity distribution of Sema7A protein across the basal cell membrane was measured using the line profile tool in ImageJ (Figure 1—figure supplement 2G–I). The lines were drawn with 1 μm length, starting from the inside of the hair cell to the outside. For each hair cell membrane, the intensity values were scaled from 0 to 1 arbitrary units. The intensity distribution of Sema7A protein along the apicobasal axis of the hair cell was measured using the line profile tool in ImageJ (Figure 1—figure supplement 2J). Because hair cell lengths differed among samples, lines were drawn with 2 μm widths and varying lengths of 6–8 μm. For each hair cell, apicobasal length was scaled from 0 to 100 arbitrary units and the intensity values from 0 to 1 arbitrary units. The average Sema7A and td-Tomato-positive sensory arbor fluorescence intensities at the hair cell bases from 2 dpf, 3 dpf, and 4 dpf neuromasts were measured by determining the mean gray level within each hair cell basal area, denoted by the region underneath the hair cell nucleus (Figure 1—figure supplement 3A–E).

The semaphorin7a^sa24691 allele harbors an A-to-T point mutation in the seventh exon of the semaphorin7a gene, which creates a premature stop codon (Figure 2—figure supplement 1A). Genomic DNA was isolated from the tail fins of adult semaphorin7a^sa24691 (sema7a^sa91) heterozygous mutant zebrafish. The mutant locus in the genome was PCR amplified using the following primers: sema7a^sa91F: 5´-AAAGCTGGAAAGCGAATCAA-3´ and sema7a^sa91R: 5´-ATATCCAAGGATCCGCCTCT-3´. The 734-base pair amplicons were sequenced, and heterozygous adults were propagated.

Freely swimming 4 dpf control and sema7a^-/- larvae were exposed to the styryl fluorophore FM 4–64 (N-(3-triethylammoniumpropyl)–4-(6-(4-(diethylamino) phenyl) hexatrienyl) pyridinium dibromide, ThermoFischer Scientific) at 5 µM for 30 s, then rinsed three times in fresh embryo water and immediately mounted in low-melting-point agarose for live imaging. The average FM 4–64 intensities within the hair cells of 4 dpf neuromasts of both genotypes were measured by determining the mean gray level within each hair cell labeled by actin-GFP expression.

The pDNR-LIB plasmid containing the sema7a^sec coding sequence was purchased from horizon (7140389, Perkin-Elmer). The coding sequence was PCR amplified using the following primers: sema7a^secF: 5´-GGGGACAAGTTTGTACAAAAAAGCAGGCTTGATGATTCGACATTATTCT-3´ and sema7a^secR: 5´-GGGGACCACTTTGTACAAGAAAGCTGGGTGCTTTGTGGAGAAAGTCACAAAGCA-3´. The italicized nucleotides denote the mutated stop codon. The amplicon was then cloned into the pDONR221 vector by BP recombination to generate the middle entry pME-sema7a^sec vector. To generate the hsp70:sema7a^sec-mKate2 construct, gateway cloning was performed by combining the plasmids p5E-hsp70 (Kwan et al., 2007), pME-sema7a^sec, p3E-mKate2-myc no-pA (Addgene, 80812), and pDESTtol2pACrymCherry (Addgene, 64023) with LR Clonase II Plus (Invitrogen). Verified constructs (25 ng/μl plasmid DNA) were injected with Tol2 Transposase mRNA (approximately 25 ng/μl) into one-cell embryos. The transiently transgenic larvae—identified by the expression of red fluorescent protein (mCherry) in their lenses—were raised till 3 dpf, heat-shocked in a water bath at 37 °C for 1 hr, incubated at 28 °C for 1 hr to allow expression, and subsequently mounted for live imaging.

The sema7a-GPI coding sequence without the stop codon (sema7a-GPI^no-stop) was synthesized using gBlocks HiFi Gene Fragments (Integrated DNA Technologies). The sema7a-GPI^no-stop coding sequence was then cloned into the pDONR221 vector by BP recombination to generate the middle entry pME-sema7a-GPI^no-stop vector, henceforth denoted as pME-sema7a-GPI. To generate the hsp70:sema7a-GPI-2A-mCherry construct, gateway cloning was performed by combining the plasmids p5E-hsp70 (Kwan et al., 2007), pME-sema7a-GPI, p3E-2A-mcherrypA (Addgene, 26031), and pDESTtol2pACryGFP (Addgene, 64022) with LR Clonase II Plus (Invitrogen). Verified constructs (25 ng/μl plasmid DNA) were injected with Tol2 Transposase mRNA (approximately 25 ng/μl) into one-cell embryos. The transiently transgenic larvae—identified by the expression of green fluorescent protein (GFP) in their lenses—were raised till 3 dpf, heat-shocked in a water bath at 37 °C for 1 hr, incubated at 28 °C for 1 hr to allow expression, and subsequently mounted for live imaging.

Living larvae were dechorionated, anaesthetized in 600  μM 3-aminobenzoic acid ethyl ester methanesulfonate in system-water with 1 μg/mL methylene blue, and mounted in 1% low-melting-point agarose on a glass-bottom MetTek dish at specific stages. The larvae were maintained at 28 °C during imaging in a temperature-controlled stage top chamber (OKO Lab UNO-T). Static images at successive focal depths were captured at 200 nm intervals and deconvolved with Microvolution software in ImageJ (NIH). For time-lapse video microscopy, the ectopic sema7a^sec integration sites were imaged at intervals of 5 min as Z-stacks acquired with 1.0 μm steps with laser excitation at 488 nm and 561 nm under a 60 X, silicone-oil objective lens of numerical aperture 1.30. The three-dimensional datasets were processed and volume-rendered with the surface evolver and filament-tracer tools in Imaris (Bitplane, Belfast, UK). The Imaris workstation was provided by the Rockefeller University Bio-imaging Resource Center.

Fluorescent intensity values of the sensory neurites and the ectopic Sema7A^sec sources were measured from the maximal-intensity projections at each time during the time-lapse video microscopy. The intensity values were normalized and binarized before summing across each timepoint for locations across the XY plane. The resulting pseudocolored trajectories represent the dwell times of the neurites and the Sema7A^sec cells through the 10 hr time-lapse video sequence.

The labeled sensory axons were traced in three dimensions with ImageJ’s semi-automated framework, SNT (Arshadi et al., 2021). Each trace depicts the skeletonized form of the posterior lateral-line nerve, the posterior lateral-line branch, and the point of arborization from which the sensory arbors radiate to contact the hair cells (Figure 2D and E; Figure 2—figure supplement 2B). The three-dimensional pixels or voxels obtained from the skeletonized sensory arbor traces were processed using custom code written in Python for visual representation and quantitative analysis.

The hair cell clusters of neuromasts from each developmental stage were aligned by the centers of their apices, which were identified by the hair bundles, and overlayed to generate combined hair cell clusters (Figure 2—figure supplement 1E and F) using custom Python code. The associated skeletonized sensory-arbor traces at each developmental stage were aligned similarly using custom Python code. The details of the procedures are available upon request and the corresponding code is available on GitHub.

The region from the center of the combined hair cell cluster to 60 μm in the post-cluster region was divided into 3600 concentric sections of equal area. Each section, as depicted with a distinct color shade, had an area of π μm². Sensory arbor density was defined as Log₁₀[(Area occupied by the arbors in each section)/ (Area of each section)]. The arbor density was then plotted as a histogram against distance from the center of the combined hair cell cluster. Each bin of the histogram represents an area of π µm² (Figure 2—figure supplement 2A). The histograms were finally represented as density trace graphs with Kernel Density Estimation (KDE) in Python.

For each hair cell cluster, the voxels of the sensory arbor traces that were inside or within 0.5 µm—the average neurite radius—of the cluster boundary were denoted as arbors in contact with their corresponding hair cell cluster (Figure 2—figure supplement 2B). The percentage of arbors in contact with each hair cell cluster was defined as: (Number of voxels that remained inside the cluster boundary/Total number of voxels in the arbor) X 100. The data were analyzed through an automated pipeline with custom Python code.

We developed a custom Python script (snt_topology.py; available on GitHub) to convert the skeletonized sensory arborization networks to network graphs (NetworkX, v3.1). We first used a custom ImageJ macro to convert SNT skeletons to csv files, then generated the network graph from the individual arbor paths. As noted previously, we defined five topological attributes: (1) nodes—junctions where arbors branch, converge, or cross; (2) loops—minimal arbor cycles forming topological holes; (3) bare terminals—arbors with free ends; (4) node degree—the number of first-neighbor nodes incident to a node; and (5) arbor curvature—the total curvature of a bare terminal (Figure 3A). To enumerate loops and quantify their lengths and node numbers, we computed the minimum cycle basis for each network graph. The relative organ height of each node in a graph was defined in 3D using Equation 1:

where ${\displaystyle \overrightarrow{\mathrm{u}}}$ is the vector from the arborization initiation point to a node and ${\displaystyle \overrightarrow{\mathrm{v}}}$ is the vector from the arborization initiation point to the hair cell cluster apex. The relative occurrence of nodes (Figure 3—figure supplement 1A–C) was normalized by both the number of hair cells per neuromast and the total number of neuromasts, for comparison across ages and genotypes. The relative number of bare terminals and loops (Figure 3B; Figure 3—figure supplement 1D, E) was normalized by the number of hair cells per neuromast and scaled between 0 and 1. The density was then normalized by the total number of neuromasts imaged for each developmental stage and genotype. For Figure 3C and Figure 3—figure supplement 1F, G, the normalized occurrence was computed as the number of arbors of a given length that formed either a bare terminal or a loop, divided by the total number of arbors of either feature of that length. Lastly, we defined bare terminals as having either high or low curvature (Figure 3F; Figure 3—figure supplement 2E and F) if their absolute total signed curvature [Equation 2] in three dimensions was either higher or lower than π/6, respectively:

where $s$ is the arc-length parameter of the bare terminal, $T\left(s\right)$ is the unit tangent vector at each point along the arbor, $N\left(s\right)$ is the unit normal vector at each point, and the integral is evaluated from the terminal’s branching node to the arbor’s tip. For a closed curve forming a full loop, this value will equal 2π. In practice, we first fit B-splines (scipy.interpolate.splprep; SciPy, v1.10.1) in 3D to the skeletons for each bare terminal before evaluating their derivatives (scipy.interpolate.splev) and integrating (scipy.integrate.simpson) over their lengths.

The sensory arbor networks were traced in three dimensions with SNT. The surface of each of the ectopically expressing Sema7A^sec-mKate2 cells was represented by a single point that was closest to the terminal point of the guided neurite. Distances between two points were calculated using standard mathematical operations.

Maximal-intensity projections of consecutive through-focus scans encompassing the entire neuromast of control and sema7a^-/- mutant larvae at 2 dpf, 3 dfp, and 4 dpf were utilized to capture the presynaptic and postsynaptic assemblies. The number of the synaptic densities across genotypes were measured using ImageJ’s cell counter framework. Area of the synaptic densities from the maximal-intensity projections were measured using ImageJ’s polygon selection tool (Figure 6—figure supplement 2A–D).

Four-day-old TgBAC(neurod1:EGFP) larvae were fixed and stained for sensory arbors and postsynaptic aggregates marked by GFP and MAGUK, respectively. The GFP-positive sensory arbors were traced in three dimensions with ImageJ’s semi-automated framework, SNT. The topological attributes of the arbor network, such as the loop and the bare terminal, were determined as described previously. The localizations of the MAGUK punctae along the three-dimensional arbor network were measured using ImageJ’s cell counter framework (Figure 6—figure supplement 2E and F). We measured 567 MAGUK punctae from 16 sensory arbor networks. Whether the MAGUK-punctae were associated with a loop or a bare terminal, we categorized them in two groups and quantified their distribution across 16 neuromasts.

The three-dimensional space surrounding each node of the arbor network were divided into concentric shells of equal volume (Figure 6—figure supplement 2E, F). For each node, we computationally generated 100,000 concentric shells that reached a 10 µm radius. We measured the number of MAGUK punctae located within each of those concentric shells from their nearest nodes.

Data visualization and statistical analysis were conducted with GraphPad Prism (Version 9) and Python (v3.10). The Mann-Whitney test was used for hypothesis testing unless otherwise noted, and the statistical details are given in the respective figure legends.

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

The authors thank Nicolas Velez and Emily Atlas for valuable discussions. Samantha Campbell and Anna Kaczynska provided expert fish husbandry. Katie Kindt kindly provided various strains of zebrafish. Image processing benefitted from facilities at the Bio-Imaging Resource Center. AD was supported by a Kavli Neural Systems Institute Postdoctoral Fellowship from The Rockefeller University. CCR is supported by National Science Foundation Graduate Research Fellowship Grant No. 1946429. AJ, SPP, and LMS were supported by Howard Hughes Medical Institute, of which AJH is an Investigator.

Experiments were performed on 1.5 to 4 dpf zebrafish larvae in accordance with the standards of Rockefeller University's Institutional Animal Care and Use Committee (IACUC) with protocol number 22081-H (PRV 19076-H).

1. Preprint posted: June 8, 2023
2. Sent for peer review: July 10, 2023
3. Reviewed Preprint version 1: September 12, 2023
4. Reviewed Preprint version 2: April 8, 2024
5. Reviewed Preprint version 3: May 17, 2024
6. Version of Record published: August 12, 2024

You can cite all versions using the DOI https://doi.org/10.7554/eLife.89926. This DOI represents all versions, and will always resolve to the latest one.

© 2023, Dasgupta et al.

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