Mechanism of barotaxis in marine zooplankton

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This fundamental study addresses the question of how certain zooplankton achieve barotaxis, directed locomotion in response to changes in hydraulic pressure. The authors provide compelling evidence that the response involves ciliary photoreceptors interacting with motoneurons. This work should be of broad interest to scientists working on mechanosensation, cilia, locomotion, and photoreceptors.

https://doi.org/10.7554/eLife.94306.3.sa0

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Abstract

Hydrostatic pressure is a dominant environmental cue for vertically migrating marine organisms but the physiological mechanisms of responding to pressure changes remain unclear. Here, we uncovered the cellular and circuit bases of a barokinetic response in the planktonic larva of the marine annelid Platynereis dumerilii. Increased pressure induced a rapid, graded, and adapting upward swimming response due to the faster beating of cilia in the head multiciliary band. By calcium imaging, we found that brain ciliary photoreceptors showed a graded response to pressure changes. The photoreceptors in animals mutant for ciliary opsin-1 had a smaller sensory compartment and mutant larvae showed diminished pressure responses. The ciliary photoreceptors synaptically connect to the head multiciliary band via serotonergic motoneurons. Genetic inhibition of the serotonergic cells blocked pressure-dependent increases in ciliary beating. We conclude that ciliary photoreceptors function as pressure sensors and activate ciliary beating through serotonergic signalling during barokinesis.

Introduction

Hydrostatic pressure increases linearly with depth in the ocean and planktonic organisms can use it as a depth cue, which is independent of light or the time of the day (Blaxter, 1978). Many marine invertebrate animals have long been known to sense and respond to changes in pressure (Knight-Jones and Qasim, 1955; Rice, 1964). The response generally consists of an increase in locomotion (barokinesis) upon an increase in pressure. Such responses could help planktonic animals retain their depth either in combination with, or independent of light cues (Forward et al., 1989). Changes in hydrostatic pressure may additionally entrain tidal rhythms in marine animals (Akiyama, 2004; Morgan, 1965; Naylor and Williams, 1984). Early studies on the barokinetic response in zooplankton have not revealed if the animals respond to relative or absolute changes in pressure. The kinematics and neuronal mechanisms of pressure responses have also not been characterized in detail for any planktonic animal. The most familiar structures for sensing changes in hydrostatic pressure are gas-filled compressible vesicles such as the swim bladder in fish (Qutob, 1963). However, barokinetic responses are seen across many animals without any identifiable gas-filled vesicles. What structures could mediate pressure sensing in these organisms? Thus far, only a few alternative structures have been proposed for pressure sensation. In the statocyst of the adult crab Carcinus maenas, millimeter-sized thread-hairs may act as a syringe plunger to sense pressure (Fraser and Macdonald, 1994). In dogfish, which lack a swim bladder, hair cells in the vestibular organ have been proposed to act as pressure detectors (Fraser and Shelmerdine, 2002). It is unknown which, if any, of these two vastly different pressure sensing mechanisms—one based on volume changes in a gas-filled vesicle and the other on deformation of sensory cilia—is used by the much smaller planktonic animals.

To understand the behavioural and neuronal mechanisms of pressure responses in marine zooplankton, we studied the planktonic ciliated larvae of the marine annelid Platynereis dumerilii (Özpolat et al., 2021). This larva uses ciliary beating to swim up and down in the water column to eventually settle on sea grass beds near coastal regions (Gambi et al., 1992). The sensory and neuronal bases of light-guided (Gühmann et al., 2015; Randel et al., 2014; Verasztó et al., 2018) and mechanically driven behaviours (Bezares-Calderón et al., 2018) in Platynereis larvae have been dissected due to the experimental tractability of this system. Its small size has allowed the entire reconstruction of the cellular and synaptic wiring map of the 3-day-old larva (Jasek et al., 2022; Verasztó et al., 2024). Here, we study Platynereis larvae to understand the cellular and neuronal bases of pressure sensation in zooplankton.

Results

Platynereis larvae respond to changes in hydrostatic pressure

To determine whether Platynereis larvae respond to changes in hydrostatic pressure, we developed a custom behavioural chamber with precise pressure control. We subjected larvae to step changes in pressure and recorded their behaviour under near-infrared illumination (Figure 1A; see Materials and methods). We used hydrocarbon-free compressed air to increase pressure in the chamber. We tested a range of pressure levels in randomized order from 3 to 1000 mbar (1 mbar equals to 1 cm water depth) (Figure 1—figure supplement 1A). We focused on 1- to 3-day-old larvae corresponding to the early and late trochophore and nectochaete stages. We used batches of >100 larvae for each experiment. Both 2- and 3-day-old larvae respond to pressure increase by swimming upwards faster and in straighter trajectories, as quantified by changes in average vertical displacement, swimming speed, the ratio of upward to downward trajectories (Figure 1B, C, Figure 1—figure supplement 1B–D; Video 1), and a straightness index (the net over total distance) (Figure 1—figure supplement 1E).

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Pressure response in *Platynereis* larvae.

(A) Schematic of the behavioural setup used to stimulate larvae with controlled pressure levels. Vertical displacement of (B) 2-day-old and (C) 3-day-old larvae (insets) as a function of time relative to different step increases in pressure. Dashed line at 60 s indicates the end of stimulation. Each data point is the average of 2–12 (B) or 4–5 (C) batches of larvae. Maximum increase in relative vertical displacement of (D) 2-day-old and (E) 3-day-old larvae for each pressure level tested. Small filled circles represent individual data points, data points from the same batch are joined by lines. Larger open circles indicate the mean across all batches. (F) Vertical displacement of 3-day-old larvae acclimated for ca. 10 min to either ambient pressure (0 mb, left), 100 mb (centre), or 500 mb (right) prior to the experiment. Lines are coloured by set fractional increments in pressure. (G) Maximum increase in relative vertical displacement of 3-day-old larvae acclimated to 0, 100, or 500 mb. The data are fitted with a saturation curve. 5 (0, 100 mb) or 6 (500 mb) batches tested in F–G. (H) Maximum ciliary beat frequency (CBF) that single larvae reached in the 30 s prior (category Before), or during the first 30 s (Stimulus) of the indicated increase in pressure. N = 18–22 larvae. Data points for the same larva are joined by lines. One-tailed paired t-test with Bonferroni correction testing for an increase in CBF; p-values <0.05 are shown. Error bars in B, C, and F show the standard error of the mean. Data in D, E, and G were fitted with a third (**D, E**) or a second (G) order polynomial function. Figure 1—source data 1 (**A, D**), Figure 1—source data 2 (**C, E**), Figure 1—source data 3 (**F, G**), Figure 1—source data 4 (H).

Figure 1—source data 1 Swimming metrics of mixed batches of 2-day-old wildtype larvae subjected to randomized step increases in pressure.: https://cdn.elifesciences.org/articles/94306/elife-94306-fig1-data1-v1.csv
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Figure 1—source data 2 Swimming metrics of mixed batches of 3-day-old wildtype larvae subjected to randomized step increases in pressure.: https://cdn.elifesciences.org/articles/94306/elife-94306-fig1-data2-v1.csv
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Figure 1—source data 3 Swimming metrics of mixed batches of 3-day-old wildtype larvae subjected to randomized step increases in pressure after acclimatization to higher than ambient pressures for 10 min.: https://cdn.elifesciences.org/articles/94306/elife-94306-fig1-data3-v1.csv
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Figure 1—source data 4 Ciliary-band dynamics of individual 2-day-old wildtype or c-opsin-1 mutant larvae subjected to randomized step increases in pressure.: https://cdn.elifesciences.org/articles/94306/elife-94306-fig1-data4-v1.csv
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Figure 1—source data 5 Swimming metrics of 1-day-old-larvae subjected to randomized step increases in pressure.: https://cdn.elifesciences.org/articles/94306/elife-94306-fig1-data5-v1.csv
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Figure 1—source data 6 Swimming metrics of 3-day-old-larvae subjected to randomized step increases in pressure using a water column of different heights.: https://cdn.elifesciences.org/articles/94306/elife-94306-fig1-data6-v1.csv
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Figure 1—source data 7 Swimming metrics of 3-day-old-larvae subjected to randomized linear increases in pressure.: https://cdn.elifesciences.org/articles/94306/elife-94306-fig1-data7-v1.csv
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Video 1

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posterframe for video — Response of 2-day-old *Platynereis* larvae to a pressure increase of 1 bar.

Larvae were placed in a pressure vessel (100 mm height) and recorded in effective darkness (with 850 nm light) with a camera placed in front of the vessel (see Figure 1A). Larvae swim towards the top of the chamber as soon as pressure is increased. Blue traces (10 s long) delineate swimming trajectories.

We used the maximal vertical displacement value (normalized to the mean displacement per trial prior to stimulus presentation) to compare the magnitude of responses as a function of pressure change. In both 2- and 3-day-old larvae, the responses were graded: higher pressure levels led to higher maximal vertical displacement (Figure 1D, E). Three-day-old larvae had a slightly lower sensitivity threshold (10–20 mbar) than 2-day-old larvae (20–30 mbar) and their response plateaued at lower pressure levels than that of 2-day-old larvae (Figure 1B–E). One-day-old larvae did not show a detectable response to even the largest pressure levels tested (Figure 1—figure supplement 2A, B). The straightness of trajectories also increased with increasing pressure changes. This was due to narrower helical swimming paths under pressure, visible in close-up videos (Video 2 and Video 3). Three-day-old larvae also showed a diving response upon the release of pressure (Figure 1B, C, F; Figure 1—figure supplement 1C, D). Two-day-old larvae stopped moving upwards when the stimulus ended, but did not show an active diving response to pressure OFF.

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To exclude the possibility that either the changes in the partial pressure of gases due to the use of compressed air, or the mechanical wave associated to the inflow of air caused the upward swimming behaviour, we also used a static column of water of different heights to change pressure levels (Figure 1—figure supplement 2C). We observed the same dependence of vertical displacement on the magnitude of pressure change in this setup (Figure 1—figure supplement 2D, E). Overall, our experiments uncovered a graded, saturable, and highly sensitive upward swimming ON response to increased pressure in Platynereis larvae and a diving OFF response (in 3-day-old larvae only).

Platynereis larvae respond to relative changes of pressure

Larvae may either detect absolute pressure levels, relative changes, or the rate at which pressure changes (Morgan, 1984). To differentiate between these possibilities, we first exposed larvae to linear increases of pressure with rates between 0.05 and 2.6 mbar s⁻¹ (Figure 1—figure supplement 3A). We used a second-degree polynomial function for rate categories 0.3–0.7 mbar s⁻¹ (analysis of variance [ANOVA], p = 6.5e−3) and 0.9–1.3 mbar s⁻¹ (ANOVA, p = 5.8e−8), as it described the relationship between vertical displacement and pressure more accurately than a simple linear model. The difference between a linear and a second-degree polynomial fit was not significant for rates >1.3 mbar s⁻¹ (ANOVA, p ~ 0.1). These results suggest that larvae compensate for the increase in pressure by a corresponding increase in upward swimming when rates of pressure increase are sufficiently high.

The linear response to a gradual increase in pressure suggests that larvae detect changes in pressure, rather than absolute pressure levels. To directly address this, we acclimated 3-day-old larvae for ca. 10 min to either 100 or 500 mbar pressure above the atmospheric level. We then tested a range of randomized step increases in pressure levels (Figure 1—figure supplement 3D). After the acclimation period, the distribution of larvae exposed to 100 or 500 mbar was not different from the larvae kept at ambient levels (two-sided Kolmogorov–Smirnoff test, 0–100 mbar: p = 0.915, 0–500 mbar: p = 0.0863) (Figure 1—figure supplement 3E, F). Upon step increase, larvae reacted with graded upward swimming even if they were pre-exposed to 100 or 500 mbar (Figure 1F). The sensitivity decreased when larvae were acclimated to 500 mbar (Figure 1G). When pressure was released at the end of the increase trials, larvae pre-exposed to 500 mbar showed a downward displacement followed by an upward displacement as soon as pressure was increased back to the corresponding basal level (Figure 1—figure supplement 3G). The downward displacement resembled the magnitude of the diving response we observed for 3-day-old larvae (Figure 1—figure supplement 3H).

Our experiments suggest that Platynereis larvae react to relative increases in pressure in a graded manner proportional to the magnitude of the increase. The response is adaptable and occurs at very different basal pressures (0 or 500 mbar—corresponding to surface or 5 m of water depth). This hints at a pressure-gauge mechanism to regulate swimming depth by compensating for vertical movements due to down-welling currents (Genin et al., 2005), sinking when cilia are arrested (Verasztó et al., 2017) or downward swimming (e.g. during UV avoidance; Verasztó et al., 2018).

Ciliary beat frequency increases with pressure

To understand the mechanism by which larvae regulate swimming in response to an increase in pressure, we analysed the effect of pressure on ciliary beating in the prototroch—the main ciliary band that propels swimming in 2- and 3-day-old Platynereis larvae. Individual 2-day-old larvae were tethered to a glass cuvette from the posterior end with a non-toxic glue previously used in Platynereis larvae (Bezares-Calderón et al., 2018). The cuvette was inserted into a custom-made pressure vessel placed under a microscope. We applied 60 s step increases in pressure in a randomized order and recorded ciliary beating in effective darkness (Figure 1—figure supplement 4A, B).

The mean ciliary beat frequency (CBF) increased as soon as a step change in pressure was applied, with larger step changes showing more noticeable increases in beat frequency (Figure 1—figure supplement 4C, Video 4). The maximum ciliary beat frequency (max. CBF) during the stimulus period showed a statistically significant increase for all but the lowest pressure steps tested relative to the period before the onset of the stimulus (85 mb p = 0.046, 237.5 mb p = 2.08E−05, 556 mb p = 8.35E−07, 988 mb p = 7.8E−07; one-tailed paired t-test with Bonferroni correction testing for an increase in CBF; Figure 1H). The related relative measure of maximum percent change (max. ∆%CBF) also showed an increase under pressure (Figure 1—figure supplement 4D). Overall, these data suggest that rapid upward swimming under pressure is due to an increase in the beating frequency of prototroch cilia that is proportional to the change in pressure.

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Brain ciliary photoreceptor cells show graded activation under increased pressure

To identify the pressure-sensitive cells in Platynereis larvae, we developed an approach to couple imaging of neuronal activity with pressure increases (Figure 2A). We injected fertilized eggs with mRNA encoding the calcium indicator GCaMP6s (Chen et al., 2013)—an indirect reporter of neuronal activity—and embedded injected larvae in low-melting agarose. Mounted larvae were introduced into a custom-built microscopy chamber, where pressure could be increased using compressed air. To provide morphological landmarks and to correct for Z-shifts during imaging, we co-injected larvae with an mRNA encoding the membrane-tagged reporter palmitoylated tdTomato.

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Ca²⁺ imaging of *Platynereis* larvae during pressure increases.

(A) Ca²⁺ imaging preparation to analyse neuronal activation upon pressure stimulation. Left: side view of the three-dimensional (3D) model of the microscopy pressure chamber. Right: larvae embedded in agarose are placed on a round coverslip. An enclosure around the embedded larva serves to keep it under water. Bottom: the enclosed larva on the coverslip is inserted in the central hole of the chamber. A screwable lid secures the coverslip to the chamber and prevents air leaks. Pressure is increased with compressed air entering from one of the inlets. (B, top) Maximum intensity projection of a Z-stack acquired before (left) or during (right) the pressure stimulus of a 2-day-old larva expressing GCaMP6s. (B, bottom) Enlarged views of the corresponding regions highlighted with dashed squares in the top panels. Asterisks mark the position of cell nuclei of the four cells activated by pressure. (C) Still images of a ciliary photoreceptor cell (cPRC) acquired at different time points relative to increase in pressure (t = 0, Video 5). The time points are indicated on the upper left of each panel. Pressure level is also indicated. (D) Maximum intensity projection of a GCaMP6s Z-stack during raised pressure (white channel) and of a Z-stack of the same larva after immunofluorescence (IF) with NIT-GC2, a marker for cPRC cilia (Jokura et al., 2023), and for serotonin (cyan channel). Anterior view in **B–D**. (E) Mean ∆R/R in cPRC_l1 across different step increases in pressure as a function of time of stimulation. Dashed line at 60 s marks the end of stimulus. N = 8 larvae. Error bars show the standard error of the mean. (F) Max. ∆R/R in cPRC_l1 as a function of pressure level. Data points from the same larva are joined by lines. Regression line fitting the data is also shown. One-tailed unpaired t-test with Bonferroni correction testing for an increase in Max. ∆R/R with pressure. p-values <0.05 are shown. Figure 2—source data 1 (**E, F**).

Figure 2—source data 1 Raw intensity values of GCaMP6s (GC) and tdTomato (Tom) channels in cPRCs of 2-day-old larvae subjected to randomized step increases in pressure. dF and dR metrics calculated from raw values are also included.: https://cdn.elifesciences.org/articles/94306/elife-94306-fig2-data1-v1.csv
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By imaging the entire larva before and during the pressure stimulus, we found a group of four cells in the dorsomedial brain that showed consistent increases in GCaMP6s fluorescence when pressure increased (Figure 2B; Figure 2—figure supplement 1). Time-lapse (TL) recordings of these cells revealed that they had prominent cilia, which become visible by the increase in GCaMP6s signal during pressure increase (Figure 2C, Figure 2—figure supplement 1, Video 1). The position, number, size, and morphology of these cells closely matched to that of the previously described brain ciliary photoreceptor cells (cPRCs) (Arendt et al., 2004; Tsukamoto et al., 2017; Verasztó et al., 2018). Immunostaining of the same larvae that were used for Ca²⁺ imaging with an antibody raised against NIT-GC2, a marker of cPRC cilia(Jokura et al., 2023), followed by image registration directly confirmed that the four cells activated under pressure were the cPRCs (Figure 2D).

To characterize the response of cPRCs to pressure, we applied a randomized set of pressure increases (Figure 2—figure supplement 2A) to 2-day-old larvae expressing GCaMP6s while recording fluorescence changes in the four cPRCs. All four cPRCs responded to the pressure levels tested (Figure 2E; Figure 2—figure supplement 2B, C). The increase in GCaMP6s signal was observed in both the cell body and in the cilia of the cPRCs (Figure 2C, Video 5). This response—like that observed at the behavioural level—was graded and increased proportionally to the pressure change (Figure 2E, F; Figure 2—figure supplement 2B, C, E). The difference in the response between pressure levels was statistically significant for some of the cPRCs (Figure 2F; Figure 2—figure supplement 2C). The calcium signal also decreased rapidly after stimulus onset. Therefore, cPRCs may be able to directly encode the intensity of the stimulation in their activity, reflected in their internal Ca²⁺ levels, and adapt to pressure levels. Their unique sensory morphology and pressure-induced Ca²⁺ dynamics make the cPRCs candidate pressure receptors.

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An additional unpaired sensory cell on the dorsal side was also activated in some of the trials (Figure 2—figure supplement 1, green asterisk). We refer to this cell here as SN^d1_unp (by position and morphology it corresponds to the neurosecretory cell SN^YFa+; Williams et al., 2017). At 750 mb, this cell responded by a transient but robust increase at stimulation onset, but ∆R/R dropped to basal values before the end of the stimulus, unlike the cPRC response (Figure 2—figure supplement 2D). SN^d1_unp (SN^YFa+) may also contribute to the pressure response, although it is less sensitive than cPRCs and has very few synapses (Williams et al., 2017).

Another indirect observation that is consistent with cPRCs being the primary pressure receptors is that 1-day-old larvae that lack differentiated cPRCs (Fischer et al., 2010) do not respond to pressure (Figure 1—figure supplement 2A).

c-opsin-1 mutants have a reduced pressure response

cPRCs express ciliary-opsin-1 (c-ops-1), which forms a UV-absorbing photopigment (Arendt et al., 2004; Tsukamoto et al., 2017; Verasztó et al., 2018). Knocking out the c-ops-1 gene abolishes a UV-avoidance response in Platynereis larvae (Verasztó et al., 2018). As cPRCs respond to pressure increases, we tested whether c-ops-1 knockout mutants (c-ops-1^∆8/∆8) also showed a defect in the pressure response.

A range of step increases in pressure were applied to single batches of either wild-type (WT) or c-ops-1^∆8/∆8 3-day-old larvae (Figure 3—figure supplement 1A). The assays were carried out in a smaller pressure vessel (height: ~4 cm), to allow consistent imaging of the fewer mutant larvae available. The swimming speed of c-ops-1^∆8/∆8 mutant larvae was not significantly different from WT larvae (p = 0.118, unpaired Wilcoxon test for lower speed in mutants; Figure 3—figure supplement 1B). c-ops-1^∆8/∆8 larvae responded in a graded manner to increases in pressure by upward swimming (Figure 3A; Figure 3—figure supplement 1C). However, their response was weaker than the response of age-matched WT larvae (Figure 3—figure supplement 1C). An ANOVA comparison showed that a model considering the genotype better explained the data than a model without this variable, for either a simple or a polynomial linear regression model (p-values = 9.98e⁻⁰⁷ and 9.92e⁻⁰⁶, respectively). Ciliary beating prior to pressure increase was not significantly different between c-ops-1^∆8/∆8 and WT larvae (p = 0.835, unpaired Wilcoxon test for lower CBF in mutants; Figure 3—figure supplement 1D). Upon pressure increases, CBF in c-ops-1^∆8/∆8 larvae showed a significant increase to the three highest pressure levels tested (237.5 mb p = 0.014, 556 mb p = 9.7E−05, 988 mb p = 6.35E−05; one-tailed paired t-test with Bonferroni correction testing for an increase in CBF; Figure 3B). c-ops-1^∆8/∆8 larvae also showed significant increases in max. ∆%CBF as the pressure stimulus was increased (Figure 3—figure supplement 1E; compare to the WT data in Figure 3B), with no significant difference in the increase to WT larvae (Figure 3—figure supplement 1F). In summary, c-ops-1^∆8/∆8 larvae can still respond to changes in pressure in a graded manner, but the response is weaker than in WT larvae, both at the population and at the single-larva levels. This indicates that c-opsin-1 is not directly required for the pressure response, but its absence leads to a weakened response to pressure.

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*c-ops-1*^∆8/∆8 larvae show a weaker response to pressure and have structural defects in ciliary photoreceptor cell (cPRC) cilia.

(A) Maximum change in vertical displacement of wild-type (WT) and *c-ops-1*^∆8/∆8 3-day-old larvae for each pressure step-change tested. Data points from the same batch are joined by lines. Larger open circles indicate the mean value. Thicker solid lines show the regression model predictions. N = 7–9 (*c-ops-1*^∆8/∆8), N = 8–10 (WT) batches. (B) Max. ciliary beat frequency (CBF) that individual *c-ops-1*^∆8/∆8 larvae reached in the 30 s prior (Before), or during the first 30 s of the indicated increase in pressure (Stimulus). Data points for the same larva are joined by lines. One-tailed paired t-test with Bonferroni correction testing for an increase in CBF; all p-values are shown. N = 10–17 larvae. (**C, D**) cPRC ciliary volume measured for each pair of cPRC cilia on the left and right body sides in 2-day-old WT and *c-ops-1*^∆8/∆8 larvae. Volumes were measured using the signal of the NIT-GC2 antibody (α-NIT-GC2). (C) Maximum intensity projections of IF stainings used for quantifying cPRC cilary volume. (D) cPRC ciliary volume distribution sorted by genotype and body side. One-tailed unpaired t-test with Bonferroni correction testing for a decrease in ciliary volume in mutant larvae. p-values <0.05 are shown. N = 9–10 (WT), 17–19 (*c-ops-1*^∆8/∆8) larvae. (E) Morphology of cPRC cilia in 3-day-old WT (top row) and *c-ops-1*^∆8/∆8 (bottom row) larvae reconstructed by serial-section electron microscopy (ssEM). Reconstructions of cPRC cilia are shown in the left-most panels. A representative micrograph of the ssEM data used for the reconstructions is shown in the adjacent panels. Dashed squares in these images mark the regions shown in the enlarged views to the right. Orange arrowheads point to cilia with more than one microtubule dublets. In the right-most panels, the branches of single cPRC cilia are coloured by its position relative to the basal body (bb): basal, internal, or terminal branches. The remaining cPRC cilia are coloured in grey. cb: cell body; n: nucleus. (F) Length of cPRC cilia of WT (N = 29 cilia) and *c-ops-1*^∆8/∆8 (N = 32 cilia) larvae measured from ssEM volume reconstructions. Unpaired Wilcoxon test for larger branches in the WT larva: p = 0.076. (G) Length distribution of basal, internal and terminal branches for each genotype shown as a percentage of the total ciliary arbor length. One-tailed unpaired Wilcoxon test with Bonferroni correction for larger basal, internal, and terminal branches in the *c-ops-1*^∆8/∆8 mutant: p = 0.0592, p = 1.26E-4, p = 6.29E-5, respectively. Figure 3—source data 1 (A), Figure 1—source data 4 (B), Figure 3—source data 2 (D).

Figure 3—source data 1 Swimming metrics of individual batches of 3-day-old wildtype or c-opsin-1 mutant larvae subjected to randomized step increases in pressure.: https://cdn.elifesciences.org/articles/94306/elife-94306-fig3-data1-v1.csv
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Figure 3—source data 2 Volume of ciliated structure of individual cPRCs of wildtype or c-opsin-1 mutant 2-day-old larvae measured by quantifying the NIT-GC2 antibody signal.: https://cdn.elifesciences.org/articles/94306/elife-94306-fig3-data2-v1.csv
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c-opsin-1 mutants have defects in cPRC cilia

The reduced response of c-ops-1^∆8/∆8 larvae to pressure may stem from morphological defects of the cPRC sensory cilia in these mutants. We used stainings with acetylated tubulin and NIT-GC2, an antibody specifically marking cPRC cilia (see Figure 2D) to measure the volume of the ciliary compartment in WT and c-ops-1^∆8/∆8 larvae (Figure 3C; Figure 3—figure supplement 1G). Volumetric imaging in 2-day-old larvae stained with these antibodies revealed that the cPRC ciliary compartment in c-ops-1^∆8/∆8 larvae was significantly smaller than in age-matched WT larvae (p = 0.044 and 0.018 for left and right ciliary compartments, one-tailed unpaired t-test with Bonferroni correction; Figure 3D). In some cases, cPRC ciliary compartments were drastically reduced, albeit never completely absent in all four cells (Figure 3—figure supplement 1F). A reduced ciliary compartment may underlie the weaker responses of c-ops-1^∆8/∆8 larvae to pressure.

To further investigate the morphological defects of cPRC sensory cilia in c-ops-1^∆8/∆8 larvae, we reconstructed the cPRC ciliary structure of a mutant larva using volume electron microscopy. We compared this reconstruction to a volume EM dataset of cPRC cilia from a 3-day-old WT larva previously reported (Verasztó et al., 2018). The two WT cPRCs reconstructed have 14 to 15 ramified cilia tightly wrapped on themselves (Figure 3E, top row, Video 6). Branching occurs close to the basal body, soon after cilia protrude from the cell body. Most branches inherit an individual microtubule doublet. The 15 to 17 cilia of the two cPRCs reconstructed in a c-ops-1^∆8/∆8 larva revealed a more sparsely packed structure (Figure 3E, bottom row; Figure 3—figure supplement 1A, Video 7). Mutant cilia are not significantly shorter than WT cPRC cilia (p = 0.076, Wilcoxon test for longer WT cilia, Figure 3F). However, we noticed when comparing cross-sections of each genotype that individual branches of mutant cPRC cilia often contained more than one microtubule doublet (arrowheads in Figure 3E; Figure 3—figure supplement 2A). This suggests that cPRC cilia of c-ops-1^∆8/∆8 larvae have alterations in branching morphology. We indeed found that the terminal branches of cilia in the mutant are significantly shorter (p = 6.29E−05 Wilcoxon test for shorter branches in the mutant, Figure 3G, right-most plot), while internal branches are longer than those in the WT (p = 1.26E−04; Wilcoxon test for longer branches in the mutant, Figure 3G, middle plot). Basal branches also tended to be larger, but no statistically significance can be concluded (p = 0.06, Wilcoxon test for longer branches in the mutant, Figure 3G, left-most plot). This result supports the former observation that branching occurs more distally to the basal body in c-ops-1^∆8/∆8 larvae. Longer internal branches in mutant cPRC cilia would also explain the presence of ciliary profiles with more than one microtubule doublet (Figure 3—figure supplement 2A).

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Overall, our physiological and genetic analyses suggest that the brain cPRCs act as graded and fast-adapting pressure receptors. In c-ops-1^∆8/∆8 larvae, the ciliary compartment is smaller and shows morphological defects, revealing a genetic requirement for c-opsin-1 in the establishment of the sensory compartment and supporting the idea that the cPRC ciliary compartment is the site of pressure transduction.

Synaptic transmission from serotonergic ciliomotor neurons mediates pressure-induced increases in ciliary beating

The complete synaptic wiring diagram of the cPRCs was previously reported from an electron microscopy volume of a 3-day-old larva (Figure 4A, B; Verasztó et al., 2018). The shortest neuronal path from cPRCs to the prototroch involves a feed-forward loop from the cPRCs to two types of postsynaptic interneurons: the INNOS and the INRGWa cells, expressing respectively Nitric Oxide Synthase (NOS) or the neuropeptide RGWamide (Figure 4B). INRGWa cells in turn synapse on a pair of head serotonergic ciliomotor neurons (Ser-h1). Ser-h1 cells directly innervate the prototroch ciliary band and synapse on the MC head cholinergic ciliomotor neuron (Figure 4B). Ser-h1 cells are thought to promote ciliary beating by directly releasing serotonin onto the ciliary band cells and indirectly by inhibiting the cholinergic MC neuron that is required to arrest ciliary beating (Verasztó et al., 2017).

Figure 4 with 1 supplement see all

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TeTxLC expression in Ser-h1 neurons inhibits ciliary beat frequency (CBF) increase during pressure.

Volume EM reconstruction (A) of the cells in the ciliary photoreceptor cell (cPRC) synaptic circuit (B). cPRCs synpase in the middle of the brain on both INNOS and INRGWa neurons. INRGWa cells synapse on Ser-h1 neurons, which directly innervate the ciliary band and also target the MC cell. Only the cells in the shortest path to the prototroch are included. (C) Schematic of the gene constructs used to test the role of Ser-h1 in the pressure response. The TPHp::tdT construct drives expression of the reporter protein Palmi-tdTomato in Ser-h1 and other serotonergic neurons. It was used as a control. The TPHp::tdT-P2A-TeTxLC construct expresses Palmi-tdTomato and the synaptic blocker TeTxLC as a fusion that gets post-translationally self-cleaved by the P2A peptide. (D) Maximum intensity projections of a larva injected with the TPHp::tdT-P2A-TeTxLC construct and stained with α-HA. Ser-h1 was labelled in this animal. Anterior view. (E) Max. CBF that single larvae reached in the 30 s prior (Before), or during the first 30 s of the indicated increase in pressure (Stimulus). Data points for the same larva are joined by lines. Larvae were injected either with the control plasmid TPHp::tdT (top row, N = 14–16 larvae), or with the TPHp::tdT-P2A-TeTxLC plasmid (bottom row, N = 13–16 larvae). One-tailed paired t-test with Bonferroni correction testing for an increase in CBF; p-values <0.05 are shown. Figure 4—source data 1 (E).

Figure 4—source data 1 Ciliary band dynamics of individual 2-day-old wildtype larvae injected with pLB316 or with pLB253 plasmid constructs and subjected to randomized step increases in pressure.: https://cdn.elifesciences.org/articles/94306/elife-94306-fig4-data1-v1.csv
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To directly test the involvement of the Ser-h1 cells in the pressure response, we used a genetic strategy to inhibit synaptic release from serotonergic neurons. We used transient transgenesis to drive the expression of the synaptic inhibitor tetanus-toxin light chain (TeTxLC) (Sweeney et al., 1995) under the promoter of tryptophan hydroxylase (TPH), a marker of serotonergic neurons (Verasztó et al., 2017). The TPH promoter labels the head Ser-h1 and other serotonergic cells including the Ser-tr1 trunk ciliomotor neurons. Of the cells labelled with this promoter, only Ser-h1 is postsynaptic to cPRCs and provides strong innervation to the prototroch cells and the MC cell (Verasztó et al., 2017).

The construct (Figure 4C) drives the expression of both TeTxLC and a hemagglutinin (HA) tagged palmitoylated tdTomato reporter, separated by P2A, a self-cleaving peptide previously used in Platynereis (Bezares-Calderón et al., 2018). The mosaic expression of this construct resulted in the labelling of different subsets of serotonergic neurons. We selected animals showing labelling in Ser-h1 (most larvae were also labelled in Ser-tr1 and other unidentified serotonergic cells). We confirmed the expression of the transgene in Ser-h1 by immunostaining against the HA-tag of Palmi-tdTomato (Figure 4D). Larvae expressing TeTxLC in Ser-h1 showed an increase in CBF only at the highest pressure used (988 mb p = 0.004; one-tailed paired t-test with Bonferroni correction testing for an increase in CBF; Figure 4E, bottom row). Larvae injected with a control plasmid expressing only Palmi-tdTomato but not TeTxLC in Ser-h1 (Figure 4C) showed a significant increase in CBF at the three highest pressure levels applied (237.5 mb p = 0.023, 556 mb p = 0.009, 988 mb p = 0.004; one-tailed paired t-test with Bonferroni correction testing for an increase in CBF; Figure 4E, top row; Figure 4—figure supplement 1A). The metric max. ∆%CBF was not significantly different between control and TeTxLC-injected larvae (Figure 4—figure supplement 1B). These results indicate that TeTxLC-mediated inhibition of Ser-h-1—albeit incomplete and in most cases limited to one of the two cells—leads to a noticeable dampening of the pressure response at the level of the ciliary band. This suggests that synaptic release from serotonergic neurons is required to increase ciliary beating upon pressure increase. Our data support the model that Ser-h1 neurons, and no other serotonergic cells, are specifically required for this response, because this cell type was labelled in all animals tested and these cells directly innervate the prototroch.

Discussion

This work provides insights into the neuronal mechanisms of pressure sensation and response in a marine planktonic larva. Our findings suggest that increases in pressure, either due to the larva’s own actions (sinking or diving) or to downwelling currents, lead to the activation of the sensory cPRCs (Figure 5). The activation is proportional to the magnitude of pressure change and leads to the activation of the downstream circuit that converges onto the Ser-h1 neurons and ultimately leads to increased ciliary beating. The Ser-h1 cells could secrete serotonin (5-HT) onto the ciliary band, which from pharmacological assays is known to increase CBF (Verasztó et al., 2017). Ser-h1 neurons may simultaneously inhibit the cholinergic ciliomotoneuron MC neuron (Verasztó et al., 2017), thereby preventing ciliary arrests while the larva tries to compensate for the increased pressure. Upon a decrease in pressure, 3-day-old (but not 2-day-old) larvae also show an off-response characterized by downward swimming. We have not analysed in detail the neuronal mechanisms of this response but it may depend on an inverted activation of the cPRC circuit, as happens during UV avoidance (Jokura et al., 2023). Pressure ‘on’ and pressure ‘off’ thus induce behavioural responses with opposite sign such that larvae move to compensate for the pressure change. The response is directional along the pressure gradient (even if larvae do not detect the gradient but temporal changes) and we refer to it as barotaxis.

Figure 5

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The pressure response as a depth-retention mechanism mediated by the ciliary photoreceptor-cell circuit.

(Top) *Platynereis* larvae maintain their position in the water column by controlling the beating dynamics of the ciliary band (t = 0, no net change in depth, ∆ depth = 0). A sufficiently rapid increase in depth (at t = 1) caused by intrinsic or extrinsic factors would lead to an increase in hydrostatic pressure. The change in depth relative to the previous state (∆ depth 1 or ∆ depth 2) will be perceived by the larva, which will try to counteract this change by increasing ciliary beat frequency (CBF) of the prototroch, leading to upward swimming (at t = 2) until the pressure returns to the level previously experienced. A smaller change in pressure (∆ depth 1) will lead to a smaller increase in CBF (CBF 1, light green arrow) than a larger change (CBF 2). (Middle) Changes in pressure are sensed by the ciliary photoreceptor cells (cPRCs). These sensory cells are activated in a graded manner according to the change in pressure. (Bottom) cPRCs signal via a postsynaptic circuit including the INRGWa and INNOS interneurons to the Ser-h1 neurons. Activation of these serotonergic neurons is required to increase ciliary beating directly on the prototroch cells and indirectly by inhibiting the MC neuron (CBF 1 or CBF2) proportional to the change in pressure.

Our proposed depth-retention model complements a previously reported spectral depth-gauge mechanism in the Platynereis larva based on its ability to respond to UV and green light (Verasztó et al., 2018). As previously hypothesized for other planktonic larvae (Sulkin, 1984), pressure sensing in Platynereis may operate through negative feedback to retain a particular depth, while light as a more variable stimulus could drive intensity and wavelength-dependent changes in the vertical position of larvae. In future, it will be interesting to explore how responses tp pressure and to directional (Randel et al., 2014) and non-directional light cues (Verasztó et al., 2018) across wavelengths and intensities interact to guide larval swimming.

Our unexpected finding that ciliary photoreceptors, previously shown to be sensitive to UV and green light (Verasztó et al., 2018), are also activated by pressure increase suggests that the integration of light and pressure could begin at the sensory level in a single-cell type. UV light activates the cPRCs to induce downward swimming while pressure increases induce upward swimming. The calcium dynamics of cPRCs depends on the stimulus applied and may therefore underlie the mechanism by which the cells decode and transmit a sensory signal to the downstream circuit. UV induces a transient increase and subsequent drop in Ca²⁺ levels followed by a sharp NO-dependent increase that persists even after UV stimulation ends (Jokura et al., 2023). In contrast, pressure induces a transient increase in Ca²⁺ levels that terminates with the stimulus. The different activation dynamics may lead to the release of different neurotransmitter and neuromodulator cocktails cPRCs are cholinergic, GABAergic, adrenergic, and peptidergic (Jokura et al., 2023; Randel et al., 2014; Tessmar-Raible et al., 2007; Williams et al., 2017) and different activation patterns in the postsynaptic cells. Similar integration can occur in Drosophila larvae, where UV/blue light and noxious mechanical stimuli are detected by the same sensory neuron that together with other cells converges onto a circuit processing multisensory stimuli (Imambocus et al., 2022). In Platynereis, other sensory cells (e.g. the SN^d1_unp cells; Figure 2—figure supplement 1) may contribute to distinguishing the nature of the stimulus. How UV and pressure signals are integrated by the cPRC and how other light responses such as phototaxis interact with pressure responses remain exciting avenues for future research.

The cellular and molecular mechanisms by which cPRCs sense and transduce changes in hydrostatic pressure deserve further enquiry. The mechanism may involve the differential compression of microtubules along each ciliary branch (Li et al., 2022; Nasrin et al., 2021), or differential displacement of fluid inside the cilium (Bell, 2008). The cPRCs have a unique multiciliated structure and are embedded in a protected environment different from fluid-exposed cilia such as hydrodynamic mechanosensory cilia (Bezares-Calderón et al., 2018). These features are hard to reconcile with current models of ciliary mechanosensation (e.g. see review by R Ferreira et al., 2019). Instead, the mechanism of pressure sensation at work in cPRCs may share more similarities with non-neuronal cells detecting pressure, such as chondrocytes (Pattappa et al., 2019) and trabecular meshwork cells (Luo et al., 2014).

The molecular mechanisms of pressure detection remain unclear. Components of the phototransduction cascade may be involved in pressure sensation. Our results indicate that the ciliary opsin required for detecting UV light is not essential for pressure sensation. This molecule rather indirectly affects the ability of cPRCs to sense pressure by contributing to the development or maintenance of a ramified ciliary sensory structure. The structural role of opsins for shaping ciliary cell morphology has also been reported in other photoreceptor and mechanoreceptor cells (Lem et al., 1999; Zanini et al., 2018). The direct transducer of pressure may be a Transient Receptor Potential (TRP) channel. TRP channels can signal downstream of opsins in phototransduction cascades and have been postulated as the ultimate integrators of sensory stimuli (Liu and Montell, 2015). Mechanosensitivity has also recently been reported for vertebrate rods (Bocchero et al., 2020), including the activation of these ciliary photoreceptors by pressure (Pang et al., 2021). The cellular and molecular mechanisms behind this sensitivity are still unclear, but TRP or Piezo channels were suggested to be involved (Bocchero et al., 2020).

The mechanism by which cPRCs detect pressure in Platynereis may also characterize the few other cases of pressure sensors based on ciliated sensory cells (the crab statocyst, and the dog hair cells) (Fraser and Macdonald, 1994; Fraser and Shelmerdine, 2002). Multiciliated pressure receptors may be widespread in zooplankton, as cells with a morphology similar to the cPRCs have been observed in species across the animal phylogeny (Hernandez-Nicaise, 1984.; Baatrup, 1982; Eakin and Kuda, 1970), reviewed in Bezares-Calderón et al., 2020. Some of these cells have long been hypothesized to function as pressure sensors. To our knowledge, this study provides the first functional evidence for the role of multiciliated sensory cells in pressure sensation.

cPRCs are part of an ancient neurosecretory centre with putative roles in circadian regulation of locomotion (Tessmar-Raible et al., 2007; Tosches et al., 2014; Williams et al., 2017). Besides light, pressure can also regularly change as a result of dial vertial migration in planktonic stages or tides in settled benthic stages. One of the original components in the first neurosecretory centres may thus have been multimodal sensory cells keeping track of and integrating changes in periodic cues such as light and pressure that allowed to coordinate physiology and behaviour with the natural rhythms.

Materials and methods

Key resources table

Reagent type (species) or resource	Designation	Source or reference	Identifiers	Additional information
Strain, strain background (Platynereis dumerilii)	Wild-type	Marine Invertebrate Culture Unit, University of Exeter	NCBITaxon:6359
Strain, strain background (P. dumerilii)	c-opsin1Δ8/Δ8 knockout	Verasztó et al., 2018	NCBITaxon:6359	Knockout generated by TALEN-induced gene editing
Transfected construct (mRNA)	GCaMP6s mRNA	Randel et al., 2014		1 mg/ml
Transfected construct (mRNA)	3xHA-Palmi-tdTomato mRNA	This paper		<0.2 ng/µl
Antibody	Monoclonal Anti-Tubulin, Acetylated antibody produced in mouse	Sigma-Aldrich	Cat# T6793, RRID:AB_477585	(1:250)
Antibody	Polyclonal NIT-GC2 antibody, produced in rabbit	Jokura et al., 2023	aNIT-GC2	5 mg/ml
Antibody	anti-5-HT, produced in rabit	ImmunoStar	Cat# 20080, RRID:AB_572263	2 mg/ml
Antibody	Monoclonal Anti-HA antibody produced in mouse	Cell Signaling Technology	HA-Tag (6E2), Cat# 2367, RRID:AB_10691311	(1:250)
Antibody	Monoclonal Anti-HA antibody produced in rabbit	Cell Signaling Technology	HA-Tag (C29F4) Cat #3724, RRID:AB_1549585	(1:250)
Antibody	Goat anti-Rabbit IgG (H+L) Cross-Adsorbed Secondary Antibody, Alexa Fluor 488 produced in rabbit	Thermo Fisher Scientific	Cat# A-11008, RRID:AB_143165	(1:250)
Antibody	F(ab’)2-Goat anti-Mouse IgG (H+L) Cross-Adsorbed Secondary Antibody, Alexa Fluor 546	Thermo Fisher Scientific	Cat# A-11018, RRID:AB_2534085	(1:250)
Commercial assay or kit	mMESSAGEmMACHINE T7ULTRA Transcription Kit	Ambion, Thermo Fisher Scientific	Cat# AM1345
Recombinant DNA reagent	pUC57-TPHp3xHA-Palmi-tdTomato-P2A-TeTxLC (plasmid)	This paper	pLB316	Injected at 250 ng/µl in water
Recombinant DNA reagent	pUC57-TPHp3xHA-Palmi-tdTomato (plasmid)	Verasztó et al., 2017	pLB253	Injected at 250 ng/µl in water
Recombinant DNA reagent	pUC57-T7-RPP2-3xHA-Palmi-tdTomato (plasmid)	This paper	pLB260	Injected at 250 ng/µl in water
Recombinant DNA reagent	pUC57-T7-RPP2-GCaMP6s (plasmid)	Randel et al., 2014	pLB112	Injected at 250 ng/µl in water
Recombinant DNA reagent	pGEMTEZ-TeTxLC	Yu et al., 2004	Addgene plasmid # 32640, RRID:Addgene_32640	Richard Axel & Joseph Gogos & C. Ron Yu
Chemical compound, drug	Ficoll PM70	Sigma-Aldrich	Cat# F2878	−20%
Chemical compound, drug	Formaldehyde Aqueous Solution (Paraformaldehyde Aqueous Solution) EM Grade	Electron Microscopy Sciences	Cat# 15710	−4%
Chemical compound, drug	Low-melting agarose	Hampton Research	LM AgaroseTM, Cat# HR8-092	(2.5–3%)
Software, algorithm	Fiji	NIH	RRID:SCR_002285
Software, algorithm	CATMAID	Saalfeld et al., 2009	RRID:SCR_006278
Other	Wormglu	GluStitch Inc

Metric	Formula
Average vertical displacement	$\frac{\sum_{j = 1}^{T r a c k s} \sum_{i = 1}^{F r a m e s} (Y_{j i + 1} - Y_{j i})}{T r a c k s \times # F r a m e s} \times F r a m e r a t e \times m m p x^{- 1}$
Vertical movement	$\frac{\sum_{j = 1}^{T r a c k s} \frac{(Y_{j f i n . p o s} - Y_{j i n i t . p o s})}{F r a m e s_{j}}}{T r a c k s} \times F r a m e r a t e \times m m p x^{- 1}$
Average speed	$\frac{\sum_{j = 1}^{T r a c k s} \sum_{i = 1}^{F r a m e s} \sqrt{{(X_{j i + 1} - X_{j i})}^{2} + {(Y_{j i + 1} - Y_{j i})}^{2}}}{T r a c k s \times F r a m e s} \times F r a m e r a t e \times m m p x^{- 1}$
Average straightness vertical path	$\frac{\sum_{j = 1}^{T r a c k s} \sum_{i = 1}^{F r a m e s} \frac{(Y_{j i + 1} - Y_{j i})}{\sqrt{{(X_{j i + 1} - X_{j i})}^{2} + {(Y_{j i + 1} - Y_{j i})}^{2}}}}{T r a c k s \times F r a m e s}$
Straightness index	$\sum_{j = 1}^{T r a c k s} \frac{\sqrt{{(X_{j f i n . p o s} - X_{j i n i t . p o s})}^{2} + {(Y_{j f i n . p o s} - Y_{j i n i t . p o s})}^{2}}}{\sum_{i = 1}^{F r a m e s} \sqrt{{(X_{j i + 1} - X_{j i})}^{2} + {(Y_{j i + 1} - Y_{j i})}^{2}} / T r a c k s}$

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Pressure response in Platynereis larvae.

Figure 1—source data 1

Figure 1—source data 2

Figure 1—source data 3

Figure 1—source data 4

Figure 1—source data 5

Figure 1—source data 6

Figure 1—source data 7

Response of 2-day-old Platynereis larvae to a pressure increase of 1 bar.

Zoomed-in view of 2-day-old larvae as they respond to an increase in pressure of 85 mb (left panel) or 500 mb.

Zoomed-in view of 3-day-old larvae as they respond to an increase in pressure of 32 mb (left panel) or 778 mb.

Representative recording of a 2-day-old larva tethered to a glass cuvette and imaged from the anterior side to quantify the effect of pressure on the ciliary dynamics.

Ca2+ imaging of Platynereis larvae during pressure increases.

Figure 2—source data 1

Time-lapse confocal microscopy recording showing the change in GCaMP6s fluorescence (fire colour scale) in a ciliary photoreceptor cell (cPRC) in response to pressure increase (750 mb).

c-ops-1∆8/∆8 larvae show a weaker response to pressure and have structural defects in ciliary photoreceptor cell (cPRC) cilia.

Figure 3—source data 1

Figure 3—source data 2

360° view of electron microscopy reconstruction of sensory cilia from one pair of ciliary photoreceptor cells in a wild-type 3-day-old larva.

360° view of electron microscopy reconstruction of sensory cilia from one pair of ciliary photoreceptor cells in a c-ops-1∆8/∆8 3-day-old larva.

TeTxLC expression in Ser-h1 neurons inhibits ciliary beat frequency (CBF) increase during pressure.

Figure 4—source data 1

The pressure response as a depth-retention mechanism mediated by the ciliary photoreceptor-cell circuit.

Swimming metrics.

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Luis Alberto Bezares Calderón

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Réza Shahidi

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Gáspár Jékely

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Ca²⁺ imaging of Platynereis larvae during pressure increases.

c-ops-1^∆8/∆8 larvae show a weaker response to pressure and have structural defects in ciliary photoreceptor cell (cPRC) cilia.

360° view of electron microscopy reconstruction of sensory cilia from one pair of ciliary photoreceptor cells in a c-ops-1^∆8/∆8 3-day-old larva.