# Pressurized plankton observatory offers a new window into deep-sea larval behavior

Rodrigo Zúñiga Mouret, Stéphane Hourdez, Molly Curran, Michelle H Dibenedetto, Susan W Mills, Costantino Vetriani, Shawn M Arellano, Johanna N J Weston, Lauren N Dykman, Ayinde C Best, et al.

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## To cite this version:

Rodrigo Zúñiga Mouret, Stéphane Hourdez, Molly Curran, Michelle H Dibenedetto, Susan W Mills, et al.. Pressurized plankton observatory offers a new window into deep-sea larval behavior. Limnology and Oceanography: Methods, 2025, 10.1002/lom3.10708. hal-05184788

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Distributed under a Creative Commons Attribution 4.0 International License
LIMNOLOGY
and
OCEANOGRAPHY: METHODS

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Limnol. Oceanogr.: Methods 2025
© 2025 The Author(s). Limnology and Oceanography: Methods published by Wiley Periodicals LLC on behalf of Association for the Sciences of Limnology and Oceanography.
doi: 10.1002/loem.3.10708

NEW METHODS

# Pressurized plankton observatory offers a new window into deep-sea larval behavior

Rodrigo Zúñiga Mouret<sup>1,2*</sup> Stéphane Hourdez,<sup>3</sup> Molly Curran,<sup>4</sup> Michelle H. DiBenedetto,<sup>5</sup>

Susan W. Mills,<sup>1</sup> Costantino Vetriani,<sup>6</sup> Shawn M. Arellano,<sup>7</sup> Johanna N. J. Weston,<sup>1</sup> Lauren N. Dykman,<sup>8</sup>

Ayinde C. Best,<sup>1</sup> Anthony Pires,<sup>1</sup> Lauren S. Mullineaux<sup>1</sup>

<sup>1</sup>Biology Department, Woods Hole Oceanographic Institution, Woods Hole, Massachusetts, USA; <sup>2</sup>Massachusetts Institute of Technology, Massachusetts, Cambridge, USA; <sup>3</sup>Observatoire Océanologique de Banyuls, Sorbonne Université-CNRS, UMR 8222, Banyuls-sur-Mer, France; <sup>4</sup>Applied Ocean Physics and Engineering, Woods Hole Oceanographic Institution, Woods Hole, Massachusetts, USA; <sup>5</sup>Civil & Environmental Engineering, University of Washington, Seattle, Washington, USA;

<sup>6</sup>Institute of Marine and Coastal Sciences, Rutgers University, New Brunswick, New Jersey, USA; <sup>7</sup>Biology Department, Western Washington University, Bellingham, Washington, USA; <sup>8</sup>University of Victoria, Victoria, BC, Canada

Abstract

The High-Pressure Plankton Observatory (HiPPO) is designed to quantify motions of zooplankton for behav-
ioral study, including swimming and metabolic responses to environmental perturbations. It builds on prior
chamber designs while filling gaps in capability for resolving orientation of small (< 1 mm) plankton, tracking their
movements over ecologically relevant spatial scales, and recording in flow-through conditions on a vessel at sea. The
HiPPO chamber has a direct light path for silhouette imaging of zooplankton as they move vertically and horizon-
tally across a 3.56 cm diameter viewing area. Seawater forced by a high-performance liquid chromatography pump is
exchanged continuously through the chamber, but flushing of zooplankton is prevented by fine mesh at the ports.
A high-resolution camera/computer setup enables sustained imaging of plankton motions for quantitative analysis.
Application of HiPPO to an investigation of larval behavior of deep-sea hydrothermal vent species revealed swim-
ming behaviors similar to those of shallow-water species, including upward and downward helices, meandering, and
short hovers. In conditions with microbial biofilm (a potential settlement cue) on a 2024 expedition, vent larvae
unexpectedly swam rapidly upward in tight helices at velocities (0.15 cm s<sup>-1</sup>) higher than those observed in prior
experiments with no biofilm (0.03 cm s<sup>-1</sup>). Many factors varied between the 2024 and earlier trials, so the difference
cannot be attributed with certainty to a cue response. This study describes key new features of HiPPO and demon-
strates the system's ability to document novel zooplankton behavior.

Zooplankton are a fundamental component of the ecology
of our oceans. They are a massive and dynamic carbon source
that can be transferred up the food web or sequestered from
the atmosphere and surface waters into the deep ocean
(Steinberg and Landry 2017). Plankton respond behaviorally
to their environment in ways that can significantly impact
their role in the ecosystem. Despite the importance of behav-
ior in broader ecological contexts, the specific drivers and
modulators of many behaviors remain either unidentified or
poorly understood. For example, holoplankton swim up and
down the water column in diel vertical migration
(Fuchs 1882). Still, it is unclear how light, food, temperature,

<footnote>*Correspondence: rodrigo.zuniga@whoi.edu</footnote>

<footnote>This is an open access article under the terms of the [Creative Commons Attribution License](https://creativecommons.org/licenses/by/4.0/), which permits use, distribution and reproduction in any medium, provided the original work is properly cited.</footnote>

<footnote>**Associate editor:** Kelly Dorgan</footnote>

<footnote>**Data Availability Statement:** Data for cruises and AT50-06 (cruise identifier https://doi.org/10.7284/909880) and AT50-20 (cruise identifier https://doi.org/10.7284/910427) have been submitted to our project (https://www.bco-dmo.org/project/851182) in the Biological and Chemical Oceanography Data Management Office (BCO-DMO).</footnote>
and tides interact to define the scale and scope of this behav-
ior (Bandara et al. 2021). Meroplankton, zooplankton that
spend the early part of their life in the water column and then
settle and remain on the seafloor as adults, are strongly
influenced by ocean currents to disperse and contribute to the
survival of their populations (Cowen and Sponaugle 2009;
Levin 2006; Metaxas and Saunders 2009). The larvae of many
species are inferred to alter their swimming behavior after
detecting chemical settlement cues, as confirmed by direct obser-
vation (Hadfield and Koehl 2004; Meyer-Kaiser et al. 2019). The
identity of these cues, how they alter behavior, or how the
behavior change facilitates settlement remains elusive for many
species of meroplankton, especially those inhabiting the deep
sea. Settlement cues are particularly important for systems that
are island-like and frequently disturbed, such as hydrothermal
vents (Adams et al. 2012).

A number of methods have proven effective for observing distributions and movement of plankton in the field. In-situ sampling of the water column, either via multinet systems (Wiebe et al. 1976) or more advanced optical or acoustic methods (Rahkola-Sorsa et al. 2014), allows for observing large-scale spatial distribution of plankton. In many cases, analysis of the environmental conditions associated with changes in distributions can be used to infer behavioral responses to cues. However, to uncover the mechanisms that drive population-wide distributions, direct observation of the swimming responses of individuals is needed. For some species, it is possible to follow larvae in the field with SCUBA equipment to observe orientation and settlement behavior (Leis et al. 1996). Still, that approach is limited in the duration of observation, time of day, and depth (Leis et al. 2015). Environmental conditions typically are not controlled in field studies but can be manipulated in mesocosm (Bollens and Frost 1991; Duarte et al. 1997; Matthews et al. 2006) and laboratory studies.

Laboratory behavioral experiments have been conducted
on many different species of marine plankton to explore the
environmental conditions that drive ecologically relevant
behaviors. Some studies have used tanks (0.8–10 m tall) to
study the effect of temperature gradients, water density
(Kamykowski and Zentara 1977; Kamykowski 1981), presence
or absence of predators (Pangle and Peacor 2006), age, and
availability of food (Huntley and Brooks 1982) on diel vertical
migrations. Behavioral studies have been conducted to iden-
tify settlement cues and characterize their effect on larvae of
shallow water species, including sponges, corals, crabs, and
gastropods (Hadfield and Paul 2001). However, all these stud-
ies were done at or near surface pressure and thus might not
be adequate for studying large vertical migrations.

Maintaining high-pressure conditions is crucial when con-
ducting behavioral experiments on deep-sea animals, as
changes in hydrostatic pressure can substantially affect the
biology of planktonic organisms (Yancey 2020). Cell and tis-
sue membranes can become more or less permeable (Pozza
et al. 2022), and proteins can denature (Balny et al. 1997)

depending on the extent of change in experienced pressure.
Decompression can induce paralysis or convulsions that may
confound behavioral or movement results (Macdonald 1997).
Quick and prolonged changes in pressure may be fatal, and
even minor changes can impact swimming behavior in
various taxa (Macdonald 1997; Epifanio and Cohen 2016).
Neuromuscular junctions have decreased signal strength in
shallow-water species when exposed to high pressures, whereas
deep-water species have evolved adaptations in these fibers to
deal with the effects of high pressure (Campenot 1975). Because
of this pressure sensitivity, behavioral observations and experi-
ments are ideally conducted at the pressure of the organism's
typical habitat, recognizing that pressure itself can be a cue
(Epifanio and Cohen 2016).

Our design was adapted from the pressure-vessel system used
in Hourdez (2018): an anodized aluminum, custom-built pressure
vessel with a single Plexiglas window (informed by the design in
Lee 2003) that allows continuous visual inspection of the animals
inside. The water flow was provided by a high-performance liquid
chromatography (HPLC) pump regulated by a pressure relief
valve. It was the closest design to our needs, with a chamber large
enough for metazoans to swim, a flow-through capability for fluid
manipulation and oxygen exchange, a viewport, and pressure tol-
erances of 500 bars or more. The Hourdez system was preceded
by a series of pressure-vessel design innovations, starting in the
Childress laboratory with pressurized flow-through capability
(Kochevar et al. 1992), followed by reinforcement for deep-sea
pressures and customization for sophisticated respirometry
(Girguis et al. 2000). For microbial investigation, Koyama et al.
(2001) created a miniaturized version with a light path for visuali-
zation under a microscope, but the chamber is too shallow to
allow movement of metazoans. A much larger, static chamber
with a light path was developed to assess the buoyancy of meta-
zoan embryos at deep-sea pressure (Marsh et al. 2001), but the
embryos are detectable only when they contact the viewports.
Pradillon et al. (2004) engineered a system that split the flow
lines, allowing for simultaneous incubations in two separate
chambers. It was adaptable to accommodate small chambers for
visualizing embryonic development but not metazoan move-
ment. Additional systems have been developed for specific adap-
tations, including metazoan thermal preferences (Girguis and
Lee 2006) and metazoan limb motion in a chamber < 1 mm thick
(Bao et al. 2010), but none with the capability of visualizing meta-
zoan vertical swimming behavior. Technological gaps remain for
visualizing and analyzing swimming responses of small plankton
at pressure, including manipulating their environmental condi-
tions and conducting observations while at sea.

Materials and procedures

System design

The objectives for this study are to develop a method to
conduct high-pressure quantitative behavioral analysis on tiny
plankton and manipulate their environmental conditions at
sea. The challenge was designing a system that remains struc-
turally sound under extreme deep-sea hydrostatic pressures
while allowing for direct observation of plankton morphology
and behavior. Plankton must be continuously supplied with
seawater during the experiments to prevent the development
of an anoxic environment in the chamber. At the same time,
the design must ensure that plankton remain visible to the
camera, which will be used for observation. Once in the cham-
ber, plankton need to be distinguished from other particles in
the video footage. Lighting and camera optics must be capable
of resolving fine details, such as swimming appendages, to
determine the plankton's orientation accurately while also
recording them over a large enough field of view to capture
ecologically relevant swimming behaviors. Additionally, for
experiments conducted on a rocking ship at sea, the camera
system must remain fixed and centered on the chamber to
track the movements of these tiny organisms reliably. Lastly,
to manipulate environmental cues, the design must include a
method for introducing those cues into the pressurized
chamber.

In response to these needs, we present the High-Pressure
Plankton Observation (HiPPO) system: a design capable of
capturing high-resolution, reliable behavioral data of deep-sea
plankton in dynamic biochemical environments. The HiPPO
pressure chamber is constructed with acrylic viewports on
either side of an anodized aluminum housing (Fig. 1), with an
O-ring seal on the inner side of each viewport that confers a
pressure rating to 401 bar or simulates 4010 m depth. For low-
pressure seals, O-rings would typically be secured by a groove
integrated into the acrylic viewport plate. However, in high-
pressure applications, the forces acting on the outer groove lip
are too high to maintain structural integrity. To address this,
our system eliminates the outer lip of the groove. Instead, the
O-ring is positioned around an inner acrylic flange and rests
against the aluminum housing for a secure seal. The camera-
side viewport is designed with a small dome protruding into
the observation pool. When this viewport seals the pool after
the plankton sample is introduced in an upright orientation,
it forces any air out from the pool that would otherwise form
bubbles (the viewport does not act as a meniscus lens because
it is curved only on one face). The two viewports create a
direct light path through the chamber from the light source to
the camera. Backlighting with an infrared (850 nm) LED flood-
light coupled with a white acrylic diffusing plate results in a
silhouette of any particle or plankton in the chamber. As for
previous chambers used for animal physiology, inlet and out-
let ports allow the flow of seawater forced by an HPLC pump.
The seawater is filtered to 0.2 µm to remove most marine
microbes and avoid clogging the smallest tubing in the flow
path (0.25 mm). To adapt the chamber so that plankton in
the size range of small (0.1–10 mm) meroplankton (e.g., larvae
of benthic invertebrates) and holoplankton (e.g., copepods)
are retained, a small piece of 63 µm Nitex mesh is fixed with
hot glue over each of the inlet and outlet ports. The mesh is

applied with minimal glue and no protruding edges to prevent
entanglement of plankton.

A custom 3D printed mount secures the pressure chamber and can be attached to an aluminum extruded rail (80/20) (Fig. 2). This keeps all components on the rig fixed relative to one another for stable recording while keeping the rig secure in rocky shipboard conditions. The aluminum-extrusion construction consists of a single optic rail and two tri-rails attached orthogonally to either side of the rail to offer additional anchoring to the mount and added stability for the rig. The single rail is centered on the mid-axis of the chamber and can support a camera via an optic mount (Thorlabs). This modular construction is easily assembled and disassembled for shipping and can accommodate different camera/lens configurations.

Solutions to introducing substratum-bound and waterborne
environmental cues involved two separate approaches. A
cradle-like insert designed to fit within the plankton observa-
tion chamber secures substratum-bound cues to the chamber's
bottom (Fig. 2). The insert is made from anodized aluminum to
prevent electrochemical interactions between metals in seawater.
Capability for introducing waterborne cues is provided by plac-
ing soluble cues in a secondary pressure chamber installed
upstream of the plankton containment chamber (Fig. 2). A
bypass line with a toggle valve allows the flow to be redirected
through the cue chamber into the primary plankton chamber,
enabling an exposure-response experiment as used commonly in
experimental pharmacology (Byiers et al. 2012). Though this fea-
ture was not implemented in our behavior experiments, in
January 2025 we demonstrated its operational capabilities with a
flow visualization study (Supplementary Fig. 1).

A high-resolution video camera with a macro lens, con-
nected to a computer with a fast processor, a solid state drive,
USB ports with high bandwidth, and sufficient RAM is all
essential for sustained imaging of motions and morphology
(including swimming appendages) of identifiable plankton. A
Basler Ace acA2040-90um NIR camera fitted with a Nikon
55 mm lens captures infrared images at up to 90 frames per
second and resolutions of up to 2048x2048 pixels. The high-
power IR light source illuminates with an array of 198 LEDs at
an 850 nm wavelength (CMVision IR130-198 LED). The com-
puter is configured to record the images to a solid state drive,
with 32 GB RAM and a 2nd Gen Intel(R) Core™ processor
(i7-12700H, 2.3 GHz), through a cable with USB-3 connectors.
In this setup, with the camera mounted 28 cm from the front
of the chamber, the image resolution is up to 798 pixels cm⁻¹,
displaying a 0.2 mm larva at the center of the chamber as
16 pixels. Calibrations are typically made by taking an image
of a ruler positioned at the center plane of the chamber and
the center of the camera's focal range. This calibration can
also be done by using the dimensions of visible components
of the chamber itself, described in further detail in the applica-
tion section. Due to the shallowness of the chamber (1.12 cm,
Fig. 1b) larvae remain in focus throughout the whole cham-
ber. Imaging must be compromised for a system with lower
<figure><img src="image_4.png" /><figcaption>**Fig. 1.** HiPPO pressure chamber. Design of HiPPO plankton chamber. (a) Upright orientation for sample introduction, showing the anodized aluminum end-caps that secure the acrylic viewport disks and the housing ring. The 3D Cartesian coordinate system shows positive z and x directions (solid lines) corresponding respectively to upward vertical and rightward horizontal motion (b) Cross-sectional detail of the camera-side acrylic viewport disk with dome and o-ring seal (upper) and the housing ring with observation pool (in blue) with inlet/outlet ports (lower). (c) Observation orientation showing the light path through the observation pool and the 63 µm mesh screens over the inlet, outlet, and pressure-release ports. Coordinate system as in (a).</figcaption></figure>

computing power to avoid dropped frames. This can be done
by reducing the size of image files (camera resolution), the
speed at which they are captured (fps), or reducing continuous
recording time and implementing breaks to allow the buffer
(RAM) to clear.

Assessment

Site background and research expeditions

We applied the HiPPO system to study larval behavior at
deep-sea hydrothermal vents. We chose this application
because larval connectivity and settlement behavior in this
transient ecosystem is a long-standing knowledge gap (Adams

et al. 2012). Regular eruptions in relatively short timescales
make vents on fast-spreading ridges ideal for studying succes-
sion and connectivity in hydrothermal vent communities
(Adams et al. 2012). Moreover, increasing interest in deep sea
mining makes studying the ecology of hydrothermal vents
paramount for making informed decisions on the extent and
methods for resource extraction (Miller et al. 2018; Van
Dover 2014). Larval transport keeps hydrothermal vent com-
munities connected and resilient to local extinction events,
such as eruptions. However, how larvae journey anywhere
between 10's and over 1000's of meters and find active vent
sites to settle on remains unknown (Adams et al. 2012). Bio-
films integrate multiple fluctuating environmental factors over
<figure><img src="image_5.png" /><figcaption>**Fig. 2.** HiPPO system diagram. The flow path of filtered seawater is from the HPLC pump, through the plankton chamber, and out the release tube. Flow can be diverted through the soluble cue chamber with a toggle valve (not used in behavior experiments but tested in 2025). The pressure relief valve regulates system pressure. The rail and mount keep the camera aligned with the plankton chamber, while the infrared floodlight and diffuser provide homogenous backlighting for the camera and lens. Mounting brackets secure the rail system to the underlying bench. Needle valves allow isolation and removal of the plankton chamber while maintaining pressure. The inset shows details of the insert that stabilizes substratum-bound cue in the plankton observation chamber.</figcaption></figure>

time and have already been known to act as chemical cues for
many shallow-water marine invertebrates (Hadfield 2011). Obser-
vations from a field colonization study at hydrothermal vents
suggest that several invertebrate species, including the founda-
tional mussel, prefer to settle on surfaces with established biofilm
(Ladd et al. 2024). We hypothesize that vent-specific biofilms act

as a settlement cue that indicates suitable vent habitats for larvae
and that larvae behave differently in the presence vs. absence of
biofilm. We chose to study gastropods due to their high regional
abundance, their biphasic life cycles, and our group's expertise in
studying the swimming behavior of their shallow water relatives
in response to settlement cues.
The HiPPO design solutions presented above enabled experiments conducted on two expeditions aboard the R/V *Atlantis* to the historically well-studied 9°N East Pacific Rise vent field (Fornari et al. 2012). Experiments were conducted without cue in the first expedition (AT50-06; December 2022), and with cue in the second (AT50-20; January 2024). Samples used in this study came from Riftia Mound, located at 9°50.297′N, 104°17.480′W, at a depth of 2510 m.

## Microbial biofilm and plankton sample collection

In this experiment, plankton were treated as two groups: those exposed to microbial biofilms (2024) and those not (2022). To collect biofilm, “Biofilm Colonizers,” sections of PVC piping covered in stainless steel mesh on either side and attached with metal band clamps, were deployed on the ben-thos by HOV *Alvin* on sites with high diffuse flow (confirmed by temperature probe reading higher than ambient tempera-tures) and left to be colonized by vent biofilm-forming bacteria for 10 d before recovery. For the 2024 (biofilm) obser-vations, segments of colonized mesh were cut to size using flame-sterilized tweezers and scissors, secured in the custom aluminum inserts, and placed in the bottom of the plankton observation chamber. This design was intended to test response to a substratum-bound cue assuming a tactile trigger, but the biofilm was flocculent and became suspended when the chamber was filled, likely bringing it into contact with larvae throughout the observation pool. For the 2022 (no biofilm) observations, the insert was not used.

Larvae were collected using a McLane Laboratories large-volume water pump (WTS-LV) with a filter assembly to capture larvae onto a large (19 cm diameter) filter of 63 µm mesh. The filter system was designed with an expanded head space and thick insulation to maximize retrieval of live specimens (Beaulieu et al. 2009). The pump was mounted on a lander to facilitate manual positioning via HOV *Alvin*. The lander was placed at about 10 m distance from the active vent site, and the inlet was positioned at 1.5 m above the seafloor, where the water temperature was 2°C and pressure was 252 bar. The pump was programmed to run at a rate of 25 L per minute for 21 h to end just before release from the seafloor to maintain larval health, capturing samples of roughly 31,000 L. The lander ascended to the surface at a rate of approximately 45 m per minute. Due to the warm surface waters of the eastern tropical Pacific, care had to be taken to recover the sample quickly, as prolonged exposure to elevated temperature can damage the larvae. At this site, our CTD data showed that temperature was below 5°C at depths greater than 1000 m, reached 10°C at about 300 m, and was above 25°C at the surface. After the pump was recovered and secured on deck, the pump filter assembly was immediately placed in chilled seawater (4°C). Gastropod larvae were sorted manually from the samples, on ice under dissecting microscopes, for 1–2 h, and identified by trained experts. All available gastropod larvae (150–400 µm) and 6–12 copepods were placed in the plankton

observation chamber. Copepods, being larger and more active swimmers, were added as a visual aid to ensure the camera remained in focus and conditions in the chamber remained habitable. The chamber was topped off with filtered bottom seawater, sealed, and fastened to the mount to prepare for recording behavior. In 2022, one of six recovered pump samples yielded actively swimming gastropod larvae, whereas in 2024, one of seven pump samples yielded swimming larvae. In 2022, three gastropod larvae were introduced into the chamber (*Lepetodrilus* sp., *Laeviphitus* sp., and “unknown”), and all revived sufficiently to swim actively in the chamber. In 2024, after the biofilm insert was placed in the chamber, nine gastropod larvae were introduced (identified with certainty as *Lepetodrilus* sp. and *Peltospira* sp., and provisionally as *Laeviphitus* sp., *Echinopelta* sp., *Clypeosectus* sp. and “unknown”); two of these individuals revived, but it was not possible to distinguish which ones.

## Plankton behavior recording and particle tracking

All video recordings were done in a 4°C cold room. To maintain darkness in the chamber while allowing for live footage, lights in the cold room were off, and the camera was connected to a laptop outside the cold room via a 3.3 m USB cable. HPLC flow rate was initially set to 10 mL min⁻¹ but lowered to 0.05–0.2 mL min⁻¹ when the pressure reached ~193 bar to avoid large pressure oscillations. Bottom pressure (252 bar) was slowly reached over about 15 min. Once up to pressure, videos were recorded as TIFF stacks using a frame rate of 30–60 fps. Occasionally, the chamber was manually flipped (four times in 2022, twice in 2024) to attempt to stimulate larvae into swimming. Experiments continued until larvae stopped swimming, between 2 and 3 h. A dye visualization, conducted in 2025 with no larvae, demonstrated that at a 0.5 mL min⁻¹ pump flow rate, flow speeds in the chamber (2.7 × 10⁻³ cm s⁻¹) are low compared to larval speeds (0.3–2.4 × 10⁻¹ cm s⁻¹) and mixing is negligible (Supplementary Fig. 1; Table 2).

We used standard two-dimensional particle tracking methods in MATLAB to track individual plankton movements (https://github.com/rzunigamou/HiPPO-behavior). After removing a mean background image, there is enough contrast to identify the plankton. Particles, treated as weighted centroids, were then filtered for minimum intensity, minimum size, and maximum speed, and larvae tracks were identified manually. Instantaneous vertical velocity was calculated using a 7-point differencing stencil and a Gaussian filter, as in Ouellette et al. (2006). Each track was initiated when a larva was first observed to move and ended when it changed behavior or reached the lower or upper surface of the chamber. While we would not expect any variation using this kind of particle tracking code, we re-tracked one of the videos from 2024 and reproduced identical swim metrics for all 14 larval tracks. We categorized the tracks into downward (downhelix, downswim, sink) and upward (uphelix, upswim, meander) motions, as described in Table 1. A brief interruption of a sinking track was
<table><caption>**Table 1.** Larval behavior categories. Number of observations for each behavior category by *n* = 3 individuals in 2022 (no-biofilm condition) and by *n* = 2 individuals in 2024 (biofilm condition).</caption><thead><tr><th>Behavior</th><th>Description</th><th>No. obs. 2022</th><th>No. obs. 2024</th></tr></thead><tbody><tr><td>Upswim</td><td>Consistent movement in an upward direction</td><td>7</td><td>17</td></tr><tr><td>Uphelix</td><td>Upward movement following a helical trajectory</td><td>4</td><td>11</td></tr><tr><td>Downswim</td><td>Consistent movement in a downward direction</td><td>2</td><td>3</td></tr><tr><td>Downhelix</td><td>Downward movement following a helical trajectory</td><td>0</td><td>1</td></tr><tr><td>Sink</td><td>Passive downward movement following a linear trajectory</td><td>3</td><td>31</td></tr><tr><td>Catch</td><td>Brief interruption in a sinking trajectory</td><td>0</td><td>5</td></tr><tr><td>Meander</td><td>Erratic, nonlinear movement pattern</td><td>1</td><td>0</td></tr><tr><td>Total</td><td></td><td>17</td><td>68</td></tr></tbody></table>

<table><caption>**Table 2.** Larval behavior metrics. Median and mean values for larval tracks from 2022 (no biofilm) and 2024 (biofilm). Means included for comparison to published observations.</caption><thead><tr><th>Behavior</th><th>Median 2022 (no biofilm)</th><th>Mean 2022 (no biofilm)</th><th>Median 2024 (biofilm)</th><th>Mean 2024 (biofilm)</th></tr></thead><tbody><tr><td>Upswim</td><td></td><td></td><td></td><td></td></tr><tr><td>Speed (cm s<sup>-1</sup>)</td><td>0.06</td><td>0.06</td><td>0.15</td><td>0.15</td></tr><tr><td>Vertical velocity (cm s<sup>-1</sup>)</td><td>0.04</td><td>0.05</td><td>0.14</td><td>0.15</td></tr><tr><td>Uphelix</td><td></td><td></td><td></td><td></td></tr><tr><td>Speed (cm s<sup>-1</sup>)</td><td>0.06</td><td>0.07</td><td>0.18</td><td>0.17</td></tr><tr><td>Vertical velocity (cm s<sup>-1</sup>)</td><td>0.03</td><td>0.03</td><td>0.15</td><td>0.14</td></tr><tr><td>Helix width (cm)</td><td>0.20</td><td>0.22</td><td>0.03</td><td>0.03</td></tr><tr><td>Downswim</td><td></td><td></td><td></td><td></td></tr><tr><td>Speed (cm s<sup>-1</sup>)</td><td>0.03</td><td>0.03</td><td>0.09</td><td>0.12</td></tr><tr><td>Vertical velocity (cm s<sup>-1</sup>)</td><td>-0.03</td><td>-0.03</td><td>-0.09</td><td>-0.11</td></tr><tr><td>Sink</td><td></td><td></td><td></td><td></td></tr><tr><td>Speed (cm s<sup>-1</sup>)</td><td>0.24</td><td>0.21</td><td>0.27</td><td>0.26</td></tr><tr><td>Vertical velocity (cm s<sup>-1</sup>)</td><td>-0.24</td><td>-0.21</td><td>-0.27</td><td>-0.26</td></tr></tbody></table>

a “catch,” and any following motion was analyzed as a separate track. Two methods were used to calibrate the scaling conversion of pixels to cm. In 2022, an image of a ruler was used, whereas in 2024 an image of the inner wall of the observation pool, where both the front face and back face were visible, was used. To calculate the center plane diameter, the diameters at the front and back faces were averaged. Swim speed was calculated by measuring a track’s total length and dividing by its duration. Vertical velocity was calculated as the mean of all the instantaneous vertical velocity values of an entire track. Mean helix width (radius) was calculated for each helical track by assuming a vertical helix orientation, measuring the horizontal distance of each half-turn, and calculating the mean of those distances over the entire track. A two-dimensional image represents all or most of the larva’s motion for sink and catch tracks. For the downswim, upswim, helical and meandering tracks, a two-dimensional image misses the horizontal motions

outside the plane orthogonal to the camera; we do not attempt to model or incorporate those motions into our calculations.

## No-biofilm (2022) vs. biofilm (2024) experiments

Since improvements were made to the system throughout this study, several notable differences between the HiPPO system setups in 2022 and 2024 influenced imaging and data analysis. First, in 2022, limitations in the computer processor restricted the image area to roughly 1/4 of the viewport area. This limited view provided a resolution of 798 pixels cm⁻¹ but required that the camera be reoriented on occasion to image regions in the chamber containing active larvae. In contrast, in 2024, a hardware upgrade allowed for imaging of the full viewport area, though fitting it all in the frame came at the sacrifice of resolution (563 pixels cm⁻¹). Second, the higher framerate (60 fps) and low data transfer speed and buffer capacity in 2022 meant that recordings were limited to
3.5 min intervals followed by ~3 min waiting periods while memory was written to disk. In 2024, an improved computing system allowed for continuous imaging of the full chamber field of view. Third, in 2022, the camera was not rigidly connected to the chamber, so larval tracks had to be smoothed (21-frame window sliding average) to reduce relative motions from the ship's vibration, which introduced high frequency noise into the data. Effective reduction of noise via smoothing was confirmed by qualitative comparisons of pre-smoothed and smoothed larval tracks. In 2024, the rigid HiPPO rail system eliminated the need for smoothing tracks. Finally, in 2022, filtered seawater from surface waters was input for the system flow, whereas filtered water from a Niskin rosette lowered to the seafloor was used as input for the system flow in 2024. Addressing our primary ecological question involves a comparison of larval behaviors between the no-biofilm condition in 2022 and the biofilm condition in 2024, but we readily acknowledge that behavioral differences between the years could be due to any of the differences in experimental set-up.

## Results

### Observed behaviors

Larval behavior tracks were generated from three gastropods in 2022 and two gastropods in 2024. Representative larval tracks in the seven track categories show the variation in larval motions in the 2022 observations with no biofilm (Fig. 3), with a few tracks from 2024 in the presence of biofilm. For the uphelices, the tracks were distinctly wider and slower in the 2022 image than in the 2024 image, and the “catch” was observed only in 2024. Quantitative analysis of track categories was conducted separately between upward and downward tracks to avoid potential bias from viewing area (as described in Methods), making them two different and non-comparable metrics. In the downward tracks, the percentages of sinks and associated catches were notably higher in 2024 than in 2022, and downhelices were observed only in 2022 (Fig. 4a). In the upward tracks, the percentages of upswims and uphelices were similar between 2024 and 2022 conditions, but meanders were observed only in 2022. We expected the number of downward and upward tracks to be equal in all observations, as larvae typically reverse direction when reaching the top or bottom of the chamber, although sometimes after a delay. However, we observed a surplus of upward tracks in the 2022 no-biofilm observations, likely due to the smaller view area available. To maximize the number of recorded tracks, we had to scan the bottom of the chamber for larvae and wait for them to swim upward. This sampling bias was not present for the biofilm condition in 2024 likely because we had the whole chamber in view, and there was no need to scan.

Differences in larval swimming metrics between 2022 and 2024 are presented qualitatively; statistical analyses were not attempted due to the low number of larvae observed. Larval

helix widths were narrower in 2024 than in 2022 (Table 2; Fig. 4b). Larval swim speeds were notably faster in 2024 than in 2022 (Table 2; Fig. 4c). Sinking speeds were similar between 2022 and 2024, as expected for similarly sized larvae. Vertical swimming velocities were greater in 2024 than in 2022, while vertical sinking velocities were similar between the years (Table 2; Fig. 4d).

## Discussion

### HiPPO system design

The HiPPO system features key capabilities enabling quantitative analysis of zooplankton behavior at high pressure in the laboratory and while at sea. Swimming and other behaviors of zooplankton as small as 0.1 mm can be visualized and tracked for sustained periods at sufficiently high resolution to distinguish their appendages and orientation. The system enables control and modification of temperature, pressure, and oxygenation of the fluid environment, as well as the introduction of other ecological influences such as food and substratum-bound cues. These innovations are made possible by a new chamber design, a reconfiguration of the camera/computer system, and structural modifications to arrange and stabilize the components of the flow-through system. The addition of a second chamber also enables the introduction of soluble chemical cues from, for example, microbial biofilms, predators, food resources, competitors, and conspecifics, including potential mates.

The redesign of the plankton observation chamber from an existing physiology chamber introduces three new characteristics critical for visualizing the swimming of plankton-sized organisms: a light path through the chamber, a customized observation pool, and barriers to prevent escape. Some previous chamber designs have a light path, but their observation pools are either too large to keep zooplankton in (Marsh et al. 2001; Girguis and Lee 2006) or too small to enable swimming (Koyama et al. 2001; Pradillon et al. 2004; Bao et al. 2010). The light path is created by adding a second acrylic viewport and replacing the solid aluminum end cap with an aluminum ring. This allows for illumination of the full chamber and backlighting of plankton to reveal silhouettes. The thickness of the observation pool (1.12 cm), set by the height of the aluminum housing ring, is thin enough to keep the swimmers in focus throughout their paths but sufficiently thick to allow unimpeded motion. The diameter of the viewing area (3.56 cm) is large enough to track motion on spatial and temporal scales relevant to zooplankton behavioral ecology. Finally, the attachment of Nitex mesh to the inside of ports in the observation pool prevents the escape of plankton while also solving a safety issue by ensuring that plankton cannot obstruct the tubing downstream in the system and over-pressurize the chamber.

Integrating the HiPPO chamber with a high-resolution camera and upgraded computer components enables zooplankton behavioral investigations that require high-resolution imaging,
<figure><img src="image_6.png" /><figcaption>**Fig. 3.** Visualization and tracking of larval behavior. Panels are arranged by cruise date: 2022 (no-biofilm) in blue and 2024 (biofilm) in yellow; the color scheme corresponds to that of Fig. 4. (a) Representative image of gastropod larva (at arrow) in 2022 view with enlarged inset showing morphology detail, image resolution in pixels, and length calibration in µm. (b–f) Representative behavior tracks in 2022 view with red dots indicating the track's starting point (origin) and duration in seconds (dur) of the track for comparison of larval speed. The coordinate system is positive z for upward vertical and positive x for rightward horizontal, as in Fig. 1. (g) The representative image of the gastropod larva (at arrow) in 2024 view shows the biofilm cradle, with an inset as for panel a. (h–i) Representative behavior tracks in 2024 view with detail as in panels b–f.</figcaption></figure>

tracking of individuals over ecologically relevant distances, and
sustained recording over time. The 2 K camera with a
55 mm lens enables high-resolution visualization of zoo-
plankton appendages and body orientation (up to
798 pixels/cm; providing 16 pixels for a 0.2 mm larva)
while maintaining focus through the full 1.12 cm depth of

field. Upgrading the computer with more RAM, a more pow-
erful processor, and a solid-state drive allows recording of
the zooplankton tracks over hundreds of body lengths and
a period of several hours. Securing and aligning the cham-
ber, camera, and lens to a rail system allows replicable con-
figurations and use on a rocking vessel at sea.
<figure><img src="image_7.png" /><figcaption>**Fig. 4.** Larval behavior in 2022 (no biofilm) and 2024 (biofilm). (a) Percentage of tracks in downward (green) and upward (purple) behavior categories for 2022 and 2024. Numbers in the bars indicate the number of tracks for each behavior. (b) Scatter dot plot of uphelix track widths observed in 2022 (in blue) and 2024 (in yellow), where each dot is the mean helix width of an individual track. Medians are displayed as a horizontal line. (c) Scatter dot plots of larval swim speeds in tracks of upswim, uphelix, downhelix, and sink behavioral categories in 2022 (in blue) and 2024 (in yellow). (d) Scatter dot plot of larval velocities in tracks of behavioral categories in 2022 (in blue) and 2024 (in yellow). Horizontal dashed line delineates positive and negative vertical velocities.</figcaption></figure>

Application to zooplankton behavior

The application of HiPPO to investigate larval behavior of
hydrothermal vent species demonstrates its capability to dis-
cover ecologically important zooplankton behaviors and open
new avenues for inquiry. The behaviors observed are new for
these species but correspond to categories described for shal-
low water species at surface pressures: most of the tracks are
oriented vertically, some with helical motion, and a few with
seemingly haphazard horizontal meanders (e.g., as in Chia
et al. 1984; Maciejewski et al. 2019). The "catch" we observed
at pressure is essentially a brief version of the hover that
has been described for many species including estuarine

gastropods (Garwood et al. 2022). The observed larval swim-
ming speeds, vertical swimming velocities, and sinking veloci-
ties are in mid-range for similarly sized shallow-water
molluscan larvae (as reported in Chan et al. 2013; Chia
et al. 1984), except for the upward swimming velocities in
2024 (in the presence of biofilm), which are at the high end of
the range.

These observations are relevant to fundamental questions
about the environmental processes driving larval settlement
and dispersal. Environmental cues for settlement are expected
to influence the direction and speed of vertical motions to
bring larvae down toward the benthic environment. Helices,
meanders, and other horizontal motions may be exploratory behaviors that allow larvae to detect and select optimal settlement sites (Maciejewski et al. 2019). Environmental cues influencing dispersal typically elicit vertical motions that move larvae into horizontal currents that transport them to new habitats or retain them locally (Epifanio and Cohen 2016).

The discovery of fast upward swimming and narrow helix radius in the presence of a potential (biofilm) settlement cue in the 2024 observations is surprising, as those motions would presumably not bring the larvae down to the seafloor or enable horizontal exploration. The catch behavior that occurred only in biofilm is also of interest as perhaps analogous to a bacterial tumble (Berg 2004) that allows an individual to optimize habitat exploration for colonization. These results are intriguing, but we are careful not to attribute them solely to the presence of biofilm, as many other factors varied between the 2024 (biofilm) and 2022 (no biofilm) trials, including larval individuals of potentially different species, the source of seawater (surface vs. bottom), and details of the imaging and analysis.

More generally, we acknowledge that rapid depressurization as a result of our sample collection methods may have affected larval behavior in all trials. However, physiological studies that used similar methods on adult deep-sea fauna reported no abnormal behavior upon repressurization (e.g., Hourdez 2018; Le Layec and Hourdez 2021). Anecdotally, we have only observed a single larval gastropod swimming in our shallow dish while sorting at surface pressure. Similarly, we have not observed any vertical swimming behavior from gastropod larvae in the chamber until fully pressurized for at least 30 min. This suggests that swimming impairment caused by depressurization can be mitigated by repressurization over time scales of tens of minutes. However, the long-term impacts of these pressure fluctuations are unknown and warrant further study, especially in light of suggestions that deep-sea larvae undergo ontogenetic vertical migrations into surface waters (Bouchet and Warén 1994). Our current observations demonstrate HiPPO's potential for discovery of planktonic behavioral responses in various pressure environments.

## Comments and recommendations

Future questions to explore with HiPPO include investigations of directional responses of holoplankton to environmental cues, physiological responses associated with phenology, and additional responses of larvae to environmental conditions. The chamber is smaller than most used for studies of diel vertical migration but is suitable for detecting vertical swimming responses of zooplankton to light at different pressures (depths) when exposed to cues for prey, predators, or other conditions that influence migration. For larval ecology, HiPPO enables investigation of larval responses to a wide

range of environmental and ecological conditions at depth and exploration of the influence of pressure on those responses. Additionally, the small size of the chamber allows one to place it under a microscope and study the real-time developmental biology of deep-sea creatures at in situ pressure. The light path through the chamber makes this system compatible with multiple optical configurations.

Specifically for vent larvae, the second chamber provides a way to study how individuals respond to soluble settlement cues by allowing cues to be introduced to, and then flushed from, a population of larvae under behavioral observation. The cradle can be refined for substratum-bound cue-choice experiments by separating the cue "bins" in the chamber with a central space for initially positioning larvae to avoid bias in their choice movement. A remaining challenge for deep-sea studies, which cannot be addressed by HiPPO refinement, is to retrieve sufficient numbers of healthy live larvae from the seafloor for replicated experimentation. Future efforts should focus on measures to reduce stress during recovery when larvae are likely to experience the most drastic change to their environment, that is, by decreasing time in warm surface waters during recovery or by increasing insulation of the filter chamber to better protect the sample. That said, larvae in similar experiments run in temperate or polar environments would need less protection from changes in temperature. We look forward to continuing to adapt the HiPPO for upcoming expeditions to solve the question of what cues activate settlement behaviors at hydrothermal vents.

## Acknowledgments

We thank the crew of the *R/V Atlantis* and the DSV *Alvin* group for their expertise and support during cruises AT50-06 and AT50-20. We benefited from technical assistance from Helen Fredericks, DC Collasius, Tim Smith, Mark St. Pierre, and Karl Helfrich on land, and Dan Fornari at sea. Stace Beaulieu advised on data science. Jeanette Wheeler and Kirsten Meyer-Kaiser contributed to MATLAB scripts for processing tracking data. OpenAI ChatGPT 4.0 mini was used for clarification and grammatical editing of text, as well as assisting with R code to generate figures in preparation of this manuscript. Funding was provided by NSF grants OCE1947735 to Lauren S. Mullineaux, OCE1948580 to Shawn M. Arellano, OCE1948623 to Costantino Vetriani, and OCE2152453 to Lauren S. Mullineaux and Stace Beaulieu. Rodrigo Zúñiga Mouret was supported in part by WHOI Discretionary Funds and Johanna N. J. Weston by a WHOI Postdoctoral Scholarship.

## Conflicts of Interest

None declared.
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**Supporting Information**

Additional Supporting Information may be found in the online version of this article.

Submitted 16 January 2025

Revised 26 June 2025

Accepted 28 June 2025