**1. Introduction**

Ectoparasitic sea lice represent the most important parasite problem to date for the salmon farming industry. The salmon louse *Lepeophtheirus salmonis* (Figure 1) is a Caligid copepod that infests both wild and farmed salmonid fish in the northern and southern hemispheres. Salmon lice are a major disease problem in the farming of Atlantic salmon, *Salmo salar*, and it has been suggested that salmon lice also play a role in the decline of wild stocks [1,2].

Severe infestations produce pathological lesions on the host that are caused by attachment and feeding of sea lice in both the adult and juvenile stages. *L. salmonis* induces stress-related responses in the host skin and gills and modulates the immune system [3]. *L. salmonis* is exceptional among parasite species in infecting adult wild Atlantic salmon (*Salmo salar*) with 100% prevalence. The infective planktonic larva is extraordinarily effective at locating and infecting wild Atlantic salmon. For this reason, *L. salmonis* has the potential to become a pest disease to salmonid fish [2].

Understanding the nature of the interactions between *L. salmonis* and its host is crucial for identifying possible ways to resolve the negative impacts of this infection [4]. *L. salmonis* is able to detect different stimuli (e.g., pressure/moving water, chemicals and light) in its habitat. However, the response thresholds to these stimuli and the role that they play in the context of host location are still unknown. Sea lice use physical (light and salinity) and chemical (kairomones) cues to locate and recognize their host. Another fundamental sensory modality to fish location is mechanoreception through sensory organs, which allows sea lice to detect and land on their host [5].

The life cycle of *L. salmonis* includes ten stages, three of which are pelagic [1]. The third, the copepodid, represents the infective stage of the salmon louse. It carries both chemosensory aesthetes and mechanosensory setae on its antennules, indicating that both mechanical and chemical signals may be important in host-finding [6]. Initial attachment

**Citation:** Solé, M.; Lenoir, M.; Fortuño, J.-M.; De Vreese, S.; van der Schaar, M.; André, M. Sea Lice Are Sensitive to Low Frequency Sounds. *J. Mar. Sci. Eng.* **2021**, *9*, 765. https:// doi.org/10.3390/jmse9070765

Academic Editor: Milva Pepi

Received: 12 June 2021 Accepted: 6 July 2021 Published: 12 July 2021

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**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

for the copepodid normally occurs on the fish fins where it hooks into the host tissue. After some time (depending on the temperature) the copepodid undergoes a moult to the first sessile stage in the life cycle, the chalimus, which attaches itself to the fish with a penetrative thread referred to as the frontal filament [7]. Adult *L. salmonis* present different types of sensitive setae on their antennae. Heuch and Karlsen [8,9] found that salmon louse copepodids are sensitive to low frequency water accelerations such as those produced by a swimming fish.

**Figure 1.** Light microscopy (LM) (**A**–**C**,**H**–**J**) and Scanning Electron Microscopy (SEM) (**C**–**G**). External morphology of *L. salmonis.* (**A**) Dorsal view of the head of an adult sea lice. (**B**) Dorsal view of pre-adult. (**C**) Dorsal view of chalimus. (**D**) Ventral view of the whole body of an adult *L. salmonis*. (**E**) Ventral view of a sea lice head. (**F**) Ventral view of an adult *L. salmonis* showing the mouth, maxilas and abdominal arms. (**G**) Detail of the ventral cavity showing the mouth and the three maxila. (**H**) String of sea lice eggs. In the low part of the string, two larvae are being extruded from the eggs. (**I**) Nauplius just hatched from the egg. (**J**) Copepodid of sea lice. Scale bar: (**A**,**C**,**E**) = 1 mm; (**D**) = 3 mm; (**B**,**F**,**H**) = 500 μm; (**I**,**J**) = 200 μm, G = 100 μm.

Recent findings on cephalopods [10], gastropods [11], crustaceans [12], bivalves [13] and cnidarians [14] have shown that exposure to anthropogenic noise: (i) had a direct consequence on the functionality and physiology of the statocysts, which are sensory organs responsible for these species equilibrium and movements (linear and angular accelerations) in the water column; and (ii) was challenging to the exposed individuals' survival. These experiments demonstrated the sensitivity of invertebrates to sound and described the associated pathological effects. Electron microscopy revealed injuries in the statocyst sensory epithelium after exposure to sound. The lesions present on the exposed animals were consistent with the manifestation of a massive acoustic trauma observed in other species.

In contrast to other invertebrate species that require statocysts to sense the water column in order to maintain balance, zooplankters (copepods, protists) use external mechanosensors for sensing spatial velocity gradients generated by preys or predators [15]. It is understood that the absence of gravity receptors (i.e., statocysts) in planktonic animals has to do with the specific gravity of the zooplankton body, which is the same or slightly higher than water. In *L. salmonis,* these external mechanoreceptors are located on the first antenna [16].

The present study aimed at addressing the problem of sea lice infestation on salmon by using acoustic and bioacoustics techniques and by evaluating the potential effects of these techniques on the parasite's sensory organs.

#### **2. Methods**

#### *2.1. Laboratory Experiments*

We initially looked at the physiological response of lice after exposure to sound in a controlled laboratory environment.

#### 2.1.1. Sound Exposure

Frequency

We tested a series of discrete frequencies, ranging from 100 Hz to 1 kHz. After determining which frequencies would trigger a stronger reaction to the sound exposure, we exposed with combinations of the optimal frequencies and compared the results with the discrete frequency experiments (Tables 1 and 2).

**Table 1.** Number of lice per frequency.




**Table 2.** Combinations of frequencies and corresponding number of lice exposed and used as control.


#### Sea Lice Specimens

Fifty (50) sets of five hundred (n = 500) copepodids from *L. salmonis* were shipped to our laboratory facilities immediately after they had moulted. In the laboratory they were held in a closed system of natural seawater (at 7–10 ◦C, salinity 35‰) consisting of plastic tanks with a capacity of 20L until required for the experiments. An air pump facilitated the copepodid movements in the water column.

Individuals were maintained in the tank system until exposure. Several specimens (see below) were used as controls and were kept in the same conditions as the experimental animals until they were exposed to noise.

#### Sound Exposure Protocol

Sequential Controlled Exposure Experiments (CEE) were conducted on copepodids (25 × 500) of *L. salmonis*. A same number of copepodids (25 × 500) was used as a control and were kept in the same conditions as the exposed ones (Table 1 andTable 2).

The sound was produced and amplified through an in-air loudspeaker while the received levels were measured by a calibrated B&K 8106 hydrophone. The sound production was tuned such that each constant tone from 100 Hz to 1000 Hz was measured at 150 dB re 1 μPa2/Hz at a fixed point in the tank.

The copepodids were exposed to sound for 4 h. The sample collection and the fixation of the lice were performed immediately after the end of the sound exposure session. The controls remained for the same time as those exposed in the isolated exposure tank (4 h) without being exposed to playback. The sacrificing process after exposure was identical for both the controls and exposed animals.

#### Amplitude

In this experiment, we tested the level of lice trauma after exposing them to different combinations of exposure duration, frequencies, and levels of exposure in order to define the required SEL that would trigger potential lesions and, accordingly, determine the combination that would best induce damage to the lice (Table 3).


**Table 3.** Frequencies and combination of frequencies and amplitudes used.

#### Sea Lice Specimens

Ten (10) sets of five hundred (n = 500) copepodids from *L. salmonis* were kept (until required for the experiments) in a closed system of natural seawater (at 7–10 ◦C, salinity 35‰) consisting of plastic tanks with a capacity of 20 L. An air pump facilitated the copepodid movements in the water column.

Individuals were maintained in the tank system until exposure. Several specimens (see below) were used as controls and were kept in the same conditions as the experimental animals until they were exposed to noise.

#### Sound Exposure Protocol

The results of the previous experiments allowed us to determine the best response from the lice and to choose the corresponding frequencies or combinations of frequencies to be used for further experimentation. Sequential Controlled Exposure Experiments (CEE) were then conducted on copepodids (5 × 500) of *L. salmonis*. Five additional sets of copepodids (5 × 500) were used as control (Table 3). The same sound exposure set up and sacrifice protocol used as in Section Frequency was followed.

#### 2.1.2. Imaging Techniques

The same imaging techniques were used as with previous experiments with cephalopods [17]. Individuals were processed according to routine Scanning (SEM) and Transmission (TEM) electron microscopy procedures.

#### Light Microscopy (LM)

Previous to preparing the samples for analysis by SEM and TEM procedures, some light microscopy images of live individuals were taken in order to clarify the morphology and location of the sensory setae of the first antenna.

#### Scanning Electron Microscopy (SEM)

All sets of *L. salmonis* copepodids (control and treatments of exposed sea lice) were used to analyse the lesions after sound exposure.

Fixation was performed in glutaraldehyde 2.5% for 24–48 h at 4 ◦C. Samples were dehydrated in graded ethanol solutions and critical-point dried with liquid carbon dioxide in a Bal-Tec CPD030 unit (Leica Mycrosystems, Austria). The dried specimens were mounted on specimen stubs with double-sided tape. The mounted samples were gold coated with a Quorum Q150R S sputter coated unit (Quorum Technologies, Laughton, East Sussex, UK) and viewed with a variable pressure Hitachi S-3500N scanning electron microscope (Hitachi High-Technologies Co., Tokio, Japan) at an accelerating voltage of 5 kV in the Institute of Marine Sciences of the Spanish Research Council (CSIC) facilities.

SEM images were used to determine the number of fused setae and to calculate the rate (%) of irregular branching tips of the first antenna that were fused after each treatment.

#### Transmission Electron Microscopy (TEM)

Ten (10) exposed and ten control *L. salmonis* copepodids were used for this study. Fixation was performed in 2.5% glutaraldehyde-2% paraformaldehyde for 24 h at 4 ◦C. Subsequently, the samples were osmicated in 1% osmium tetroxide, dehydrated in acetone, and embedded in Spurr. To orient the specimens properly, semithin sections (1 mm) were cut transversally or tangentially with a glass knife, stained with methylene blue, covered with Durcupan, and observed on an Olympus CX41. Ultrathin (around 100 nm) sections of the samples were then obtained by using a diamond knife (Diatome) with an Ultracut Ultramicrotome from Reichert-Jung. Sections were double-stained with uranyl acetate and lead citrate and viewed with a Jeol JEM 1010 at 80 kV. Images were obtained with a Bioscan camera model 792 (Gatan) at the University of Barcelona technical services.

#### *2.2. Sea Trials*

Sea trials were performed at an experimental fish farm located in Averøy (Norway). Two isolated test cages were exposed to sound and four cages at different distances from the test cages were used as a control (Figure 2).

**Figure 2.** Control and test cages distribution.

#### 2.2.1. Sound Exposure

Sound Exposure Level (SEL)

The protocol refers to the term "Sound Exposure Level (SEL)" as the total cumulative squared sound pressure that an organism is exposed to, expressed in decibels. In other words, it is equivalent to exposing the target organisms to a certain dose of sound. Here, the duration of the exposure represents the cumulative time interval that is necessary to induce lesions.

Sound Pressure Level (SPL)

To expose the target organisms to the necessary SEL, the protocol also refers to the term "Sound Pressure Level (SPL)" as the average sound pressure ina1s time interval expressed in decibel, which is produced by an underwater speaker, called a transducer.

### Acoustic and Time Parameters for the Sea Trials

The protocol was based on a method and system (Figures 3 and 4) where lice are exposed to continuous acoustic signals over time until a target Sound Exposure Level (SEL) is achieved for the organism: the SEL is chosen at a level that induces sufficient lesions in the sensory organs of the lice to disrupt vital functions necessary for survival, and particularly to detect (and attach) salmon.

**Figure 3.** Sound exposure system.

**Figure 4.** Drawing of the experimental setup. Note that the depth of the structure that holds the loud speakers was modified along the duration of the experiments. M9 loud speakers were lowered to −5 m.

Based on the output of the previous laboratory experiments (see Section 3, Results) where the lice showed sensitivity to a rather broad range of frequencies (and particularly to continuous exposure to individual 350 Hz and 500 Hz signals) during, respectively, a cumulative cycle of 2 h and 1 h, this combination was initially played back every 4 h (Figure 5). Due to the presence of continuous free-swimming lice from external sources, it was decided to increase the exposure time as well as the SPL at one cage so that the required SEL could be reached faster (see below).

**Figure 5.** Sound exposure protocol. Note that this cycle was modified along the duration of the experiments favouring the exposure to 500 Hz to produce higher SPL.

#### Hardware

Under the sea trial protocol, the system and method included producing the sounds using calibrated transducers capable of reproducing sound covering the essential part of the sensitivity range for the lice, particularly from 300 Hz to 600 Hz. The transducers had a source level of at least up to 160 dB re 1 μPa<sup>2</sup> at 1 m for individual frequencies and 180 dB re 1 μPa<sup>2</sup> at 1 m for each selected third octave band. The transducers were driven by amplifiers that could reach the required voltages to reach these levels. Typical peak voltage levels were below 100 V. The sound production system was calibrated as a whole and for each individual frequency.

Calibrated hydrophones recorded the acoustic pressure in a given frequency range with maximum sound pressure levels at least up to 180 dB re 1μPa2 without saturation. The hydrophone system was arranged to provide digitized data to a sound exposure control system.

#### 2.2.2. Scanning Electron Microscopy (SEM)

Ten (10) exposed and ten control *L. salmonis* for each pre-adult and adult stage were used for this study. The fish in sea conditions were naturally exposed to lice infestation. The lice specimens were collected after two weeks from the start of the sound exposure experiments from the fishes in the cages and processed immediately for the analysis. Individuals were processed according to routine SEM procedures (See Section Scanning Electron Microscopy (SEM)).

#### 2.2.3. Transmission Electron Microscopy (TEM)

Ten (10) exposed and ten control *L. salmonis* for each chalimus, pre-adult and adult stage were used for this study. The specimens were collected after three and six weeks (chalimus), after six weeks (adults and pre-adults) from the start of the sound exposure experiments, and from the fishes in the cages and were processed immediately for the analysis. Individuals were processed according to routine TEM procedures (See Section Transmission Electron Microscopy (TEM)).

#### **3. Results**

#### *3.1. Laboratory Experiments*

#### 3.1.1. *L. salmonis* Copepodids Sensory Setae Morphology

Copepodids (length 0.7 ± 0.01; width 0.2 ± 0.01) (Figure 6) present 10 pairs of setules arranged symmetrically about the medial longitudinal axis (6 pairs of simple setules, 4 pairs bifurcates) on the dorsal shield of the cephalothorax and 2 simple setules near the base of the rostrum.

**Figure 6.** SEM. Copepodid setae morphology. Control animals: (**A**) Dorsal and (**B**) Ventral view of a *L. salmonis* copepodid. (**C**) Cephalothorax dorsal view showing some paired setae distributed along the body (arrows). (**D**) Detail from C shows the structure of a birrame setae (arrow). (**E**) Dorsal view of the abdomen showing some paired setae (arrows). (**F**) Mouth of the copepodid. (**G**) Ventral arms showing pinnate setae (arrow). (**H**) Caudal ramus showing the distal setae (arrow). (**I**) First antenna. The irregular branching tips are visible (arrowheads). (**J**–**L**) Detail of the first antenna setae showing their irregularly branching tips. Scale bar: (**A**,**B**) = 300 μm; (**C**,**G**,**H**,**I)** = 50 μm; (**E**) = 30 μm; (**F**) = 20 μm; (**J**,**K**) = 10 μm; (**D**,**L**)=5 μm.

The first antenna (Figure 6) presents a proximal segment with 3 unramed setae and a distal segment with 5 setae with irregularly branching tips, 7 unramed setae, and 1 aesthete. The second antenna exhibits 3 segments with a spiniform process.

The 3 thoracic legs (Figure 6) present plumose setae, semipinnate setae, pinnate setae, spines, spiniform process, and fine setules. Caudal ramus (Figure 6) shows both short and long pinnate setae and aesthete.

#### 3.1.2. Ultrastructural Analysis of Copepodids Setae after Noise Exposure

Ultrastructural changes took place on *L. salmonis* copepodid setae following acoustic exposure. In the control animals, the first antenna presented completely free setae with irregularly branching tips on the distal segment (Figure 7A–D). All the exposed copepodids presented different degrees of fusion of the irregularly branching tips of the setae on the distal segment of the first antenna (Figure 7E–T).

**Figure 7.** SEM: (**A**) *L. salmonis* copepodids. Setae on distal segment of first antenna; (**A**–**D**) Normal setae on control animal. The tips on the setae distal segments are entirely free (not fused); (**E**–**H**) Different views of exposed animals showing fusion (arrowheads) on the basal segment of the setae on the distal segment of the first antenna; (**I**–**N**) Different views of exposed animals showing the almost entirely fused (arrowheads) distal segment of the first antenna; (**O**–**T**) Different views of exposed animals showing completely fused distal segment of the first antenna. Scale bar: (**A**–**I**,**K**,**L**,**N**,**Q**) = 10 μm; (**M**,**O**,**P**,**R**)=5 μm; (**J**,**S**,**T**)=3 μm.

3.1.3. Exposure Parameters vs. Lesions Frequency

After sound exposure and the analysis of the first antenna setae of the sea lice, we found maximum fusion at 350 Hz (95.5%). However, with frequencies between 300 Hz and 550 Hz, we achieved a percentage of setae fusion higher than 90% (Figure 8).

**Figure 8.** Setae fusion on sea lice first antenna (%) in function of frequency. 350 Hz achieved the maximum percentage of setae fusion. Between 350 Hz and 550 Hz the fusion percentage was higher than 90% (red bar).

> After exposure to combinations of frequencies that previously had achieved the maximum fusion and the analysis of the first antenna setae of the sea lice, we found that 350 Hz–450 Hz and 350 Hz–550 Hz were the two combinations that achieved the maximum percentage of setae fusion (95.2%) (Figure 9).

> We appointed that there was not a significant increase in the level of fusion with the combination of two frequencies.

**Figure 9.** Setae fusion on sea lice first antenna (%) in function of frequency combinations of 350 Hz–450 Hz and 350 Hz. 550 Hz are the combination that achieve the maximum percentage of setae fusion (95.2%).

#### Amplitude

We found maximum setae fusion with the combination of 350 Hz-2 h-65 V and 500 Hz-2 h-65 V (93.02%). Other combinations (e.g., 350 Hz-2 h-65 V and 500 Hz-1 h-65 V, 350 Hz-3 h-65 V) achieved a percentage of setae fusion higher than 90% (Table 4).

**Table 4.** Setae fusion on sea lice first antenna (%) in function of frequency, time, and level of exposure in our tank conditions.


3.1.4. Determination of Ultrastructural Lesions in Inner Tissues of Sea Lice Copepodids

Sound exposure affected the nervous system and A/B cells, which are responsible for the precursor secretions of the frontal filament. The lesions present in all the samples of exposed sea lice copepodids (but not in any of the control animals) were characterized by an exuberant accumulation of dark material and by cell cytoplasm vacuolization (Figures 10 and 11). These pathological features suggest the involvement of massive autophagy processes.

**Figure 10.** TEM. Sagittal section of the copepodid anterior cephalotorax showing the copepodid eye: (**A**,**C**) Control copepodid; (**B**,**D**); Exposed copepodid. (**A**) In control animals the dark inclusions around the eye are scarce. (**B**) In exposed copepdids a large amount of dark inclusions are visible in the axons of the central nervous system surrounding the eye (arrowhead). Vacuolization is visible on the tissue surrounding the eye (arrow); (**C**) Detail of A showing the optic nerve. Note the low quantity of dark inclusions around the eye; (**D**) Detail from B. Arrowhead point to the large amount of dark inclusions. Scale bar: (**A**,**B**) = 20 μm; (**D**)=5 μm; and (**C**)=2 μm.

**Figure 11.** TEM. Frontal medial section of the copepodid anterior cephalotorax showing A and B Cells involved in Frontal Filament production: (**A**,**B**,**D**) Control; (**C**,**E**,**F**) Exposed: In control animals A and B cells do not show inner dark inclusions. (**B**) Axons sited next to A and B cells present normal aspect. (**C**) In exposed copepodids dark inclusions are visible in the axons of the nervous system (arrow). (**D**) Normal aspect of cells without dark inclusions. (**E**,**F**) Dark inclusions in the cells of exposed copepodids (arrows). Scale bar: (**A**) = 20 μm; (**C**,**D**)=5 μm; and (**B**,**E**,**F**)=2 μm.
