Dissertation for the degree of philosophiae doctor (PhD)

at the University of Bergen

Dissertation date:

(Lepeophtheirus salmonis)

© Copyright Liv Sandlund

The material in this publication is protected by copyright law.

Year: 2016

Title: Functional investigations of the Ecdysone Receptor and production of

ecdysone in the salmon louse (Lepeophtheirus salmonis)

Author: Liv Sandlund

Print: A T AS / University of Bergen

i Bjerch

2

Scientific environment

The work of this thesis was carried out at the Sea Lice Research Centre (SLRC),

Department of Biology, University of Bergen, Bergen, Norway, in the period

February 2012- January 2016. The project was financially supported by: the Research

Council of Norway: Centre for Research-Based Innovation project number

203513/O30. The education was formally administered by the Department of

Biology, University of Bergen.

3

4

Dedicated to my nieces Tuva and Eira

“Much to learn, you still have”

-Yoda-

5

6

Acknowledgements

I would like to give a special thank to my master supervisor Sussie Dalvin: Thank

you for providing help and guidance throughout the last four years and for correcting

my manuscripts a.s.a.p!! I would also like to thank my co-supervisors Frank Nilsen

and Rune Male for always keeping your office doors open for a desperate PhD

student and input along the way.

I would also like to thank Heidi Kongshaug for all the lab help, mountain hikes and

encouraging words in the late hours. Thank you Lars Hamre and Per Gunnar Espedal

for providing The Lice and letting me feed the fish so often during the weekends and

holidays.

Furthermore, I would like to thank my colleagues at the SLRC and the fish health

group for creating a good and fun work environment!! A special thanks to Ewa

Harasimczuk, Steffen Blindheim and Arnfinn Lodden for entertaining coffee breaks

and a lot of good laughs! Thank you Anna Komisarczuk and Aina Øvergård for

general PhD help ☺. Thanks to all my friends (spesielt Lene, Hanne og Tone) for

your support and cheers ☺ and to Sir David Attenborough who`s documentaries are a

true inspiration.

To my family: Mamma, Pappa & Nina tusen takk for all støtte gjennom hele

utdannelsesløpet! Hadde ikke greid det uten dere, gratis middager og

overraskelsespakker i posten!! ☺

Last, but not least: To my Bond, thanks for always saying “You can do it!”, for

picking me up and make me smile on the roughest days and for always believing in

me ☺ You are the best!

7

8

Table of Contents ACKNOWLEDGEMENTS ....................................................................................... 6

ABBREVIATIONS ................................................................................................... 10

ABSTRACT ............................................................................................................... 12

1. GENERAL INTRODUCTION ......................................................................... 14

1.1 BACKGROUND ................................................................................................. 14

1.2 SALMON LOUSE ............................................................................................... 15

1.2.1 Salmon louse; biology, behaviour and host interactions ......................... 16

1.2.2 General anatomy of the salmon louse ...................................................... 18

1.3 NUCLEAR RECEPTORS ..................................................................................... 20

1.4 DISCOVERY OF THE ECDYSONE RECEPTOR ..................................................... 22

1.4.1 Structural domains of the Ecdysone Receptor ......................................... 23

1.4.2 EcR mediated ecdysone signalling ........................................................... 25

1.5 BIOSYNTHESIS OF ECDYSTEROIDS ................................................................... 28

1.6 RNA INTERFERENCE AS AN EXPERIMENTAL TOOL .......................................... 32

2. AIMS OF THE STUDY ..................................................................................... 34

LIST OF PUBLICATIONS ..................................................................................... 35

3. ABSTRACT OF PAPERS ................................................................................. 36

3.1 PAPER I: ........................................................................................................... 36

3.2 PAPER II: ......................................................................................................... 36

3.3 PAPER III: ........................................................................................................ 37

4. GENERAL DISCUSSION ................................................................................. 39

4.1 IDENTIFICATION OF ECDYSTEROIDOGENIC GENES AND THE ECDYSONE

RECEPTOR ................................................................................................................ 40

4.1.1 Characterization of the ecdysone biosynthetic genes: neverland,

disembodied and shade in L. salmonis ............................................................... 40

4.1.2 Characterisation of the L. salmonis EcR .................................................. 40

4.2 FUNCTIONAL ASSESSMENT OF LSECR DURING MOLTING AND DEVELOPMENT 41

4.3 FUNCTIONAL ASSESSMENT OF LSECR DURING REPRODUCTION ...................... 44

9

4.4 BIOSYNTHESIS OF ECDYSTEROIDS IN L. SALMONIS ........................................... 47

4.4.1 Functional assessment of ecdysteroid biosynthetic enzymes in L. salmonis

early development ............................................................................................... 47

4.4.2 Functional assessment of L. salmonis Oct3βR during early development48

4.5 THE ECDYSONE PATHWAY AND LICE CONTROL ............................................... 49

5. CONCLUDING REMARKS AND FUTURE PERSPECTIVES ................... 51

REFERENCES .......................................................................................................... 53

10

Abbreviations

20E 20-hydroxyecdysone

aa amino acid

AF-1 activation function 1

AF-2 activation function 2

Ab/G abdomen/genital segment

CT cephalothorax

DBD DNA binding domain

Dib disembodied

E ecdysone

EcR ecdysone receptor

EcRE ecdysone response elements

HRE hormone response elements

LBD ligand binding domain

Nvd neverland

Oct3βR β3-Octopamine receptor

PonA ponasterone A

PCD programmed cell death

PG prothoracic gland

RT-qPCR Real-time quantitative PCR

RXR retinoid X receptor

Shd shade

USP ultraspiracle YO Y-organ

11

12

Abstract

The salmon louse Lepeophtheirus salmonis is a marine ectoparasitic copepod

naturally infecting salmonid fishes in the Northern hemisphere. At present, salmon

louse infections are the most severe disease problem in the salmon farming industry

causing significant economical losses. Salmon louse infections on farmed fish have

largely been treated with chemotherapeutants which in recent years have lead to the

development of resistance towards the majority of available treatment methods. Cases

of multi-resistance are reported as increasing, underlining the severity of the

situation. Although non-therapeutic methods such as the use of cleaner fish have been

implemented into the management of lice infestations, it is clear that new alternative

methods are essential to gain control of the parasite in the future.

The ecdysone hormone system and the ecdysone receptor/retinoid X receptor

(EcR/RXR) complex are key regulators of molting, development and growth in

arthropods. A wide range of studies has demonstrated the importance of this

endocrine system in insects and to some extent in malacostraca, but knowledge about

ecdysone signalling in copepods is limited. Therefore, we aimed to increase our

knowledge about this hormone system in the salmon louse. In this study, the

ecdysone receptor (EcR) and the key ecdysteroidogenic genes neverland (nvd),

disembodied (dib) and shade (shd) were identified and functionally assessed using

RNA interference studies. LsEcR transcripts were expressed in all life stages and in

most tissues in both the copepodid (i.e. brain, muscle and immature intestine) and the

adult female (i.e. ovaries, sub cuticula, intestine, oocytes and glandular tissue). The

wide tissue distribution is expected due to the numerous physiological and biological

processes that are regulated by EcR signalling. Interestingly, knock-down of LsEcR

in nauplia I larvae did not cause immediate molting arrest, but developed into viable

copepodids, indicating another partner of RXR. However, further incubation of the

LsEcR knock-down copepodids on salmon resulted in severe tissue damage and

increased mortality. During metamorphosis, an extensive range of tissues is

remodelled concurrently with the molting process in order to adapt to the new life

13

stage. The results obtained from the studies indicate that LsEcR is a key regulator of

developmental processes associated with tissue remodelling and molting in L.

salmonis.

Ecdysteroids regulate reproductive processes in arthropods such as vitellogenesis and

oocyte maturation. In the salmon louse, vitellogenin and yolk production takes place

in the subcuticula before being incorporated into the oocyte. These processes showed

to be indirectly affected by knock-down of LsRXR which resulted in abnormal egg

chambers and no egg-production. This is supported by knock-down studies of LsEcR

in pre-adult females resulted in hypotrophy of tissues associated with yolk and

vitellogenin synthesis, degeneration of the oocytes and termination of egg production.

This demonstrates that LsEcR plays a key role in the reproduction of the salmon

louse. We have not proven that ecdysteroid signalling has a direct effect on the

oocytes. However, LC/MS/MS analysis of the ecdysteroid content during oocyte

maturation in adult females showed higher levels of ecdysteroids in the

abdomen/genital segment (Ab/G) compared to the cephalothorax (CT), suggesting

that ecdysteroids may directly affect oocyte maturation and embryogenesis.

The biosynthesis of the ecdysteroids is performed by members of the cytochrome

P450 family through several enzymatic reactions. The last step in the pathway is the

conversion of ecdysone to the active metabolite by Shd. Knock-down of the

biosynthetic genes in other species is associated with embryonic defects and

mortality. Interestingly, functional assessment of LsNvd, LsDib and LsShd in L.

salmonis nauplia I larvae resulted in molting arrest only in the LsShd knock-down

animals, while the LsNvd, LsDib developed normally into copepodids. Based on the

knock-down results and the biology of the salmon louse, we hypothesize that

ecdysone are incorporated into the oocytes during maturation and is further converted

by LsShd into the active metabolite during development of the lecitotrophic stages.

To our knowledge, no-loss of function phenotype has been observed in nvd and dib

knock-down animals during early development in other arthropods, indicating a

possible novel regulation pathway in L. salmonis.

14

1. General introduction

1.1 Background

Norway is the largest producer of Atlantic salmon (Salmo salar) in the world. The

production has more than doubled the last decade from 550 000 metric tons in 2004

to more than 1.2 metric million tons in 20141 (www.fao.org; www.ssb.no). With intensive

cultivation and a growing aquaculture industry, challenges concerning fish pathogens

have become an increasing issue in terms of economical loss and negative

environmental interactions. In Norway, the salmon louse (Lepeophtheirus salmonis)

is one of the most severe pathogens in salmon farming estimated to cost the farming

industry as much as 3-4 billion NOK in 2014 http://nofima.no/nyhet/2015/08/kostnadsdrivere-i-oppdrett/.

Since the beginning of large-scale salmon farming in the 1970`s, salmon lice

infestations have mainly been controlled using anti-sea lice medicaments. However,

the efficacy of medical treatments has decreased due to development of lice

populations with reduced sensitivity and resistance towards one or more of the anti-

parasitic drugs available [4-7]. In the south- and mid-west of Norway where the lice

problems are most severe, occurrences of multi-resistance towards all available

medicines, including hydrogen peroxide, have been reported [8]. The result has been

an increase in the use of chemotherapeutants (e.g. chitinase inhibitors

(fluobezurones), nerve toxins (emamectin benzoate, pyrthroids), organophosphates

affecting the cholinergic nerve system (azamethiphos) and hydrogen peroxide)

causing concerns due to potential negative impact on non-target organisms and the

surrounding environment of the fish farm [9]. Additionally, lice larvae originating

from fish farms can infect wild salmonids and influence post-smolt survival during

costal and oceanic migration. Currently, high sea lice densities as well as escaped

farmed salmon have been suggested to be the two most significant threats to the wild

salmon populations in Norway. Due to the significant lice problems and reduced

efficacy of the existing medicines available, alternative non-medical methods are

under development and some of these methods are currently being implemented to

15

facilitate lice control in salmon farms. The use of cleaner wrasse [10], mechanical

delousing and lice-skirts to reduce lice infestations are just a few alternative methods

that have been applied by the aqua-culture industry the last few years. Regardless,

salmon louse infestations remain persistent making it imperative to develop novel

treatment methods to control the parasite population in order to sustain both farmed

and wild salmon populations and a growing industry. However, most of these

methods are not sufficiently efficient (or the capacity is too low) to bring the lice

levels down to the limits enforced by the authorities alone and must be used in

combination with other tools. Medical treatment of salmon louse has been the most

important tool for parasite control. The last new medicine was introduced in 1999

(SLICE®) and there is a strong demand for new efficient medicine. Increasing basic

knowledge about key biological processes in the salmon louse is a long-term effort

towards new treatment tools. In arthropods, ecdysone hormones regulate key steps in

development, growth and reproduction. Hence, a highly relevant topic for research

based innovation towards future sea lice control.

1.2 Salmon louse

Lepeophtheirus salmonis (Krøyer, 1837) is a marine ectoparasitic copepod on

salmonid fishes from the genera Oncorhynchus (O. mykiss (Walbaum, 1792)),

Salvelinus (S. alpinus (Linnaeus, 1758)) and Salmon (S. salar (Linnaeus, 1758) and

S. trutta (Linnaeus, 1758) [11]. Two allopatric subspecies of Lepeophtheirus

salmonis has been identified; Lepeophtheirus salmonis salmonis (Atlantic) and

Lepeophtheirus salmonis onchorynchii (Pacific) [12]. The salmon louse has a

northern circumpolar distribution and is naturally found on wild salmon populations

[13]. The parasitic stages feed on mucus, skin and blood [14], causing local erosion in

the epidermal tissue of the host, often on and near the head and dorsal fins [15]. At

high infestation rates, chronic stress, lesions in the dermis and subcutaneous tissue

can occur, compromising the osmoregulation of the host in which can lead to host

mortality, especially to post-smolts [13, 15-17]. Salmon louse infestations has

negative effects on the host`s reproduction, growth and quality. Damage to the skin

16

makes the fish more susceptible to secondary infections as the skin acts as their first

line of defence against pathogens [18]. The salmon louse itself has also been

suggested to be an important vector in transferring diseases between fish [19].

1.2.1 Salmon louse; biology, behaviour and host interactions

The life cycle of the salmon louse (Fig. 1.) includes both a planktonic and a parasitic

phase. It consists of eight developmental stages where each instar is separated by a

molting event where the exoskeleton is shed [20-22]. The developmental rate is

temperature dependent and at 10°C, it takes around 40(♂)-52(♀) days to complete the

lifecycle on S. salar [23]. The planktonic stages after hatching, nauplius I and II and

the free-living copepodid are lecithotrophic (feeding on yolk reserves only [24, 25].

Planktonic stages are passively dispersed by the ocean currents that enables them to

spread over great geographic distances and infect farming sites up to 30 km away [26,

27]. To increase host-parasite encounter, the infectious copepodid displays both

positive phototactic and rheotaxic qualities. This allows the copepodid to seek

towards the upper water column (> 27 ‰ salinity) where they react to pressure waves

generated by a nearby swimming fish [28]. It is believed that the copepodids use

chemo- and mechano sensors located on their frontal antennas to identify the right

host [29, 30]. The copeodid attach to the host using the 2nd antennae which has hook

like structures [13] and starts to feed instantly after attachment [31]. During molting

into and through the two attached chalimus stages (Fig. 1.), the lice generate and

remains attached to the host by a frontal filament [32]. Sexual dimorphism is apparent

in the chalimus II stage [33]. After the chalimus stages, the louse molts into the

mobile pre-adult I and II and, finally, the adult stage. In the mobile stages, the lice are

able to feed over a larger area of the host and thereby increasing the virulence

significantly [16]. Males become adults prior to the females and once males are adult,

they locate immature pre-adult II female and engage in precopula. The male deposit

spermatophores onto the genital complex of the female [34]. Despite deposition of

the spermatophore and guarding of the females, polyandry is known to occur which

can lead to multiple paternities [35, 36]. A female louse is thought to be able to

17

produce eggs throughout her lifetime (≤ 15.5 months in the lab) and each pair of egg

strings can result in as much as 1200 eggs [36, 37].

L. salmonis infestations and the degree of damage that is inflicted upon the host

largely depend on the amount of lice present and host size leaving post-smolts

particularly vulnerable to the parasite [15, 38]. In addition, the effects of salmon louse

significantly vary between species due to natural resistance. Juvenile pink (O.

gorbuscha) and coho (O. kisutch) salmon rejects L. salmonis copepodids faster than

chum (O. keta) and Atlantic salmon [39, 40]. Atlantic salmon has been shown to

Fig. 1. Representation of the life cycle of Lepeophtheirus salmonis. The generation time

of adult male and females are 40-52 days. Nauplia I/II and the free-swimming copepodid

are nourished by yolk-reserved until the copepod becomes parasitic. The adult female

generates a new pair of egg strings every ten days. Both development and reproduction is

temperature dependent and the given time-points correspond to 10 °C. The figure is from

Schram 1993 and adapted by The Marine Institute of Galtway.

18

possess a limited inflammatory response and epithelial hyperplasia compared to the

coho and chinook (O. nerka) salmon [40]. Recent gene expression analysis of skin

from susceptible and resistant species has found that the susceptible species have a

lower expression of pro-inflammatory genes compared to more resistant species [39,

41-44]. Additionally, Atlantic salmon has compared to the more resistant salmonids a

lower concentration of protective mucosal lysozymes and proteases due to thinner

epidermis and less mucosal cells [45]. In addition, some studies have indicated that

the mucosa of susceptible hosts stimulates secretion of trypsin-like proteases and

prostaglandin E2 (PGE2) from the salmon louse, which is thought to modulate host

immunity [46-49].

1.2.2 General anatomy of the salmon louse

Cuticle

The exoskeleton of arthropods consists of chitin, sclerotin and calcium carbonate,

which pose a unique challenge for growth due to its rigid structural arrangement. In

order to grow, arthropods must form a new skeleton before discarding the old cuticle

(molt). Molting requires a series of physiological steps, which is initiated by

separation of the old cuticula from the underlying epidermal layers. The newly

created gap (exuvial space) is filled with molting fluid where the old exoskeleton is

digested from underneath, and the protein components are absorbed and reused to

build the new exoskeleton. The underlying tissue starts to secrete the new soft

exoskeleton that is convoluted in order to expand when old exoskeleton is shed. After

shedding, the new soft exoskeleton is inflated due to influx of water and hardening of

the new exoskeleton (sclerotinization) begins. The new exoskeleton is folded

allowing the animal to increase in size during the instar phase [33].

Subcuticular tissue

The subcuticular tissue (Fig. 2. black frame) is located underneath the exoskeleton

(cuticula) and is found throughout the louse. The tissue is thought to perform

functions similar to those of the liver and is the site of vitellogenin and yolk-

19

production in adult females [25, 50]. In addition, genes involved in fatty-acid

metabolism and amino acid degradation is associated with the tissue [51] Different

types of glandular tissues are present with in the subcuticular tissue [32].

Alimentary canal

No functional gut is present in the nauplius stage but intestinal tissue fully develops

during the copepodid stage. The gut stretches all the way from the mouth located

anteriorly in the cephalothorax to the rectum of the animal located posterior in the

abdomen (Fig. 2.) [52]. Not surprisingly, expression of digestive enzymes i.e.

proteases and lysosomal genes are present in the intestine [51, 53]. It has been shown

that pancreatic function is located to the intestine in salmon louse [53], which is

different from members of the malacostracan, which have hepatopancreas where liver

and pancreas function is co-localized.

Reproductive organs

The ovaries (Fig. 2. white frame) and testes are paired organs located on each side of

the coalesced eyes in the anterior part of the cephalothorax. The ovaries continuously

produce oocytes that are transported via the oviduct to the genital segment where the

oocyte matures. Vitellogenesis takes place in the genital segment. Maturation of the

oocytes is temperature dependent and takes approximately 10 days at 10 °C.

Spermatozytes are produced in the male testes and are transferred to the female via

spermatophores that are deposited on the female genital segment [34].

20

1.3 Nuclear receptors

Nuclear receptors (NRs) make up an ancient superfamily of mostly ligand-dependent

transcription factors for cell growth and differentiation, metabolism, homeostasis and

embryonic development by directly linking extracellular signals and transcriptional

response [54-56]. Mutations in NRs are associated with many common and lethal

disorders hence extensive research focus on understanding and modulating the NR

functions in order to develop pharmaceuticals that target NRs. Members of the NR

superfamily are believed to be present in all metazoans and are classified into six

subfamilies based on multiple sequence alignments and phylogenetic analysis of

conserved domains [54, 57-61]. NRs like the thyroid receptor, retinoic X receptor and

steroid receptors (i.e. glucocorticoid receptor and the oestrogen receptor) are well

studied in vertebrates and have given important knowledge to the nature of NRs and

their ability to directly regulate gene expression. In insects, the ecdysone receptor

(EcR) and its partner ultraspiracle (USP) has been subjected to intensive studies.

However, much remains to be learned about the role of NRs physiological pathways

Fig. 2. L. salmonis adult female. The sub-cuticular tissue (black frame) is found underneath the cuticula around the edges of the cephalothorax (CT), abdomen/genital segment (Ab/G). The alimentary canal is blood filled and stretches from the CT to the anterior of the Ab/G. The ovaries (white frame) are situated in the front of the coalesced eye. Testes are found in the same position in male lice. The oviducts reach from the ovaries to the Ab/G. Maturing oocytes can be seen in the Ab/G (marked with asterisk). Scalebar = 5 mm

*

21

such as development and reproduction in other arthropods. In the human genome, 48

different NRs are identified compared to the Drosophila genome containing 21 [62],

while 23 NR or NR-like sequences are predicted from the L. salmonis genome

(Licebase.org: unpublished). The primary feature distinguishing the NRs from other

transcription factors is their ability to bind ligands. The ligands include a vide range

of small hydrophobic compounds that are derivatives of vitamins, retinoids, fatty

acids, lipophilic hormones and cholesterol.

The NR genes presumably evolved from a common ancestor more than 700 million

years ago and diversified and duplicated into the subfamilies known today. The

evolved ability to bind ligands as well as the ability to homo- and heterodimerize and

bind DNA, increased the possibilities and complexities of signal transduction and is

considered a potential driving force in the evolution of higher organisms.

In classical signal transduction, an external ligand bind to a membrane-bound

receptor that initiates a cascade of events in the cytoplasm, eventually activating

nuclear transcription factors. In contrast to these very complex and “time-consuming”

pathways, the NRs can shorten the time of signal transduction by their simultaneous

ligand and DNA binding ability. This capability allows for signals to be transferred in

a one-step response that directly affects the expression of the target gene. Generally,

the NR signalling pathway is initiated by diffusion of hydrophobic ligands through

the nuclear membrane followed by receptor binding or binding of ligand to the

receptor in the cytoplasm, dependent on the receptor. Following binding of a ligand,

the receptor complex will translocate to the nucleus or, if already present in the

nucleus, bind to specific hormone response elements (HRE) (reviewed in [55]).

Regulation of gene expression of NRs is enabled by recruitment of co-activators and

co-repressors, which modulate gene transcription by modifying the chromatin

architecture 2 (Fig. 3.) [3].

2 Chromatin structure is modified by ATP-dependent chromatin remodelling complexes and histone modifying complexes. Enzymatic modification includes acetylation, methylation, phosphorylation and ubiquitinylation (e.g. acetylation relax chromatin structure and recruits the transcription machinery in contrast to methylation which condense the chromatin structure and prevents transcription).

22

1.4 Discovery of the Ecdysone Receptor

In 1974, Ashburner and his colleagues postulated a hierarchical genetic response

model for the puffing of polytene chromosomes induced by ecdysone based on the

work performed earlier by Peter Karlson [63]. The model was based on studies of

salivary glands from Drosophila where “puffs” occurred at specific loci on the

chromosomes when treated with ecdysone. The “early puff” response was rapid and

peaked at four hours in the presence of protein synthesis inhibitors, suggesting the

ecdysteroids to act directly on the chromosomes. A set of “late puffs” was observed

to follow the “early puff” response, however, the late puffs did not occur in the

presence of protein inhibitors. The observations lead Ashburner and his colleagues to

suggest that the ecdysone bound to a cognate receptor protein that directly activated

“early puff” expression and that the protein product of the initial response induced a

larger set of “late puffs” expression [64].

Fig. 3. Co-activator and co-repressor complexes are required for nuclear receptor-mediated transcriptional regulation. The figure illustrates the complexity of eukaryotic transcription. Copied from [3].

23

Based on the work of Ashburner and his colleagues, Koelle et al., [65] isolated and

characterized the ecdysone receptor (EcR) from Drosophila by screening cDNA

libraries for members of the steroid receptor family. Although the gene product

acquired the properties consistent with an ecdysone receptor binding both active

steroid and DNA, the receptor was later found only to be fully functional when

dimerized to a second NR, ultraspiracle (USP) a homolog of the retinoid X receptor

(RXR) found in vertebrates and crustaceans [66, 67]. The EcR was initially

recognized as the molting receptor but considerable research the last few decades has

shown that it is also a central regulator of major developmental and biological

processes across the arthropod phyla. A number of EcR orthologs have also been

identified in nematodes, molluscs, leeches, squid and polychaete worms [68]. The

discovery of the ecdysone receptor as a nuclear receptor and target for ecdysone did

not only revolutionize the field of arthropod endocrinology but also showed that the

NRs evolved prior to the divergence of protostomes and deuterostomes.

1.4.1 Structural domains of the Ecdysone Receptor

The EcR is the ortholog to the vertebrate farnesoid X receptor (FXR) [69] and shares

a similar organisation of domains and core modular architecture common to the NRs

(Fig. 4.) [70]. Flanked between a highly variable N-terminal (A/B-domain) that

harbour a ligand-independent-activation function-1 (AF-1) and the hinge region (D-

domain) that plays a role in nuclear translocation [71] and subcellular distribution, is

the central DNA-binding domain (DBD; C-domain). The DBD is highly conserved

maintaining about 50 % identity between all NRs in the superfamily. The domain

contains two zinc-finger motifs that facilitate both sequence-specific interaction with

DNA and receptor complex-DNA dimerization [72]. Most importantly, in terms of

function, is the ligand binding domain (LBD; E-domain) that includes the ligand-

dependent transcription activation function-2 (AF-2). Moreover, some NRs contain a

highly variable C-terminal F-domain that may be involved in the in co-factor

recruitment [73].

24

Ligand binding

To date, the structure of five insect EcR-LBD/USP-LBD in complex with

ecdysteroids or inhibitors have been determined by X-ray crystallography [74-77]. As

expected, all EcR-LBD tertiary structures displays a canonical shape made up of 12

α-helices and an anti-parallel β-sheet that pack together and facilitate the formation

of a hydrophobic ligand binding pocket (LBP). Although EcR is capable of ligand-

binding in the absence of USP, the affinity for ecdysteroid binding increase

significantly in the presence of its heterodimerization partner [78]. After binding of

the ligand, dissociation of ligand-receptor complex is prevented by folding of Helix-

12 (also called AF-2) across the pocket. The conformational change of the helix

exposes an interactive surface enabling recruitment of co-activator proteins and

members of the transcription initiation complex beginning transcription. The LBD of

EcR is flexible and capable of adapting the LBP to ligands with different chemistries

[79]. This feature explains how some arthropods can utilize various ecdysteroids to

regulate development at different life stages. The absence or presence of ligand

determines how the ecdysone receptor binds to DNA and associates with either co-

activators or co-repressors (Fig. 3.) [80-82]. The USP receptor in insects is defined as

an orphan receptor because it is locked in an antagonistic conformation preventing

Fig. 4. Schematic overview of the primary structure of NRs- important properties is listed under the corresponding domain. The A/B domain (AF1) is associated with both ligand dependent and ligand independent transcriptional activation mediating crosstalk between signaling pathways. The C-domain (DNA-binding domain (DBD)) is primarily involved in DNA-dimerization and initiates binding of co-factors. The highly variable D-domain links the DBD and E/F region with the conserved ligand binding domain (LBD) which contributes to dimerization and recruitment of co-regulatory factors. The F part of the domain is highly variable and can be absent in some nuclear receptors.

25

binding of ligand [83]. In contrast, crustacean RXRs has the ability to bind ligands

including 9-cis retinoic acid, methyl farnesoate and neurotransmitters [84, 85].

EcR/USP complex-DNA interactions

The DBDs of the EcR/USP binds as a heterodimer to specific ecdysteroid response

elements (EcREs) which are half-sites with a one base pair spaced inverted repeat

(palindrome; IR1; 5`-AGGTCA-3`) located in the regulatory regions of target genes

[86-88]. Upon binding of DNA, the EcR/USP heterodimer adopts an asymmetrical

organization that induces a conformational change in the LBD of USP, which

stabilises the EcR/USP/DNA complex and aid in the fine-tuning of gene regulation

[89].

1.4.2 EcR mediated ecdysone signalling

Ecdysone and ecdysone signalling has mostly been studied in holometabolous insects

like the model organism Drosophila melanogaster due to the major transitional

changes they undergo during metamorphosis. However, some crustacean species like

shrimp, crabs, lobster and very recently the salmon louse has been increasingly

investigated due to their commercial importance and, therefore, understanding of

mechanisms regulating growth and reproduction has been the topics of investigation.

Insects and crustaceans differ in many ways e.g. growth, sexual differentiation,

reproduction and life cycles but common for both groups are that these biological

events are regulated by ecdysteroids [90]. Additional neurohormones and peptide

hormones such as hyperglycemic hormones (e.g. molt-inhibiting hormone among

others) and farnesoic acids (e.g. methyl farnesoate) are important regulatory factors in

addition to external environmental factors (e.g. temperature, nutrition and salinity).

Crosstalk between signalling pathways regulated by these factors allows for

adaptation of the hormonal response to meet the functional requirements of the target

tissue (for more information please see reviews [90-93]). However, as these

regulatory functions are beyond the scope of this study they will not be further

addressed.

26

Just like the holometabolous insects, the molt cycle in crustaceans results in extensive

physiological changes in addition to changes in the integument. Molting is not just

restricted to shedding of the exoskeleton but also the series of events required for

synthesis, degradation and exchange of the old exoskeleton to facilitate growth and

metamorphosis3 [94-96]. During metamorphosis, an extensive remodelling takes

place in organs like the hepatopancreas, muscles and neurological and adipose tissue

in order to morphologically adapt into the new life stage. During this remodelling,

some tissue is triggered to undergo programmed cell death and histolysis and some

tissue will grow and differentiate while others will not respond at all [97]. All the

processes associated with molting and metamorphosis, are triggered by pulses and

fluctuating levels of circulating ecdysone (reviewed in [55]). During the molting

cycle, the concentration of ecdysone can fluctuate dramatically (e.g. between < 10

ng/ul and > 350 ng/ul) in juvenile lobster Homarus americanus [98] in a time-

dependent manner. The titer remains low during intermolt and postmolt stages but a

peak is reached in the premolt stage with an abrupt drop in ecdysteroid concentration

that triggers the shedding of the exoskeleton [55, 99].

The extensive physiological and biological changes that take place during the life

cycle is achieved by binding of ecdysone to the EcR/RXR complex, which results as

proposed by Ashburner et al., [64] in the regulation of a conserved hierarchical

cascade of hundreds of ecdysone-responsive early genes and early-late genes. The

ecdysone derivative 20E has in both insects and found to be biologically active

hormone during molting [100] in addition to ponasterone A (PonA) present in

crustaceans. The gene products include but are not limited to the transcription factors

E74, E75, Drosophila hormone receptor 3 (DHR3), Broad-Complex (Br-C) and FTZ

transcription factor-1 (FTZ-F1) [101-105]. Products of the early genes subsequently

regulate ecdysone responsive late-genes that determine the phenotypic effects of the

ecdysteroids in a time and tissue specific manner [106]. In addition, in response to the

3 Not all crustaceans go through complete metamorphosis. In addition, molting pattern may vary between species.

27

increase in ecdysone level, EcR provides an auto-regulatory loop and activate its own

transcription thereby increasing its own expression [65].

EcR is typically found in different forms that allows for differential regulation within

time and space. The key role that EcR plays in this comprehensive diversity of

physiological and morphological processes is partially due to the various isoforms.

This allows for differences in the receptor`s ability to repress and activate expression

of down-stream genes and hence influence separate physiological functions. Most of

the EcR variants differ mainly in the N-terminal region which is associated with

regulation of transcription [107-109], however, splice variations of the hinge region

and the LBD has been identified in some crustaceans including, but not exclusive to

the fiddler crab Uca pugilator [110], the kuruma prawn Marsupenaeus japonicus

[111], the freshwater prawn Macrobrachium nipponense [112] and the water flea D.

magna [113]. In Drosophila and Manduca sexta, three EcR isoforms (EcRA, EcRB1

and EcRB2) differing in the length of their N-terminal have been identified with

varying biological effects at different time-points [109, 114]. Mutations that block all

three variants of EcR cause embryonic lethality while removal of one isoform cause

effects in specific physiological processes. In Drosophila, EcRA is predominantly

expressed in cells that proliferate and differentiate during metamorphosis of adult

stages, whereas isoform EcRB1 and B2 are essentially expressed during larval stages

in cells that enter apoptosis [115]. During arthropod development, the neuronal tissue

undergo crucial remodelling of the mushroom bodies (MBs: the brain memory

centre), olfactory circuits and neuromuscular junctions where axons and dendrites are

pruned and regrown to fit their new functions. All neuronal remodelling events

depend on the EcR, however, in an isoform-specific manner (reviewed in [116]).

Growth and reproduction are two processes that are tightly regulated and connected

in arthropods and ecdysteroids play a key role in both of them. The role of

ecdysteroids in insect reproductive processes is well established and has shown to be

important in vitellogenesis [117, 118], follicle development [119] and ovarian and

oocyte development [120]. Depletion of the EcR level is associated with oogenic

defects such as the presence of abnormal egg chambers and loss of vitellogenic stages

28

[120-124]. Although ecdysteroids are primarily considered to be molting hormones in

crustaceans, it has become evident from recent studies that they also play a role in

reproductive processes (e.g. vitellogenesis and ovarian maturation) [125, 126].

The significant effect of ecdysteroids on gene regulation is also evident through

several transcriptomic studies performed in Drosophila Kc cells. Consistent with the

morphological changes during metamorphosis, genes encoding proteins involved in

cell movement and organization associated with the cytoskeleton has shown to be

regulated by 20E [127]. In addition, several members of the cytochrome P450 family,

stress-response genes, lipid transporters, starvation-genes (i.e. peptidases) and Toll-

ligand response genes were regulated by 20E in an EcR- dependent manner [127,

128]. These observations indicate that ecdysone signalling regulates many metabolic

processes as well as the immune response. This is in accordance with microarray

analysis of L. salmonis RXR knock-down lice where genes involved in fatty acid

metabolism and transport, steroid biosynthesis and genes involved din different

metabolic pathways (e.g. chitin metabolism and digestion) were regulated [129].

These studies demonstrate that ecdysteroids are associated with a large gene-

regulatory network, which illustrates the complexity involved in endocrine signalling.

1.5 Biosynthesis of ecdysteroids

Arthropods are incapable of synthesizing cholesterol de novo and are dependent on

uptake of cholesterol or alkylated plant sterols through the diet for ecdysteroid

synthesis. Ecdysteroids are polyhydroxylated steroid hormones that are synthesized

by steroidogenic enzymes classified as members of the cytochrome P450 (CYP)

family commonly known as the Halloween genes [1, 2, 130-133]. The biosynthesis of

ecdysteroid hormones takes place in specific hormone producing tissues or glands

such as the PG of insect larva [134], the ovarian follicle cells of adult insects and the

Y-organ (YO) in decapods crustaceans [135]. The ecdysteroids are then secreted into

the circulatory system and transported to the peripheral tissue where conversion into

the active metabolite by shade takes place. No tissue like the Y-organ has so far been

identified in microcrustaceans such as the copepods.

29

The biosynthetic pathway of ecdysteroids is complex and can according to Mykles et

al., [1] be divided into two stages: (1) conversion of cholesterol to 5β-diketol and (2)

hydroxylation of 5β-diketol to the secreted steroid (Fig. 5.). The first part of the

biosynthetic pathway where cholesterol is converted into 5β-diketol by the 7,8-

dehydrogenase neverland is similar in insects and crustaceans but the second part is

more complex in crustaceans as they produce a broader range of ecdysteroids [136-

140]. The enzymatic conversion of 5β-diketol to the active metabolite is performed

by the Halloween genes phantom (CYP306A1), disembodied (CYP302A1), shadow

(CYP315A1) and shade (CYP314A1) where each gene is believed to perform one

specific hydroxylation as mutations in these genes have resulted in low ecdysteroid

levels, abnormalities in cuticula formation and embryonic death [130, 141-143].

Additional enzymes are contributing to the biosynthetic steps called “the Black box”

which are a series of hypothetical reactions that results in the conversion of 7-

dehydrocholesterol to ketodiol. The precise intermediates in these steps are currently

unknown, due to their chemical instability, however, the four enzymes

CYP307A1/spook (spo) [144], CYP307A2/spookier (spok) [144, 145], CYP6T3

[146] and non-molting glossy/shroud (nm-g/sro) [147] have been characterized and

are considered to act during these steps. In decapod crustaceans, it has been suggested

that the biosynthetic pathway has branching points at the conversion of diketol and

ketodiol (see [1] for extensive description) resulting in four final ecdysteroid products

that are converted to either 3-dehydro-20-hydroxyecdysone, 20E or PonA by shade

dependent on the precursor steroid. For simplicity, only one pathway is presented in

Fig. 5.

Although the biosynthesis of ecdysteroids have mainly been investigated in insects

and to some degree in decapod crustaceans, orthologs of the Halloween genes have

been identified in microcrustaceans such as the branchiopod water flea Daphnia

magna [148], the copepod Calanus finmarchicus [149] and the salmon lice Caligus

rogercresseyi [150] and Lepeophtheirus salmonis (present study).

30

Octopamine receptor

It has been suggested that ecdysteroids are capable of interacting with G protein-

coupled receptors (GPCRs) thereby activating a broad range of signalling pathways.

The GPCRs is recognized by a structural motif containing seven transmembrane

domains that show considerable diversity in their sequences [151]. One such GPCR

is the octopamine receptor (OctR), which is known to bind the biogenic amids

octopamine and tyramine, which acts as neurotransmitters, neuromodulators and

neurohormones in both vertebrates and invertebrates (reviewed in [152]). The OctR

mediates attenuation of adenylyl cyclase, which induce responses of secondary

messengers such as cyclic nucleotides (cAMP, cGMP), calcium ions (Ca2+) and

inositol-1,4,5-triphosphate (IP3) that in turn regulates the activities of enzymes and

nonenzymatic proteins in a wide variety of signalling pathways. Four different

octopamine receptors (Oamb, Oct1βR, Oct2βR, Oct3βR) have been identified in

Drosophila which all are expressed in the central nervous system but differ in their

expression pattern in the peripheral tissues [153]. The presence of several octopamine

receptors contributes to many behavioural and physiological reactions [154, 155] and

more importantly for this thesis, the biosynthesis of ecdysteroids [156].

31

Pona

ster

one A

(P

onA

)

7-de

hydr

ocho

lest

erol

ch

oles

tero

l nv

d ke

todi

ol

Bla

ck b

ox

2,25

-deo

xyec

dyso

ne

ecdy

sone

20-h

ydro

xyec

dyso

ne

(20E

)

shd

dib

keto

triol

ph

m

25-d

eoxy

ecdy

sone

sad

Nm

g/Sr

o sp

o

Fig.

5. S

impl

ified

pre

sent

atio

n of

the

ecdy

ster

oid

bios

ynth

etic

pat

hway

in

crus

tace

ans.

In

mac

rocr

usta

cean

s, al

l hy

drox

ylat

ion

step

s by

nev

erla

nd (

nvd)

, th

e bl

ack-

box

gene

s no

n-

mol

ting

glos

sy/sh

roud

and

spo

ok (

CY

P307

A1)

, pha

ntom

(ph

m; C

YP3

06A

1), d

isem

bodi

ed

(CY

P302

A1)

and

sha

dow

(C

YP3

15A

1) t

ake

plac

e in

the

mol

ting

glan

d (Y

-org

an)

with

exce

ptio

n fo

r th

e co

nver

sion

of e

cdys

one

and

25-d

eoxy

ecdy

sone

to th

e ac

tive

met

abol

ites

20-h

ydro

xyec

dyso

ne (

20E)

and

pon

aste

rone

A (

PonA

), by

sha

de (

shd;

CY

P314

A1)

whi

ch

occu

r in

the

perip

hera

l tis

sue.

The

bio

synt

hetic

pat

hway

in c

rust

acea

ns b

ranc

h po

int a

t the

conv

ersi

on o

f dik

etol

and

ket

odio

l, w

hich

giv

e ris

e to

four

diff

eren

t pat

hway

s th

at g

ener

ate

then

act

ive

met

abol

ites.

The

read

er i

s ad

vise

d to

rev

iew

[1]

for

a m

ore

com

preh

ensi

ve

desc

riptio

n. M

odifi

ed fr

om [2

].

32

1.6 RNA interference as an experimental tool

RNA interference (RNAi) is a natural biological process in which small RNA

molecules inhibit gene expression. It was first described by Fire et al., [157] in the

nematode worm Caenorhabditis elegans and has since then been applied as an

experimental tool to study the gene function in cell cultures and in vivo in different

organisms. This is achieved by introducing double-stranded RNA (dsRNA) into an

organism to manipulate gene expression. In this process, the complementary strand of

the dsRNA becomes part of the RNA-induced silencing complex (RISC) a multi-

protein complex that identifies the corresponding target mRNA and cleaves it at a

specific site. Next, the cleaved mRNA is targeted for degradation, which results in the

loss of protein expression. The double-stranded RNA can be introduced in several

ways such as RNAi vectors, soaking, through food or injection [157, 158].

Since the discovery of RNAi as an analytical tool, a plethora of studies using dsRNA

for gene knock-down has been applied in a variety of metazoan species to study the

functionality of genes essential in development, growth and reproduction [50, 159-

161]. This includes genes involved in ecdysteroidogenesis and ecdysteroid signal

transduction such as the EcR and RXR genes [123, 129, 162-164].

Most RNAi studies in crustaceans have been performed on commercially important

decapods but have very recently been employed to microcrustaceans such as the

branchiopod Daphnia pulex [165, 166] and the copepod Lepeophtheirus salmonis

[25, 167]. In the L. salmonis, protocols have been developed in order to perform

functional studies both in larval [167] and pre-adult stages [25]. In larval stages, gene

silencing is achieved by soaking nauplius I larva in dsRNA during their molt into the

nauplius II stage. The experiments are normally terminated when the control animals

reach the copepodid stage and potential phenotypes can be determined. Pre-adult

animals, on the other hand, are injected with dsRNA in the CT and put on salmon for

approximately 35 - 40 days. The knock-down effect has been detected two days after

both treatment methods, however, the degree of knock-down decrease during the time

33

after treatment and has shown to cease between 14 - 40 days in the adult animals

dependent on the gene [167, 168].

RNAi provides an efficient tool to functionally assess genes within a genome and

evaluate their role in signalling pathways or physiological processes. Dependent on

the target gene, phenotypic traits be a direct effect of gene knock-down [25] however,

it is important to keep in mind that phenotypes can be caused by indirect effects of

decreased gene expression as reduced expression of some genes can affect many

molecular processes. Moreover, it is crucial to distinguish between gene knock-down

where gene expression is reduced as opposed to gene knock-out where gene

expression is eliminated.

34

2. Aims of the study

The overall objective of this research was to gain knowledge of the endocrine system

in the salmon louse on a molecular level focusing on the ecdysone receptor (EcR).

Ecdysone signalling through the EcR/RXR nuclear complex is well known to play

vital roles in a multitude of biological processes in all arthropod species. Interference

of the ecdysone signalling pathway is associated with molting arrest, embryonic death

and disruption of reproductive processes. Therefore, it is of great interest to gain

knowledge of the ecdysone signalling pathway in L. salmonis. Using RNA

interference (RNAi) techniques we can get an in-depth understanding of the

functionality of genes involved in the pathway, which in the future can be used in

parasite control. The specific objectives for the present study were:

• To characterize the LsEcR gene, describe the transcript expression pattern and

study its functional role in reproduction using RNAi in adult female lice

• To study the transcript pattern and function of the EcR during molting and

development in salmon louse larvae through knock-down studies

• To investigate the temporal expression pattern of the ecdysteroids: ecdysone,

20-hydroxyecdysone and ponasterone A during molting and oocyte maturing

using LC/MS/MS

• To identify genes involved in the biosynthesis of ecdysteroids and investigate

their function during early developmental stages

35

List of publications

Paper I: Liv Sandlund, Frank Nilsen, Rune Male, Sindre Grotmol, Heidi Kongshaug

and Sussie Dalvin (2015). Molecular characterisation of the salmon louse,

Lepeophtheirus salmonis salmonis (Krøyer, 1837), ecdysone receptor with emphasis

on functional studies of female reproduction. International Journal for Parasitology,

45:175-185

Paper II: Liv Sandlund, Frank Nilsen, Rune Male and Sussie Dalvin (2016). The

Ecdysone Receptor (EcR) is a Major Regulator of Tissue Development and Growth

in the Marine Salmonid Ectoparasite, Lepeophtheirus salmonis (Copepoda,

Caligidae). Molecular and Biochemical Parasitology

Accepted for publication after minor reviews

Paper III: Liv Sandlund, Rune Male, Tor Einar Horsberg, Frank Nilsen, Heidi

Kongshaug and Sussie Dalvin (2016). Identification and functional assessment of the

ecdysone biosynthetic genes neverland, disembodied and shade in the salmon louse

Lepeophtheirus salmonis (Copepoda, Caligidae).

(Manuscript)

36

3. Abstract of papers

3.1 Paper I:

The salmon louse Lepeophtheirus salmonis (Copepoda, Caligidae) is an important

parasite in the salmon farming industry in the Northern Hemisphere causing annual

losses of hundreds of million US dollars worldwide. To facilitate development of a

vaccine or other novel control measures to gain control of the parasite, knowledge

about molecular biological functions of L. salmonis is vital. In arthropods, a nuclear

receptor complex consisting of the ecdysone receptor (EcR) and the retinoid X

receptor, ultraspiracle (USP) are well known to be involved in a variety of both

developmental and reproductive processes. To investigate the role of the ecdysone

receptor in the salmon louse, we isolated and characterized cDNA with the

5´untranslated region of the predicted L. salmonis EcR (LsEcR). The LsEcR cDNA

was 1608 bp encoding a 536 aa sequence that demonstrated high sequence

similarities level to other arthropod EcRs including Tribolium castaneum and Locusta

migratoria. Moreover, in situ analysis of adult female louse revealed LsEcR transcript

to be localized in a wide variety of tissues such as ovaries, sub cuticula and oocytes.

Knock down studies of LsEcR, using RNA interference, terminated egg production

indicating that the LsEcR plays important roles in reproduction and oocyte

maturation. This is the first report of on the ecdysone receptor in the economically

important parasite L. salmonis.

3.2 Paper II:

The function of the ecdysone receptor (EcR) during development and molting has

been thoroughly investigated in in some arthropods such as insects but rarely in

crustacean copepods such as the salmon louse Lepeophtheirus salmonis (L. salmonis)

(Copepoda, Caligidae). The salmon louse is an ectoparasite on Atlantic salmon that

cause major economical expenses in aquaculture due to the cost of medical treatment

37

methods to remove lice from the fish. Handling of salmon louse infestations is further

complicated by development of resistance towards available medicines.

Understanding of basic molecular biological processes in the salmon louse is

essential to enable development of new tools to control the parasite. In this study, we

found L. salmonis EcR (LsEcR) transcript to be present in the neuronal somata of the

brain, nuclei of muscle fibers and the immature intestine. Furthermore, we explored

the function of LsEcR during development using RNA interference mediated knock-

down and through infection trials. Our results show that knock-down of LsEcR is

associated with hypotrophy of several tissues, delayed development and mortality. In

addition, combined knock-down of LsEcR/LsRXR resulted in molting arrest during

early larval stages.

3.3 Paper III:

The salmon louse is a marine ectoparasitic copepod on salmonis fishes. Its lifecycle

consists of eight developmental stages, each separated by a molt. In crustaceans and

insects, molting and reproduction is controlled by circulating steroid hormones such

as 20-hydroxyecdysone (20E). Steroid hormones are synthesized from cholesterol

through catalytic reactions involving a 7,8-dehydrogenase neverland and several

cytochrome P450 genes collectively called the Halloween genes. In this study, we

have isolated and identified orthologs of neverland (nvd), disembodied (dib) and

shade (shd) in the salmon louse L. salmonis genome. Tissue-specific expression

analysis showed that the genes are expressed in intestine and reproductive tissue.

Furthermore, knock-down studies using RNA interference in adult females showed

that only shd terminates molting in larval stages. However, knock-down of nvd

affected development of the ovaries and oocyte maturation. In addition, we

performed knock-down studies of an ortholog of the Drosophila octopamine receptor

(Oct3βR) a regulator of the Halloween genes, to determine its role during early

development. Depletion of the Oct3βR ortholog in L. salmonis resulted in molting

arrest, but did not down-regulate expression of all of the identified Halloween genes.

38

Our results show that ecdysone biosynthetic genes are present in L. salmonis and that

ecdysteroids is necessary for both molting and reproduction in the lice.

“The published papers are reprinted with permission from Elsevier. All rights reserved.”

39

4. General Discussion

Steroid hormones have an essential role in regulating biological processes within all

animals. In arthropods, one of the main steroid hormones is ecdysone (with its

metabolic variants). Ecdysteroid signalling is crucial in arthropod physiology

regulating a wide diversity of biological processes. Biologically active ecdysteroid

hormones are synthesized from cholesterol via several enzymatic steps before it

transducing its signal by binding to the EcR/RXR nuclear receptor complex. A wide

range of studies have shown the importance of EcR/RXR signalling in hexapod

insects but for many groups of arthropods like the copepods, very limited information

exist about this basic biological system. In insects, it has been demonstrated that the

ecdysteroid pathways has a key role in developmental transitions [123, 169-171] and

reproduction [115, 119, 120, 124, 172, 173]. From arthropods other than hexapods

the literature is more limited but some studies have been conducted in decapods,

ticks, daphnia and copepods [110, 148, 174, 175] (present study).

In model organisms like Drosophila, detailed information exists about ecdysteroid

function through a wide range of studies. This includes knock-out studies where it has

been demonstrated that mutation and depletion of EcR are associated with embryonic

lethality and disruption of reproductive processes. Hence, uncovering the functions of

EcR as well as regulation of ecdysone production is an important step towards

understanding basic physiological processes in the salmon louse. Emphasis is given

to the function of the EcR during molting, development and reproduction through a

range of experiments and functional studies (see paper I/II). To further understand

ecdysteroid function, key genes involved in ecdysteroid biosynthesis and regulation

(octopamine receptor) have been identified and functionally assessed (paper III).

40

4.1 Identification of ecdysteroidogenic genes and the ecdysone receptor

4.1.1 Characterization of the ecdysone biosynthetic genes: neverland, disembodied and shade in L. salmonis

Ecdysteroids are synthesized through several enzymatic steps performed by

dehydrogenases and members of the cytochrome P450 family proteins coded by the

so-called Halloween genes. In paper III, orthologs of the 7,8 dehydrogenase

neverland (nvd) and transcripts from the Halloween genes disembodied (dib) and

shade (shd) were cloned and sequenced from the L. salmonis genome. The amino

acid (aa) sequence of all three enzymes showed low sequence identity to their

orthologs, which is commonly seen in the CYP family. Only 33 – 40 % sequence

identity is found between lepidopteran and dipterian Halloween gene orthologs but

their functions are conserved [176]. However, it should be noted that one amino acid

substitution of essential residues could change the catalytic specificity. As mentioned

in the introduction, the ecdysteroid synthetic pathway is far more complex in

crustaceans than insects. Both dib and shd can hydroxylate several different

compounds (reviewed in [1]) indicating that the Halloween genes identified in L.

salmonis could acquire the same competence. Only one transcript encoding one

single ORF has so far been identified for each gene. However, successful sequencing

of the 5`UTR was not accomplished in this study, hence, it cannot be excluded that

several transcripts exist for the three genes.

4.1.2 Characterisation of the L. salmonis EcR

In paper I, three mRNA transcripts with highly different 5’ UTRs and alternative

splicing, but encoding only one ORF of the L. salmonis EcR was identified based on

sequencing and identity of conserved domains. Comparisons of L. salmonis EcR with

other EcRs showed primary structural conservation across phyla (Fig. 2., paper I).

Due to high degree of sequence similarities between the DBDs and to some extent the

LBDs, we can assume that the LsEcR share similar functions as its homologs. Salmon

41

louse has one copy of EcR and RXR (paper I and [129]) however, a minimum of

three different ORF isoforms was identified for LsRXR.

In mammals, variations of the 5`UTR of genes are relatively common and can either

be expressed using different promoters [177, 178] or by alternative splicing within

the UTR [179]. In L. salmonis it is evident that different levels of the EcR mRNA

variants are present at different life stages. This suggests that EcR transcription is

regulated by different promoters and/or alternative splicing during lice ontogeny.

This could allow for rapid changes in gene expression in response to different stimuli

e.g. during development and cell differentiation in a spatial and temporal manner

(reviewed in [180-182]. However, in order to verify this, targeted studies

investigating promoter function should be undertaken.

4.2 Functional assessment of LsEcR during molting and

development

Developmental processes from embryogenesis to life stage transitions are under the

influence of ecdysteroids in arthropods. In order to gain a deeper understanding of

ecdysteroid signalling during molting of the salmon louse, attempts were made to

measure the level of the three main steroid hormones known to be present in

crustaceans: E, 20E and PonA, during the molting cycle of the copepod stage (paper

III). This was challenging due to the combination of small sample sizes of the larval

stages and technical difficulties with the LC/MS/MS. However, from the preliminary

experiments, it was established that ecdysteroids are key regulators of molting in L.

salmonis. PonA was present at higher levels compared to the other measured

ecdysteroids in the pre-molting stage, which is in accordance with results from the

shore crab Carcinus maenas [183]. Although we have not been able to successfully

perform time specific measurements of ecdysteroid levels throughout the molting

cycle in copepodids, the obtained data gives information of the ecdysteroid content in

L. salmonis and provides valuable information to our understanding of the

ecdysteroid regulatory system in the salmon louse.

42

Ecdysteroids exert their effect through the EcR/RXR receptor complex and reduced

expression of EcR during early life stages is associated with molting defects and

lethality [123, 169, 184, 185]. Surprisingly, RNAi knock-down of LsEcR in nauplia I

larva did not cause immediate molting arrest, but resulted in viable copepodids

identical to the control group even though significant knock-down of LsEcR was

achieved (paper II). No deviation in swimming behaviour or histological aberrations

(in sections) was detected in the LsEcR knock-down (LsEcRkd) copepodids.

However, it should be noted that detection of LsEcR protein in the salmon louse has

not been successful in this study and, therefore, we cannot rule out the possibility that

residual protein sufficient to mediate ecdysone response is still present. Interestingly,

similar results were observed in LsRXR knock-down larvae [167]. Although the

ecdysone receptor is normally thought to act in a heterodimer with RXR, ligand

binding without RXR has been reported [186]. These findings indicate that both

members of the EcR/RXR complex can act as a receptor and function under certain

conditions without its partner in other species. The same situation may apply in the

salmon louse.

However, further incubation of LsEcR knock-down animals (on fish) resulted in high

mortality and the surviving lice exhibited severe tissue damage in their pre-adult

stage (paper II). During Drosophila metamorphosis, extensive neuronal remodelling

by pruning and regrowth of axons and neuronal cell death are essential in order to

establish the fully mature brain architecture and connectivity between motor neurons

and muscles necessary for muscle growth [187, 188]. Ecdysteroids play a key role in

the regulation of neuronal remodelling through the EcR/USP receptor complex [189]

hence, it is reasonable to assume that the severe degeneration of both neuronal and

muscle tissue found in the LsEcRkd lice, is caused by silencing of LsEcR. In a recent

study, Eichner et al., [129] showed that both an ortholog of advillin, a Ca2+-regulated

actin binding protein important in nervous system development and motor neuron

protein precursors involved in regulating motor neuron differentiation and survival

was down-regulated in LsRXR knock-down lice. In addition, genes regulating muscle

growth such as tropomyosin-2, twitchin and myosin heavy chain were down-regulated

43

accordingly, indicating that also muscle development is regulated, either directly or

indirectly, by the ecdysteroid pathway in L. salmonis.

Programmed cell death (PCD) has shown to be involved in both neuronal remodelling

and organogenesis in insects in response to ecdysone [116, 119, 190]. RNAi studies

in T. castaneum showed that EcRA plays a critical role in the 20E regulation of

midgut remodelling through the EcR/RXR complex [191]. Midgut reorganisation is

required in holometabolous insects and amphibians due to dietary changes between

larval and adult stages [192]. The salmon louse shift from using yolk as the only

energy source in the free-living stages to digesting host mucosa, skin and blood when

they infect the host and initiate the parasitic phase of the life cycle. It is highly likely

that differentiation of intestinal tissue occurs in order to adapt to the new diet and it is

possible that ecdysteroids have a significant role in this process. The midgut

epithelium in Drosophila and T. castaneum is replaced during metamorphosis by

PCD and activation of caspases is triggered by induction of BR-C and E93 through

ecdysone activation [190]. Eichner et al., [129] found that apoptosis regulating factors

(inhibitor of apoptosis 2 protein) are down-regulated after knock-down of LsRXR.

Considering that EcR and RXR to a large extent govern the same pathways, it is not

unlikely that LsEcR knock-down could influence midgut development (e.g. through

PCD) explaining the large developmental abnormalities found in pre-adults when

LsEcR was silenced (paper II). However, to confirm if abnormal PCD activity

creates these developmental aberrations further studies need to be performed (e.g.

assess expression levels of caspases and other key genes important in PCD in

LsEcRkd lice).

We observed a striking difference in phenotype between individual knock-down of

LsEcR and LsRXR compared to the combined knock-down of the receptors. The

double knock-down animals (LsEcR/LsRXR) failed to enter the copepodid stage

whereas single knock-downs successfully molted into apparently healthy infectious

copepodids (paper II). In order to molt, arthropods are strictly dependent on the

ability to remodel chitinous structures and malfunction in chitin metabolism leads to

developmental disorders. Ecdysteroids have shown to be both positive and negative

44

regulators of chitin synthesis [193]. In Drosophila, it is indicated that ecdysone

through the activation of EcR/RXR has a direct regulatory role of the chitin synthase

genes as EcREs are found in their promoter region [194]. The promoter region for the

two chitin synthases present in L. salmonis has not been examined for EcREs.

However, down-regulation of sub-cuticular LsCHS2 in both LsEcR and LsEcR/LsRXR

knock-down lice strongly suggests that they are regulated by the ecdysteroid pathway

through the EcR/RXR complex (paper II). RNAi experiments in Drosophila embryo

showed that the CHS1 gene krotzkopf-verkehrt (kkv) is essential for maintaining the

structure of procuticula and stabilisation of the epicuticula as well as epidermal

morphology. Sclerotization and melanization were additionally impaired in these

animals, suggesting that the activity of chitin synthases regulate several enzymes in

chitin metabolism [195]. It is indeed possible that the molting arrest of

LsEcRkd/LsRXRkd animals is caused by disruption of chitin metabolism.

Furthermore, even though no visible phenotype was observed in the LsEcRkd animals

in the copepodid stage, several enzymes important in chitin metabolism (LsCP1,

LsCHS2, LsChs1, LsChs2) were affected which could cause discrepancies in the

molting process. This could account for the high mortality observed for the LsEcRkd

lice during the infection trial (paper II).

4.3 Functional assessment of LsEcR during reproduction

Ecdysteroid signalling is essential for reproduction in L. salmonis. Transcript knock-

down of the ecdysone biosynthetic enzyme LsNvd (paper III) as well as the

transcription factor LsEcR (paper I) inhibits the development of reproductive tissues,

but more severely for the latter where egg production is completely abolished and

females do not extrude any eggs.

In D. magna, ovarian ecdysteroids are transferred into the oocytes as free

ecdysteroids or as polar (mostly phosphate esters) or apolar (e.g. long chained fatty

acid esters) ecdysteroid conjugates. It is suggested that the purpose of the conjugates

is to act as inactive storage forms of maternally derived hormones that can be

45

hydrolyzed into active ecdysteroids during embryogenesis [125]. Transcripts of both

LsNvd and very low levels of the two Halloween genes are found in the ovaries (In-

situ hybridization and Licebase.org; unpublished) of L. salmonis (paper III)

suggesting that either complete and/or partial biosynthesis of ecdysteroids take place

in the reproductive tissue. This is supported by the observed irregularities of the

ovaries and follicular epithelium lining the oocytes in the adult female lice when

LsNvd transcripts are significantly reduced (paper III). A complex network of

signalling events acts to establish the lining of the follicular cell layer of the oocyte

during maturation. In both Drosophila and T. castaneum, depletion of EcR disrupts

development of the follicular cell layer necessary for oocyte maturation and loss of

vitellogenic stages thereby preventing embryogenesis. Blocking of EcR signalling in

follicular cells prevents proper organisation of the oocyte membrane presumably

causing anomalies in the actin cytoskeleton in the microvilli [196]. Microvilli are

important in the assembly of the vitelline membrane [197] which stored information

of embryo patterning in Drosophila. Disruption of the vitelline membrane aborts

embryonic development due to loss of eggshell function resulting in sterile females

[198]. The only invertebrate where this is described in detail is for Drosophila and

although the evolutionary distance to L. salmonis is large it is tempting to speculate

that the lack of normal egg chamber generation and egg string formation in adult

female LsEcRkd lice (paper I) is caused by aberrations in the cytoskeleton. This

speculation is further supported by up-regulation of actin depolymerisation factors in

LsRXRkd lice [129] that indicates impaired cytoskeleton function by increased

depolymerisation. Dysfunction of the cytoskeletal components interferes with a

cascade of events such as cell differentiation, vesicle/organelle trafficking and

synaptic signalling. Based on this, it is possible that the extensive tissue damage

observed in LsEcRkd animals in the infection trial (paper II) is linked to alterations

in the actin filaments. In contrast to knock-down of LsEcR, knock-down studies of

total LsRXR transcript in adult females showed that the females were able to generate

and protrude egg strings, which either did not hatch or produced offspring that was

not viable [129]. Even though LsEcR and LsRXR transcripts locate to the same tissue

and the same degree of knock-down is achieved (~ 60 %), it appears that the absence

46

of LsEcR results in larger consequences for the reproductive processes than the

absence of LsRXR. Histological assessment of the genital segment of both LsEcR and

LsRXR knock-down lice shows the presence of individual oocytes in the LsRXRkd

lice, whereas the oocytes are completely disintegrated in the LsEcRkd lice. As

mentioned earlier, the presence of residual LsRXR protein or homo/

heterodimerization with an alternative partner has to be taken into account.

The last molt of the L. salmonis life cycle occurs between the pre-adult II and adult

stage hence possible variations in ecdysteroid level in the adult female lice is

presumably related to reproductive processes. Since ecdysteroids are key regulators

of arthropod reproduction, investigation of the ecdysteroid level in adult female lice

was performed. In Paper III we demonstrated that the ecdysteroid level is

significantly different between the CT and the Ab/G segment of gravid adult females.

One explanation for this is that ecdysteroids in CT are important for oocyte

production and maturation as well as yolk production while the high levels in the

genital segments could serve as a source for maternally provided ecdysteroids.

Presence of LsNvd, LsDib and LsShd mRNA transcripts in unfertilised oocytes

suggests that the oocytes are capable of de novo synthesis from cholesterol,

supporting the high level of ecdysteroids present at the end of L. salmonis oocyte

maturation in. The rise in the ecdysteroid level confirms previous reports from other

crustaceans where ecdysteroids are associated with promotion of the ovary [125, 175,

199]. In adult Drosophila, ecdysteroids stimulate both proliferation and maintenance

of germline stem cells through EcR signalling. It has also been shown that ecdysone

signalling is required in follicle formation in somatic cells (review in [122]), which is

the site for vitellogenin synthesis in adult insects. In L. salmonis, yolk production

takes place in the sub-cuticular tissue. The yolk proteins are taken up by oocytes in

the genital segment [25, 50] and are either directly or indirectly dependent on EcR-

transduced ecdysteroid signalling to occur (paper I). Knock-down of LsRXR had a

similar effect on yolk protein production [129]. Since the salmon louse is

lecitotrophic, the eggs must be supplemented with sufficient nutrition that ensures

proper embryogenesis and development until the parasitic copepodid can take up

47

external food, which occurs after successful host settlement. Energy conservation by

incorporation of nutrients (i.e. aa, lipids, carbohydrates, proteins and hormonal

metabolites) into the maturing oocyte is, therefore, beneficial in order to prolong the

free-living lifetime of the lice. The presence of ecdysteroids in the oocytes could also

explain why only LsShd gives a significant phenotype upon larval knock-down

whereas LsNvd and LsDib gave no phenotype (paper III: see below).

4.4 Biosynthesis of ecdysteroids in L. salmonis

Several enzymes are involved in the biosynthesis of ecdysteroids. In this study, three

enzymes neverland, disembodied and shade were investigated that represents three

parts of the biosynthetic pathway; the initial, middle and the last steps (paper III).

The data obtained give us insight into the localisation and function of the enzymes

during specific physiological processes. However, it should be mentioned that no

functional enzyme assay was conducted in the present study.

4.4.1 Functional assessment of ecdysteroid biosynthetic enzymes in L. salmonis early development

In other species, knock-down of Nvd, Dib and Shd are associated with embryonic

defects resulting in mortality [130, 141]. Functional assessment of the genes in L.

salmonis indicates that, of the investigated ecdysteroidigenic genes, only the shade

gene product is necessary for development during the lecitotrophic stages. The results

obtained (paper III) indicate that the regulation of ecdysteroid synthesis in L.

salmonis might differ from that described in previous literature.

As mentioned, the salmon louse free-living larvae rely solely on yolk nutrition that is

deposited into the eggs until it becomes parasitic during the copepodid stage. Nauplia

I larvae with significant knock-down of LsNvd and LsDib molted through the

nauplius II stage and into the copepodid stage and did not show any morphological

aberrations. However, depletion of LsShd in nauplia I resulted in immediate molting

arrest. We hypothesize that sufficient ecdysone is synthesized in the maturing oocytes

and are stored as part of the food package. The ecdysone reserves are then converted

48

by LsShd into the active metabolite that is used by the LsEcR/LsRXR complex

during embryogenesis and development from the free-swimming to the parasitic

stage. The hypothesis is supported by the increase in ecdysteroid level observed at the

end of oocyte maturation (paper III) prior to the extrusion of the eggs and initiation

of embryogenesis. In addition, preliminary RNAi knock-down studies of the L.

salmonis shadow (LsSad) (CYP315A1) ortholog showed the same results as those

obtained by knock-down LsNvd and LsDib (data not shown). To the best of our

knowledge, the no-loss of function phenotype observed in the LsNvd, LsDib and

LsSad, has not been previously reported in arthropods and shows a novel regulation

pathway that is most likely an adaptation to the lecitotrophic life stages of the parasite

larvae. Whether this regulatory mechanism is unique to L. salmonis or similar in

other lecitotrophic parasite larvae remains a topic for future studies.

4.4.2 Functional assessment of L. salmonis Oct3ββR during early development

Octopamine receptors are known to modulate an extensive variety of physiological

processes including learning and memory responses, neuronal development [154],

ovulation [200] and ecdysteroidogenesis [156]. We searched the salmon louse

genome using the Drosophila melanogaster Oct3βR (DmOct3βR) sequence and

identified a putative ortholog sequence (paper III). RNAi knock-down of LsOct3βR

gene in nauplia I resulted in immediate molting arrest resembling the phenotype

observed for the LsShdkd animals.

Recent studies of DmOct3βR showed that depletion of the gene negatively regulated

the ecdysone biosynthetic genes present in the PG [156]. Based on this study, we

performed RT-qPCR on the identified biosynthetic genes LsNvd, LsDib and LsShd

and the LsEcR and LsRXR nuclear receptors. Surprisingly, in the LsOct3βR RNAi

animals, LsShd transcripts were significantly up-regulated while LsDib remained

unregulated. These results contradict the observations from Drosophila, where dib

was significantly down-regulated [156]. LsNvd was however regulated in a similar

49

manner in both L. salmonis and Drosophila. The findings in this study indicate that

the ecdysteroid biosynthetic pathway in L. salmonis is different compared to that of

insects, which is in agreement with observations from other crustaceans (review in

[1]). Interestingly, the LsOct3βR appears to only regulate LsEcR and not its partner

LsRXR, which remained unaffected by the LsOct3βR knock-down in the salmon

louse. Originally, EcR and RXR are thought to act as heterodimeric partners.

However, this study (Paper I, [129]) indicate that they are regulated in different ways

dependent on the physiological/biological conditions. It is evident from the present

study that the LsOct3βR is necessary for proper ecdysteroidogenesis and ecdysone

signalling in L. salmonis.

4.5 The ecdysone pathway and lice control

From the aspect of applied biology, knowledge of the ecdysone receptor and the

ecdysone biosynthetic enzymes may contribute to the development of novel treatment

methods against the salmon louse. In insects, ecdysteroid-controlled mechanisms like

molting and growth has been used as targets for development of pesticides with low

environmental impact and vertebrate toxicity, therefore, there is a potential for novel

medicines targeting the ecdysone hormone system in L. salmonis. The best prospect

in the area is to exploit the susceptibility of the ecdysone receptor using

antagonists/agonists, administered either by oral or bath treatment, that is specific to

the LsEcR. However, this requires investigation of the shape and structure of ligand-

binding pocket of the LsEcR. Teflubenzuron and diflubenzuron are today used as an

anti-lice medicine and targets the molting process, by interfering with the chitin

synthesis. These chemicals are species unspecific and have shown to have a negative

impact on other arthropods such as shrimp, lobster and copepods in the environment

[201]. In addition, teflubenzuron and diflubenzuron only work on molting stages

hence adult lice and egg strings are not affected. LsEcR specific of

antagonists/agonists would reduce the probability of influencing other important

marine arthropods in addition to targeting all life stages including reproductive

processes.

50

Members of the P450 family are present in both vertebrates and invertebrates and

even though their P450 enzyme systems differ, some xenobiotics (e.g. phenobarbital)

can induce metabolism in both groups (reviewed in [202]). The Halloween genes are

specific to arthropods but are related to vertebrate CYPs. Hence, it could be

conflicting to target these enzymes in the salmon louse as the xenobiotics or their

metabolites (formed through biotransformation) could have a negative effect on the

salmon and other vertebrates.

51

5. Concluding remarks and future perspectives

The present study has extended our knowledge of EcR-mediated ecdysone signalling

in the salmon louse Lepeophtheirus salmonis with respect to its key role in the

regulation of developmental and reproductive processes.

In papers I-II, we identified and functionally assessed the LsEcR during early

developmental and reproductive stages. From the results, it is concluded that the

LsEcR play key roles during oocyte maturation and reproduction in adult female

louse and loss of LsEcR function terminates egg production. Depletion of LsEcR did

however, not cause molting arrest between nauplia II and the copepodid stage but

resulted in high mortality and severe tissue damage in later stages of development,

showing that EcR is essential in growth and survival of the parasitic life stages.

Future studies should investigate the interaction between ecdysone biosynthesis and

signalling and other potential hormonal regulators of salmon louse development.

Future studies of the LsEcR should aim to determine potential subcellular localisation

of the protein outside the nucleus. Detection of protein would also be beneficial in

order to determine the half-life of protein after gene knock-down and the presence of

residue protein. To get a better understanding of EcR-mediated ecdysone signalling in

L. salmonis, protein-protein and protein-ligand interactions should also be

investigated. This work has been initiated by master students at the Sea Lice Research

Centre (SLRC), using a mammalian two-hybrid luciferase system. Although this is an

artificial in vitro system it will undoubtedly give information of the L. salmonis

EcR/RXR dimerization and ligand binding properties. Alternative partners for LsEcR

(and LsRXR) should be investigated to give information about possible alternative

regulation of ecdysone signalling pathways. Moreover, possible LsEcR

agonist/antagonist can be tested for future parasite control.

From the functional studies of the biosynthetic genes LsNvd, LsDib and LsShd in

nauplia larvae (paper III), only reduction of LsShd transcripts caused developmental

defects. This lead us to hypothesize that precursors of the active steroid hormones

52

20E and PonA is synthesized and stored in the maturing oocyte for utilization during

embryogenesis and development of the lecitotrophic stages until entry into the

parasitic stage. In order to verify our hypothesis, it would be necessary to measure the

levels of the metabolites (e.g. cholesterol and diketol) converted by LsNvd and LsDib

during embryogenesis (i.e. from the point of fertilization to hatching of the egg

string) as well as the ecdysone level. In addition, infection trials using LsNvd and

LsDib knock-down animals would be beneficial to determine the phenotype and

viability of the lice in the parasitic stages when these genes are decreased. At present

time, only the three genes involved in ecdysteroidogenesis of L. salmonis have been

partially investigated. It is apparent that other enzymes of this pathway have to be

functionally assessed in order to gain a better understanding of the ecdysone

biosynthetic pathway in the salmon louse.

Biogenic amines such as octopamine and tyramine have shown to play a role in the

upstream regulation of ecdysteroidogenesis in insects through GPCRs. Recent studies

by Ohhara et al., [156] showed that ecdysone biosynthesis is regulated by the GPCR

Oct3βR, in Drosophila. From our LsOct3βR knock-down studies, it is apparent that

the octopamine receptor also play a role in the regulation of ecdysoteroid biosynthesis

in L. salmonis. Therefore, further studies of the LsOct3βR should, first and foremost,

include proper identification and sequencing analysis as well as identification of

possible paralogs present in the salmon louse genome. It would be beneficial to

perform functional studies of LsOct3βR and/or possible paralogs using RNAi in adult

female louse in addition to the RNAi in the nauplia stage. The octopamine receptor is

well known to be a target for insecticides (e.g. amitraz), hence, the LsOct3βR is of

great interest as a target for future salmon louse control.

53

References

1. Mykles, D.L., Ecdysteroid metabolism in crustaceans. Journal of Steroid Biochemistry and Molecular Biology, 2011. 127: p. 8.

2. Warren, J.T., et al., Molecular and biochemical characterization of two P450

enzymes in the ecdysteroidogenic pathway of Drosophila melanogaster. Proceedings of the National Academy of Sciences of the United States of America, 2002. 99(17): p. 11043-11048.

3. Perissi, V. and M.G. Rosenfeld, Controlling nuclear receptors: The circular logic of

cofactor cycles. Nature Reviews Molecular Cell Biology, 2005. 6(7): p. 542-554. 4. Fallang, A., et al., Evidence for occurrence of an organophosphate-resistant type of

acetylcholinesterase in strains of sea lice (Lepeophtheirus salmonis Kroyer). Pest Management Science, 2004. 60(12): p. 1163-1170.

5. Kaur, K., et al., Mechanism behind Resistance against the Organophosphate

Azamethiphos in Salmon Lice (Lepeophtheirus salmonis). Plos One, 2015. 10(4). 6. Jones, M.W., C. Sommerville, and R. Wootten, Reduced Sensitivity of the Salmon

Louse, Lepeophtheirus-Salmonis, to the Organophosphate Dichlorvos. Journal of Fish Diseases, 1992. 15(2): p. 197-202.

7. Espedal, P.G., et al., Emamectin benzoate resistance and fitness in laboratory reared

salmon lice (Lepeophtheirus salmonis). Aquaculture, 2013. 416: p. 111-118. 8. Grøntvedt, R., et al., The surveillance programme for resistance to

chemotherapeutants in salmon lice (Lepeophtheirus salmonis) in Norway 2014. Surveillance programmes for terrestrial and aquatic animals in Norway. 2015.

9. Samuelsen, O.B., et al., Mortality and deformities in European lobster (Homarus

gammarus) juveniles exposed to the anti-parasitic drug teflubenzuron. Aquatic Toxicology, 2014. 149: p. 8-15.

10. Treasurer, J.W., A review of potential pathogens of sea lice and the application of

cleaner fish in biological control. Pest Manag Sci, 2002. 58(6): p. 546-58. 11. Kabata, Z., Parasitic Copepoda of British Fishes. The Ray Society, London, 1979: p.

468 12. Skern-Mauritzen, R., O. Torrissen, and K.A. Glover, Pacific and Atlantic

Lepeophtheirus salmonis (Kroyer, 1838) are allopatric subspecies: Lepeophtheirus salmonis salmonis and L. salmonis oncorhynchi subspecies novo. Bmc Genetics, 2014. 15.

54

13. Pike, A.W. and S.L. Wadsworth, Sealice on salmonids: Their biology and control, in Advances in Parasitology, Vol 44, J.R. Baker, R. Muller, and D. Rollinson, Editors. 2000. p. 233-337.

14. Brandal, P.O., E. Egidius, and I. Romslo, Host Blood - Major Food Component for

Parasitic Copepod Lepeophtheirus-Salmonis Kroyeri, 1838 (Crustacea-Caligidae). Norwegian Journal of Zoology, 1976. 24(4): p. 341-343.

15. Finstad, B., et al., Laboratory and field investigations of salmon lice [Lepeophtheirus

salmonis (Kroyer)] infestation on Atlantic salmon (Salmo salar L.) post-smolts. Aquaculture Research, 2000. 31(11): p. 795-803.

16. Grimnes, A. and P.J. Jakobsen, The physiological effects of salmon lice infection on

post-smolt of Atlantic salmon. Journal of Fish Biology, 1996. 48(6): p. 1179-1194. 17. Wagner, G.N., M.D. Fast, and S.C. Johnson, Physiology and immunology of

Lepeophtheirus salmonis infections of salmonids. Trends in Parasitology, 2008. 24(4): p. 176-183.

18. Easy, R.H. and N.W. Ross, Changes in Atlantic salmon (Salmo salar) epidermal

mucus protein composition profiles following infection with sea lice (Lepeophtheirus salmonis). Comparative Biochemistry and Physiology D-Genomics & Proteomics, 2009. 4(3): p. 159-167.

19. Nylund, et al., The possible role of Lepeophtheirus salmonis (Krøyer) in the transmission of infectious salmon anaemia, in Pathogens of wild and farmed fish: sea lice, G. Boxshall and D. Defaye, Editors. 1993, Ellis Horwood Ltd.: London. p. 367-373.

20. Hamre, L.A., et al., The Salmon Louse Lepeophtheirus salmonis (Copepoda: Caligidae) Life Cycle Has Only Two Chalimus Stages. Plos One, 2013. 8(9).

21. Albright, S.C.J.a.L.J., The developmental stages of Lepeophtheirus salmonis.

Canadian Journal of Zoology, 1991. 69. 22. Schram, T.A., Supplementary descriptions of the develompemntal stages of

Lepeophtheirus salmonis (Krøyer 1837) (Copepoda: Caligidae), in Pathogens of wild and farmed fish: sea lice., G.B.D. Defaye, Editor. 1993, New York: Ellis Horwood. p. 30-47.

23. Boxaspen, K., A review of the biology and genetics of sea lice. Ices Journal of Marine

Science, 2006. 63(7): p. 1304-1316. 24. Tucker, C.S., C. Sommerville, and R. Wootten, An investigation into the larval

energetics and settlement of the sea louse, Lepeophtheirus salmonis, an ectoparasitic copepod of Atlantic salmon, Salmo salar. Fish Pathology, 2000. 35(3): p. 137-143.

25. Dalvin, S., et al., Functional characterisation of the maternal yolk-associated protein

(LsYAP) utilising systemic RNA interference in the salmon louse (Lepeophtheirus salmonis) (Crustacea: Copepoda). International Journal for Parasitology, 2009. 39(13): p. 1407-1415.

55

26. Amundrud, T.L. and A.G. Murray, Modelling sea lice dispersion under varying environmental forcing in a Scottish sea loch. Journal of Fish Diseases, 2009. 32(1): p. 27-44.

27. Salama, N.K.G., et al., Development and assessment of a biophysical dispersal model

for sea lice. Journal of Fish Diseases, 2013. 36(3): p. 323-337. 28. Bron, J., C. Sommerville, and G. Rae, Aspects of the behaviour of copepodid larvae

of the salmon louse Lepeophtheirus salmonis (Krøyer, 1837). Pathogens of wild and farmed fish: sea lice, 1993: p. 125-142.

29. Hull, M.Q., et al., Patterns of Pair Formation and Mating in an Ectoparasitic

Caligid Copepod Lepeophtheirus Salmonis (Kroyer 1837): Implications for Its Sensory and Mating Biology. Philosophical Transactions: Biological Sciences, 1998. 353(1369): p. 753-764.

30. Fields, D.M., M.J. Weissburg, and H. Browman, Chemoreception in the salmon

louse Lepeophtheirus salmonis: an electrophysiology approach. Diseases of Aquatic Organisms, 2007. 78(2): p. 161-168.

31. Bron, J.E., et al., The Settlement and Attachment of Early Stages of the Salmon

Louse, Lepeophtheirus-Salmonis (Copepoda, Caligidae) on the Salmon Host, Salmo-Salar. Journal of Zoology, 1991. 224: p. 201-212.

32. Bell, S., J.E. Bron, and C. Sommerville, The distribution of exocrine glands in

Lepeophtheirus salmonis and Caligus elongatus (Copepoda : Caligidae). Contributions to Zoology, 2000. 69(1-2): p. 9-20.

33. Eichner, C., L.A. Hamre, and F. Nilsen, Instar growth and molt increments in

Lepeophtheirus salmonis (Copepoda: Caligidae) chalimus larvae. Parasitol Int, 2015. 64(1): p. 86-96.

34. Ritchie, G., et al., Morphology and ultrastructure of the reproductive system of

Lepeophtheirus salmonis (Kroyer, 1837) (Copepoda: Caligidae). Journal of Crustacean Biology, 1996. 16(2): p. 330-346.

35. Todd, C.D., et al., Polyandry in the ectoparasitic copepod Lepeophtheirus salmonis

despite complex precopulatory and postcopulatory mate-guarding. Marine Ecology Progress Series, 2005. 303: p. 225-234.

36. Hamre, L.A., K.A. Glover, and F. Nilsen, Establishment and characterisation of

salmon louse (Lepeophtheirus salmonis (Kroyer 1837)) laboratory strains. Parasitology International, 2009. 58(4): p. 451-460.

37. Costello, M.J., Ecology of sea lice parasitic on farmed and wild fish. Trends

Parasitol, 2006. 22(10): p. 475-83. 38. Jones, S., E. Kim, and W. Bennett, Early development of resistance to the salmon

louse, Lepeophtheirus salmonis (Kroyer), in juvenile pink salmon, Oncorhynchus gorbuscha (Walbaum). Journal of Fish Diseases, 2008. 31(8): p. 591-600.

56

39. Jones, S.R.M., et al., Differential rejection of salmon lice by pink and chum salmon: disease consequences and expression of proinflammatory genes. Diseases of Aquatic Organisms, 2007. 75(3): p. 229-238.

40. Johnson, S.C. and L.J. Albright, Comparative Susceptibility and Histopathology of

the Response of Naive Atlantic, Chinook and Coho Salmon to Experimental-Infection with Lepeophtheirus-Salmonis (Copepoda, Caligidae). Diseases of Aquatic Organisms, 1992. 14(3): p. 179-193.

41. Fast, M.D., et al., Differential gene expression in Atlantic salmon infected with

Lepeophtheirus salmonis. Journal of Aquatic Animal Health, 2006. 18(2): p. 116-127.

42. Tadiso, T.M., et al., Gene expression analyses of immune responses in Atlantic

salmon during early stages of infection by salmon louse (Lepeophtheirus salmonis) revealed bi-phasic responses coinciding with the copepod-chalimus transition. Bmc Genomics, 2011. 12.

43. Braden, L.M., et al., Comparative defense-associated responses in salmon skin

elicited by the ectoparasite Lepeophtheirus salmonis. Comparative Biochemistry and Physiology D-Genomics & Proteomics, 2012. 7(2): p. 100-109.

44. Braden, L.M., B.F. Koop, and S.R.M. Jones, Signatures of resistance to

Lepeophtheirus salmonis include a T(H)2-type response at the louse-salmon interface. Developmental and Comparative Immunology, 2015. 48(1): p. 178-191.

45. Fast, M.D., et al., Skin morphology and humoral non-specific defence parameters of

mucus and plasma in rainbow trout, coho and Atlantic salmon. Comparative Biochemistry and Physiology a-Molecular and Integrative Physiology, 2002. 132(3): p. 645-657.

46. Fast, M.D., et al., Lepeophtheirus salmonis secretory/excretory products and their

effects on Atlantic salmon immune gene regulation. Parasite Immunology, 2007. 29(4): p. 179-189.

47. Fast, M.D., et al., Lepeophtheirus salmonis: characterization of prostaglandin E-2 in

secretory products of the salmon louse by RP-HPLC and mass spectrometry. Experimental Parasitology, 2004. 107(1-2): p. 5-13.

48. Fast, M.D., N.W. Ross, and S.C. Johnson, Prostaglandin E-2 modulation of gene

expression in an Atlantic salmon (Salmo salar) macrophage-like cell line (SHK-1). Developmental and Comparative Immunology, 2005. 29(11): p. 951-963.

49. Fast, M.D., et al., Enzymes released from Lepeophtheirus salmonis in response to

mucus from different salmonids. J Parasitol, 2003. 89(1): p. 7-13. 50. Dalvin, S., et al., Characterisation of two vitellogenins in the salmon louse

Lepeophtheirus salmonis: molecular, functional and evolutional analysis. Diseases of Aquatic Organisms, 2011. 94(3): p. 211-224.

57

51. Edvardsen, R.B., et al., Gene expression in five salmon louse (Lepeophtheirus salmonis, Krøyer 1837) tissues. Marine Genomics, 2014(0).

52. Nylund, A., S. Okland, and B. Bjorknes, Anatomy and Ultrastructure of the

Alimentary Canal in Lepeophtheirus-Salmonis (Copepoda, Siphonostomatoida). Journal of Crustacean Biology, 1992. 12(3): p. 423-437.

53. Kvamme, B.O., et al., Molecular characterisation of five trypsin-like peptidase

transcripts from the salmon louse (Lepeophtheirus salmonis) intestine. International Journal for Parasitology, 2004. 34(7): p. 823-832.

54. Laudet, V., Evolution of the nuclear receptor superfamily: early diversification from

an ancestral orphan receptor. J Mol Endocrinol, 1997. 19(3): p. 207-26. 55. King-Jones, K. and C.S. Thummel, Nuclear receptors - A perspective from

Drosophila. Nature Reviews Genetics, 2005. 6(4): p. 311-323. 56. Laudet, V. and H. Gronemeyer, eds. The Nuclear Receptor Factsbook. ed. V. Laudet

and H. Gronemeyer. 2002, Academic Press: London. 57. Escriva, H., S. Bertrand, and V. Laudet, The evolution of the nuclear receptor

superfamily. Essays Biochem, 2004. 40: p. 11-26. 58. Escriva, H., F. Delaunay, and V. Laudet, Ligand binding and nuclear receptor

evolution. Bioessays, 2000. 22(8): p. 717-27. 59. Escriva, H., et al., Evolution and diversification of the nuclear receptor superfamily.

Ann N Y Acad Sci, 1998. 839: p. 143-6. 60. Laudet, V., et al., Evolution of the nuclear receptor gene superfamily. EMBO J,

1992. 11(3): p. 1003-13. 61. Auwerx, J., et al., A unified nomenclature system for the nuclear receptor

superfamily. Cell, 1999. 97(2): p. 161-163. 62. Thomson, S.A., et al., Annotation, phylogenetics, and expression of the nuclear

receptors in Daphnia pulex. Bmc Genomics, 2009. 10. 63. Karlson, P., The hormonal control of insect metamorphosis. A historical review.

International Journal of Developmental Biology, 1996. 40(1): p. 93-96. 64. Ashburner M, Sequential Gene Activation by Ecdysone in Polytene Chromosomes of

Drosophila-Melanogaster .2. Effects of Inhibitors of Protein-Synthesis. Developmental Biology, 1974. 39(1): p. 141-157.

65. Koelle, M.R., et al., The Drosophila EcR Gene Encodes an Ecdysone Receptor, a

New Member of the Steroid-Receptor Superfamily. Cell, 1991. 67(1): p. 59-77.

58

66. Thomas, H.E., H.G. Stunnenberg, and A.F. Stewart, Heterodimerization of the Drosophila Ecdysone Receptor with Retinoid-X Receptor and Ultraspiracle. Nature, 1993. 362(6419): p. 471-475.

67. Yao, T.P., et al., Functional Ecdysone Receptor Is the Product of Ecr and

Ultraspiracle Genes. Nature, 1993. 366(6454): p. 476-479. 68. Laguerre, M. and J.A. Veenstra, Ecdysone receptor homologs from mollusks, leeches

and a polychaete worm. Febs Letters, 2010. 584(21): p. 4458-4462. 69. Enmark, E. and J.-Å. Gustafsson, Comparing nuclear receptors in worms, flies and

humans. Trends in Pharmacological Sciences, 2001. 22(12): p. 611-615. 70. Giguere, V., et al., Functional Domains of the Human Glucocorticoid Receptor. Cell,

1986. 46(5): p. 645-652. 71. Zhou, Z.X., et al., A ligand-dependent bipartite nuclear targeting signal in the

human androgen receptor. Requirement for the DNA-binding domain and modulation by NH2-terminal and carboxyl-terminal sequences. J Biol Chem, 1994. 269(18): p. 13115-23.

72. Mangelsdorf, D.J., et al., The nuclear receptor superfamily: the second decade. Cell,

1995. 83(6): p. 835-9. 73. Hill, R.J., et al., Ecdysone Receptors: From the Ashburner Model to Structural

Biology. Annual Review of Entomology, Vol 58, 2013. 58: p. 251-271. 74. Carmichael, J.A., et al., The X-ray structure of a hemipteran ecdysone receptor

ligand-binding domain - Comparison with a Lepidopteran ecdysone receptor ligand-binding domain and implications for insecticide design. Journal of Biological Chemistry, 2005. 280(23): p. 22258-22269.

75. Iwema, T., et al., Structural and functional characterization of a novel type of ligand-

independent RXR-USP receptor. Embo Journal, 2007. 26(16): p. 3770-3782. 76. Billas, I.M.L., et al., Structural adaptability in the ligand-binding pocket of the

ecdysone hormone receptor. Nature, 2003. 426(6962): p. 91-96. 77. Browning, C., et al., Critical role of desolvation in the binding of 20-

hydroxyecdysone to the ecdysone receptor. Journal of Biological Chemistry, 2007. 282(45): p. 32924-32934.

78. Lezzi, M., et al., Ligand-induced heterodimerization between the ligand binding

domains of the Drosophila ecdysteroid receptor and ultraspiracle. European Journal of Biochemistry, 2002. 269(13): p. 3237-3245.

79. Billas, I.M.L., Browning, C., Lawrence, M.C., Graham, L.D., Moras. D., Hill. R.J, ,

The structure and function of ecdysone receptors. . In: Smagghe, G. (Ed.), Ecdysteroids. Structures and Functions., 2009(Springer, Berlin): p. 335-375.

59

80. Schubiger, M., et al., Isoform specific control of gene activity in vivo by the Drosophila ecdysone receptor. Mechanisms of Development, 2003. 120(8): p. 909-918.

81. Ghbeish, N., et al., The dual role of ultraspiracle, the Drosophila retinoid X receptor,

in the ecdysone response. Proceedings of the National Academy of Sciences of the United States of America, 2001. 98(7): p. 3867-3872.

82. Cherbas, L., K. Lee, and P. Cherbas, Identification of Ecdysone Response Elements

by Analysis of the Drosophila Eip28/29 Gene. Genes & Development, 1991. 5(1): p. 120-131.

83. Billas, I.M., et al., Crystal structure of the ligand-binding domain of the ultraspiracle

protein USP, the ortholog of retinoid X receptors in insects. J Biol Chem, 2001. 276(10): p. 7465-74.

84. Hopkins, P.M., D. Durica, and T. Washington, RXR isoforms and endogenous

retinoids in the fiddler crab, Uca pugilator. Comparative Biochemistry and Physiology a-Molecular & Integrative Physiology, 2008. 151(4): p. 602-614.

85. Nagaraju, G.P., et al., Computational analysis of the structural basis of ligand

binding to the crustacean retinoid X receptor. Comp Biochem Physiol Part D Genomics Proteomics, 2010. 5(4): p. 317-24.

86. Riddihough, G. and H.R.B. Pelham, An Ecdysone Response Element in the

Drosophila Hsp27 Promoter. Embo Journal, 1987. 6(12): p. 3729-3734. 87. Antoniewski, C., M. Laval, and J.A. Lepesant, Structural Features Critical to the

Activity of an Ecdysone Receptor-Binding Site. Insect Biochemistry and Molecular Biology, 1993. 23(1): p. 105-114.

88. Martinez, E., F. Givel, and W. Wahli, A Common Ancestor DNA Motif for

Invertebrate and Vertebrate Hormone Response Elements. Embo Journal, 1991. 10(2): p. 263-268.

89. Maletta, M., et al., The palindromic DNA-bound USP/EcR nuclear receptor adopts

an asymmetric organization with allosteric domain positioning. Nat Commun, 2014. 5.

90. Covi, J.A., E.S. Chang, and D.L. Mykles, Neuropeptide signaling mechanisms in

crustacean and insect molting glands. Invertebrate Reproduction & Development, 2012. 56(1): p. 33-49.

91. Katayama, H., et al., The solution structure of molt-inhibiting hormone from the

Kuruma prawn Marsupenaeus japonicus. J Biol Chem, 2003. 278(11): p. 9620-3. 92. Qu, Z., et al., How Did Arthropod Sesquiterpenoids and Ecdysteroids Arise?

Comparison of Hormonal Pathway Genes in Noninsect Arthropod Genomes. Genome Biology and Evolution, 2015. 7(7): p. 1951-1959.

60

93. Dubrovsky, E.B., Hormonal cross talk in insect develoment. TRENDS in Endocrinology and Metabolism, 2005. 16(1).

94. Skinner, D.M., Molting and Regeneration. The Biology of Crustacea. 1985, New

York: Academic Press. 95. Chang, E.S., Endocrine Regulation of Molting in Crustaceans and Insects -

Variations on Common Themes. Abstracts of Papers of the American Chemical Society, 1995. 210: p. 21-Agro.

96. Chang, E., Endocrine regulation of molting in Crustacea. Reviews in Aquatic

science, 1989. 1: p. 131-157. 97. Quinn L., L.J., Cranna. N, Er J., Lee A., Mitchell N. and Hannan R. , Steroid

Hormones in Drosophila: How Ecdysone Coordinates Developmental Signalling with Cell Growth and Division, in Steroids - Basic Science, P.H. Abduljabbar, Editor. 2012: InTech.

98. Chang, E.S. and M.J. Bruce, Ecdysteroid titers of juvenile lobsters following molt

induction. Journal of Experimental Zoology, 1980. 214(2): p. 157-160. 99. Cheng, J.H. and E.S. Chang, Ecdysteroid Treatment Delays Ecdysis in the Lobster,

Homarus-Americanus. Biological Bulletin, 1991. 181(1): p. 169-174. 100. Riddiford, L.M. and J.W. Truman, Hormone Receptors and the Regulation of Insect

Metamorphosis. American Zoologist, 1993. 33(3): p. 340-347. 101. Henrich, V.C. and N.E. Brown, Insect Nuclear Receptors - a Developmental and

Comparative Perspective. Insect Biochemistry and Molecular Biology, 1995. 25(8): p. 881-897.

102. Qian, Z.Y., et al., Identification of ecdysteroid signaling late-response genes from

different tissues of the Pacific white shrimp, Litopenaeus vannamei. Comparative Biochemistry and Physiology a-Molecular & Integrative Physiology, 2014. 172: p. 10-30.

103. Riddiford, L.M., Hormone Receptors and the Regulation of Insect Metamorphosis.

Receptor, 1993. 3(3): p. 203-209. 104. Thummel, C.S., Dueling orphans - interacting nuclear receptors coordinate

Drosophila metamorphosis. Bioessays, 1997. 19(8): p. 669-672. 105. Thummel, C.S., Flies on steroids - Drosophila metamorphosis and the mechanisms

of steroid hormone action. Trends in Genetics, 1996. 12(8): p. 306-310. 106. Thummel, C.S., Eedysone-regulated puff genes 2000. Insect Biochemistry and

Molecular Biology, 2002. 32(2): p. 113-120.

61

107. Bortolin, F., et al., Cloning and expression pattern of the ecdysone receptor and retinoid X receptor from the centipede Lithobius peregrinus (Chilopoda, Lithobiomorpha). General and Comparative Endocrinology, 2011. 174(1): p. 60-69.

108. Gaertner, K., et al., Identification and expression of the ecdysone receptor in the

harpacticoid copepod, Amphiascus tenuiremis, in response to fipronil. Ecotoxicology and Environmental Safety, 2012. 76: p. 39-45.

109. Bender, M., et al., Drosophila ecdysone receptor mutations reveal functional

differences among receptor isoforms. Cell, 1997. 91(6): p. 777-788. 110. Durica, D.S., A.C.K. Chung, and P.M. Hopkins, Characterization of EcR and RXR

gene homologs and receptor expression during the molt cycle in the crab, Uca pugilator. American Zoologist, 1999. 39(4): p. 758-773.

111. Asazuma, H., et al., Molecular cloning and expression analysis of ecdysone receptor

and retinoid X receptor from the kuruma prawn, Marsupenaeus japonicus. Comparative Biochemistry and Physiology B-Biochemistry & Molecular Biology, 2007. 148(2): p. 139-150.

112. Shen, H., et al., Ecdysone receptor gene from the freshwater prawn Macrobrachium

nipponense: identification of different splice variants and sexually dimorphic expression, fluctuation of expression in the molt cycle and effect of eyestalk ablation. Gen Comp Endocrinol, 2013. 193: p. 86-94.

113. Kato, Y., et al., Cloning and characterization of the ecdysone receptor and

ultraspiracle protein from the water flea Daphnia magna. Journal of Endocrinology, 2007. 193(1): p. 183-194.

114. Fujiwara, H., et al., Cloning of an Ecdysone Receptor Homolog from Manduca-Sexta

and the Developmental Profile of Its Messenger-Rna in Wings. Insect Biochemistry and Molecular Biology, 1995. 25(7): p. 845-856.

115. Bender, M., et al., Mutational dissection of the Drosophila ecdysone receptor (EcR)

gene: EcR is required maternally For normal oogenesis and EcR-B isoforms are required for neuronal remodeling during metamorphosis. Developmental Biology, 1998. 198(1): p. 221-221.

116. Boulanger, A. and J.M. Dura, Nuclear receptors and Drosophila neuronal

remodeling. Biochimica Et Biophysica Acta-Gene Regulatory Mechanisms, 2015. 1849(2): p. 187-195.

117. Raikhel, A.S., K. Miura, and W.A. Segraves, Nuclear receptors in mosquito

vitellogenesis. American Zoologist, 1999. 39(4): p. 722-735. 118. Martin, D., S.F. Wang, and A.S. Raikhel, The vitellogenin gene of the mosquito

Aedes aegypti is a direct target of ecdysteroid receptor. Molecular and Cellular Endocrinology, 2001. 173(1-2): p. 75-86.

62

119. Romani, P., et al., Cell Survival and Polarity of Drosophila Follicle Cells Require the Activity of Ecdysone Receptor B1 Isoform. Genetics, 2009. 181(1): p. 165-175.

120. Carney, G.E. and M. Bender, The Drosophila ecdysone receptor (EcR) gene is

required maternally for normal oogenesis. Genetics, 2000. 154(3): p. 1203-1211. 121. Hodin, J. and L.M. Riddiford, The ecdysone receptor and ultraspiracle regulate the

timing and progression of ovarian morphogenesis during Drosophila metamorphosis. Development Genes and Evolution, 1998. 208(6): p. 304-317.

122. Belles, X. and M.D. Piulachs, Ecdysone signalling and ovarian development in

insects: from stem cells to ovarian follicle formation. Biochimica Et Biophysica Acta-Gene Regulatory Mechanisms, 2015. 1849(2): p. 181-186.

123. Cruz, J., et al., Functions of the ecdysone receptor isoform-A in the hemimetabolous

insect Blattella germanica revealed by systemic RNAi in vivo. Developmental Biology, 2006. 297(1): p. 158-171.

124. Parthasarathy, R., et al., Ecdysteroid regulation of ovarian growth and oocyte

maturation in the red flour beetle, Tribolium castaneum. Insect Biochemistry and Molecular Biology, 2010. 40(6): p. 429-439.

125. Subramoniam, T., Crustacean ecdysteriods in reproduction and embryogenesis.

Comp Biochem Physiol C Toxicol Pharmacol, 2000. 125(2): p. 135-56. 126. Brown, M.R., D.H. Sieglaff, and H.H. Rees, Gonadal Ecdysteroidogenesis in

Arthropoda: Occurrence and Regulation. Annual Review of Entomology, 2009. 54: p. 105-125.

127. Gonsalves, S.E., et al., Genome-wide examination of the transcriptional response to

ecdysteroids 20-hydroxyecdysone and ponasterone A in Drosophila melanogaster. Bmc Genomics, 2011. 12.

128. Beckstead, R.B., G. Lam, and C.S. Thummel, The genomic response to 20-

hydroxyecdysone at the onset of Drosophila metamorphosis. Genome Biology, 2005. 6(12).

129. Eichner, C., et al., Characterization of a novel RXR receptor in the salmon louse

(Lepeophtheirus salmonis, Copepoda) regulating growth and female reproduction. BMC Genomics, 2015. 16(1): p. 81.

130. Chavez, V.M., et al., The Drosophila disembodied gene controls late embryonic

morphogenesis and codes for a cytochrome P450 enzyme that regulates embryonic ecdysone levels. Development, 2000. 127(19): p. 4115-4126.

131. Niwa, R., et al., CYP306A1, a cytochrome P450 enzyme, is essential for ecdysteroid

biosynthesis in the prothoracic glands of Bombyx and Drosophila. J Biol Chem, 2004. 279(34): p. 35942-9.

63

132. Petryk, A., et al., Shade is the Drosophila P450 enzyme that mediates the hydroxylation of ecdysone to the steroid insect molting hormone 20-hydroxyecdysone. Proceedings of the National Academy of Sciences of the United States of America, 2003. 100(24): p. 13773-13778.

133. Warren, J.T., et al., Phantom encodes the 25-hydroxylase of Drosophila

melanogaster and Bombyx mori: a P450 enzyme critical in ecdysone biosynthesis. Insect Biochem Mol Biol, 2004. 34(9): p. 991-1010.

134. Gilbert, L.I., Q. Song, and R. Rybczynski, Control of ecdysteroidogenesis: activation

and inhibition of prothoracic gland activity. Invert Neurosci, 1997. 3(2-3): p. 205-16. 135. Spaziani, E., et al., Signaling pathways for ecdysteroid hormone synthesis in

crustacean Y-organs. American Zoologist, 1999. 39(3): p. 496-512. 136. Lachaise, F., et al., The Molting Gland of Crustaceans - Localization, Activity, and

Endocrine Control (a Review). Journal of Crustacean Biology, 1993. 13(2): p. 198-234.

137. Grieneisen, M.L., Recent Advances in Our Knowledge of Ecdysteroid Biosynthesis in

Insects and Crustaceans. Insect Biochemistry and Molecular Biology, 1994. 24(2): p. 115-132.

138. Rees, H.H., Ecdysteroid Biosynthesis and Inactivation in Relation to Function.

European Journal of Entomology, 1995. 92(1): p. 9-39. 139. Gilbert, L.I. and K.F. Rewitz, The function and evolution of the halloween genes: the

pathway to the arthropod molting hormone, in Ecdysone: Structures and Functions. 2009, Springer. p. 231-269.

140. Hopkins, P.M., Crustacean ecdysteroids and their receptors, in Ecdysone: structures

and functions, G. Smagghe, Editor. 2009, Springer. p. 73-97. 141. Yoshiyama, T., et al., Neverland is an evolutionally conserved Rieske-domain protein

that is essential for ecdysone synthesis and insect growth. Development, 2006. 133(13): p. 2565-2574.

142. Sumiya, E., et al., Roles of ecdysteroids for progression of reproductive cycle in the

fresh water crustacean Daphnia magna. Frontiers in Zoology, 2014. 11. 143. Jia, S., et al., Knockdown of a putative Halloween gene Shade reveals its role in

ecdysteroidogenesis in the small brown planthopper Laodelphax striatellus. Gene, 2013. 531(2): p. 168-174.

144. Ono, H., et al., Spook and Spookier code for stage-specific components of the

ecdysone biosynthetic pathway in Diptera. Developmental Biology, 2006. 298(2): p. 555-570.

64

145. Namiki, T., et al., Cytochrome P450CYP307A1/spook: A regulator for ecdysone synthesis in insects. Biochemical and Biophysical Research Communications, 2005. 337(1): p. 367-374.

146. Ou, Q.X., A. Magico, and K. King-Jones, Nuclear Receptor DHR4 Controls the

Timing of Steroid Hormone Pulses During Drosophila Development. Plos Biology, 2011. 9(9).

147. Niwa, R., et al., Non-molting glossy/shroud encodes a short-chain

dehydrogenase/reductase that functions in the 'Black Box' of the ecdysteroid biosynthesis pathway. Development, 2010. 137(12): p. 1991-1999.

148. Rewitz, K.F. and L.I. Gilbert, Daphnia Halloween genes that encode cytochrome

P450s mediating the synthesis of the arthropod molting hormone: Evolutionary implications. Bmc Evolutionary Biology, 2008. 8.

149. Hansen, B.H., et al., Expression of ecdysteroids and cytochrome P450 enzymes

during lipid turnover and reproduction in Calanus finmarchicus (Crustacea: Copepoda). General and Comparative Endocrinology, 2008. 158(1): p. 115-121.

150. Gonçalves, A.T., R. Farlora, and C. Gallardo-Escárate, Transcriptome survey of the

lipid metabolic pathways involved in energy production and ecdysteroid synthesis in the salmon louse Caligus rogercresseyi (Crustacea: Copepoda). Comparative Biochemistry and Physiology Part B: Biochemistry and Molecular Biology, 2014. 176(0): p. 9-17.

151. Baldwin, J.M., G.F.X. Schertler, and V.M. Unger, An alpha-carbon template for the

transmembrane helices in the rhodopsin family of G-protein-coupled receptors. Journal of Molecular Biology, 1997. 272(1): p. 144-164.

152. Roeder, T., Octopamine in invertebrates. Progress in Neurobiology, 1999. 59(5): p.

533-561. 153. El-Kholy, S., et al., Expression analysis of octopamine and tyramine receptors in

Drosophila. Cell Tissue Res, 2015. 361(3): p. 669-84. 154. Grohmann, L., et al., Molecular and functional characterization of an octopamine

receptor from honeybee (Apis mellifera) brain. J Neurochem, 2003. 86(3): p. 725-35. 155. Menzel, R. and U. Muller, Learning and memory in honeybees: from behavior to

neural substrates. Annu Rev Neurosci, 1996. 19: p. 379-404. 156. Ohhara, Y., et al., Autocrine regulation of ecdysone synthesis by beta3-octopamine

receptor in the prothoracic gland is essential for Drosophila metamorphosis. Proc Natl Acad Sci U S A, 2015. 112(5): p. 1452-7.

157. Fire, A., et al., Potent and specific genetic interference by double-stranded RNA in

Caenorhabditis elegans. Nature, 1998. 391(6669): p. 806-811.

65

158. Montgomery, M.K., S.Q. Xu, and A. Fire, RNA as a target of double-stranded RNA-mediated genetic interference in Caenorhabditis elegans. Proceedings of the National Academy of Sciences of the United States of America, 1998. 95(26): p. 15502-15507.

159. Kamath, R.S. and J. Ahringer, Genorne-wide RNAi screening in Caenorhabditis

elegans. Methods, 2003. 30(4): p. 313-321. 160. Quan, G.X., et al., Characterization of the kynurenine 3-monooxygenase gene

corresponding to the white egg 1 mutant in the silkworm Bombyx mori. Molecular Genetics and Genomics, 2002. 267(1): p. 1-9.

161. Sharabi, O., et al., Dual function of a putative epidermal growth factor receptor in

the decapod crustacean Macrobrachium rosenbergii. Integrative and Comparative Biology, 2013. 53: p. E195-E195.

162. Das, S. and D.S. Durica, Ecdysteroid receptor signaling disruption obstructs

blastemal cell proliferation during limb regeneration in the fiddler crab, Uca pugilator. Molecular and Cellular Endocrinology, 2013. 365(2): p. 249-259.

163. Priya, T.A.J., et al., Molecular characterization and effect of RNA interference of

retinoid X receptor (RXR) on E75 and chitinase gene expression in Chinese shrimp Fenneropenaeus chinensis. Comparative Biochemistry and Physiology B-Biochemistry & Molecular Biology, 2009. 153(1): p. 121-129.

164. Priya, T.A.J., et al., Molecular characterization of an ecdysone inducible gene E75 of

Chinese shrimp Fenneropenaeus chinensis and elucidation of its role in molting by RNA interference. Comparative Biochemistry and Physiology B-Biochemistry & Molecular Biology, 2010. 156(3): p. 149-157.

165. Schumpert, C.A., J.L. Dudycha, and R.C. Patel, Development of an efficient RNA

interference method by feeding for the microcrustacean Daphnia. Bmc Biotechnology, 2015. 15.

166. Kato, Y., et al., Development of an RNA interference method in the cladoceran

crustacean Daphnia magna. Development Genes and Evolution, 2011. 220(11-12): p. 337-345.

167. Eichner, C., et al., A method for stable gene knock-down by RNA interference in

larvae of the salmon louse (Lepeophtheirus salmonis). Experimental Parasitology, 2014. 140: p. 44-51.

168. Sandlund, L., et al., Molecular characterisation of the salmon louse, Lepeophtheirus

salmonis salmonis (Kroyer, 1837), ecdysone receptor with emphasis on functional studies of female reproduction. Int J Parasitol, 2015. 45(2-3): p. 175-85.

169. Schubiger, M., et al., Drosophila EcR-B ecdysone receptor isoforms are required for

larval molting and for neuron remodeling during metamorphosis. Development, 1998. 125(11): p. 2053-2062.

66

170. Carney, G.E., et al., DHR3, an ecdysone-inducible early-late gene encoding a Drosophila nuclear receptor, is required for embryogenesis. Proceedings of the National Academy of Sciences of the United States of America, 1997. 94(22): p. 12024-12029.

171. Kozlova, T. and C.S. Thummel, Essential roles for ecdysone signaling during

Drosophila mid-embryonic development. Science, 2003. 301(5641): p. 1911-4. 172. Buszczak, M., et al., Ecdysone response genes govern egg chamber development

during mid-oogenesis in Drosophila. Development, 1999. 126(20): p. 4581-4589. 173. Xu, J.J., J.A. Shu, and Q. Zhang, Expression of the Tribolium castaneum

(Coleoptera: Tenebrionidae) hsp83 gene and its relation to oogenesis during ovarian maturation. Journal of Genetics and Genomics, 2010. 37(8): p. 513-522.

174. Durica, D.S., et al., Characterization of crab EcR and RXR homologs and expression

during limb regeneration and oocyte maturation. Molecular and Cellular Endocrinology, 2002. 189(1-2): p. 59-76.

175. Gong, J., et al., Ecdysone receptor in the mud crab Scylla paramamosain: a possible

role in promoting ovarian development. Journal of Endocrinology, 2015. 224(3): p. 273-287.

176. Rewitz, K.F., et al., Identification, characterization and developmental expression of

Halloween genes encoding P450 enzymes mediating ecdysone biosynthesis in the tobacco hornworm, Manduca sexta. Insect Biochemistry and Molecular Biology, 2006. 36(3): p. 188-199.

177. Zhang, T., P. Haws, and Q. Wu, Multiple variable first exons: A mechanism for cell-

and tissue-specific gene regulation. Genome Research, 2004. 14(1): p. 79-89. 178. Trinklein, N.D., et al., Identification and functional analysis of human

transcriptional promoters. Genome Research, 2003. 13(2): p. 308-312. 179. Carninci, P., et al., The transcriptional landscape of the mammalian genome.

Science, 2005. 309(5740): p. 1559-1563. 180. Jackson, R.J., C.U.T. Hellen, and T.V. Pestova, The mechanism of eukaryotic

translation initiation and principles of its regulation. Nature Reviews Molecular Cell Biology, 2010. 11(2): p. 113-127.

181. Sonenberg, N. and A.G. Hinnebusch, Regulation of Translation Initiation in

Eukaryotes: Mechanisms and Biological Targets. Cell, 2009. 136(4): p. 731-745. 182. Araujo, P.R., et al., Before It Gets Started: Regulating Translation at the 5 ' UTR.

Comparative and Functional Genomics, 2012. 183. Styrishave, B., et al., Variations in ecdysteroid levels and Cytochrome p(450)

expression during moult and reproduction in male shore crabs Carcinus maenas. Marine Ecology Progress Series, 2004. 274: p. 215-224.

67

184. Li, T.R. and M. Bender, A conditional rescue system reveals essential functions for the ecdysone receptor (EcR) gene during molting and metamorphosis in Drosophila. Development, 2000. 127(13): p. 2897-2905.

185. Mirth, C.K., J.W. Truman, and L.M. Riddiford, The Ecdysone receptor controls the

post-critical weight switch to nutrition-independent differentiation in Drosophila wing imaginal discs. Development, 2009. 136(14): p. 2345-2353.

186. Nakagawa, Y., et al., Molecular cloning of the ecdysone receptor and the retinoid X

receptor from the scorpion Liocheles australasiae. Febs Journal, 2007. 274(23): p. 6191-6203.

187. Hegstrom, C.D. and J.W. Truman, Steroid control of muscle remodeling during

metamorphosis in Manduca sexta. Journal of Neurobiology, 1996. 29(4): p. 535-550. 188. Hegstrom, C.D., L.M. Riddiford, and J.W. Truman, Steroid and neuronal regulation

of ecdysone receptor expression during metamorphosis of muscle in the moth, Manduca sexta. Journal of Neuroscience, 1998. 18(5): p. 1786-1794.

189. Lee, T., et al., Cell-autonomous requirement of the USP/EcR-B ecdysone receptor for

mushroom body neuronal remodeling in Drosophila. Neuron, 2000. 28(3): p. 807-818.

190. Lee, C.Y., B.A. Cooksey, and E.H. Baehrecke, Steroid regulation of midgut cell

death during Drosophila development. Dev Biol, 2002. 250(1): p. 101-11. 191. Parthasarathy, R., et al., Transcription factor broad suppresses precocious

development of adult structures during larval-pupal metamorphosis in the red flour beetle, Tribolium castaneum. Mechanisms of Development, 2008. 125(3-4): p. 299-313.

192. Shi, Y.B. and A. Ishizuya-Oka, Biphasic intestinal development in amphibians:

embryogenesis and remodeling during metamorphosis. Curr Top Dev Biol, 1996. 32: p. 205-35.

193. Spindler, K.D., S. Przibilla, and M. Spindler-Barth, Moulting hormones of

arthropods: Molecular mechanisms. Zoology-Analysis of Complex Systems, 2001. 103(3-4): p. 189-201.

194. Tellam, R.L., et al., Insect chitin synthase - cDNA sequence, gene organization and

expression. European Journal of Biochemistry, 2000. 267(19): p. 6025-6042. 195. Moussian, B., et al., Involvement of chitin in exoskeleton morphogenesis in

Drosophila melanogaster. Journal of Morphology, 2005. 264(1): p. 117-130. 196. Romani, P., G. Gargiulo, and V. Cavaliere, The ecdysone receptor signalling

regulates microvilli formation in follicular epithelial cells. Cell Mol Life Sci, 2016. 73(2): p. 409-25.

68

197. Schlichting, K., et al., Cadherin Cad99C is required for normal microvilli morphology in Drosophila follicle cells. Journal of Cell Science, 2006. 119(6): p. 1184-1195.

198. Elalayli, M., et al., Palisade is required in the Drosophila ovary for assembly and

function of the protective vitelline membrane. Developmental Biology, 2008. 319(2): p. 359-369.

199. Tiu, S.H.K., S.M. Chan, and S.S. Tobe, The effects of farnesoic acid and 20-

hydroxyecdysone on vitellogenin gene expression in the lobster, Homarus americanus, and possible roles in the reproductive process. General and Comparative Endocrinology, 2010. 166(2): p. 337-345.

200. Lee, H.G., S. Rohila, and K.A. Han, The Octopamine Receptor OAMB Mediates

Ovulation via Ca2+/Calmodulin-Dependent Protein Kinase II in the Drosophila Oviduct Epithelium. Plos One, 2009. 4(3).

201. Samuelsen, O.B., et al., Distribution and persistence of the anti sea-lice drug

teflubenzuron in wild fauna and sediments around a salmon farm, following a standard treatment. Science of the Total Environment, 2015. 508: p. 115-121.

202. Rewitz, K.F., et al., Marine invertebrate cytochrome P450: Emerging insights from

vertebrate and insect analogies. Comparative Biochemistry and Physiology C-Toxicology & Pharmacology, 2006. 143(4): p. 363-381.

69

I

Molecular characterisation of the salmon louse, Lepeophtheirus salmonissalmonis (Krøyer, 1837), ecdysone receptor with emphasis on functionalstudies of female reproduction

Liv Sandlund a,⇑, Frank Nilsen a, Rune Male b, Sindre Grotmol a, Heidi Kongshaug a, Sussie Dalvin c

a Sea Lice Research Centre, Department of Biology, University of Bergen, Thormøhlensgt. 55, 5008 Bergen, Norwayb Sea Lice Research Centre, Department of Molecular Biology, University of Bergen, Thormøhlensgt. 55, 5008 Bergen, Norwayc Sea Lice Research Centre, Institute of Marine Research, 5817 Bergen, Norway

a r t i c l e i n f o

Article history:Received 5 June 2014Received in revised form 8 October 2014Accepted 17 October 2014Available online 20 November 2014

Keywords:CopepodSea liceRNA interferenceIn situVitellogenesisReproduction

a b s t r a c t

The salmon louse Lepeophtheirus salmonis (Copepoda, Caligidae) is an important parasite in the salmonfarming industry in the Northern Hemisphere causing annual losses of hundreds of millions of dollars(US) worldwide. To facilitate development of a vaccine or other novel measures to gain control of the par-asite, knowledge about molecular biological functions of L. salmonis is vital. In arthropods, a nuclearreceptor complex consisting of the ecdysone receptor and the retinoid X receptor, ultraspiracle, are wellknown to be involved in a variety of both developmental and reproductive processes. To investigate therole of the ecdysone receptor in the salmon louse, we isolated and characterised cDNA with the50untranslated region of the predicted L. salmonis EcR (LsEcR). The LsEcR cDNA was 1608 bp encoding a536 amino acid sequence that demonstrated high sequence similarities to other arthropod ecdysonereceptors including Tribolium castaneum and Locusta migratoria. Moreover, in situ analysis of adult femalelice revealed that the LsEcR transcript is localised in a wide variety of tissues such as ovaries, sub-cuticulaand oocytes. Knock-down studies of LsEcR using RNA interference terminated egg production, indicatingthat the LsEcR plays important roles in reproduction and oocyte maturation. We believe this is the firstreport on the ecdysone receptor in the economically important parasite L. salmonis.� 2014 The Authors. Published by Elsevier Ltd. on behalf of Australian Society for Parasitology Inc. This isan open access article under the CCBY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/3.0/).

1. Introduction

In arthropods, steroid hormones such as 20-hydroxyecdysone(20-E) and ponasterone A (PonA) (hereafter referred to collectivelyas ecdysone) initiate signalling through a multitude of pathwaysthat regulate different aspects of biological processes such asdevelopment and reproduction. The effect of ecdysone is generallymediated by binding to a nuclear receptor (NR) complex consistingof two transcription factors; the ecdysone receptor (EcR, NR1H1)and the retinoid X receptor homolog ultraspiracle (USP, NR2B)(Yao et al., 1992, 1993; Thomas et al., 1993). The ligand–receptorcomplex regulates the transcription of ecdysone-responsive earlyand early-late genes such as E74, E75 and Broad Complex (Br-C)(Thummel and Chory, 2002; Riddiford et al., 2003) by binding toecdysone response elements (EcREs) in the promoter region oftheir DNA sequence. Activation of these transcription factors fur-ther trigger the expression of ecdysone-responsive late genes,

which define the phenotypic effects of the steroid hormones in aspatial and tissue-specific manner (Thummel, 2002; Qian et al.,2014).

The EcR belongs to the NR protein superfamily that is character-ised by five typical NR domains (Evans, 1988; Billas et al., 2009): (i)a highly variable N-terminal (domain A/B) important in activationof transcription, (ii) a highly conserved DNA binding domain (DBD)(domain C) containing two C2C2 zinc finger motifs important inheterodimerisation and recognition of EcREs, (iii) a flexible andvariable hinge region (domain D) involved in EcRE recognitionand heterodimerisation, (iv) a moderately conserved ligand bind-ing domain (LBD) (domain E) including 12 a-helices and twob-sheets making up a complex tertiary structure that is subjectedto conformational changes which enable involvement in ligandbinding and dimerisation with other transcription factors, and (v)a highly variable C-terminal of unknown function (F domain)(Hill et al., 2013). Different isoforms of the EcR are found in a selec-tion of arthropods such as the marine copepod Amphiascus tenuire-mis (Gaertner et al., 2012) and the freshwater decapodMacrobrachium nipponense (Shen et al., 2013) that have three andfour isoforms, respectively. The temporal and spatial expression

http://dx.doi.org/10.1016/j.ijpara.2014.10.0030020-7519/� 2014 The Authors. Published by Elsevier Ltd. on behalf of Australian Society for Parasitology Inc.This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/3.0/).

⇑ Corresponding author.E-mail address: liv.sandlund@bio.uib.no (L. Sandlund).

International Journal for Parasitology 45 (2015) 175–185

Contents lists available at ScienceDirect

International Journal for Parasitology

journal homepage: www.elsevier .com/locate / i jpara

differ between isoforms, however the biological functions areunknown. Recent studies on the Chinese freshwater prawn, M. nip-ponense, showed that isoforms MnEcR-S1 and MnEcR-S2 weremainly found in testes while isoforms L1 and L2 were predomi-nantly detected in the ovaries, suggesting a sex-specific expressionpattern for the different isoforms. Knock-down studies of EcR per-formed in Tribolium castaneum resulted in impairment of ovariangrowth and oocyte maturation as well as possible induction ofapoptosis in the follicular cells (Parthasarathy and Palli, 2007). Inaddition, functional analysis performed in the fruit fly, Drosophilamelanogaster, revealed defects in ovarian differentiation whenEcR levels were reduced (Hodin and Riddiford, 1998).

The EcR sequence has been identified in crustacean species suchas the decapods Uca pugilator (Hopkins et al., 2008) and Homarusamericanus (Tarrant et al., 2011), the branchiopod Daphnia magna(Kato et al., 2007), the copepods Tigriopus japonicus (Hwanget al., 2010) and A. tenuiremis (Gaertner et al., 2012), and the mys-ids Americamysis bahia (Hirano et al., 2009) and Neomysis integer(De Wilde et al., 2013). Even though the receptor has been identi-fied and sequenced in several crustacean species, few functionalstudies have been performed, leaving the action of the EcR in spe-cies other than insects poorly understood.

The endocrine system has been extensively studied in hexapodswhere ecdysteroids are produced and secreted from the protho-racic glands during metamorphosis (Gilbert et al., 1997) and fromovarian follicle cells after adult female eclosion (reviewed by Bellesand Piulachs, 2014). In many crustaceans such as the Americanlobster H. americanus, the hormones are produced and secretedfrom the Y-organ (Mykles, 2011). In copepods, however, such anorgan has yet to be identified which renders the origin of steroidsecretion and distribution pathways, for example in the salmonlouse Lepeophtheirus salmonis (Krøyer, 1837), unknown. Onehypothesis suggests the ecdysone steroid is secreted from oneorgan and transported with vitellogenins to the oocytes where itis stored for use in embryogenesis.

The salmon louse, L. salmonis, is a marine ecto-parasite ofsalmonid fishes (Salmo and Onchorhyncus) in the Northern Hemi-sphere (Kabata, 1979). In the salmon farming industry, the salmonlouse has become an increasing problem due to the high number ofhosts available, which facilitates continuous re-infestation and thespread of lice between farming sites (Heuch et al., 2005). A majorconcern is the development of resistance to the currently approvedpesticides (Fallang et al., 2004; Espedal et al., 2013), which leads tohigher consumption of these drugs followed by the spread of resis-tance within louse populations, thereby creating a loop of negativeeffects.

The EcR has long been known as a site of action for ecdysteroidagonists such as the bisacylhydrazines (BAH). Ligand bindingassays using recombinant EcRs have demonstrated that thesechemicals attain large variations in binding affinities between dif-ferent phylogenetic groups, thus making them target-specific.Their selective specificity and their non-toxic effect on vertebrateshave made these agonists important tools in integrated pest man-agement as they have reduced the risk of affecting non-pest spe-cies and of causing negative environmental effects (Dhadiallaet al., 1998; Hill et al., 2012). Understanding the EcR/USP heterodi-mer complex and the endocrine signalling pathways in L. salmoniscould be of great importance for development of vaccines and/ornovel medicines against this important parasite.

Here we show that the L. salmonis EcR (LsEcR) gene codes for EcRfrom a single exon but contains several alternative 50 untranslated(UTR) exons that may determine in which organs of the adultfemale louse the gene is expressed. Moreover, in female lice genesilencing using RNA interference (RNAi) targeted to LsEcR gave adistinct phenotype with no production of egg strings. This suggeststhat signalling mediated by LsEcR, either directly or indirectly,

plays a key role in oogenesis and that disruption of this signallingpathway may provide a means by which to control louse reproduc-tion and, consequently, infestation.

2. Materials and methods

2.1. Animal culture and sampling

Eggs from the Atlantic salmon louse strain Lepeophtheirus sal-monis salmonis (Skern-Mauritzen et al., 2014) were hatched andcultivated to copepodid stage in flow-through incubators beforeinfection of Atlantic salmon Salmo salar (Hamre et al., 2009). Bothlice and fish were kept in seawater with a salinity of 34.5‰ and atemperature of approximately 10 �C. The lice were kept on the fishuntil they reached the desired developmental stage. Prior to sam-pling, the salmon were either killed with a blow to the head oranaesthetised in a mixture of methomidate (5 mg/l) and benzo-caine (60 mg/l); thereafter lice were removed with forceps. Salmonwere held and treated in accordance with the Norwegian legisla-tion for animal welfare.

2.2. Cloning and sequencing of LsEcR

For all stages of salmon lice, total RNA was isolated using TRIReagent� (Sigma–Aldrich, St Louis, MO, USA) according to themanufacturer’s protocol. The total RNA was treated with Amp-Grade DNase I (Invitrogen, Carlsbad, CA, USA) and reverse tran-scribed for preparation of template cDNA using SMARTscribeReverse Transcriptase (Clontech, Takara Bio, CA, USA). 50-RACEwas performed using the SMARTer™RACE cDNA Amplification kit(Clontech, TaKaRa) with kit primers and an EcR-specific primer(EcR_specific_P1; Table 1), according to the manufacturer’s recom-mendations (Sigma–Aldrich). The following PCR program wasused: initial denaturation step 94 �C for 2 min and subsequent 35cycles of amplification (94 �C, 30 s; 68 �C, 30 s; 72 �C, 2 min). ThePCR products were run on a 1% agarose gel, purified using a GelE-lute™ Gel Extraction Kit (Sigma–Aldrich), sub-cloned using apCR�4-TOPO� vector system (Invitrogen) and transformed intoEscherichia coli TOP10 cells. Clones were verified by PCR withM13_f and M13_r primers (Table 1), grown overnight and purifiedusing a Miniprep Nucleospin� Plasmid Purification Kit (Macherey–Nagel, Duren, Germany). Plasmids were sequenced using a BigDye�

Terminator v3.1 Cycle sequencing kit (Applied Biosystems�, FosterCity, CA, USA) and analysed in MacVector (MacVector Inc., NorthCarolina, USA).

2.3. Sequence comparison and phylogenetic analysis

To investigate the phylogenetic position of the LsEcR protein,homologous proteins were found by basic local alignment searchtool (BLAST) searches performed in GenBank (National Center forBiotechnology Information (NCBI), Bethesda, USA). A total of 30EcR protein sequences or EcR-like sequences from different speciescovering the phyla Annelida, Arthropoda, Chordata, Mollusca,Nematoda and Platyhelminthes were chosen. GenBank accessionnumbers of selected sequences are listed in Table 2. Multiplesequence alignment was performed using ClustalX2 (Thompsonet al., 1997) with the multiple alignment parameter settings of10 for gap opening and 0.2 for gap extension. The alignment wastrimmed in MacVector by removal of parts of the highly variable50-A/B domain and converted to Nexus format using Mesquite(Maddison, W.P., Maddison, D.R., 2004. Mesquite: a modular sys-tem for evolutionary analysis. v2.5. http://mequiteproject.org).Phylogenetic analysis was performed using MrBayes v3.2(Huelsenbeck and Ronquist, 2001; Ronquist et al., 2012) with the

176 L. Sandlund et al. / International Journal for Parasitology 45 (2015) 175–185

General time-reversal inverted gamma (GTR + I + G) amino acid(aa) substitution matrix. The Monte Carlo Markov Chain (MCMC)was run with two simultaneous runs and four simultaneous chainsfor 1,000,000 generations to approximate the posterior probability.The MCMC temperature was set to 0.5. FigTree v1.4 (A. Rambaut,2007, http://tree.bio.ed.ac.uk/software/figtree/) was used to evalu-ate the consensus tree with percent posterior probability valuesestimated on each branch node. To root the tree, sequences fromvertebrates, S. salar, Xenopus tropicalis and Crotatus adamantus,were used as outgroups.

2.4. Analysis of expression levels of the 50UTR mRNA splice variants ofLsEcR at different life stages using real-time quantitative PCR (RTq-PCR)

Five parallels of different life stages of the salmon louse weresampled prior to ontogenetic analysis; nauplia I/II (n � 150),free-living copepodids (n � 150), parasitic copepodids (n = 10),chalimus I (n = 10), chalimus II (n = 10), pre-adult male I/II(n = 1), pre-adult female I/II (n = 1), adult male (n = 1), immatureadult female lice (n = 1) and gravid female lice (n = 1), and storedon RNAlater™ (Ambion Inc., Austin, TX, USA). Total RNA was iso-lated using TRI Reagent� (Sigma–Aldrich) according to the manu-facturer’s protocol. Concentration and purity of RNA wasdetermined using a NanoDrop ND-1000 spectrophotometer (Nano-Drop Technologies Inc., Thermo Fisher Scientific, Wilmington, DE,USA). RNA quantity and quality was checked by standard O.D.

260/280 and O.D. 260/230. The normalised stocks (500 ng/ll) weretreated with DNase I (Amplification Grade, Invitrogen). Two paral-lel cDNA synthesis reactions were set up using an AffinityScriptcDNA Synthesis Kit (Agilent Technologies, Santa Clara, CA, USA)to a final concentration of 10 ng/ll. PCR was performed using2.5 lg of cDNA, 5 lM LsEcR-specific TaqMan� probe (Table 1)and 2� TaqMan� Universal PCR mix (Applied Biosystems�) in atotal volume of 10 ll. The RTq-PCR of the mRNA LsEcR variantswas carried out independently but simultaneously with the house-keeping gene, elongation factor 1 alpha (EF1a; Frost and Nilsen,2003) as the reference. RTq-PCR was performed with parallel seriesof each sample. Standard curves (cycle at threshold (Ct) versus logquantity), slope evaluation and transcription levels of the mRNALsEcR variants were compared with EF1a using the Applied Biosys-tems 7500 Real-Time PCR System (Applied Biosystems�). Resultswere analysed by the 2^�DDCt approach and presented with the95% confidence interval calculated from the 2^�DDCt values.

2.5. Localisation of LsEcR transcript

Localisation of LsEcR mRNA in adult female lice was accom-plished using in situ hybridisation carried out according toKvamme et al. (2004) with some modifications. PCR product withT7 promoters generated from LsEcR-specific cDNA was used as atemplate for a single stranded digoxigenin (DIG)-labelled RNAprobe (667 bp) synthesis (Table 1, primers: LsEcR_specific_f,LsEcR_specific_r). Probe concentration and quality was determined

Table 1Primer sequences and Taqman� assaysa used in this study.

Primer nameb Sequence (50–30) Method

EcR_specific_P1 GTTGATCCCTAAGGATCGAAGCTCAGTA 50-RACEEcR_specific_P2 GAAAGTCGATAACGCAGAATACGCTCTCM13_f GTAAAACGACGGCCAG TOPO cloningM13_r CAGGAAACAGCTATGACLsEcR_specific_P3 CCGATTTGCCATTACGTAGGCTTGTAGAGC 30RACE/in situ/dsRNALsEcR_specific_P4 CCGCAGCTGCAGCCGACACAACTGTAGAT in situ/dsRNALsEcR_specific_P5 CGAGCGTTTCCACTTACTTGCCAT dsRNALsEcR_specific_P6 CGCCAACAACGACGACCC TCCACCAACAGCACT dsRNACod_specific_T7f ATAGGGCGAATTGGGTACCG dsRNACod_specific_T7r AAAGGGAACAAAAGCTGGAGC dsRNALsEF1a_f CATCGCCTGCAAGTTTAACCAAATT RTq-PCRLsEF1a_r CCGGCATCACCAGACTTGA RTq-PCRLsEF1a_TaqMan� ACGTACTGGTAAATCCAC RTq-PCRmRNA LsEcR total_f TCGGGAGAAAGTCCCTCTTCT RTq-PCRmRNA LsEcR total_r ACAGCTCCAGTAGGTGTTAAAGGA RTq-PCRmRNA LsEcR total TaqMan� TCGCAGTCCATTCTC RTq-PCRmRNA LsEcRa_f GTGTAGATGTGTTGTTGAAAGGGAAAAA RTq-PCRmRNA LsEcRa_r CCTATCAATGCACCCTTTAATTTTCCAA RTq-PCRmRNA LsEcRa TaqMan� AAACACGGCAAATATG RTq-PCRmRNA LsEcRb_f AACGAAACAAAAAAGACAAGTGGAATGT RTq-PCRmRNA LsEcRb_r TCACCCGTTGAGTGACTTCTTT RTq-PCRmRNA LsEcRb TaqMan� CATCTCCGCAGAACTT RTq-PCRmRNA LsEcRc_f CATCATCAGAGTCTCTGCAATCAAT RTq-PCRmRNA LsEcRc_r TTTTGGACCAATCGTTCTAGAAAACTTTTT RTq-PCRmRNA LsEcRc TaqMan� CCTCACCCACTTTTGC RTq-PCRLsE75_f CCTTGACCAATTTTCAGAACGGTTT RTq-PCRLsE75_r AATCCAGGGATCCGCTTGG RTq-PCRLsE75_TaqMan� CACGTTCGCCAAGTTT RTq-PCRLsBR-C_f CTCCATTGTACATAAAACAGAGTAGTGACT RTq-PCRLsBR-C_f CAGTACCTCATCAACATCCTTTGCT RTq-PCRLsBR-C_TaqMan� AATGCCTCGCAAATAG RTq-PCRLsVit-1_P1 ACATCGACTACAAAGGAACTCAGAAC RTq-PCRLsVit-1_P2 GGAAGCATGTAACGAATGAACTCA RTq-PCRLsVit-1_TaqMan� AGATTTTCTTTAGCTTCTGGATACAAACCTGCTCCA RTq-PCRLsVit-2_P1 AATGAGCAATTTAGTTGAGAAAACTTGT RTq-PCRLsVit-2_P2 CAATCTCGCTTTGAGCATCACA RTq-PCRLsVit-2_TaqMan� TGGATAAATCACGTCAAGTTACTTACCCTACCGC RTq-PCR

RACE, rapid amplification of cDNA ends; TOPO, DNA topoisomerase I; dsRNA, double-stranded RNA; RTq-PCR, real-time quantitative PCR.a Taqman� assays were provided by Applied Biosystems, Branchurg, NJ, USA.b All general primers were purchased from Sigma–Aldrich, St Louis, MO, USA.

L. Sandlund et al. / International Journal for Parasitology 45 (2015) 175–185 177

by spectrometry (Nanodrop ND-1000) and a spot test, respectively.Briefly, paraffin sections were baked at 60 �C for a minimum of20 min and treated with Histoclear (National Diagnostics, Atlanta,GA, USA) prior to rehydration of tissue and proteinase K treatmentfor 10 min, followed by tissue fixation in 4% formaldehyde in PBS,acetic anhydride treatment and dehydration. Hybridisation mix(100 ll) containing 20 ng of DIG-labelled RNA was added to the tis-sue and left overnight in a vacuum chamber at 60 �C. DIG-labelledprobes were visualised using secondary antibody labelled with ananti-DIG alkaline phosphatase-conjugated FAB fragment and achromogen substrate containing nitroblue tetrazolium (NBT)(Roche Diagnostics GMbH, Mannheim, Germany) and 5-bromo-4-chloro-3-indolyl phosphate (BCIP) (Roche Diagnostics). SenseRNA was used as a negative control.

2.6. LsEcR knock-down using RNAi

Two primer pairs with and without a 50 T7 promoter extensionwere used to generate PCR products of the LsEcR open readingframe (ORF). Fragment 1 (667 bp; Table 1, primers: LsEcR_spe-cific_P3 and LsEcR_specific_P4) and fragment 2 (815 bp; Table 1,primers LsEcR_specific_P5 and LsEcR_specific_P6) localised to thehinge and A/B region, respectively. An Atlantic cod (Gadus morhua)gene fragment, CPY185 (850 bp), was used as a control (Table 1,primers: Cod_specific_T7f and Cod_specific_T7r). The PCR productswere used as templates with T7 RNA polymerase to synthesisedsRNA fragments as described by the MEGAscript� RNAi Kit(Ambion Inc.). The concentrations of sense and anti-sense strandswere measured by spectrometry (NanoDrop Technologies Inc.)before equimolar amounts of each strand were pooled to generatedsRNA. A solution containing 50 ll of dsRNA was added to 5 ll ofsaturated Trypan blue to the final concentration of 600 ng/ll ofdsRNA. Pre-adult female and male lice were collected with forcepsfrom anesthetised salmon. Pre-adult II female lice were then

injected with 1 ll of the dsRNA solution in the cephalothorax usingcustom-made injection needles. These were pulled by utilising a1 mm Borosilicate glass tube with an inner diameter of 0.5 mm(Sutter Instrument, Novato, CA, USA) on the P-2000 laser-basedmicropipette puller system (Sutter Instrument). Needles wereground and opened using a Micropipette Grinder EG-44 (TritechResearch, Los Angeles, CA, USA), and coupled to a microinjectorbefore use. By blowing air into the needle, the dsRNA fragmentswere dispersed in the louse, visualised by dispersion of blue colourwithin the cephalothorax. After injection, the lice were kept in sea-water for 6 h before they were placed on anesthetised fish togetherwith male lice, in a 1:1 ratio (female n = 13). Three parallel exper-iments were set up for each gene. Lice were kept on one salmon,each in single fish tanks (50 L) with seawater for either 2, 4 or12 days, or until the female adults had produced a second set ofegg strings (approximately 38 days), when the remaining lice wereremoved from the fish. Lice were harvested at different time pointsin order to detect any reduction in mRNA levels and to study thefunction in sexually mature lice. Egg strings, when present, werecollected and placed in individual incubators for hatching. Live licewere transferred and stored on either RNAlater™ (Ambion Inc.) forRTq-PCR, fixed in phosphate buffered 4% formaldehyde at 4 �Covernight for in situ hybridisation or fixed for light microscopy(see Section 2.7). To confirm hatching, egg strings were observeddaily. Phenotypes were evaluated throughout nauplia and thecopepodid stages. The number of recovered lice from each experi-ment is listed in Table 3.

2.7. Histology

Specimens for light microscopy were fixed by immersion in amixture of 10 ml of 10% formaldehyde (fresh from paraformalde-hyde), 10 ml of 25% glutaraldehyde, 20 ml of 0.2 M cacodylatebuffer and 60 ml of PBS, and the pH was adjusted to 7.35. Thereaf-

Table 2List of amino acid (aa) sequences from all species used to determine the phylogenetic relationship of Lepeophtheirus salmonis ecdysone receptor (LsEcR).

Classification Species EMBL Accession No. Product size (aa)

Annelida Platynereis dumerilii ACC94156 496Chelicerata Liocheles australasia (Australian rainforest scorpion) AB297929 539

Agelena silvatica GQ281317 533Ornithodoros moubata AB191193 567Amblyomma americanum (Lone star tick) isoform1 AF020187 560

Crustacea L. salmonis KP100057 536Tigriopus japonicus ADD82902.1 546Penacus japonicus AB295492 499Uca pugilator (Sand fiddler crab) AF034086 518Amphiascus tenuiremis JF926564 458Homarus americanus (American lobster) HQ335007 541Daphnia magna (Water flee) isoform1 AB274821 693Portunus trituberculatus (Gazami crab) JQ250795 503

Hexapoda Locusta migratoria (Migratory locust) AF049136 541Gryllus firmus (Sand cricket) GU289704 416Apis mellifera (Honey bee) AB267886 567Drosophila melanogaster (Fruit fly) isoformB1 NP_724460 878Tribolium castaneum (Flour beetle) isoformA CM000284 549Aedes aegyptii (Yellow fever mosquito) AY345989 776Diploptera punctata isoformA JQ229679 538

Mollusca Crassostrea gigas (Pacific oyster) EKC19773.1 471Lymnaea stagnalis (Great pond snail) ADF43963.1 478

Nematoda Caenorhabditis elegans NP_492615.2 373Trichinella spiralis XP_003376657.1 573Ascaris suum ADY42534.1 496

Plathyhelminths Schmidtea mediterranea AFF18489 655Schistosoma mansoni ARR29357.1 715

Vertebrata Salmo salar (Atlantic salmon) FJ470290 462Xenopus tropicalis NP_001072853.1 441Crotatus adamantus (Pit viper) AFJ50856.1 435

178 L. Sandlund et al. / International Journal for Parasitology 45 (2015) 175–185

ter specimens were rinsed in PBS and dehydrated in a series of eth-anol solutions (50%, 70% and 96%), before being embedded in Tech-novit 7100 (Heraeus Kulzer GmbH & Co, Germany). Sections (1–2 lm) were stained with Toluidine blue.

Digital micrographs were acquired with a ColorView III camera(Soft Imaging System GmbH, Münster, Germany) mounted on anOlympus BX61 Microscope (Olympus, Tokyo, Japan), and processedusing Adobe Photoshop CS6 (Adobe Systems, San Jose, California,USA).

2.8. Detection of transcript levels in dsRN- treated lice by RTq-PCR

RTq-PCRs using TaqMan� probes (Table 1) were used to detecttotal expression of LsEcR, Vitellogenin-1 (LsVit-1), Vitellogenin-2(LsVit-2), ecdysone induced protein 75 (LsE75) and Broad-Complex(LsBr-C) from dsRNA-treated lice harvested from the RNAi experi-ments (samples listed in Table 3). Total RNA was isolated and sam-ples prepared as described in Section 2.4. Two micrograms of cDNAfrom RNAi lice were added to the RTq-PCR mix (Applied Biosys-tems�) to a total volume of 10 ll. Each louse was analysed sepa-rately as described in Section 2.4. The number of lice analysedfrom each RNAi experiment is listed in Table 3.

2.9. Statistical analysis

From the RNAi experiments, significant differences between thecontrol groups and the treated groups were determined using theKolmogrov–Smirnov test (non-parametric, un-paired: comparedcumulative distributions) by employing Prism6 software (Graph-Pad Software, Inc., La Jolla, CA, USA). Statistical evaluation of themRNA LsEcR splice variant at specific life stages was performedby two-way ANOVA analysis utilising SPSS software V. 21 (IBM�

SPSS� Statistics, Armonk, NY, USA).

3. Results

3.1. Sequence analysis and molecular phylogeny of the LsEcR

In order to obtain full-length LsEcR cDNA, 50 and 30 Rapid ampli-fication of cDNA ends (RACE) PCR was run using EcR-specific prim-ers (Table 1, EcR specific_P1 and EcR specific_P3) based onexpressed sequence tag (EST) sequences. A 2932 bp cDNA wasretrieved with a 50 UTR of 1044 bp, a 280 bp 30 UTR and a1608 bp ORF consisting of one exon, encoding 536 aa. The pre-dicted molecular weight was 60.4 kDa (Expasy, ProtParam Tool,http://web.expasy.org/protparam/). Cloning and sequencing ofthe RACE products revealed the existence of three mRNA variants,LsEcRa, LsEcRb and LsEcRc, differing in their 50UTR (Fig. 1). A BLASTsearch revealed the deduced protein sequence encodes the EcR of L.salmonis and exhibits 61% identity to the full-length aa sequenceand 82% and 77% for the DBD and LBD, respectively, with the cope-pod T. japonicus (ADD82902.1). The deduced aa sequence of LsEcRcontained domains characteristic of nuclear receptors, namely anA/B domain associated with transcriptional activation, DBD(C-domain, aa 170–261), a hinge region (D-domain) and a LBD(E/F-domain, aa 303–535) containing a short aa sequence(ATGMRA) recognised as the activation factor-2 domain (AF-2; aa235–240). Alignments of the domains to conserved domains inNCBI (Marchler-Bauer et al., 2011) proved the retrieved cDNAsequence form L. salmonis encodes the nuclear receptor LsEcR.

Phylogenetic analysis of the aa sequence of LsEcR was per-formed by conducting a Bayesian analysis of a full-length aa align-ment of EcR and EcR-like receptors from a variety of species (listedin Table 2). From the rooted bootstrap tree (Fig. 2), LsEcR groupedtogether with the copepods T. japonicus and A. tenuiremis and wasseparated from the decapods. The water flea D. magna (Branchio-poda) EcR form a separate clade and is the closest sister group tothe copepods, followed by Hexapoda and Chelicerata.

Table 3Summary of recovered lice and phenotypic traits observed using RNA interference experiments.

Recovered female lice Blood in intestine Lice producing egg strings RTq-PCRd

Control: Fragment 1: 2 daysa 10 Not registered – 9dsRNA: Fragment 1: 2 daysa 10 Not registered – 6Control: Fragment 1: 4 daysa 7 Not registered – 6dsRNA: Fragment 1: 4 daysa 8 Not registered – 7Control: Fragment 1: 12 daysa 13 11 – 10dsRNA: Fragment 1: 12 daysa 19 6 – 17Control: Fragment 1: 38 days 23 23 23 10dsRNA: Fragment 1: 38 days 14 10 (7b) 1c 13Control: Fragment 2: 38 days 16 16 16 10dsRNA: Fragment 2: 38 days 16 5 (3b) 0 11

dsRNA, double-stranded RNA.a Lice had not reached mature adult stage, hence no egg string production.b Barely visible blood in intestine.c Egg strings did not hatch.d Number of lice submitted to real-time quantitative PCR (RTq-PCR).

ORF1608 197 294

3`UTR

2616 Ex 5 Ex 1 Ex 2 Ex 3 Ex 4

373 14403 6143 14426

134 113

5`UTR

187

START

Fig. 1. Schematic representation of the genomic sequence of the Lepeophtheirus salmonis ecdysone receptor (LsEcR). Cloning and sequencing revealed the presence of threedifferent mRNA variations. The exons (Ex; depicted in light grey boxes) were mapped to the genomic DNA and show the gene to extend over 38.5 kbp. The open reading frame(ORF) is depicted in dark grey and the translation start site is marked with an arrow. Introns are depicted as lines between exons with lengths in numbers of nucleotides. Thethree mRNA variants are represented with connecting lines; mRNA LsEcRa consists of exons 1, 2 and 5; mRNA LsEcRb of exons 3 and 5 and mRNA LsEcRc of exons 4 and 5. AllmRNA variations share a common exon (Ex 5) linked directly to the coding sequence consisting of only one exon and no introns.

L. Sandlund et al. / International Journal for Parasitology 45 (2015) 175–185 179

3.2. Expression pattern analysis of 50UTR mRNA splice variants ofLsEcR

The expression pattern of the different mRNA splice variants ofLsEcR was evaluated in different developmental stages in L. salmo-nis. Thus, ontogenetic analysis was performed using RTq-PCR onRNA extracted from nauplia I/II, free-living copepodids, parasiticcopepodids, chalimus I, chalimus II, pre-adult male I/II, pre-adultfemale I/II, adult male and immature adult female lice, and gravidfemale lice. Specific Taqman� assays (Table 1) were designed to dis-criminate between the three 50UTRmRNA splice variants (Fig. 3). Ingeneral, the highest relative expression was detected in the naupliaI/II and free-living copepodids for all three splice variants with LsE-cRa and LsEcRc significantly more highly expressed compared withLsEcRb. The expression pattern decreased from copepodid to thechalimi stages before an increased expression occurred in the pre-adult and adult stages, with the relative expression of LsEcRa beingsignificantly higher compared with LsEcRb and LsEcRc. The expres-sion of LsEcRawas significantly higher in immature and gravid adultfemale lice compared with pre-adult female/male and adult malelice.

3.3. LsEcR transcript is expressed in a variety of tissues

In situ hybridisation analysis performed on paraffin sections ofan adult female louse demonstrated that LsEcR transcript waspresent in most tissues except for the muscle tissue (Fig. 4B–E).Expression was observed in the ovaries, immature/mature eggspresent in the genital segment (Fig. 4B), different glandular tissuesof unknown function present in the legs (Fig. 4C) and the anteriorpart of the cephalothorax, intestine (Fig. 4D) and in the sub-cuticular tissue (Fig. 4E). Unspecific colouring of the outer cuticulartissue was observed both for the sense and anti-sense probes.

3.4. Down-regulation of LsEcR by RNAi inhibits the production ofoffspring

Functional studies using RNAi were performed to assess theeffect of LsEcR in reproduction of the salmon louse. First, an exper-iment was set up using fragment 1 (Table 1) to determine thedegree of down-regulation in L. salmonis. In total, 39 pre-adult IIfemale lice were injected with a dsLsEcR fragment and 39 wereinjected with dsRNA from a cod and left on the fish for 2, 4 and

Schmidtea mediterranea HNF4

77

Caenorhabditis elegans NRH69

Platynereis dumerilii EcR Schistosoma mansoni NR1Lymnaea stagnalis EcR

Diploptera punctata EcRA Gryllus firmus EcR Locusta migratoria EcR Tribolium castaneum EcRA Apis mellifera EcRA Aedes aegyptii EcR Drosophila melanogaster EcRB1Daphnia magna EcRA1

Liocheles australasia EcRAmblyomma americanum EcRA1

Omithodoros moubata EcR

Agelena silvatica EcR

Trichinella spiralis EcR Ascaris suum EcR

Crassostrea gigas EcR

Portunus trituberculatus EcR Uca pugilator EcR Homarus americanus EcRsplice Penacus japonicus EcR

Lepeophtheirus salmonis EcR Tigriopus japonicus EcR Amphiascus tenuiremis EcR

Salmo salar LXR Xenopus tropicalis NR1H

Crotatus adamantus LXRbetalike

97

91

62

55

77

99

5388

Chelicerata

Hexapoda

Vertebrata

Copepoda

Crustacea

0.2

Fig. 2. Phylogenetic analysis of Lepeophtheirus salmonis ecdysone receptor (LsEcR). A rooted phylogenetic tree of amino acid sequences of full-length EcR and EcR-likereceptors from different species was generated using Bayesian methods. Branch length is proportional to sequence divergence. Branch numbers and bars represent bootstrapvalues in percent and 0.2 substitutions per site, respectively. LsEcR is marked with an arrow. Salmo salar LXR, Xenopus tropicalis NR1H and Crotatus adamantus LXRbetalikereceptors were used as an outgroup. The GenBank accession numbers of all EcR sequences used are listed in Table. 2.

180 L. Sandlund et al. / International Journal for Parasitology 45 (2015) 175–185

0.0

10.0

20.0

30.0

40.0

50.0

60.0

70.0

80.0

90.0

100.0

110.0

Rel

ativ

e ex

pres

sion

LsEcRa LsEcRb

LsEcRc

Fig. 3. Quantitative real-time PCR (RTq-PCR) analysis of relative expression of the three mRNA Lepeophtheirus salmonis ecdysone receptor (LsEcR) variants (a, b, c) in differentdevelopmental stages. Each point represents the mean and confidence intervals (n = 5 parallels of approximately 150 animals for the nauplia and free-living copepodid (Free.Cop.) stages, 10 animals for the parasitic copepodid (Par. Cop.), chalimus I (Chal. I) and chalimus II (Chal. II.) and one animal for each of the pre-adult male (Pre-A. M.), pre-adult female (Pre-A. F.), adult male (Adult M.), adult female (Adult F.) and gravid adult female (Adult F. gravid)) stages. The relative expression of LsEcRb at the chalimus IIstage was set to 1.

Fig. 4. Localisation of Lepeophtheirus salmonis ecdysone receptor (LsEcR) transcripts in an adult female louse. (A) Light microscope image of gravid adult female louse. Lettersand asterisks are guides to the corresponding photos of individual tissues. A part of the sub-cuticular tissue is framed to better visualise localisation. (B–E) In situhybridisation using LsEcR-specific anti-sense RNA was used for detection of transcript. Negative controls (sense RNA) are shown (insets). Positive staining was seen in matureeggs (B), unidentified glandular and surrounding tissue in the legs (C), intestine (D) and sub-cuticular tissue (E). Unspecific colouring of the outer cuticular layer was seenusing both sense and anti-sense probes. Scale bar = 5 mm (A); 200 lM (B, D, E) and 100 lM (C).

L. Sandlund et al. / International Journal for Parasitology 45 (2015) 175–185 181

12 days. No phenotype or reduced survival rate was observed forthe immature female animals compared with the control group;however, RTq-PCR showed that the LsEcR-gene was significantlyknocked down (by 53% at day 12; Fig. 5B). At days 2 and 4, how-ever, no significant knock-down was observed (Fig. 5A). A secondexperiment was set up using two different fragments in order toexclude any non-target effects. Fragments 1 and 2 were injectedand run in two separate experiments for 38 days under the samecriteria as the first experiment. RTq-PCR analysis of the secondexperiments terminated at 38 days and did not detect any signifi-cant regulation of the LsEcR (Fig. 5C). However, at 38 days the dsL-sEcR-treated lice showed a characteristic phenotype where noproduction of offspring was observed. Female lice injected withdsRNA from cod had no phenotype and produced viable offspring(Fig. 6A, D). We also observed that dsLsEcR-treated lice were foundwith less blood in the intestine (Table 3 and Fig. 6D), which

deviates from what is observed to be normal in adult female licein our laboratory system, where most females have a blood-filledgut (Table 3 and Fig. 6A). Histological sections from dsLsEcR-treated lice revealed that the oocytes did not display a normalstacking pattern like the control lice and an individual ova couldnot be detected (Fig. 6B, E). The lining of the developing oocyteswas disintegrated, leaving the area filled with a mesh of fat andyolk granules. The cellular structure of the sub-cuticular tissuewas observed to be hypotrophic compared with the control lice(Fig. 6C, F) giving an impression of a reduction in tissue.

The binding of ecdysone to the EcR/USP complex is known toregulate several down-stream genes. The expression level of theknown down-stream genes LsE75 and LsBr-C was evaluated in liceafter dsRNA from LsEcR was injected. No significant regulation wasdetected for LsE75 or LsBr-C from the dsLsEcR-treated lice after12 days (Fig. 5B). In contrast, both LsE75 and LsBr-C were up-regulated in dsLsEcR-treated lice after 38 days (Fig. 5C) (Kolmogo-rov–Smirnov, P < 0.05). RTq-PCR analysis was also conducted onLsVit-1 and LsVit-2 from lice treated for 38 days and both geneswere significantly knocked down (Fig. 5C) (Kolmogorov–Smirnov,P < 0.05). The expression of the two vitellogenins was only evalu-ated in lice treated for 38 days as LsVit-1 and LsVit-2 are onlyexpressed in mature female lice (Eichner et al., 2008).

4. Discussion

In the present study, we isolated a cDNA for the EcR in L. salmo-nis. The genetic composition of LsEcR proved to be similar to theEcR gene found in T. japonicus (Hwang et al., 2010) with only oneexon spanning the ORF and with introns only detected in the UTRs.Putative full-length protein sequence alignment (Table 1) and phy-logenetic analysis (Fig. 2) cluster the LsEcR together with the cope-pods T. japonicus and A. tenuriemis in the Malacostraca group, withthe water flee D. magna (Branchiopoda) as the closest sister group.Identical aa found in the LBD of EcRs between species areconsistent with the widespread use of ecdysone as the hormoneinitiating developmental processes. Identity searches and determi-nation of phylogenetic position of the retrieved L. salmonis cDNAsequence classify it as an ecdysone receptor.

The 50UTR region of the retrieved cDNA revealed the existenceof three LsEcR mRNA splice variants, all starting from differentexons. This suggests that those are regulated by different promoterregions. Selective promoter regions are well known from steroidhormone receptors such as the human oestrogen receptor (ER)(Kwak et al., 1993; Bockmuhl et al., 2011) and have been shownto possess different tissue specificity and to be activated by differ-ent signals (Ayoubi and VanDeVen, 1996). The mechanismsinvolved in 50UTR mediated regulation is poorly understood andhas to our knowledge not been studied in crustaceans. It is possiblethat the LsEcR mRNA splice variants are expressed in different tis-sues or regulated by specific signals in the salmon louse. However,further studies are required in order to understand how the differ-ent LsEcR mRNA splice variants are regulated.

The existence of multiple EcR isoforms that differ in their spatialand temporal expression are common in many crustacean species(Durica et al., 1999; Tarrant et al., 2011; Verhaegen et al., 2011).Expression profiling using RTq-PCR, performed on embryos andadults of the water flea (D. magna), revealed that the EcRB isoformwas expressed at a higher level during embryogenesis comparedwith EcRA, while the opposite expression pattern was observedin adult fleas during molting (Kato et al., 2007). In the salmonlouse, of the three variants of LsEcR mRNA transcripts present,the LsEcRb variant had relatively low expression throughout all lifestages compared with LsEcRa and LsEcRc that were observed tohave the highest relative expression in the nauplia I/II and the

0

0.2

0.4

0.6

0.8

1

1.2

1.4

LsEcR LsE75 LsBr-C

Rel

ativ

e ex

pres

sion

Transcript level 12 d. post dsRNA treatment

Control

RNAi

*

0.0

1.0

2.0

3.0

4.0

5.0

6.0

LsEcR LsE75 LsBr-C LsVit-1 LsVit-2

Rel

ativ

e ex

pres

sion

Transcript level 38 d. post dsRNA treatment

Control

RNAi

* *

*

C

*

B

0

0.2

0.4

0.6

0.8

1

1.2

1.4

LsEcR 2 .d.p.i LsEcR 4. d.p.i.

Rel

ativ

e ex

pres

sion

Transcript level 2 and 4 d. post dsRNA treatment

Control

RNAi

A

Fig. 5. Transcript level of selected ecdysone receptor (EcR) target genes afterinjection of double-stranded (ds)RNA. Lepeophtheirus salmonis EcR (LsEcR) adultfemale lice were removed from anesthetised fish and analysed after (A) 2, 4, (B) 12or (C) 38 days (d.) post treatment (i.e. dsEcR injection; d.p.i.). Quantitative real-timePCR (RTq-PCR) analysis of the relative expression of LsEcR and selected downstreamgenes LsE75 and LsBroad-complex (LsBr-C) (B, C) LsVitellogenin-1 (LsVit-1) andLsVitellogenin-2 (LsVit-2) (C) was evaluated. (C) The graph is representative of twoexperiments. The expression levels of the respective genes in the control groupswere set to 1. Mean ± confidence interval of treated lice is shown. Each louse wasanalysed separately and confidence intervals represent individual differences.Numbers of lice analysed are listed in Table. 3. ⁄Statistically significant (P < 0.05).Statistical analysis was performed using a Kolmogorov–Smirnov test.

182 L. Sandlund et al. / International Journal for Parasitology 45 (2015) 175–185

free-living copepodid stage (Fig. 3). A similar expression patternwas observed for the EcR in the free-living copepod T. japonicus(Hwang et al., 2010). A second peak in expression was observedfor LsEcRa in immature (T1) and mature females (T6) (classificationof maturing female louse after Eichner et al. (2008)) which couldindicate that the LsEcRa transcript is used more predominantly infemale maturation and reproduction. Overall, the differentialexpression of the three LsEcR mRNAs could suggest that those playdifferent roles in different biological processes.

To investigate the spatial distribution of LsEcR transcript in theadult female louse, we performed in situ hybridisation. With ourprotocol, the presence of LsEcR transcript was, with the exceptionof muscle tissue, evident in most tissues such as glandular andsub-cuticular tissues and oocytes (Fig. 4A–E). The wide distributionof EcR transcripts has similarly been demonstrated in the kurumaprawn Marsupenaeus japonicus (Mj) and the soft tick Ornithodorosmoubata (Om) using RTq-PCR and RT-PCR, respectively (Asazumaet al., 2007). In contrast to our results, MjEcR and OmEcR were alsodetected, in low quantities, in muscle tissue (Asazuma et al., 2007),however this may be explained by the difference in sensitivitybetween the methods.

From our knock-down studies of LsEcR in reproducing femalelice, it has become apparent that the nuclear receptor eitherdirectly or indirectly affects a variety of biological processes. Inthe salmon louse, the sub-cuticular region has been demonstratedto be an active tissue with functions similar to the liver (Edvardsenet al., in press). Yolk proteins such as the vitellogenins are pro-duced in the sub-cuticular tissue before they are incorporated intothe oocyte during oocyte maturation (Dalvin et al., 2009, 2011).The reduction of vitellogenin 1 and 2 transcripts observed in dsL-sEcR knock-down lice (Fig. 5C) could most likely be explained bythe major changes occurring in the sub-cuticular tissue (Fig. 6F).At the same time, when depriving the female lice of LsEcR, repro-duction was halted and eggs failed to mature in the genital seg-ment. Similar observations were reported from EcR knock-downstudies of T. castaneum, where a 50–75% reduction in the vitello-genin transcript level resulted in a decrease in egg development

(Xu et al., 2010). Moreover, development of a follicular cell layernecessary for oocyte maturation was disrupted, resulting in anarrest of the oocyte in the pre-vitellogenic stage (Parthasarathyet al., 2010). The same observations had previously been recordedin D. melanogaster where EcR deficiency resulted in abnormal eggchamber development and loss of vitellogenic stages (Carney andBender, 2000). Loss of egg production in L. salmonis is presumablynot a function of reduced yolk production, but either a direct orindirect of effect of LsEcR depletion in the oocytes.

In insects, eggs mature in the ovaries to gametes that contain allthe proteins and maternal mRNA needed to initiate and maintainmetabolism and development of the eggs before fertilisation. Fromwork performed in D. melanogaster and T. castaneum, it was shownthat components of the ecdysone hierarchy such as EcR wereexpressed and required in germline cells for progression throughoogenesis (Buszczak et al., 1999; Carney and Bender, 2000;Freeman et al., 1999). The observation of LsEcR transcript in theoocyte implies the presence of maternal transcript in the eggs.The absence of normal egg development in dsLsEcR-treated liceprovides a good indication that the Ec-EcR pathway plays animportant role in reproduction and development of offspring inthe salmon louse. The specific mechanism for loss of egg develop-ment is currently unknown and further studies are necessary tounderstand the complexity of the Ec-EcR hormonal pathway.

RNAi is a well established genetic tool for functional studies indifferent organisms. However, with the exception of plants and thenematode Caenorhabditis elegans, little is known about the sys-temic RNAi response mechanisms in non-traditional model organ-isms (Miller et al., 2012). In the dsLsEcR-treated lice, significantdown-regulation of LsEcR was not observed until 12 days afterinjection. Our results deviate from knock-down studies performedon the putative prostaglandin E synthase 2 (LsPGES2) in L. salmonis,where reduction in the transcript level was most prominent after48–72 h (Campbell et al., 2009). However, it should be noted thatthe optimal requirements for knock-down differ among genesdepending their locations and turnover rates. After 38 days, theRNAi effect had ceased but a distinct phenotype, only observed

Fig. 6. Functional assessment of the Lepeophtheirus salmonis ecdysone receptor (LsEcR) by RNA interference (RNAi). The control lice produced normal egg strings (A) thathatched and produced viable offspring. LsEcR dsRNA-treated lice (D) showed a distinct phenotype with no production of eggs. It was also observed that the LsEcR dsRNA-treated lice attained a thicker genital segment as well as less blood in the intestine (marked with arrow, D), compared with the control (marked with arrow, A). (B and C, E andF) Toluidine stained sections showed the normal stacking pattern of the eggs seen in the control (B) which was lost in the dsRNA LsEcR-treated lice (E). The sub-cuticulartissue was hypotrophic in the LsEcR dsRNA-treated lice (F) compared with the control lice (C). Scale bar = 5 mm (A, D), 200 lM (B, E) and 1000 lM (C, F).

L. Sandlund et al. / International Journal for Parasitology 45 (2015) 175–185 183

after the prolonged period of LsEcR knock-down in adult females,was evident. At the same time, the LsEcR response genes LsE75and LsBr-C were significantly up-regulated in the dsLsEcR-treatedlice compared with the control lice. The increased expression ofthe response genes could be a secondary response as a cause ofLsEcR depletion and disruption of several biological processes.However, as these genes naturally have very irregular expressionpatterns, further studies are necessary in order to determine therelation between ecdysone pathway and the response genes in L.salmonis.

In summary, we report the identification of an EcR from the sal-mon louse L. salmonis and demonstrate the presence of the LsEcRtranscript in all life stages of the parasite. In situ hybridisation,together with functional knock-down studies, indicates that theLsEcR plays a key role in regulation of female reproduction andoocyte maturation. The Ec-EcR hierarchy is a very complex systemwith a multitude of factors interacting through different pathways.The essential role EcR plays in this hierarchy makes it a good targetfor pesticide development, as knock-down of EcR results in severephysiological changes in the animal, including the termination ofegg production. However, further studies are necessary in orderto elucidate the functional role of LsEcR and to fully understandthe complexity of the Ec-EcR hierarchy in the salmon louse.

Acknowledgements

We would like to thank Lars Hamre, Per Gunnar Espedal andTeresa Cieplinska for excellent technical assistance in the labora-tory. This research has been funded by the Research Council ofNorway, SFI-Sea Lice Research Center, Grant Number 203513.

References

Asazuma, H., Nagata, S., Kono, M., Nagasawa, H., 2007. Molecular cloning andexpression analysis of ecdysone receptor and retinoid X receptor from thekuruma prawn, Marsupenaeus japonicus. Comp. Biochem. Physiol. B 148, 139–150.

Ayoubi, T.A.Y., VanDeVen, W.J.M., 1996. Regulation of gene expression byalternative promoters. FASEB J 10, 453–460.

Belles, X., Piulachs, M.-D., 2014. Ecdysone signalling and ovarian development ininsects: from stem cells to ovarian follicle formation. Biochim. Biophys. Acta.http://dx.doi.org/10.1016/j.bbagrm.2014.05.025.

Billas, I.M.L., Browning, C., Lawrence, M.C., Graham, L.D., Moras, D., Hill, R.J., 2009.The structure and function of ecdysone receptors. In: Smagghe, G. (Ed.),Ecdysteroids. Structures and Functions. Springer, Dordrecht, Netherlands, pp.335–375.

Bockmuhl, Y., Murgatroyd, C.A., Kuczynska, A., Adcock, I.M., Almeida, O.F., Spengler,D., 2011. Differential regulation and function of 50-untranslated GR-exon 1transcripts. Mol. Endocrinol. 25, 1100–1110.

Buszczak, M., Freeman, M.R., Carlson, J.R., Bender, M., Cooley, L., Segraves, W.A.,1999. Ecdysone response genes govern egg chamber development during mid-oogenesis in Drosophila. Development 126, 4581–4589.

Campbell, E.M., Pert, C.C., Bowman, A.S., 2009. RNA-interference methods for gene-knockdown in the sea louse, Lepeophtheirus salmonis: studies on a putativeprostaglandin E synthase. Parasitology 136, 867–874.

Carney, G.E., Bender, M., 2000. The Drosophila ecdysone receptor (EcR) gene isrequired maternally for normal oogenesis. Genetics 154 (1203–1), 211.

Dalvin, S., Frost, P., Biering, E., Hamre, L.A., Eichner, C., Krossoy, B., Nilsen, F., 2009.Functional characterisation of the maternal yolk-associated protein (LsYAP)utilising systemic RNA interference in the salmon louse (Lepeophtheirussalmonis) (Crustacea: Copepoda). Int. J. Parasitol. 39, 1407–1415.

Dalvin, S., Frost, P., Loeffen, P., Skern-Mauritzen, R., Baban, J., Ronnestad, I., Nilsen, F.,2011. Characterisation of two vitellogenins in the salmon louse Lepeophtheirussalmonis: molecular, functional and evolutional analysis. Dis. Aquat. Organ. 94,211–224.

De Wilde, R., Swevers, L., Soin, T., Christiaens, O., Rouge, P., Cooreman, K., Janssen,C.R., Smagghe, G., 2013. Cloning and functional analysis of the ecdysteroidreceptor complex in the opossum shrimp Neomysis integer (Leach, 1814). Aquat.Toxicol. 130–131, 31–40.

Dhadialla, T.S., Carlson, G.R., Le, D.P., 1998. New insecticides with ecdysteroidal andjuvenile hormone activity. Annu. Rev. Entomol. 43, 545–569.

Durica, D.S., Chung, A.C.K., Hopkins, P.M., 1999. Characterization of EcR and RXRgene homologs and receptor expression during the molt cycle in the crab, Ucapugilator. Am. Zool. 39, 758–773.

Edvardsen, R.B., Dalvin, S., Furmanek, T., Malde, K., Mæhle, S., Kvamme, B.O., Skern-Mauritzen, R., in press. Gene expression in five salmon louse (Lepeophtheirussalmonis, Krøyer 1837) tissues. Mar. Genomics. http://dx.doi.org/10.1016/j.margen.2014.06.008.

Eichner, C., Frost, P., Dysvik, B., Jonassen, I., Kristiansen, B., Nilsen, F., 2008. Salmonlouse (Lepeophtheirus salmonis) transcriptomes during post molting maturationand egg production, revealed using EST-sequencing and microarray analysis.BMC Genomics 9, 126. http://dx.doi.org/10.1186/1471-2164-9-126.

Espedal, P.G., Glover, K.A., Horsberg, T.E., Nilsen, F., 2013. Emamectin benzoateresistance and fitness in laboratory reared salmon lice (Lepeophtheirussalmonis). Aquaculture 416, 111–118.

Evans, R.M., 1988. The steroid and thyroid hormone receptor superfamily. Science240, 889–895.

Fallang, A., Ramsay, J.M., Sevatdal, S., Burka, J.F., Jewess, P., Hammell, K.L., Horsberg,T.E., 2004. Evidence for occurrence of an organophosphate-resistant type ofacetylcholinesterase in strains of sea lice (Lepeophtheirus salmonis Kroyer). PestManag. Sci. 60, 1163–1170.

Freeman, M.R., Dobritsa, A., Gaines, P., Segraves, W.A., Carlson, J.R., 1999. The daregene: steroid hormone production, olfactory behavior, and neural degenerationin Drosophila. Development 126, 4591–4602.

Frost, P., Nilsen, F., 2003. Validation of reference genes for transcription profiling inthe salmon louse, Lepeophtheirus salmonis, by quantitative real-time PCR. Vet.Parasitol. 118, 169–174.

Gaertner, K., Chandler, G.T., Quattro, J., Ferguson, P.L., Sabo-Attwood, T., 2012.Identification and expression of the ecdysone receptor in the harpacticoidcopepod, Amphiascus tenuiremis, in response to fipronil. Ecotoxicol. Environ. Saf.76, 39–45.

Gilbert, L.I., Song, Q., Rybczynski, R., 1997. Control of ecdysteroidogenesis:activation and inhibition of prothoracic gland activity. Invert. Neurosci. 3,205–216.

Hamre, L.A., Glover, K.A., Nilsen, F., 2009. Establishment and characterisation ofsalmon louse (Lepeophtheirus salmonis (Kroyer 1837)) laboratory strains.Parasitol. Int. 58, 451–460.

Heuch, P.A., Bjorn, P.A., Finstad, B., Holst, J.C., Asplin, L., Nilsen, F., 2005. A review ofthe Norwegian ‘National Action Plan Against Salmon Lice on Salmonids’: theeffect on wild salmonids. Aquaculture 246, 79–92.

Hill, R.J., Graham, L.D., Turner, K.A., Howell, L., Tohidi-Esfahani, D., Fernley, R.,Grusovin, J., Ren, B., Pilling, P., Lu, L., Phan, T., Lovrecz, G.O., Pollard, M., Pawlak-Skrzecz, A., Streltsov, V.A., Peat, T.S., Winkler, D.A., Lawrence, M.C., 2012.Chapter Four – structure and function of ecdysone receptors—interactions withecdysteroids and synthetic agonists. Adv. Insect Physiol. 43, 299–351.

Hill, R.J., Billas, I.M.L., Bonneton, F., Graham, L.D., Lawrence, M.C., 2013. Ecdysonereceptors: from the Ashburner model to structural biology. Annu. Rev. Entomol.58 (58), 251–271.

Hirano, M., Ishibashi, H., Kim, J.W., Matsumura, N., Arizono, K., 2009. Effects ofenvironmentally relevant concentrations of nonylphenol on growth and 20-hydroxyecdysone levels in mysid crustacean, Americamysis bahia. Comp.Biochem. Physiol. C: Toxicol. Pharmacol. 149, 368–373.

Hodin, J., Riddiford, L.M., 1998. The ecdysone receptor and ultraspiracle regulate thetiming and progression of ovarian morphogenesis during Drosophilametamorphosis. Dev. Genes. Evol. 208, 304–317.

Hopkins, P.M., Durica, D., Washington, T., 2008. RXR isoforms and endogenousretinoids in the fiddler crab, Uca pugilator. Comp. Biochem. Physiol. A 151, 602–614.

Huelsenbeck, J.P., Ronquist, F., 2001. MRBAYES: Bayesian inference of phylogenetictrees. Bioinformatics 17, 754–755.

Hwang, D.S., Lee, J.S., Lee, K.W., Rhee, J.S., Han, J., Lee, J., Park, G.S., Lee, Y.M., Lee, J.S.,2010. Cloning and expression of ecdysone receptor (EcR) from the intertidalcopepod, Tigriopus japonicus. Comp. Biochem. Physiol. C 151, 303–312.

Kabata, Z., 1979. Parasitic Copepoda of British Fishes. The Ray Society, London.Kato, Y., Kobayashi, K., Oda, S., Tatarazako, N., Watanabe, H., Iguchi, T., 2007. Cloning

and characterization of the ecdysone receptor and ultraspiracle protein fromthe water flea Daphnia magna. J. Endocrinol. 193, 183–194.

Kvamme, B.O., Skern, R., Frost, P., Nilsen, F., 2004. Molecular characterisation of fivetrypsin-like peptidase transcripts from the salmon louse (Lepeophtheirussalmonis) intestine. Int. J. Parasitol. 34, 823–832.

Kwak, S.P., Patel, P.D., Thompson, R.C., Akil, H., Watson, S.J., 1993. 50-Heterogeneityof the mineralocorticoid receptor messenger-ribonucleic-acid – differentialexpression and regulation of splice variants within the rat hippocampus.Endocrinology 133, 2344–2350.

Marchler-Bauer, A., Lu, S., Anderson, J.B., Chitsaz, F., Derbyshire, M.K., DeWeese-Scott, C., Fong, J.H., Geer, L.Y., Geer, R.C., Gonzales, N.R., Gwadz, M., Hurwitz, D.I.,Jackson, J.D., Ke, Z., Lanczycki, C.J., Lu, F., Marchler, G.H., Mullokandov, M.,Omelchenko, M.V., Robertson, C.L., Song, J.S., Thanki, N., Yamashita, R.A., Zhang,D., Zhang, N., Zheng, C., Bryant, S.H., 2011. CDD: a Conserved Domain Databasefor the functional annotation of proteins. Nucleic Acids Res. 39, D225–D229.

Miller, S.C., Miyata, K., Brown, S.J., Tomoyasu, Y., 2012. Dissecting systemic RNAinterference in the red flour beetle Tribolium castaneum: parameters affectingthe efficiency of RNAi. PLoS ONE 7 (10), e47431. http://dx.doi.org/10.1371/journal.pone.0047431.

Mykles, D.L., 2011. Ecdysteroid metabolism in crustaceans. J. Steroid Biochem. Mol.Biol. 127, 196–203.

Parthasarathy, R., Palli, S.R., 2007. Stage- and cell-specific expression of ecdysonereceptors and ecdysone-induced transcription factors during midgutremodeling in the yellow fever mosquito, Aedes aegypti. J. Insect Physiol. 53,216–229.

184 L. Sandlund et al. / International Journal for Parasitology 45 (2015) 175–185

Parthasarathy, R., Sheng, Z.T., Sun, Z.Y., Palli, S.R., 2010. Ecdysteroid regulation ofovarian growth and oocyte maturation in the red flour beetle, Triboliumcastaneum. Insect Biochem. Molec. 40, 429–439.

Qian, Z.Y., He, S.L., Liu, T., Liu, Y.J., Hou, F.J., Liu, Q., Wang, X.Z., Mi, X., Wang, P., Liu,X.L., 2014. Identification of ecdysteroid signaling late-response genes fromdifferent tissues of the Pacific white shrimp, Litopenaeus vannamei. Comp.Biochem. Physiol. A 172, 10–30.

Riddiford, L.M., Hiruma, K., Zhou, X.F., Nelson, C.A., 2003. Insights into the molecularbasis of the hormonal control of molting and metamorphosis from Manducasexta and Drosophila melanogaster. Insect Biochem. Molec. 33, 1327–1338.

Ronquist, F., Teslenko, M., van der Mark, P., Ayres, D.L., Darling, A., Hohna, S., Larget,B., Liu, L., Suchard, M.A., Huelsenbeck, J.P., 2012. MrBayes 3.2: efficient Bayesianphylogenetic inference and model choice across a large model space. Syst. Biol.61, 539–542.

Shen, H.S., Zhou, X., Bai, A.X., Ren, X.F., Zhang, Y.P., 2013. Ecdysone receptor genefrom the freshwater prawn Macrobrachium nipponense: identification ofdifferent splice variants and sexually dimorphic expression, fluctuation ofexpression in the molt cycle and effect of eyestalk ablation. Gen. Comp.Endocrinol. 193, 86–94.

Skern-Mauritzen, R., Torrissen, O., Glover, K.A., 2014. Pacific and AtlanticLepeophtheirus salmonis (Kroyer, 1838) are allopatric subspecies:Lepeophtheirus salmonis salmonis and L. salmonis oncorhynchi subspecies novo.BMC Genet. 15 (1), 32.

Tarrant, A.M., Behrendt, L., Stegeman, J.J., Verslycke, T., 2011. Ecdysteroid receptorfrom the American lobster Homarus americanus: EcR/RXR isoform cloning andligand-binding properties. Gen. Comp. Endocrinol. 173, 346–355.

Thomas, H.E., Stunnenberg, H.G., Stewart, A.F., 1993. Heterodimerization of theDrosophila ecdysone receptor with retinoid-X receptor and ultraspiracle. Nature362, 471–475.

Thompson, J.D., Gibson, T.J., Plewniak, F., Jeanmougin, F., Higgins, D.G., 1997. TheCLUSTAL_X windows interface: flexible strategies for multiple sequencealignment aided by quality analysis tools. Nucleic Acids Res. 25, 4876–4882.

Thummel, C.S., 2002. Ecdysone-regulated puff genes 2000. Insect Biochem. Molec.32, 113–120.

Thummel, C.S., Chory, J., 2002. Steroid signaling in plants and insects – commonthemes, different pathways. Gene Dev. 16, 3113–3129.

Verhaegen, Y., Parmentier, K., Swevers, L., Renders, E., Rouge, P., De Coen, W.,Cooreman, K., Smagghe, G., 2011. The heterodimeric ecdysteroid receptorcomplex in the brown shrimp Crangon crangon: EcR and RXR isoformcharacteristics and sensitivity towards the marine pollutant tributyltin. Gen.Comp. Endocrinol. 172, 158–169.

Xu, J.J., Tan, A.J., Palli, S.R., 2010. The function of nuclear receptors in regulation offemale reproduction and embryogenesis in the red flour beetle, Triboliumcastaneum. J. Insect Physiol. 56, 1471–1480.

Yao, T.P., Segraves, W.A., Oro, A.E., Mckeown, M., Evans, R.M., 1992. Drosophilaultraspiracle modulates ecdysone receptor function via heterodimer formation.Cell 71, 63–72.

Yao, T.P., Forman, B.M., Jiang, Z.Y., Cherbas, L., Chen, J.D., Mckeown, M., Cherbas, P.,Evans, R.M., 1993. Functional ecdysone receptor is the product of Ecr andultraspiracle genes. Nature 366, 476–479.

L. Sandlund et al. / International Journal for Parasitology 45 (2015) 175–185 185

II

Accepted for publication after minor revisions

The Ecdysone Receptor (EcR) is a Major Regulator of Tissue Development and

Growth in the Marine Salmonid Ectoparasite, Lepeophtheirus salmonis

(Copepoda, Caligidae).

Authors and affiliations

Liv Sandlunda*, Frank Nilsena, Rune Maleb, Sussie Dalvinc

aSea Lice Research Centre, Department of Biology, University of Bergen, Norway

bSea Lice Research Centre, Department of Molecular Biology, University of Bergen,

Norway

cSea Lice Research Centre, Institute of Marine Research, Norway

*Corresponding author.

E-mail address: liv.sandlund@bio.uib.no (Liv Sandlund)

Abstract

The function of the ecdysone receptor (EcR) during development and molting has

been thoroughly investigated in in some arthropods such as insects but rarely in

crustacean copepods such as the salmon louse Lepeophtheirus salmonis (L. salmonis)

(Copepoda, Caligidae). The salmon louse is an ectoparasite on Atlantic salmon that

cause major economical expenses in aquaculture due to the cost of medical treatment

methods to remove lice from the fish. Handling of salmon louse infestations is further

complicated by development of resistance towards available medicines.

Understanding of basic molecular biological processes in the salmon louse is essential

to enable development of new tools to control the parasite. In this study, we found L.

salmonis EcR (LsEcR) transcript to be present in the neuronal somata of the brain,

nuclei of muscle fibers and the immature intestine. Furthermore, we explored the

function of LsEcR during development using RNA interference mediated knock-down

and through infection trials. Our results show that knock-down of LsEcR is associated

with hypotrophy of several tissues, delayed development and mortality. In addition,

combined knock-down of LsEcR/LsRXR resulted in molting arrest during early larval

stages.

Keywords: Copepod; Sea lice; Parasite; RNA interference; Ecdysone Receptor;

Molting

Abbreviations: L. salmonis, Lepeoptheirus salmonis; LsEcR, Lepeoptheirus salmonis

ecdysone receptor; LsRXR, Lepeoptheirus salmonis retinoid X receptor; RT-qPCR,

real-time quantitative PCR; RNAi, RNA interference; dsRNA, double stranded RNA;

1. Introduction

The salmon louse (Lepeophtheirus salmonis) is an ectoparasitic copepod on salmonid

fish with a life cycle that includes both free-living and parasitic stages. The louse

feeds on the skin, mucosa and blood of its host and can cause skin erosion and open

wounds that may lead to death if the parasite infection becomes severe. Due to its

high fecundity, salmon louse has become an increasing threat to the salmon farming

industry where high fish densities and all year round production provide favourable

conditions large numbers of the parasite [1, 2]. The free-living stages of the parasite

can spread between pens, different farming sites and also to wild salmonid fish

causing environmental concerns. Infestation of wild fish with salmon louse reared on

fish in aquaculture has been argued to cause reduction in wild populations nearby

aquaculture sites [3-7].

The salmon louse life cycle consists of eight developmental stages named nauplia I

and II, copepodid, chalimi I and II, pre-adult and the adult stage [8, 9] that are

separated by a molt. During molt a new exoskeleton is synthesized and synchronized

with shedding of the old exoskeleton. The molting process is a cyclic event that is

carefully regulated and requires the precise coordination of specific hormones to be

successful. The hormone considered as the main determinant of the molting process in

crustaceans is the steroid hormone 20-hydroxyecdysone (20E), reviewed by Chang et

al., 2011 [10]. The effect of 20E is exerted through a nuclear receptor complex

consisting of the ecdysone receptor (EcR) and the retinoid X receptor (RXR) referred

to as ultraspiracle (USP) in insects [11, 12]. This heterodimer also plays a key role in

the regulation of many other physiological processes such as development,

reproduction and limb generation [13-16]. Upon activation by ecdysteroids, the

EcR/RXR complex directly regulates a small set of “early” genes that further

regulates a larger set of “late” genes in a genetic hierarchy [17, 18]. Multiple splice

variants with different expression patterns of EcR and RXR have been identified in

both insect and crustaceans such as Drosophila melanogaster [19], Homarus

americanus [20], Daphnia magna [21] and Marsupenaeus japonicus [14]. Differences

in spatial and temporal expression patterns between the isoforms have shown to

correlate with specific responses and have distinct functions during development [22-

24]. In the salmon louse, three LsEcR transcripts differing only in their 5`UTR region

has been characterized [25] whereas several LsRXR splice variants of the open reading

frame were identified by Eichner et al., 2015 [26]. During metamorphic

reorganisation of the insect central nervous system, 20E regulates a wide variety of

cellular responses such as neuronal proliferation, maturation, cell death and

remodelling of larval neurons into their adult forms. Inactivation of the two

Drosophila EcR-B1 and EcR-B2 isoforms gave defects in the early stages of neuronal

remodelling during metamorphosis [22], while heat shock induced EcR null mutants

were arrested in late embryogenesis or unable to go through molt and metamorphosis

dependent on their developmental stage [23, 27].

The role of the EcR in developmental processes and molting is very well established

in insects and to some extent in decapods, however, molecular data regarding the

specific function of EcR in copepods is still limited [21]. RNA interference (RNAi)

has been successfully applied to investigate gene function in salmon louse and serves

as an important tool to understand the biology of the parasite [28-31]. Knock-down of

LsEcR in adult female salmon lice resulted in degeneration of vitellogenin producing

tissue and egg strings and therefor abolished of reproduction, proving the an

important role of LsEcR in female reproduction [25]. Due to the important role of

EcR in many vital biological processes as a ligand activated transcription factor, it is

important to explore the function of LsEcR in molting and development and as a

possible target for pesticides. In this study, we show that knock-down of LsEcR

expression in L. salmonis larvae results in comprehensive tissue damage and mortality

in later developmental stages. In addition, we show that simultaneous knock-down of

LsEcR and its partner LsRXR in nauplia interrupts the molting process. In addition,

knock-down of LsEcR indirectly or directly affects the expression of genes important

in chitin synthesis. This suggests that LsEcR plays a key role in molting, growth and

tissue development of the salmon louse.

2. Materials and methods

2.1. Animals

Eggs from the Atlantic salmon louse Lepeophtheirus salmonis [32] were hatched in

wells with flow-through seawater and kept in culture on Atlantic salmon (Salmo

salar) as previously described [33]. The salmon were kept in single tanks with

seawater (34.5 ppt) at approximately 10°C [34]. All experiments were conducted in

accordance with the Norwegian legislation for animal welfare.

2.2. In situ hybridisation

Copepodids were fixed in 4 % paraformaldehyde in phosphate buffered saline for 24

h and transferred to 70 % ethanol for a minimum of 24 h before being embedded in

paraffin wax. Localisation of LsEcR transcript in copepodids was performed as

described by Trösse et al., 2014 [31] with modifications described in Sandlund et al.,

2015 [25]. LsEcR specific cDNA was used as template to generate PCR product with

T7 promoter extensions followed by synthesis of single stranded digoxigenin-labelled

RNA probes (667 bp) (Primers listed in Table 1.). Probe concentration and quality

was determined by spectrometry, employing a Nanodrop ND 1000 (Thermo Fisher

Scientific) and a spot test, respectively. In brief, sections were dewaxed in Histoclear

(National Diagnostics) before rehydration of tissue and proteinase K treatment (10

min) followed by fixation in 4 % formaldehyde in PBS, acetic anhydride treatment

and dehydration. 100 μl hybridization mix containing 20 ng digoxigenin-labelled

RNA were added to the tissue and incubated in a vacuum chamber at 60 °C overnight.

Chromogenesis was carried out using nitroblue tetrazolium (NBT) (Roche

Diagnostics GMbH) and 5-bromo-4-chloro-3-indolyl phosphate (BCIP) (Roche).

Sense RNA was used as a negative control.

2.3. Protein detection

Rabbit polyclonal antibodies (pAb) were raised against a synthetic LsEcR peptide that

corresponding to a unique region of the LsEcR, residues 264-277

(AADTTVDPKSNNNG), by GenScript (Piscataway, NJ, USA). The pAb

successfully detected the LsEcR protein at the expected Mw of 60.3 kDa by Western

blot analysis. Several trials of immunofluorescence studies using the same antibody

were not successful.

2.4. Functional studies using RNAi

Double stranded RNA (dsRNA) fragments for LsEcR and LsRXR (LsEcR fragment:

position 782-1458, Ac. no. KP100057.1; LsRXR fragment: position in 323-945, Ac.

No. KJ361516) were generated by T7 RNA polymerase by MEGAscript® RNAi Kit

(Ambion inc) from PCR templates with T7 promoter regions. An 850 bp gene

fragment from Atlantic cod (Gadus morhua) was used as negative control (Primers

are listed in Table 1.). RNAi on salmon louse larvae was performed as described [30].

Five biological parallels of nauplia I (n = 5 x 30) were soaked in either 1.5 μg of

control dsRNA fragment or LsEcR dsRNA alone or together with equal amounts of

LsRXR dsRNA, in 150 μl of seawater and left for incubation overnight at 9 ± 1°C.

The next day, samples were checked for the presence of exuvia, shed during the molt

from nauplia I to nauplia II stage, and nauplia II was transferred to flow through

incubators and left for 7 days post treatment (d.p.t.) when the control had molted into

copepodids. Copepodids were inspected under a binocular microscope (Olympus SZX

0.5 and 1.6x Olympus objective) and photographed (Canon EOS 600D). The animals

were photographed, collected and stored on RNAlater™ (Ambion) or fixed in

Karnovsky`s reagent for embedding in plastic.

2.5. Infection trial

Copepodids (100/fish) were released in tanks holding one salmon/tank. The water

level was held low for the first 10 min of the infection after which water flow was

returned to normal. Water from the tanks was filtered through a double filter system

(20 μm) to collect copepodids that did not successfully attach to the fish. Three fish

were infected with control lice and three fish were infected with LsEcR dsRNA

treated lice (knock-down of LsEcR was verified before infection). The experiment

was terminated 27 days post infection when the control lice had developed to the pre-

adult/adult stage. The fish was anaesthetised in a mixture of methomidate (5 mg/l) and

benzocaine (60 mg/l) and lice were carefully removed with forceps, counted,

inspected under a microscope and documented as described in section 2.4. Both

cephalothorax length (CL) and total length (TL) of the control and LsEcRkd lice were

measured as described by Eichner et al., 2015 [35] in order to determine any

differences in size.

2.6. Extraction of RNA and cDNA synthesis

For RNAi experiments, 30 copepodids were homogenised using 1.4 mm zirconium

oxide beads (Bertin) in TRI Reagent® (Sigma-Aldrich) TRI Reagent® (Sigma-

Aldrich), according to the manufacturers protocol. RNA was extracted from the water

phase using RNeasy micro kit (Qiagen) as described by the manufacturer. Samples

were stored at -80 °C. Concentrations of RNA was determined by NanoDrop ND-

1000 spectrophotometer at 260 nm (NanoDrop Technologies Inc.) Two parallel

reactions of cDNA synthesis were achieved using AffinityScript qPCR cDNA

synthesis Kit (Agilent Technologies) according to the protocol. A five fold dilution of

cDNA was stored at -20 °C.

2.7. Detection of transcript level in dsRNA treated lice by real time quantitative-PCR

(RT-qPCR)

RT-qPCR was carried out using Applied Biosystems 7500 Real-Time PCR System

(Applied Biosystems®) under standard conditions (initiation: 50 °C 2 min, holding:

95 °C 10 min, 40 cycles of 95 °C 15 sec followed by 60 °C 1 min) as previously

described in Sandlund et al., 2015 [25]. Transcription levels of all genes investigated

were normalised to the established salmon louse standard gene eEF1α [36]. Eight

dilutions of RNA were used to create a standard curve and all RT-qPCRs were carried

out in duplicates. Samples were analysed independently but simultaneously as eEF1α

using the same cDNA and mastermix (TaqMan® Universal PCR Master Mix;

Applied Biosystems). Five biological parallels of copepodids (n = 5 x 30) were run

for each dsRNA treatment group. Results were analysed by the 2^-∆∆Ct approach and

presented with 95 % confidence interval calculated from the 2^-∆∆Ct values. T-tests

were used to determine differences in expression between the control group and

dsRNA treated animals. A p-value of 0.05 was set as threshold.

2.8. Histology

Copepodids for light microscopy were fixed in Karnovsky`s fixative, rinsed in PBS

and dehydrated in a series of ethanol solutions (50 %, 70 % and 96 %), before

embedding in Technovit 7100 (Heraeus Kulzer Teqhnique) as described by [35].

Sections (1–2 μm) were stained with Toluidine blue (1 % in 2 % borax). Sections

were mounted using Histomount (Invitrogen).

Microscopy and imaging were acquired with an Axio Scope. A1 light microscope

connected to Axiocam 105 color camera (Zeiss International) and processed using

Adobe Photoshop CS6 (Adobe Systems).

3. Results

3.1. LsEcR transcript expression in L. salmonis copepodids

Sections of free-living copepodids (7 days old) were used for to in situ hybridisation

in order to identify the localisation of LsEcR transcripts in copepodids. Expression of

LsEcR transcript was detected in the neuronal somata of the brain (Fig. 1B; white

frame) as well as in myonuclei throughout the animal (Fig. 1B; muscle fibres are

marked with asterisks). In addition, a weak staining of the immature intestine could

also be observed (Fig. 1B; black frame).

3.2. Knock-down of LsEcR by RNAi did not inhibit molting from nauplia II to

copepodid larvae

To assess the function of LsEcR during early development and molting, RNAi

experiments were performed by soaking nauplia I larvae in dsRNA fragment specific

to LsEcR. The experiment was terminated seven d.p.t. when control animals had

molted to the copepodid stage. Two separate experiments were run showing similar

results. The dsLsEcR treated group developed normally from nauplia II to the

copepodid stage and did not show any loss-of-function phenotype (Fig. 2B) compared

to the control animals (Fig. 2A) seven d.p.t. RTq-PCR analysis was performed in

order to determine the degree of LsEcR knock-down (LsEcR) and confirmed a

significant decrease of LsEcR transcript by 57 % (Fig. 3A). Histological sections of

the dsLsEcR treated lice were examined to identify any differences in internal

morphology compared to the control, however no abnormalities were observed

(results not shown).

The binding of ecdysone hormones to the EcR/USP complex is known to directly and

indirectly regulate several other genes including the transcription factors LsE75 and

LsE74 and the, hormone receptor HR38. Consequently RT-qPCR was performed in

dsRNA treated lice on these genes and several genes directly involved in the molting

process: LsChitinase1 (LsChi1), LsChitinase2 (LsChi2), LsChitin synthase1

(LsCHS1), LsChitin synthase2 (LsCHS2), LsZinc carboxypeptidase (LsCP1) were

evaluated in dsRNA treated lice. A significant down-regulation was observed for

LsCP1, LsCHS2 and LsHR38 while a significant up-regulation were detected for

LsChi1 in the LsEcRkd animals (Fig. 3A) (T-test, P < 0.05).

3.3. Simultaneous knock-down of LsEcR and LsRXR isoforms resulted in molting

arrest of nauplia II larvae

An additional experiment was set up to investigate the effect of simultaneous knock-

down of LsEcR and LsRXR in salmon louse larvae, as described above. In contrast to

the dsLsEcR treated lice, molting of the LsEcR/LsRXR knock-down lice was

interrupted and the treated lice did not molt from nauplia II (Fig. 2D) to the copepodid

stage (Fig. 2A). The dsLsEcR/dsLsRXR treated lice had developed some features

similar to that of the control copepodid such as segmentation of the abdomen (Fig.

2C; marked by white arrow) but they were not able to emerge from the old cuticula

(Fig. 2C; edge of exuvia marked by black arrow). RT-qPCR confirmed a significant

knock-down of both LsEcR and LsRXR by 46 % and 57 %, respectively (Fig. 3B) (T-

test, P < 0.05). In addition, the same genes mentioned in section 3.2 were analysed by

RT-qPCR analysis in the dsLsEcR/dsLsRXR treated animals. Similar to the LsEcR

knock-down animals, LsCP1, LsCHS2 and LsHR38 were significantly down-regulated

in the dsLsEcR/dsLsRXR treated animals (Fig. 3B) (T-test, P < 0.05). However, in

contrast to the dsLsEcR treated lice no significant regulation was observed for the

LsChi1 or LsChi2 in the dsLsEcR/dsLsRXR treated animals (Fig. 3B) (T-test, P <

0.05).

3.4. Infection trial using LsEcR knock-down animals

An infection trial with LsEcR knock-down copepodids was performed. Atlantic

salmon were infected with either control or LsEcRkd copepodids and cultivated on the

host fish until the control group had developed into pre-adult I and II females and pre-

adult II and adult males (27 d.p.i.). At termination, no abnormalities could be

observed by light microscopy in the dsLsEcR lice. However, high mortality was

evident and a significantly lower number of LsEcRkd lice (n = 29) were recovered

from the host compared to the control group (n = 96). The LsEcRkd males were

significantly shorter in CL and TL (Fig. 4E, F; T-test, P < 0.05) compared to the

control group (Fig. 4D, F) with a few individuals that were even shorter than normally

developed pre-adult I males [9]. In contrast, the female LsEcRkd lice could be divided

into two groups. One group that had significantly smaller CL and TL (Fig. 4B, C; T-

test, P < 0.05) compared to the control and one group with CL and TL size similar to

the control group (Fig. 4C; T-test, P > 0.05). Histological sections from female and

male dsLsEcR treated lice (Fig. 5F-J) both revealed abnormal morphology compared

to the control (Fig. 5A-E). Most striking was the severe reduction of muscle tissue of

the dsLsEcR treated animals (Fig. 5F). The intestine of the dsLsEcR treated animals

(Fig. 5H) resembled the intestinal tissue observed in copepodids rather then the

anatomy described for adult animals (Fig. 5C) [37]. Hypotrophy of the cellular

structure of the sub-cuticula (Fig. 5G), the neurological tissue (Fig. 5J framed), female

ovaries (Fig. 5I) and male testes (Fig. 5J marked with asterisks) was apparent in the

knock-down animals compared to the control (Fig. 5B-E). Histological examinations

of LsEcRkd females that were the same size as the control group showed similar

morphology as the other LsEcRkd animals.

4. Discussion

While extensive knowledge of the EcR function in a variety of developmental and

physiological processes in insects has been obtained, the functional importance of this

receptor in copepods remains largely unstudied. Here we report on the transcriptional

distribution and function of the newly characterized L. salmonis EcR (LsEcR) [25]

during molting and development of the salmon louse. Not surprisingly, analysis of the

spatial distribution of LsEcR in copepodids using in situ demonstrated a wide

expression pattern the intestine, neuronal somata of the brain and in the nuclei of

muscle cells (Fig. 1B). A similar wide distribution pattern of LsEcR transcript was

observed in the adult female salmon lice [25] and the copepod Marsupenaeus

japonicus [14]. The extensive expression of LsEcR is in accordance with the broad

range of biological processes involving the nuclear receptor such as molting,

remodelling of neurons [38, 39] and muscle tissue during metamorphosis [40].

Knock-down studies of LsEcR using RNAi caused near 60 % down-regulation (Fig.

3A) of LsEcR transcript in the salmon lice larvae, but did not result in molting arrest

or any observable morphological abnormalities in the copepodids (7 d.p.t.). Similar

results has been reported in LsRXR knock-down copepodids after dsRNA bath

treatment [30]. The lack of phenotype was surprising as depletion of both EcR and

RXR transcripts in larvae from several insect and crustacean species both cause severe

developmental defects and molting arrest that eventually lead to death of the animal

[27, 41-43]. Studies have shown that both EcR and USP are able to interact with DNA

either as homodimers or monomers [44, 45]. In addition, USP alone has shown to

bind ligand and exert its function without EcR [46]. These results from our

experiments indicate that LsEcR and LsRXR have other partners or manage to retain

its function alone. The apparent normal development of the LsEcRkd copepodids

could be due to insufficient knock-down resulting from remaining LsEcR protein.

However, due to unsuccessful detection of LsEcR protein, we cannot determine if the

extent of protein depletion corresponds to the decrease in LsEcR transcript. Molting

arrest was however achieved by simultaneous knock-down of both LsEcR and LsRXR.

To our knowledge, not many knock-down studies of the nuclear receptor complex

have been performed in crustaceans. Studies of EcR/RXR depletion in the fiddler crab

Uca pugilator showed similar results with arrest of growth during early blastemal

development and during limb generation. In addition, the dsEcR/dsRXR injected crabs

failed to molt and subsequently died [43].

Despite the lack of observable abnormalities in the dsLsEcR treated copepodids (7

d.p.t), the infection trial using the same copepodids resulted in 70 % mortality and

developmental defects that obstructed the surviving lice from normal development.

Whether high mortality is caused by disruption of the molting process or other

physiological mechanisms is uncertain. In addition to the high mortality, the CL and

TL of the recovered animals were significantly shorter for the males and one group of

the females (T-test: P < 0.05) compared to the control. This could either reflect a

decrease in size and/or a delay in development. Similar results were observed in the

cotton mirid bug Apolygus lucorum where the duration of development increased in

the third, fourth and fifth instar after siRNA treatment targeted to EcR. Interference of

AlEcR additionally caused a lethal phenotype where mortality increased from the

fourth nymph instar to adult animals [47]. A significant increase in developmental

times was also observed for larva and pupal stages of Spodoptera exigua after

injection of dsRNA in the fifth instar [48]. During metamorphosis of holometabolous

insects, neurons undergo a crucial remodelling process dependent on EcR (reviewed

in Boulanger et al., 2015) [49]. The ecdysone titer and the pattern of EcR isoform

expression have also shown to be a fundamental factor during regrowth of muscle

during metamorphosis in Manduca [40] and midgut morphogenesis during

embryogenesis in Drosophila [50]. Histological examinations showed lack of muscle

tissue and hypotrophy of neuronal-, sub-cuticular and gonadal tissue in the LsEcRkd

lice (Fig. 5F-J). Our results indicate that LsEcR play a similar role in the salmon louse

and is required either directly or indirectly in the development of different tissues.

Knock-down of LsEcR in pre-adult II females resulted in failure in normal egg

development and offspring generation [25]. The hypotrophic gonads found in both

female and male dsLsEcR treated lice in this study suggests that the animals would

not be able to successfully reproduce and that knock-down of LsEcR transcript in

larval stages affects maturation of the gonadal tissue of the treated animals and

thereby the ability to reproduce. Similar destruction of the gonadal tissues was

observed in adult female LsRXR knock-down lice [26]. Due to the significant reduced

development of internal tissues, reduced growth or even mortality is expected. This is

in accordance with present results.

In crustaceans, chitin synthases, chitinases and carboxypeptidases are important

enzymes in chitin metabolism that is crucial for reorganisation and exchange of the

exoskeleton [51, 52]. A significant decrease of LsCP1 and LsCHS2 transcript (Fig. 3)

were evident in the LsEcR and LsEcR/LsRXR knock-down animals suggesting them to

play a role in molting. The results are supported by previous expression studies in L.

salmonis larval stages [30] and RNA-seq data (unpublished; Licebase.org) that

confirms expression of LsCP1 and LsCHS2 in all molting stages. No down-regulation

was observed for LsCHS1 that is mainly expressed in the adult female intestine

(unpublished; Licebase.org). A similar difference in CHS transcript expression was

demonstrated in the shrimp Pandalopsis japonica [53]. Ontogenetic analysis of

LsChi1 and LsChi2 showed that expression varies within the intermolt and postmolt

stages and peaks between premolt nauplius I and postmolt nauplius II [35]. LsChi1

were significantly up-regulated (Fig. 3.), respectively in the LsEcRkd animals.

Previous RNAi studies of LsChitinases demonstrated them to affect remodelling of

the exoskeleton [35], supporting our results that regulation of the LsChi1 gene in

LsEcRkd copepodids may contribute to the high mortality rate and phenotype

observed in animals from the infection trial. No regulation was observed for the LsChi

genes in the LsEcRkd/LsRXRkd group. However, due to the physiological differences

between the LsEcRkd and the LsEcRkd/LsRXRkd groups (Fig. 2B-C) a direct

comparison of transcript expression of down-stream genes is not possible. Of the

early response genes E74, E75 and HR38 (Fig. 3.) only the latter was significantly

down-regulated in both treatment groups. This could be because of secondary

responses caused by depletion of LsEcR and LsEcR/LsRXR and disruption of several

biological processes. Comparison of expression of the response genes between

different LsEcR/LsRXR knock-down experiments show that the expression of these

genes are dependent on the degree of LsEcR/LsRXR knock-down indicating that the

response genes are tightly regulated by the nuclear receptor complex.

In summary, we found that LsEcR transcript is expressed in several tissues in the L.

salmonis copepods. This is in accordance with its expected role in many molecular

biological processes. Functional studies using RNAi showed that knock down of

LsEcR and LsEcR/LsRXR in L. salmonis larva results in increased mortality, severe

histological changes and molting arrest, respectively. This shows that EcR clearly has

an important function in molting and development of the salmon louse L. salmonis.

The results achieved in this study may be valuable in the development of new tools to

fight the parasite.

Acknowledgements

We would like to thank Heidi Kongshaug, Per Gunnar Espedal, Lars Hamre and

Teresa Cieplinska for excellent help in the laboratory.

Funding

This research has been funded by the Research Council of Norway, SFI-Sea Lice

Research Centre, Grant Number 203513/O30

References

1. Westcott, J.D., K.L. Hammell, and J.F. Burka, Sea lice treatments, management practices and sea lice sampling methods on Atlantic salmon farms in the Bay of Fundy, New Brunswick, Canada. Aquaculture Research, 2004. 35(8): p. 784-792.

2. Johnson, S.C., et al., A review of the impact of parasitic copepods on marine

aquaculture. Zoological Studies, 2004. 43(2): p. 229-243. 3. Costello, M.J., How sea lice from salmon farms may cause wild salmonid

declines in Europe and North America and be a threat to fishes elsewhere. 2009.

4. Jackson, D., et al., Impact of Lepeophtheirus salmonis infestations on

migrating Atlantic salmon, Salmo salar L., smolts at eight locations in Ireland with an analysis of lice-induced marine mortality. Journal of Fish Diseases, 2013. 36(3): p. 273-281.

5. Krkosek, M., et al., Comment on Jackson et al. 'Impact of Lepeophtheirus

salmonis infestations on migrating Atlantic salmon, Salmo salar L., smolts at eight locations in Ireland with an analysis of lice-induced marine mortality'. Journal of Fish Diseases, 2014. 37(4): p. 415-417.

6. Taranger, G.L., et al., Risk assessment of the environmental impact of

Norwegian Atlantic salmon farming. Ices Journal of Marine Science, 2015. 72(3): p. 997-1021.

7. Torrissen, O., et al., Salmon lice - impact on wild salmonids and salmon

aquaculture. Journal of Fish Diseases, 2013. 36(3): p. 171-194. 8. Hamre, L.A., et al., The Salmon Louse Lepeophtheirus salmonis (Copepoda:

Caligidae) Life Cycle Has Only Two Chalimus Stages. Plos One, 2013. 8(9). 9. Schram, T.A., Supplementary descriptions of the develompemntal stages of

Lepeophtheirus salmonis (Krøyer 1837) (Copepoda: Caligidae), in Pathogens of wild and farmed fish: sea lice., G.B.D. Defaye, Editor. 1993, New York: Ellis Horwood. p. 30-47.

10. Chang, E.S. and D.L. Mykles, Regulation of crustacean molting: A review and

our perspectives. General and Comparative Endocrinology, 2011. 172(3): p. 323-330.

11. Thummel, C.S. and J. Chory, Steroid signaling in plants and insects - common

themes, different pathways. Genes & Development, 2002. 16(24): p. 3113-3129.

12. Kozlova, T. and C.S. Thummel, Steroid regulation of postembryonic development and reproduction in Drosophila (vol 11, pg 276, 2000). Trends in Endocrinology and Metabolism, 2000. 11(10): p. 436-436.

13. Ogura, T., et al., Molecular cloning, expression analysis and functional

confirmation of ecdysone receptor and ultraspiracle from the Colorado potato beetle Leptinotarsa decemlineata. Febs Journal, 2005. 272(16): p. 4114-4128.

14. Asazuma, H., et al., Molecular cloning and expression analysis of ecdysone

receptor and retinoid X receptor from the kuruma prawn, Marsupenaeus japonicus. Comparative Biochemistry and Physiology B-Biochemistry & Molecular Biology, 2007. 148(2): p. 139-150.

15. Durica, D.S., et al., Characterization of crab EcR and RXR homologs and

expression during limb regeneration and oocyte maturation. Molecular and Cellular Endocrinology, 2002. 189(1-2): p. 59-76.

16. Hopkins, P.M., Ecdysteroids and Regeneration in the Fiddler Crab Uca-

Pugilator. Journal of Experimental Zoology, 1989. 252(3): p. 293-299. 17. King-Jones, K. and C.S. Thummel, Nuclear receptors - A perspective from

Drosophila. Nature Reviews Genetics, 2005. 6(4): p. 311-323. 18. Riddiford, L.M., et al., Insights into the molecular basis of the hormonal

control of molting and metamorphosis from Manduca sexta and Drosophila melanogaster. Insect Biochemistry and Molecular Biology, 2003. 33(12): p. 1327-1338.

19. Talbot, W.S., E.A. Swyryd, and D.S. Hogness, Drosophila Tissues with

Different Metamorphic Responses to Ecdysone Express Different Ecdysone Receptor Isoforms. Cell, 1993. 73(7): p. 1323-1337.

20. Tarrant, A.M., et al., Ecdysteroid receptor from the American lobster

Homarus americanus: EcR/RXR isoform cloning and ligand-binding properties. General and Comparative Endocrinology, 2011. 173(2): p. 346-355.

21. Kato, Y., et al., Cloning and characterization of the ecdysone receptor and

ultraspiracle protein from the water flea Daphnia magna. Journal of Endocrinology, 2007. 193(1): p. 183-194.

22. Schubiger, M., et al., Drosophila EcR-B ecdysone receptor isoforms are

required for larval molting and for neuron remodeling during metamorphosis. Development, 1998. 125(11): p. 2053-2062.

23. Bender, M., et al., Drosophila ecdysone receptor mutations reveal functional

differences among receptor isoforms. Cell, 1997. 91(6): p. 777-788. 24. Davis, M.B., et al., Phenotypic analysis of EcR-A mutants suggests that EcR

isoforms have unique functions during Drosophila development. Developmental Biology, 2005. 282(2): p. 385-396.

25. Sandlund, L., et al., Molecular characterisation of the salmon louse,

Lepeophtheirus salmonis salmonis (Kroyer, 1837), ecdysone receptor with emphasis on functional studies of female reproduction. Int J Parasitol, 2015. 45(2-3): p. 175-85.

26. Eichner, C., et al., Characterization of a novel RXR receptor in the salmon

louse (Lepeophtheirus salmonis, Copepoda) regulating growth and female reproduction. BMC Genomics, 2015. 16(1): p. 81.

27. Li, T. and M. Bender, A conditional rescue system reveals essential functions

for the ecdysone receptor (EcR) gene during molting and metamorphosis in Drosophila. Development, 2000. 127(13): p. 2897-905.

28. Campbell, E.M., C.C. Pert, and A.S. Bowman, RNA-interference methods for

gene-knockdown in the sea louse, Lepeophtheirus salmonis: studies on a putative prostaglandin E synthase. Parasitology, 2009. 136(8): p. 867-874.

29. Dalvin, S., et al., Functional characterisation of the maternal yolk-associated

protein (LsYAP) utilising systemic RNA interference in the salmon louse (Lepeophtheirus salmonis) (Crustacea: Copepoda). International Journal for Parasitology, 2009. 39(13): p. 1407-1415.

30. Eichner, C., et al., A method for stable gene knock-down by RNA interference

in larvae of the salmon louse (Lepeophtheirus salmonis). Experimental Parasitology, 2014. 140: p. 44-51.

31. Trosse, C., F. Nilsen, and S. Dalvin, RNA interference mediated knockdown of

the KDEL receptor and COPB2 inhibits digestion and reproduction in the parasitic copepod Lepeophtheirus salmonis. Comparative Biochemistry and Physiology B-Biochemistry & Molecular Biology, 2014. 170: p. 1-9.

32. Skern-Mauritzen, R., O. Torrissen, and K.A. Glover, Pacific and Atlantic

Lepeophtheirus salmonis (Kroyer, 1838) are allopatric subspecies: Lepeophtheirus salmonis salmonis and L. salmonis oncorhynchi subspecies novo. Bmc Genetics, 2014. 15.

33. Hamre, L.A., K.A. Glover, and F. Nilsen, Establishment and characterisation

of salmon louse (Lepeophtheirus salmonis (Kroyer 1837)) laboratory strains. Parasitology International, 2009. 58(4): p. 451-460.

34. Hamre, L.A. and F. Nilsen, Individual fish tank arrays in studies of

Lepeophtheirus salmonis and lice loss variability. Diseases of Aquatic Organisms, 2011. 97(1): p. 47-56.

35. Eichner, C., et al., Molecular characterisation and functional analysis of

LsChi2, a chitinase found in the salmon louse (Lepeophtheirus salmonis salmonis, Kroyer 1838). Exp Parasitol, 2015. 151-152: p. 39-48.

36. Frost, P. and F. Nilsen, Validation of reference genes for transcription profiling in the salmon louse, Lepeophtheirus salmonis, by quantitative real-time PCR. Veterinary Parasitology, 2003. 118(1-2): p. 169-174.

37. Nylund, A., S. Okland, and B. Bjorknes, Anatomy and Ultrastructure of the

Alimentary Canal in Lepeophtheirus-Salmonis (Copepoda, Siphonostomatoida). Journal of Crustacean Biology, 1992. 12(3): p. 423-437.

38. Robinow, S., et al., Programmed Cell-Death in the Drosophila Cns Is

Ecdysone-Regulated and Coupled with a Specific Ecdysone Receptor Isoform. Development, 1993. 119(4): p. 1251-1259.

39. Truman, J.W., et al., Ecdysone Receptor Expression in the Cns Correlates

with Stage-Specific Responses to Ecdysteroids during Drosophila and Manduca Development. Development, 1994. 120(1): p. 219-234.

40. Hegstrom, C.D., L.M. Riddiford, and J.W. Truman, Steroid and neuronal

regulation of ecdysone receptor expression during metamorphosis of muscle in the moth, Manduca sexta. Journal of Neuroscience, 1998. 18(5): p. 1786-1794.

41. Tan, A. and S.R. Palli, Edysone receptor isoforms play distinct roles in

controlling molting and metamorphosis in the red flour beetle, Tribolium castaneum. Molecular and Cellular Endocrinology, 2008. 291(1-2): p. 42-49.

42. Bender, M., et al., Drosophila ecdysone receptor mutations reveal functional

differences among receptor isoforms. Cell, 1997. 91(6): p. 777-88. 43. Das, S. and D.S. Durica, Ecdysteroid receptor signaling disruption obstructs

blastemal cell proliferation during limb regeneration in the fiddler crab, Uca pugilator. Molecular and Cellular Endocrinology, 2013. 365(2): p. 249-259.

44. Niedziela-Majka, A., M. Kochman, and A. Ozyhar, Polarity of the ecdysone

receptor complex interaction with the palindromic response element from the hsp27 gene promoter. European Journal of Biochemistry, 2000. 267(2): p. 507-519.

45. Grad, I., et al., Analysis of Usp DNA binding domain targeting reveals critical

determinants of the ecdysone receptor complex interaction with the response element. European Journal of Biochemistry, 2001. 268(13): p. 3751-3758.

46. Jones, G. and D. Jones, Considerations on the structural evidence of a ligand-

binding function of ultraspiracle, an insect homolog of vertebrate RXR. Insect Biochemistry and Molecular Biology, 2000. 30(8-9): p. 671-679.

47. Tan, Y.A., et al., Ecdysone receptor isoform-B mediates soluble trehalase

expression to regulate growth and development in the mirid bug, Apolygus lucorum (Meyer-Dur). Insect Mol Biol, 2015.

48. Yao, Q.O., et al., Identification of 20-Hydroxyecdysone Late-Response Genes in the Chitin Biosynthesis Pathway. Plos One, 2010. 5(11).

49. Boulanger, A. and J.M. Dura, Nuclear receptors and Drosophila neuronal

remodeling. Biochimica Et Biophysica Acta-Gene Regulatory Mechanisms, 2015. 1849(2): p. 187-195.

50. Chavoshi, T.M., B. Moussian, and A. Uv, Tissue-autonomous EcR functions

are required for concurrent organ morphogenesis in the Drosophila embryo. Mech Dev, 2010. 127(5-6): p. 308-19.

51. Rocha, J., et al., Cuticular chitin synthase and chitinase mRNA of whiteleg

shrimp Litopenaeus vannamei during the molting cycle. Aquaculture, 2012. 330: p. 111-115.

52. Sui, Y.P., et al., Characterization and influences of classical insect hormones

on the expression profiles of a molting carboxypeptidase A from the cotton bollworm (Helicoverpa armigera). Insect Mol Biol, 2009. 18(3): p. 353-63.

53. Kim, K.R., et al., cDNAs encoding chitin synthase from shrimp (Pandalopsis japonica): molecular characterization and expression analysis. Integrative and Comparative Biology, 2015. 55: p. E284-E284.

Figure legends

Fig. 1. Localisation of LsEcR transcript in Lepeophtheirus salmonis copepodid larvae.

A) Light microscope image of a copepodid larvae (7 days) to show the outline of the

animal. B) In situ hybridisation using LsEcR-specific probes was used for detection of

transcript. Negative control (sense RNA) is shown (insert). Transcript was detected in

the neuronal somata of the brain (white frame) and myonuclei throughout the

copepodid (asterisk). A weak positive staining was also observed in the immature

intestine (black frame). Weak unspecific colouring of the outer cuticular layer was

seen using both the sense and the anti-sense probes. C) A histological section of a

copepodid was shown for better visualisation of selective tissues. White and black

frames are used to better visualize the localisation of neuronal and intestinal tissue,

respectively and asterisks are used to mark muscle tissue. Scale bars = 200 μm (A, B,

C).

Fig. 2. Selection of representative animals obtained from the RNAi experiment using

molting nauplius I larvae. A) Control group. B) Larvae treated with double-stranded

RNA (dsRNA) targeted to LsEcR alone developed into copepodids and did not show

any abnormal phenotype compared to the control group (A). C) Larvae treated with

fragments targeting both LsEcR and LsRXR, was observed to have developed

copepodid like features (marked with white arrow) inside the exuvia (marked with

black arrow) and was viable at day 7 post soaking. However, the molting from nauplia

II to copepodid was arrested and the animals started to die after 12 days. Morphology

of a normally developed nauplia II larvae is shown in D (for reference only). Scale

bars = 200 μm (A-D).

Fig. 3. Relative expression of the ecdysone receptor (EcR) and the retinoid X

receptor (RXR) together with possible target genes regulated by the LsEcR/LsRXR

complex, after treatment with double-stranded RNA (dsRNA). Lepeophtheirus

salmonis EcR (LsEcR) and LsEcR/LsRXR treated copepodids were collected 7 days

post treatment. Quantitative real-time PCR (RTq-PCR) of LsEcR, LsRXR and the

possible target genes LsE75, LsE74, LsChi1, LsChi2, LsCHS1, LsCHS2, LsCP1and

LsHR38 were evaluated. GenBank Accession Numbers: LsEcR: KP100057, LsRXR:

KJ361516, LsChi1: AJD87505.1, LsChi2: AIE45495.1, LsCHS1: submitted, LsCHS2:

submitted, LsE75: submitted LsE74: submitted, LsCP1: BT078319.1, LsHR38:

submitted. The graphs are representatives of two individual experiments. The

expression levels of the respective genes in the control groups were set to 1. Values

are expressed as mean ± confidence intervals of five biological replicates (n = 30

copepodids/sample). *Statistically significant (P < 0.05). Statistical analysis was

performed using a T-test.

Fig. 4. Representative animals obtained after infection trial using LsEcR knock-down

(LsEcRKd) copepodids. A, D) shows pre-adult II female (A) and male (D) control

animals 27 days after infection. B, E) represents LsEcRkd female (B) and male (E)

animals. C) The female LsEcRkd lice could be divided into two groups that were

either significantly smaller or the same size as the control group. F) All male LsEcRkd

lice were significantly smaller compared to the control. Statistical analysis was

performed using a T-test; P < 0.05. Scalebar = 1 mm A-B, D-E.

Fig. 5. Histological examination of Lepeophtheirus salmonis EcR knock-down

(LsEcRkd) animals. Toluidine stained control sections and sections from LsEcRkd

animals are shown in A-E and F-J, respectively. A and F) shows sections of the whole

animal with letters as guides to the corresponding photos of individual tissues. E and

J) are sections of males where testes are marked with asterisks and brain tissue is

framed with a black circle. Testes and ovaries are located at the same position in their

respective gender (A, F marked with asterisk). Muscle tissue was observed throughout

the control animal (A) in contrast to the LsEcRkd animals where a severe reduction of

muscle tissue was evident (F). The sub-cuticular tissue (G) ovaries (I), testes (J*) and

neuronal tissue (J framed) were hypotrophic compared to the control (B, D, E*, E

framed) while the intestinal tissue of the LsEcRkd animals appeared to resemble that

found in coepodid larva (H) rather then the control (C). Scale bar = 300 μm (A, F) and

100 μM (B-E, G-H).

a Taqman® assays were provided by Applied Biosystems, Branchurg, NJ, USA. b All general primers were purchased from Sigma-Aldrich, St Louis, MO, USA. RACE, rapid amplification of cDNA ends; TOPO, DNA topoisomerase I; dsRNA, double-stranded RNA; RTq-PCR, real-time quantitative PCR

Primer nameb Sequence (5`-3`) Method

LsEcR_specific_P1 CCGATTTGCCATTACGTAGGCTTGTAGAGC in situ/ dsRNA

LsEcR_specific_P2 CCGCAGCTGCAGCCGACACAACTGTAGAT in situ/dsRNA

LsEcR_specific_P3 CGAGCGTTTCCACTTACTTGCCAT dsRNA

LsEcR_specific_P4 CGCCAACAACGACGACCC TCCACCAACAGCACT dsRNA

LsRXR_specific_P1 CGGAATTGGGATGTCTACGAGCCATCATA dsRNA

LsRXR_specific_P2 CTTCCTCTGACTCACTATAGAAGCATA dsRNA

Cod_specific_T7f ATAGGGCGAATTGGGTACCG dsRNA

Cod_specific_T7r AAAGGGAACAAAAGCTGGAGC dsRNA

LsEF1α_f CATCGCCTGCAAGTTTAACCAAATT RTq-PCR

LsEF1α_r CCGGCATCACCAGACTTGA RTq-PCR

LsEF1α_TaqMan® ACGTACTGGTAAATCCAC RTq-PCR

LsEcR total_f TCGGGAGAAAGTCCCTCTTCT RTq-PCR

LsEcR total_r ACAGCTCCAGTAGGTGTTAAAGGA RTq-PCR

LsEcR total TaqMan® TCGCAGTCCATTCTC RTq-PCR

LsRXR total_f CCTAGTTGAACTCATCGCCAAAATG RTq-PCR

LsRXR total_r TGAAGAGTATGATGGCTCGTAGACA RTq-PCR

LsRXR total TaqMan® CCGCTTTGTCCATTTGCAT RTq-PCR

LsHR38_f CCGTTCTCACAACTTCCTTTACCAT RTq-PCR

LsHR38_r CGTCGAAATCGATGTCAGTTTTGC RTq-PCR

LsHR38 TaqMan® CCCACGGCAAGACAT RTq-PCR

LsE74_f GGTCACGTTAAAGATGGGTCAATTT RTq-PCR

LsE74_r CCAACAGGAGTACTAACACAACTGAT RTq-PCR

LsE74_TaqMan® CAGCGCCTCGTTCAC RTq-PCR

LsE75_f CCTTGACCAATTTTCAGAACGGTTT RTq-PCR

LsE75_r AATCCAGGGATCCGCTTGG RTq-PCR

LsE75_TaqMan® CACGTTCGCCAAGTTT RTq-PCR

LsCP1_f TGTCAAATAAGCGTGAGGTTAAGGAA RTq-PCR

LsCP1_f GCGCGTGAATACCACAATCC RTq-PCR

LsCP1_ TaqMan® ACCCAAACGATCTTCC RTq-PCR

LsChi1_f TCCATTCATTTGTACACATGTGGCTTA RTq-PCR

LsChi1_r CATTGTAAGGGTCAAGGAGTCGAAT RTq-PCR

LsChi1 TaqMan® CAGACCAGCAAATCCA RTq-PCR

LsChi2_f GTTAATTTCTTAAACAAGTATAACTTCGACGGTT RTq-PCR

LsChi2_r GGAACACCACCTCGTTTAGCA RTq-PCR

LsChi2 TaqMan® TTCCCAGTCAATATCC RTq-PCR

LsCHS1_f TTTCGAGGTAAAGCACTTATGGATGAT RTq-PCR

LsCHS1_R AGCCATCTATCTTCTCCTTGATCGT RTq-PCR

LsCHS1_TaqMan® ATGCCTTGGTTCTTCC RTq-PCR

LsCHS2_f AGCAGTGACTATGCTTTTAGATTTGATGA RTq-PCR LsCHS2_r CTGAGCCTAAAGGATGAATTCTTCCA RTq-PCR LsCHS2_TaqMan® CACTGCGCCGGTTTT RTq-PCR

Table. 1. Primer sequences and Taqman® assaysa used in this study

Fig. 1.

Fig. 2.

Fig. 3

Fig. 4.

Fig. 5.

III

1

Manuscript

Identification and functional assessment of the ecdysone biosynthetic genes

neverland, disembodied and shade in the salmon louse Lepeophtheirus salmonis

(Copepoda, Caligidae)

Authors and affiliations

Liv Sandlunda*, Heidi Kongshauga Tor Einar Horsbergb, Rune Malec, Frank Nilsena,

Sussie Dalvind

a Sea Lice Research Centre, Department of Biology, University of Bergen, Norway b Sea Lice Research Centre, Norwegian University of Life Sciences, Norway c Sea Lice Research Centre, Department of Molecular Biology, University of Bergen,

Norway d Sea Lice Research Centre, Institute of Marine Research, Norway

*Corresponding author.

E-mail address: liv.sandlund@bio.uib.no (Liv Sandlund)

Abstract

The salmon louse is a marine ectoparasitic copepod on salmonis fishes. Its lifecycle

consists of eight developmental stages, each separated by a molt. In crustaceans and

insects, molting and reproduction is controlled by circulating steroid hormones such

as 20-hydroxyecdysone (20E). Steroid hormones are synthesized from cholesterol

through catalytic reactions involving a 7,8-dehydrogenase neverland and several

cytochrome P450 genes collectively called the Halloween genes. In this study, we

have isolated and identified orthologs of neverland (nvd), disembodied (dib) and

shade (shd) in the salmon louse L. salmonis genome. Tissue-specific expression

analysis showed that the genes are expressed in intestine and reproductive tissue.

Furthermore, knock-down studies using RNA interference in adult females showed

that only shd terminates molting in larval stages. However, knock-down of nvd

affected development of the ovaries and oocyte maturation. In addition, we performed

knock-down studies of an ortholog of the Drosophila octopamine receptor (Oct3βR) a

2

regulator of the Halloween genes, to determine its role during early development.

Depletion of the Oct3βR ortholog in L. salmonis resulted in molting arrest, but did not

down-regulate expression of all of the identified Halloween genes. Our results show

that ecdysone biosynthetic genes are present in L. salmonis and that ecdysteroids is

necessary for both molting and reproduction in the lice.

Abbreviations

RT-qPCR: real-time quantitative PCR, LC/MS/MS: Liquid chromatography tandem

mass spectrometry

1. Introduction

The salmon louse, Lepeophtheirus salmonis, is an ectoparasittic copepod (Copepoda,

Caligida) infecting salmonid fishes. The parasite feeds on the mucosa, skin and blood

of the host creating open wounds that increase the chance of secondary infections and

mortality. The lice cause a serious threat to farming of Atlantic salmon in Norway,

UK, USA and Canada and are estimated to contribute to an annual loss of more than

500 million USD in Norway alone http://nofima.no/nyhet/2015/08/kostnadsdrivere-i-oppdrett/.

The life cycle consists of eight developmental stages: nauplia I and II, copepodid,

chalimus I and II, pre-adult and adult stage, that are separated by a molt (Albright,

1991; Hamre et al., 2013; Schram, 1993). Oocytes are generated as syncytia in coiled

tubular structures that form the ovaries in the anterior part of the animal and are

transported to the genital segment through long oviducts where they become

convoluted (Ritchie et al., 1996). The oocytes mature inside the genital segment

where lipids and vitellogenins are incorporated (Dalvin et al., 2009; Dalvin et al.,

2011; Pike and Wadsworth, 2000). The eggs are fertilized and extruded as long

strings of several hundred eggs from the female and the embryo begins to develop.

Both oocyte maturation in the genital segment and embryogenesis take approximately

10 days at 10 °C in L. salmonis (Dalvin et al., 2011).

In arthropods, embryogenesis and molting are regulated by ecdysteroid hormones, a

group of cholesterol derived poly-hydroxylated ketosteroids that in addition to

3

molting, are responsible for regulating essential endocrine regulated processes such as

embryogenesis, growth and reproduction. Ecdysteroidogenesis takes place in special

molting glands known as the prothoracic glands (PG) and Y-organ (YO) in insects

and decapod crustaceans, respectively. No such organ has been identified in

crustacean like the salmon louse but it has been suggested that these species possibly

synthesise ecdysteroids in the epidermis (Hopkins, 2009; Lachaise et al., 1993; Lafont

and Mathieu, 2007). Regulation of ecdysteroid synthesis is complex and is controlled

by peptide hormones. While the prothoracicotropic hormone (PTTH) stimulates

ecdysteroid synthesis in insects, the Y-organ is under inhibitory control of the molt-

inhibiting hormone (MIH) (Mykles, 2011). However, the synthesis pathway of

ecdysteroids is similar in insects and crustaceans suggesting that the pathway was

present in their common ancestor (Grieneisen, 1994; Lachaise et al., 1993; Rees,

1995; Rewitz and Gilbert, 2008).

Arthropods are unable to synthesise cholesterol de novo and uptake of exogenous

cholesterol through the diet is nescessary (Gilbert et al., 2002; Goad, 1981; Spaziani

et al., 1999). In Drosophila a number of enzymes known to be involved in the

synthesis has been identified and orthologs of these have in recent years been

identified in crustaceans (Asazuma et al., 2009; Niwa et al., 2010; Sumiya et al.,

2014). These include neverland (nvd) and the cytochrome P450 mono-oxygenases

(CYP450), coded by what is collectively called the Halloween genes: spook (spo;

CYP307A1), phantom (phm: CYP306A1), disembodied (dib: CYP302A1), shadow

(sad: CYP315A1) and shade (shd: CYP314A1). Recently it has been shown that

ecdysteroidogenesis driving metamorphosis in Drosophila is regulated by

monoaminergic autocrine signalling through the G protein-coupled receptor β3-

octopamine receptor (Octβ3R) by activating prothoracicotropic hormone (PTTH) and

insulin-like peptides (Ilps) (Ohhara et al., 2015). The initial reaction in the conversion

of dietary cholesterol to 7-dehydrocholesterol (7dC) is performed by the 7,8-

dehydrogenase nvd (Yoshiyama et al., 2006; Yoshiyama-Yanagawa et al., 2011).

Loss of nvd function in the prothoracic gland (PG) of Drosophila melanogaster and

Bombyx mori results in an arrest in both growth and molting (Yoshiyama et al., 2006).

7dC is converted to another intermediate 5β-diketol by enzymes coded from non-

molting glossy (Nm-g)/shroud (sro), spook (spo) and spookier (spok) in several steps

4

called “the black box” (Iga and Kataoka, 2012; Namiki et al., 2005; Niwa et al., 2010;

Ono et al., 2006). 5β-diketol is, through sequential hydroxylations, modified into the

secreted steroid, which in crustaceans include ecdysone (E), 3-dehydroecdysone

(3DE), 25-deoxyecdysone (25DE) and ponasterone A (PonA, 25-Deoxyecdysterone)

(Baldwin et al., 2009; Feyereisen, 2006, 2011; Mykles, 2011; Rewitz et al., 2007).

These products are released into the hemolymph and transported to peripheral tissues

where they are converted into the biologically active metabolite 20-hydroxyecdysone

(20E) by shd (Gilbert and Warren, 2005; Lachaise et al., 1993; Subramoniam, 2000).

20E transduce the signal by binding to a heterodimer complex consisting of the two

nuclear transcription factors the ecdysone receptor (EcR) and retinoid X receptor

(RXR). In L. salmonis, EcR and RXR are expressed in many tissues including the

intestine, sub-cuticular tissue, ovaries and oocytes (Eichner et al., 2015; Sandlund et

al., 2015). The sub-cuticular tissue in L. salmonis has been demonstrated to have

functions similar to the liver (Edvardsen et al., 2014) and is the production site for

yolk proteins (Dalvin et al., 2009; Dalvin et al., 2011). Synthesis of yolk proteins has

been associated with an increase in 20E level in the isopod Porcellio dilatatus (Gohar

and Souty, 1984).

In crustaceans and insects, the 20E level in the hemolymph peak right before molting

followed by an immediate decline in concentration after ecdysis (Martin-Creuzburg et

al., 2007). Hormone levels seem to directly mimic the expression level of their

synthesizing enzymes. The temporal expression patterns of Halloween genes

identified in B. mori and Manduca sexta are correlated with changes in ecdysteroid

titer during development. Mutations in the Halloween genes result in low ecdysteroid

level and decreased expression of 20E responsive genes, reduced deposition of

embryonic cuticle and embryonic lethality (Chavez et al., 2000; Niwa et al., 2004;

Niwa and Niwa, 2011; Petryk et al., 2003; Warren et al., 2002; Warren et al., 2004).

Although the pathway of ecdysteroid synthesis is well established in insects,

functional analysis of this pathway has not been reported in copepods. Recently,

orthologs of nvd and the CYP450 genes spo, phm, dib, sad and shd were discovered in

the water flea Daphnia (Markov et al., 2009; Rewitz and Gilbert, 2008; Sumiya et al.,

2014). In malacostraca, cDNA encoding CYP306A1 (phm) has been cloned form the

5

Kuruma Prawn Marsupenaeus japonicus (Asazuma et al., 2009) and four CYPs (spo,

phm, dib and sad) and nvd was identified in the crayfish Pontastacus leptodactylus

(Tom et al., 2013).

The aims of the present study were to identify and assess the expression pattern of

neverland and the Halloween genes disembodied and shade and investigate their

significance in the biosynthesis of ecdysteroids in the salmon louse. These genes were

cloned and sequenced and ontogenetic expression analysis using RT-qPCR were

performed. In addition, RNA interference was applied to knock-down each these

genes and LsOct to further examine their significance in growth and molting in L.

salmonis larvae.

2. Materials and methods

2.1. Salmon lice

A laboratory strain of the Atlantic salmon louse Lepeophtheirus salmonis salmonis

(Skern-Mauritzen et al., 2014) was maintained and farmed on Atlantic salmon (Salmo

salar) (Hamre et al., 2009). Both lice and fish were kept in seawater with salinity of

34.5 ppt and with temperature at 10 ± 0.2 °C. Eggs were hatched and cultivated to

copepodid stage in flow-through incubators before infection of salmon (Hamre et al.,

2009). The lice were kept on the fish until they reached the desired developmental

stage. Prior to sampling of lice, the salmon were either killed with a blow to the head

or anaesthetized in a mixture of methomidate (5 mg/l) and benzocaine (60 mg/l);

thereafter lice were removed with forceps. All experimental procedures were in

accordance with the Norwegian legislation for animal welfare.

2.2. Cloning, sequence analysis and alignment

Candidate genes from the cytochrome P450 enzymes CYP314a1/shade (shd) and

CYP302a1/disembodied (dib) and the 7,8 dehydrogenase neverland (nvd) known to

be involved in the synthesis of ecdysteroids were identified in the salmon louse

genome (unpublished data; Licebase.org) by homology to known sequences from

6

insects and crustaceans. Sequences with the lowest e-value were chosen. Gene

specific 5`and 3`RACE primers were deigned (Table. 1.) and RACE was performed

using the SMARTer™RACE cDNA Amplification kit (Clontech, TaKaRa) according

to manufacturer’s recommendations (Sigma-Aldrich). Sub-cloned was performed

using a pCR®4-TOPO® vector system (Invitrogen) that were transformed into

Escherichia coli TOP10 cells. Clones were verified by PCR with M13_f and M13_r

primers (Table. 1.), grown overnight and purified using a Miniprep Nucleospin®

Plasmid Purification Kit (Macherey-Nagel, Duren, Germany). Plasmids were

sequenced using a BigDye® Terminator v3.1 Cycle sequencing kit (Applied

Biosystems®, Foster City, CA, USA) and analyzed in MacVector (MacVector Inc.,

North Carolina, USA). Sequencing was performed by the Sequencing Facility at the

Molecular Biological Institute, University of Bergen.

2.3. Collection of salmon lice

2.3.1 Lice for ontogenetic analysis

Five parallels of each developmental stage of the L. salmonis were collected prior to

analysis; nauplia I/II and free-living copepodids (n ≈ 5 x 150), parasitic copepodids,

chalimus I and chalimus II (n = 5 x 10), pre-adult male I/II, pre-adult female I/II, adult

male and immature adult female lice (n = 5 x 1) and stored on RNAlater™ (Ambion

inc.).

2.3.2 Lice for LC/MS/MS analysis

From the moment an adult female is fertilized, it takes 10 days to produce a pair of

egg strings at 10 °C. In order to determine the presence of ecdysteroid and the

ecdysteroid levels during the maturation cycle of the egg strings, adult female louse

were sampled and their corresponding egg strings were transferred to individual

incubators with flowing seawater at 10 °C. The time point of hatching for each egg

string were noted and used to determine the exact point of the egg maturation cycle

for each female. Five biological samples for each time point (10 days) and all together

50 female lice were collected and dry frozen at – 80 °C. In addition, we collected

7

nauplia II (n = 2 x 200), free-swimming copepodids (n = 2 x 200) and parasitic

copepodids (n = 3 x 30) two and four days after host infection, in order to determine

the ecdysteroid content during larval stages.

2.4. Detection of transcript using in situ hybridisation

Salmon louse were incubated in 4 % paraformaldehyde in phosphate buffered saline

(PBS) for 24 hours and kept in 70 % ethanol at 4 °C for at least 24 hour before

paraffin embedding. In situ hybridization was carried out according to (Kvamme et

al., 2004), with the same modifications as described in (Sandlund et al., 2015).

Digoxigenin (DIG)-labelled anti-sense and sense RNA probes (LsNvd: 261 bp, LsShd:

327 bp, LsDib: 475 bp) were generated from PCR templates (Table. 1.) with T7

promoters using DIG RNA labelling kit (Roche). 10 and 25 ng/μl probe was used in

the hybridisation mix for detection of LsShd, LsDib and LsNvd, respectively.

Chromogenesis was carried out using nitroblue tetrazolium (NBT) (Roche

Diagnostics GMbH) and 5-bromo-4-chloro-3-indolyl phosphate (BCIP) (Roche).

Sense RNA was used as a negative control.

2.5. RNA interference

Generally for all RNAi experiments, double stranded RNA (dsRNA) fragments for

LsNvd, LsDib and LsShd was produced using T7 RNA polymerase by MEGAscript®

RNAi Kit (Ambion inc) from PCR templates with T7 promoter regions according to

the supplier´s description. An Atlantic cod (Gadus morhua) trypsin gene fragment

(850 bp) was used as negative control. Primers and fragment sizes are listed in Table.

1. Live lice were transferred and stored on either RNAlater™ (Ambion inc.) for

qRTPCR or fixed in Karnovsky`s fixative before embedding in plastic. Lice were

inspected using a binocular microscope (Olympus SZX12, 0.5 and 1.6x Olympus

objective) and photographed (Canon EOS 600D). RNAi on adult female lice was

performed as described by (Dalvin et al., 2009). Pre- adult II female lice were then

injected with approximately 1 μl of the dsRNA (600 ng/μl) solution in the

cephalothorax using custom made injection needles coupled to a microinjector

(Sandlund et al., 2015). After injection, the lice were kept in seawater for 4 h before

8

they were placed on anesthetised fish together with male lice, in a 1:1 ratio (female n

= 13). Three parallels were set up for each gene. Lice were removed from the salmon

after the female adult control lice had produced a second set of egg strings (approx. 40

days). Egg strings were collected and placed in individual incubators for hatching.

RNAi on larvae was performed as previously described by (Eichner et al., 2014). For

each gene fragment, five biological parallels of nauplia I larvae (n = 5 x 30) were

soaked in 1.5 μg of dsRNA fragment for the respective gene in 150 μl seawater and

incubated for 20-24 h at 9 ± 1°C. The next day, samples were checked for the

presence of exuvia, shed during the molt from nauplia I to nauplia II stage, and

nauplia II was transferred to flow through incubators and left for 7 days post treatment

(d.p.t.) when the control had molted into copepodids.

2.6 RNA extraction and cDNA synthesis

Total RNA from animals collected from RNAi experiments and for ontogenetic

analysis was isolated using TRI Reagent® (Sigma-Aldrich) as previously described

by (Sandlund et al., 2015), according to the manufacturers protocol. Animals were

homogenised in Tri Reagent® and added chloroform. After extraction of the water

phase, RNA from nauplia and copepodid samples were extracted using RNeasy micro

kit (Qiagen) after the manufacturer`s protocol. RNA was treated with DNase I

(Amplification Grade, Invitrogen). Samples were stored at -80 °C. cDNA synthesis

was achieved using AffinityScript qPCR cDNA synthesis Kit (Agilent Technologies)

according to the protocol. cDNA was diluted 5 or 10 times and stored at -20 °C.

2.7. Real Time quantitative PCR (RT-qPCR)

Real-time quantitative PCR (RT-qPCR) using SyBR® green assays (Table. 1.) were

applied to detect total expression of genes from dsRNA treated lice harvested from

the RNAi experiments as well as genes submitted to ontogenetic analysis. Primers

were designed using MacVector (MacVector inc.). Primers were designed to localise

the respective gene independently of their specific dsRNA fragments. Eight dilutions

(from 100 – 0.78 ng/ul) of RNA from L. salmonis were used as template to generate a

9

dilution curve and verify the efficiency (1.87-2.1) for each assay. Reaction specificity

was determined when only a single peak in the dissociation curve was present. All

assays were run in parallel series together with the reference gene eEF1α (Frost and

Nilsen, 2003). One reaction without reverse transcriptase was run to exclude

contamination of genomic DNA. All samples were added 2 ng/ul cDNA containing,

2X SyBR® Select Master Mix with ROX (Applied Biosystems) and 10 μM of each

primer to the total reaction volume of 10 μl. Thermal cycling and quantification were

performed using Applied Biosystems 7500 Real-Time PCR System (Applied

Biosystems®) under the following conditions: enzyme activation step; 95 °C 15 min,

followed by 40 cycles of denaturation of 95 °C for 15 sec and extension; 50-60 °C for

1 min (dependent on the gene). All samples were normalised to eEF1α by the 2^-∆∆Ct

approach and relative expression was calculated using controls from each experiment

as standard.

2.8. Chemicals for LC/MS/MS analysis

Ecdysone (E), 20-hydroxyecdysone (20-E) and ponasterone A (PonA) were purchased

from Sigma-Aldrich (St Louis, MO, USA). All stock solutions were made in the

laboratory.

2.9. Ecdysteroid extraction

Animal were divided into the cephalothorax (CT) and abdomen/genital (Ab/G)

segment and samples were added 1 ml acetonitrile and homogenized using a 5 mm

stainless steel beads (Qiagen) in a Tissuelyser LT (Qiagen) for 3 x 2 min with 1 min

cooling on ice between each homogenization step followed by 10 min of vortexing.

The homogenate was centrifuged at 16500G for 5 min at 4 °C and the supernatant was

transferred to a 15 ml polypropylene (PP) tube and steam dried with N2 (12-15 psi) at

35 °C using a TurboVap LV evaporator (Zymark). Extracts were re-dissolved in 50 μl

MeOH/H2O (50/50 v/v), vortexed and transferred to High Performance Liquid

Chromatography (HPLC) vials and subjected to LC/MS/MS analysis. Five biological

parallels were subjected to analysis for each life cycle stage analysed.

10

2.10. LC/MS/MS analysis

Samples were analysed in an LC/MS/MS system consisting of a PerkinElmer series

200 HPLC system (Shelton, CT) and an API4000 triple quadruple mass spectrometer

(AB SCIEX; Foster City, CA) containing an electrospray ionization source. HPLC

separation was performed using a Discovery® C18 HPLC Column (15 cm x 2.1 mm,

5 μm; Supelco, Sigma-Aldrich), 300 μl/min flow rate at room temp. Analytes were

separated by methanol and 0.1 % acetic acid under the gradient conditions presented

in Table. 2. Injection volume of sample was 30 μl. MS/MS analysis was performed

under the following conditions: Curtain gas (CUR), 30 L/min; Collision gas (CAD),

4.0 L/min; Ion Source Gas (GS1), 20 V; Ion Source Gas (GS2), 70 V; Ion spray

voltage (IS) 5000; Temperature (TEM) 400°C; Declustering potential (DP), 95.0 V;

Entrance potential (EP), 9 V; Collision energy (CE), 20.0 V; Collision cell exit

potential (CXP), 11.0 V. Selected reaction monitoring (SRM) was executed using the

transitions specific for E, 20-E and PonA (Table. 3). Obtained results were analysed

using Analyst® 1.6.1 Software (AB SCIEX; Foster City, CA). Concentrations of the

compounds in each sample were estimated using the peak areas of the SRM

chromatogram on the basis of a calibration curve constructed using a standard. For the

20E analyte, two fraction ions were detected due to an unspecific interfering retention

top. A sample of the analytes (1ng/mL) and a diluent sample (MeOH:H2O) was run

every ten samples to track any degradation of analytes or contamination during the

run.

2.11 Statistical analysis

Students T-tests were used to determine differences in expression between control

group and dsRNA treated animals. One-way ANOVA tests were used to determine

differences in steroid levels between days. A p-value of 0.05 was set as threshold for

both tests.

11

3. Results

3.1. Identification and sequencing of neverland (nvd) and the two Halloween genes

disembodied (dib) and shade (shd)

To verify the sequences of the selected genes, primers were designed from predicted

gene sequences from the L. salmonis genome (Licebase.org ; unpublished) and 5´and

3´RACE were performed (see Table. 1. for list of primers). We obtained partial

sequences of L. salmonis disembodied (LsDib; Genbank ACO13011.1) shade (LsShd;

accession no; submitted) and neverland (LsNvd; accession no; submitted). The

analysis of the genes revealed an open reading frame (ORF) of LsDib 1510 bp and

470 aa, LsNvd 1417 bp and 399 aa and LsShd 1590 bp and 530 aa. Alignment with

amino acid sequences from other species (listed in Table. 2) obtained through Blast

search (NCBI) revealed that LsNvd contain the characteristic Rieske [2Fe-2s] and the

non-heme Fe (II) domains (Fig. 1A). The five conserved cytochrome P450 domains; a

P/G rich motif, Helix-C, Helix-I, Helix-K, PERF and a Heme-binding motif were

identified in the obtained sequences for LsDib and LsShd (Fig. 1B, C).

3.2. Ontogenetic analysis of LsNvd, LsDib and LsShd using RT-qPCR

Expression levels bfor the different life stages of L. salmonis for LsNvd, LsDib and

LsShd were obtained using RT-qPCR (see Fig. 2A, B and C, respectively). The

relative expression of LsNvd was around 5 fold higher in adult females compared to

the other life stages where transcript was low (Fig. 2A.). A significantly higher

expression of LsNvd was also detected in adult pre-adult males compared to the adult

males (Fig. 2A; ANOVA, P < 0.05). In contrast to LsNvd, transcript expression of

both LsDib and LsShd was significantly higher in the early life stages of the life cycle

compared to the pre-adult and adult stages (Fig. 2B and C; ANOVA, P < 0.05). The

relative expression of LsNvd in was more than 18 fold higher than LsDib and LsShd in

the adult female stage, but only between 4 and 1.5 fold higher in the larval stages,

respectively.

12

3.3. Localisation of transcripts

In situ hybridisation was performed in order to localise the transcripts of LsNvd,

LsDib and LsShd (Fig. 3A-F). Sections of L. salmonis adult male and female louse

and copepodids (7 days post molting) were used based on the results from the

expression analysis (section 3.2). Parallel sections were incubated with sense probe as

a negative control. Transcript of LsNvd was detected in the ovaries of the adult female

(Fig. 3A). A weak staining in the ovaries was also observed for LsShd (data not

shown) but stronger signal was observed in the intestine (Fig. 3E). A strong transcript

signal was detected for LsShd in specific but unidentified cells in the male

spermatophore (Fig. 3D) and in most tissues of the copepodid stage including nerve

tissue (Fig. 3F). LsDib transcript was weakly detected throughout the copepodid

tissues (Fig 3C) and more prominently in the intestine of the adult stages (Fig 3B).

3.4. Functional studies of LsOctβ3R, LsNvd, LsDib and LsShd

To assess the function of the LsOctβ3R, LsNvd, LsDib and LsShd during early

development and molting, RNAi studies were performed by soaking nauplia I larvae

in dsRNA fragment specific for each gene (Eichner et al., 2014). The experiments

were terminated seven d.p.t. when control animals had molted to the copepodid stage.

Two or three different fragments were selected from the 5` and 3` regions of each of

the four transcripts and separately used in experiments in order to exclude any non-

target effects of the dsRNA treatment. The LsNvd and LsDib knock-down (LsNvdkd,

LsDibkd) animals all molted from the nauplia II to the copepodid stage and did not

show any morphological defects or abnormalities compared to the control animals. In

contrast, about 70 - 75 % of the LsShd knock-down (LsShdkd) animals did not molt

into copepodid stage (Fig. 4Cb-c). The remaining LsShdkd animals (Fig. 4Cd) were

similar to the control animals (Fig. 4Ca). Similar results were achieved in the

LsOctβ3R animals, where 85 - 90 % of the treated larvae were not able to molt into

the copepodid stage (Fig. 4Db-c). A small percentage of the LsOctβ3R knock-down

(LsOctβ3Rkd) animals had molted into copepodid but all of them was lying on the

back (Fig. 4Dd) and did not show normal swimming behaviour. The degree of knock-

13

down was assessed by RTq-PCR analysis. A significant decrease of LsOctβ3R,

LsNvd, LsDib and LsShd transcripts by 79 %, 95 %, 60 % and 74 %, respectively (Fig.

4A, T-test, P < 0.05) was confirmed. Due to the no-loss of function phenotype

obtained after LsNvd and LsDib knock-down, an experiment with collective knock-

down of both genes was performed. Although a significant transcript knock-down was

achieved (data not shown) no phenotype was observed for the copepodids.

3.4.1. LsOctβ3R regulates the expression of ecdysone biosynthetic genes and the

ecdysone receptor

It has recently been shown that Octβ3R is essential for expression of biosynthetic

genes Drosophila, we investigate if loss of LsOctβ3R expression affected the

transcript level of the three ecdysteroid biosynthesis genes as well as the EcR/RXR

receptor complex. RT-qPCR revealed a significant down-regulation for LsNvd (68 %),

while LsShd and LsEcR were up-regulated by as much as 161 and 131 %,

respectively, at 7 d.p.t. (Fig. 4B, T-test: P < 0.05). LsDib and LsRXR did not show any

significant change in expression compared to the control group.

3.5 Down-regulation of LsNvd in adult female lice resulted in aberrations in the

ovaries and oocytes

Ontogenetic transcript analysis showed that expression of LsNvd was highest in the

adult female animals (section 3.2). Based on this result we wanted to investigate the

effect of LsNvd knock-down (LsNvdkd) in adult females using RNAi. In total, 36-39

pre-adult II female lice were injected with double-stranded LsNvd (dsLsNvd) or

control fragment encoding a trypsin from Atlantic cod (Gadus morhua). When the

experiment were terminated, no gross morphological differences or increase in

mortality was observed for the LsNvdkd lice compared to the control. The number of

recovered LsEcRkd adult female lice from the two separate experiments was 16 and

15 compared to 11 and 19 recovered lice from the control group. The first set of egg-

strings collected from the LsNvdkd lice (Fig. 5D) hatched and molted from the nauplia

to the copepodid stage similar to the control (Fig. 5A). However, histological

examination in the adult LsNvdkd lice showed that the lining of germinal vesicle of

14

the oocyte, the ovary and the maturing oocyte inside the genital segment was irregular

in shape (Fig. 5E-F) compared to the control (Fig. 5B-C). In addition, aberrations

were observed in the intestinal tissue (data not shown). RT-qPCR analysis confirmed

a 69 % knock-down of LsNvd 35 days after injection (T-test, P < 0.05).

3.6. Ecdysteroid titer fluctuates during egg maturation

LC/MS/MS in SRM mode (Table. 3.) was used for ecdysteroid analysis. The

corresponding peaks in the SRM chromatograms of the reference and extracted

samples were used to identify ecdysteroids present. We applied the method to identify

the ecdysteroids E, 20E and PonA in adult female lice during oocyte maturation (10

days cycle). As shown in Fig. 6A, the adult female lice were divided in two parts: the

cephalothorax (CT) and the abdomen/genital segment (Ab/G) containing primarily

maturing oocytes in order to detect any differences in ecdysteroid content and level

between the two segments. A significant difference was detected between E and 20E

expression in both the CT (P < 0.05) and the Ab/G (P < 0.01) (Fig. 6B-C). In addition,

the total ecdysteroid content in the Ab/G segment was significantly higher compared

to the CT (P < 0.05). In the CT, E concentration peaked at day two at 2.56 ng/g before

a significantly drop to 0.87 ng/g was detected at day three (P < 0.05) (Fig. 6B-C). The

E concentration remained stable between days 3-6 until it increased and peaked at the

end of oocyte maturation (Fig. 3B, day 8, 3.01 ng/g). The same trend was observed

for 20E where 20E level peaked at 0.54 ng/g (Fig. 6B, day 10) just before the oocytes

are excreted from the female. A similar pattern in the ecdysteroid titer was observed

in the Ab/G segment where the E concentration dropped from 8.35 to 1.43 ng/g (P <

0.01) between days two and three before it increased from 3.78 ng/g at day nine and

peaked at 8.78 ng/g day ten (Fig. 6C, P < 0.05). In addition, PonA was observed at

low levels around the detection limit in the extract of the Ab/G but not in the CT.

PonA concentration remained low (0.061 - 0.131 ng/g) through oocyte maturation and

no significant difference in PonA levels could be detected (Fig. 6C).

3.7 Ponasterone A is the predominant ecdysteroid in the pre-molt stage of L. salmonis

copepodids

15

LC/MS/MS analysis was performed under the same conditions as described in section

2.10 (Table. 3.) to investigate the ecdysteroid content and levels during larval stages. .

It is evident that E, 20E and PonA are present in nauplia II and free-swimming and

parasitic copepodids. The total ecdysteroid levels increase from the free-swimming

copepodid stage to the parasitic copepodid stage two days after infection, before a

drop is detected between day two and four post infection when molting is initiated

(Fig. 6D). PonA was detected as the main hormone in the parasitic copepodid and

peaked at ~50 ng/animal, while the E level (~25 ng/animal) was higher compared to

20E ~12 ng/animal (2 days post infection). Since only two or three biological parallels

were run for each stage, no statistics was performed on the data.

4. Discussion

The steroid hormone 20-hydroxyecdysone is known to coordinate the execution of

embryonic and post-embryonic development in many arthropods. Through a series of

enzymatic steps, cholesterol is converted to the biologically active 20E that binds to

the EcR/RXR nuclear complex and initiates a range of physiological processes. The

structure of the enzymes involved in ecdysteroid biosynthesis are highly conserved

between insects and crustaceans, however, the physiological function in marine

invertebrates has received little attention. In this study, we identified orthologs of the

insect neverland (nvd) and the two cytochrome P450 enzymes disembodied (dib) and

shade (shd) in L. salmonis, which is a crustacean belonging to the copepoda subclass,

family caligida. Sequence alignment showed that the amino acid sequence of LsNvd

exhibited the typical Rieske [2Fe-2S] binding motif and a C-terminal non-heme iron-

binding domain (Fig. 1A.) while the conserved P450 motifs (Helix-C, Helix-I, Helix-

K, a PERF-motif and a heme-binding domain) were identified in the primary structure

of LsDib and LsShd (Fig. 1B-C). This indicates that the three genes identified in L.

salmonis are functionally equivalent to their insect orthologs and are part of the

ecdysteroid synthesis in the salmon louse. Although the biosynthesis of ecdysteroids

has not been studied in details in crustaceans these results support the view that the

biosynthetic pathway in L. salmonis is similar to the pathway present in insects.

In situ hybridisation was performed to identify the tissues where the

ecdysteroidogenic genes are transcribed. Low transcript levels of the genes made the

16

localisation studies challenging and RNA-seq data from Licebase.org (unpublished)

was used as support. In adulfemales, LsNvd was found in the ovaries using in situ,

however, tissue specific RNA-seq data showed additional expression of LsNvd

transcript in the unfertilised oocytes, intestine and male testes (unpublished;

Licebase.org). These results are similar to the expression pattern found for the two

nvd paralogs identified in Daphnia magna (DmNvd) ref. Both LsDib and LsShd

transcript were detected in intestine and reproductive tissue of adult lice (Fig. 3 B, D,

E) and in most tissue of the copepodid larvae (Fig. 3C, F In crustaceans, the activity

of Halloween gene products are typically detected in ecdysone biosynthetic tissue

such as the Y-organ and ovaries (Mykles, 2011) with the exception of shd, which is

found in a variety of peripheral tissue such as the fat body, malphigian tubules and

midgut (Iga and Smagghe, 2010; Sumiya et al., 2014). In insects, ecdysteroid

synthesis takes place in the prothoracic glands of larval stages and the ovaries of the

adults (Iga and Smagghe, 2010; Rewitz et al., 2007; Rewitz et al., 2006). The

presence of both CYP genes LsDib and LsShd in reproductive tissue and intestine of

L. salmonis suggests that both the biosynthesis of ecdysone and its conversion to the

active metabolite 20E primarily takes place in the gonads and intestine before likely

being excreted into the hemolymph and distributed to target tissues. This is supported

by microarray analysis of the L. salmonis intestine where all 28 cytochrome P450

genes identified in the L. salmonis genome were expressed (Edvardsen et al., 2014).

The widespread transcript distribution of the Halloween genes in copepodids indicates

that ecdysteroidogenesis is not restricted to specific tissues like the PG in immature

insects.

Having established the existence of a biosynthetic pathway for production of

ecdysones, further work was performed to measure the ecdysteroid level in L.

salmonis. Specifically we wanted to investigate the ecdysteroid titer during molting.

Unfortunately, due to technical difficulties we could not perform proper

measurements during the complete molting cycle. However, it is indicated from our

preliminary studies that PonA is the main hormone present during pre-molt in

copepodid stages, which is in accordance with prior studies performed in crustaceans

(Lachaise et al., 1993). The drop in total ecdysteroid level in copepodids between day

two and four before initiation of ecdysis, suggests that ecdysteroids play a key role in

the regulation of molting in L. salmonis (Fig. 6D) as well known from other

17

arthropods. Furthermore, we wanted to explore the ecdysteroid titer during oocyte

maturation and determine any differences in ecdysteroid content between the

maturing oocytes and the adult female. Here, measurements of ecdysteroids, clearly

demonstrated the existence of E, 20E and PonA in L. salmonis. Our study established

that the ecdysteroid levels significant change during maturation of the oocytes both in

the CT and the Ab/G segment of female lice (Fig. 6B-C). The level of ecdysteroids is,

however, significantly higher in the Ab/G segment compared to the CT. In addition,

PonA was present in the Ab/G segment at very low levels but below the detection

level in the CT, which implies that this steroid is utilized by the vitellogenic oocytes

but not by the ovaries in the adult female. A drop in E and 20E level is evident at the

start of vitellogenesis before both ecdysteroids increase in concentration at the end of

oocyte maturation. A similar increase in ecdysteroids in the oocytes was confirmed in

the shore crab Carcinus maenas (Styrishave et al., 2008) where ecdysteroids are

necessary for initiation of oocyte maturation. Higher concentrations of ecdysteroid

level were also observed in female Calanus finmarchicus with large egg sacs

indicating that ecdysteroids are involved in egg maturation and reproduction (Hansen

et al., 2008). RNA-seq data from L. salmonis (unpublished; Licebase.org) shows that

several members of the CYP450 genes involved in ecdysteroid synthesis are

expressed in the unfertilized eggs. This suggests that the oocytes are capable of de

novo synthesis of ecdysteroids in L. salmonis oocytes.

Due to the high expression of LsNvd in the ovaries of adult female lice, functional

assessment of LsNvd was performed using RNAi. Even though significant LsNvd

knock-down was achieved, the collected egg strings hatched into viable nauplia that

continued to molt into the copepodid stage. However, histological analysis revealed

irregularities in ovary and ooginia morphology and abnormal segmentation of the

oocyte maturing in the genital segment resembling the phenotype (Fig. 5) observed in

LsRXR knock-down lice (Eichner et al., 2015). This suggests that LsNvd plays a role

in ecdysteroid synthesis either directly or indirectly affect oocyte maturation in L.

salmonis.

In Drosophila, mutations in the Halloween genes interrupts the biosynthesis of

ecdysteroids and cause embryonic lethality (Gilbert and Warren, 2005). In light of

this, we wanted to explore the function of LsNvd, LsDib and LsShd using RNAi

18

during larval development of the salmon louse. Significant knock-down was achieved

for each gene (Fig. 4A), however, both LsNvdkd and LsDibkd animals molted from

nauplia II to the copeodid stage and no aberrant morphology was observed. The

results differ from findings in Drosophila where mutations in both nvd and dib

transcript disrupt development of the embryo and cause mortality due to a reduction

in ecdysteroid synthesis (Chavez et al., 2000; Yoshiyama et al., 2006) and may be

caused by activity of residual protein. In contrast to LsNvd and LsDib knock-down,

LsShdkd resulted in molting arrest in the nauplia II stage (Fig. 4C). Similar results

were observed in Drosophila where homozygous shd mutants were disrupted during

late embryonic morphogenesis and displayed the same phenotype as the nvd and dib

mutants (Petryk et al., 2003). From the results obtained from our knock-down studies

of LsNvd, LsDib and LsShd we propose the hypothesis that ecdysteroids are

synthesized in high levels in the oocytes during maturation and is converted to either

PonA or 20E by LsShd during embryogenesis and development of copepodids. While

expression of LsNvd and LsDib remains stable during larval stages, the expression of

LsShd is significantly higher in the nauplia and the parasitic copepodids compared to

the free-swimming copepodids (Fig. 2C). It is possible that after molting into the free-

swimming copepodid the ecdysteroid biosynthetic enzymes are not needed until the

copepodid becomes parasitic and starts feeding on the host. At this stage, cholesterol

is taken up through the diet and converted to ecdysteroids needed for essential

physiological processes. Infection trials using LsNvd and LsDib knock-down

copepodids would be necessary to support the theory as increased mortality of the

copepodids after infection would confirm the necessity of these enzymes in

conversion of dietary cholesterol during the parasitic stages.

G-coupled protein receptor has long been proposed as a potential regulator of

ecdysteroid biosynthesis both in insects and crustaceans (Gilbert et al., 2002; Smith

and Sedlmeier, 1990). In Drosophila, recent RNAi knock-down studies of Octβ3R in

the PG significantly reduced the expression of nvd and several Halloween genes. The

results showed that Octβ3R is required for the expression of ecdysone biosynthetic

genes (Ohhara et al., 2015). Knock-down of Octβ3R in L. salmonis resulted in an

arrest of molting or defects in the cuticula and secondary antenna of the copepodid

(Fig. 4D). The phenotype resembled that detected in Drosophila larvae however, it is

19

indicated from the RT-qPCR analysis of LsOctβ3Rkd animals (Fig. 4B) that the

ecdysteroid biosynthetic genes are regulated in a different manner in L. salmonis then

that of Drosophila.

Acknowledgements

We would like to thank former technical assistant at NMBU Rune Landsem,

technical assistant Lars Hamre and Per Gunnar Espedal at the SLRC in Bergen for

excellent technical assistance in the laboratory. This work was supported by the

Research Council of Norway, SFI-Sea Lice Research Centre, grant number

203513/O30.

20

References

Albright, S.C.J.a.L.J., 1991. The developmental stages of Lepeophtheirus salmonis. Canadian Journal of Zoology 69.

Asazuma, H., Nagata, S., Nagasawa, H., 2009. Inhibitory effect of molt-inhibiting

hormone on phantom expression in the y-organ of the kuruma prawn, Marsupenaeus japonicus. Arch Insect Biochem 72, 220-233.

Baldwin, W.S., Marko, P.B., Nelson, D.R., 2009. The cytochrome P450 (CYP) gene y

in Daphnia pulex. Bmc Genomics 10, 169. Chavez, V.M., Marques, G., Delbecque, J.P., Kobayashi, K., Hollingsworth, M., Burr,

J., Natzle, J.E., O'Connor, M.B., 2000. The Drosophila disembodied gene controls late embryonic morphogenesis and codes for a cytochrome P450 enzyme that regulates embryonic ecdysone levels. Development 127, 4115-4126.

Dalvin, S., Frost, P., Biering, E., Hamre, L.A., Eichner, C., Krossoy, B., Nilsen, F.,

2009. Functional characterisation of the maternal yolk-associated protein (LsYAP) utilising systemic RNA interference in the salmon louse (Lepeophtheirus salmonis) (Crustacea: Copepoda). Int J Parasitol 39, 1407-1415.

Dalvin, S., Frost, P., Loeffen, P., Skern-Mauritzen, R., Baban, J., Ronnestad, I.,

Nilsen, F., 2011. Characterisation of two vitellogenins in the salmon louse Lepeophtheirus salmonis: molecular, functional and evolutional analysis. Dis Aquat Organ 94, 211-224.

Edvardsen, R.B., Dalvin, S., Furmanek, T., Malde, K., Mæhle, S., Kvamme, B.O.,

Skern-Mauritzen, R., 2014. Gene expression in five salmon louse (Lepeophtheirus salmonis, Krøyer 1837) tissues. Marine Genomics.

Eichner, C., Dalvin, S., Skern-Mauritzen, R., Malde, K., Kongshaug, H., Nilsen, F.,

2015. Characterization of a novel RXR receptor in the salmon louse (Lepeophtheirus salmonis, Copepoda) regulating growth and female reproduction. Bmc Genomics 16, 81.

Eichner, C., Nilsen, F., Grotmol, S., Dalvin, S., 2014. A method for stable gene

knock-down by RNA interference in larvae of the salmon louse (Lepeophtheirus salmonis). Exp Parasitol 140, 44-51.

Feyereisen, R., 2006. Evolution of insect P450. Biochem Soc T 34, 1252-1255. Feyereisen, R., 2011. Arthropod CYPomes illustrate the tempo and mode in P450

evolution. Bba-Proteins Proteom 1814, 19-28. Frost, P., Nilsen, F., 2003. Validation of reference genes for transcription profiling in

the salmon louse, Lepeophtheirus salmonis, by quantitative real-time PCR. Vet Parasitol 118, 169-174.

21

Gilbert, L.I., Rybczynski, R., Warren, J.T., 2002. Control and biochemical nature of the ecdysteroidogenic pathway. Annu Rev Entomol 47, 883-916.

Gilbert, L.I., Warren, J.T., 2005. A molecular genetic approach to the biosynthesis of

the insect steroid molting hormone. Vitamins and Hormones 73, 31-57. Goad, L.J., 1981. Sterol Biosynthesis and Metabolism in Marine-Invertebrates. Pure

Appl Chem 53, 837-852. Gohar, M., Souty, C., 1984. Action temporelle d'ecdystéroïdes sur la synthèse

protéique ovarienne in vitro chez le Crustacé isopode terrestre Porcellio dilatatus (Brandt). Reproduction Nutrition Développement 24, 137-145.

Grieneisen, M.L., 1994. Recent Advances in Our Knowledge of Ecdysteroid

Biosynthesis in Insects and Crustaceans. Insect Biochem Molec 24, 115-132. Hamre, L.A., Eichner, C., Caipang, C.M.A., Dalvin, S.T., Bron, J.E., Nilsen, F.,

Boxshall, G., Skern-Mauritzen, R., 2013. The Salmon Louse Lepeophtheirus salmonis (Copepoda: Caligidae) Life Cycle Has Only Two Chalimus Stages. Plos One 8.

Hamre, L.A., Glover, K.A., Nilsen, F., 2009. Establishment and characterisation of

salmon louse (Lepeophtheirus salmonis (Kroyer 1837)) laboratory strains. Parasitol Int 58, 451-460.

Hansen, B.H., Altin, D., Hessen, K.M., Dahl, U., Breitholtz, M., Nordtug, T., Olsen,

A.J., 2008. Expression of ecdysteroids and cytochrome P450 enzymes during lipid turnover and reproduction in Calanus finmarchicus (Crustacea: Copepoda). Gen Comp Endocr 158, 115-121.

Hopkins, P.M., 2009. Crustacean ecdysteroids and their receptors, in: Smagghe, G.

(Ed.), Ecdysone: structures and functions. Springer, pp. 73-97. Iga, M., Kataoka, H., 2012. Recent Studies on Insect Hormone Metabolic Pathways

Mediated by Cytochrome P450 Enzymes. Biol Pharm Bull 35, 838-843. Iga, M., Smagghe, G., 2010. Identification and expression profile of Halloween genes

involved in ecdysteroid biosynthesis in Spodoptera littoralis. Peptides 31, 456-467.

Kvamme, B.O., Skern, R., Frost, P., Nilsen, F., 2004. Molecular characterisation of

five trypsin-like peptidase transcripts from the salmon louse (Lepeophtheirus salmonis) intestine. Int J Parasitol 34, 823-832.

Lachaise, F., Leroux, A., Hubert, M., Lafont, R., 1993. The Molting Gland of

Crustaceans - Localization, Activity, and Endocrine Control (a Review). J Crustacean Biol 13, 198-234.

Lafont, R., Mathieu, M., 2007. Steroids in aquatic invertebrates. Ecotoxicology 16,

109-130.

22

Markov, G.V., Tavares, R., Dauphin-Villemant, C., Demeneix, B.A., Baker, M.E.,

Laudet, V., 2009. Independent elaboration of steroid hormone signaling pathways in metazoans. P Natl Acad Sci USA 106, 11913-11918.

Martin-Creuzburg, D., Westerlund, S.A., Hoffmann, K.H., 2007. Ecdysterold levels in

Daphnia magna during a molt cycle: Determination by radioimmunoassay (RIA) and liquid chromatography-mass spectrometry (LC-MS). Gen Comp Endocr 151, 66-71.

Mykles, D.L., 2011. Ecdysteroid metabolism in crustaceans. Journal of Steroid

Biochemistry and Molecular Biology 127, 8. Namiki, T., Niwa, R., Sakudoh, T., Shirai, K., Takeuchi, H., Kataoka, H., 2005.

Cytochrome P450CYP307A1/spook: A regulator for ecdysone synthesis in insects. Biochemical and Biophysical Research Communications 337, 367-374.

Niwa, R., Matsuda, T., Yoshiyama, T., Namiki, T., Mita, K., Fujimoto, Y., Kataoka,

H., 2004. CYP306A1, a cytochrome P450 enzyme, is essential for ecdysteroid biosynthesis in the prothoracic glands of Bombyx and Drosophila. J Biol Chem 279, 35942-35949.

Niwa, R., Namiki, T., Ito, K., Shimada-Niwa, Y., Kiuchi, M., Kawaoka, S.,

Kayukawa, T., Banno, Y., Fujimoto, Y., Shigenobu, S., Kobayashi, S., Shimada, T., Katsuma, S., Shinoda, T., 2010. Non-molting glossy/shroud encodes a short-chain dehydrogenase/reductase that functions in the 'Black Box' of the ecdysteroid biosynthesis pathway. Development 137, 1991-1999.

Niwa, R., Niwa, Y.S., 2011. The fruit fly Drosophila melanogaster as a model system

to study cholesterol metabolism and homeostasis. Cholesterol 2011. Ohhara, Y., Shimada-Niwa, Y., Niwa, R., Kayashima, Y., Hayashi, Y., Akagi, K.,

Ueda, H., Yamakawa-Kobayashi, K., Kobayashi, S., 2015. Autocrine regulation of ecdysone synthesis by beta3-octopamine receptor in the prothoracic gland is essential for Drosophila metamorphosis. Proc Natl Acad Sci U S A 112, 1452-1457.

Ono, H., Rewitz, K.F., Shinoda, T., Itoyama, K., Petryk, A., Rybczynski, R., Jarcho,

M., Warren, J.T., Marques, G., Shimell, M.J., Gilbert, L.I., O'Connor, M.B., 2006. Spook and Spookier code for stage-specific components of the ecdysone biosynthetic pathway in Diptera. Dev Biol 298, 555-570.

Petryk, A., Warren, J.T., Marques, G., Jarcho, M.P., Gilbert, L.I., Kahler, J., Parvy,

J.P., Li, Y.T., Dauphin-Villemant, C., O'Connor, M.B., 2003. Shade is the Drosophila P450 enzyme that mediates the hydroxylation of ecdysone to the steroid insect molting hormone 20-hydroxyecdysone. P Natl Acad Sci USA 100, 13773-13778.

23

Pike, A.W., Wadsworth, S.L., 2000. Sealice on salmonids: Their biology and control, in: Baker, J.R., Muller, R., Rollinson, D. (Eds.), Advances in Parasitology, Vol 44, pp. 233-337.

Rees, H.H., 1995. Ecdysteroid Biosynthesis and Inactivation in Relation to Function.

Eur J Entomol 92, 9-39. Rewitz, K.F., Gilbert, L.I., 2008. Daphnia Halloween genes that encode cytochrome

P450s mediating the synthesis of the arthropod molting hormone: Evolutionary implications. Bmc Evol Biol 8.

Rewitz, K.F., O'Connor, M.B., Gilbert, L.I., 2007. Molecular evolution of the insect

Halloween family of cytochrome P450s: Phylogeny, gene organization and functional conservation. Insect Biochem Molec 37, 741-753.

Rewitz, K.F., Rybczynski, R., Warren, J.T., Gilbert, L.I., 2006. Identification,

characterization and developmental expression of Halloween genes encoding P450 enzymes mediating ecdysone biosynthesis in the tobacco hornworm, Manduca sexta. Insect Biochem Molec 36, 188-199.

Sandlund, L., Nilsen, F., Male, R., Grotmol, S., Kongshaug, H., Dalvin, S., 2015.

Molecular characterisation of the salmon louse, Lepeophtheirus salmonis salmonis (Kroyer, 1837), ecdysone receptor with emphasis on functional studies of female reproduction. Int J Parasitol 45, 175-185.

Schram, T.A., 1993. Supplementary descriptions of the develompemntal stages of

Lepeophtheirus salmonis (Krøyer 1837) (Copepoda: Caligidae), in: Defaye, G.B.D. (Ed.), Pathogens of wild and farmed fish: sea lice. New York: Ellis Horwood, pp. 30-47.

Skern-Mauritzen, R., Torrissen, O., Glover, K.A., 2014. Pacific and Atlantic

Lepeophtheirus salmonis (Kroyer, 1838) are allopatric subspecies: Lepeophtheirus salmonis salmonis and L. salmonis oncorhynchi subspecies novo. Bmc Genet 15.

Smith, W.A., Sedlmeier, D., 1990. Neurohormonal Control of Ecdysone Production -

Comparison of Insects and Crustaceans. Invertebr Reprod Dev 18, 77-89. Spaziani, E., Mattson, M.P., Wang, W.N.L., McDougall, H.E., 1999. Signaling

pathways for ecdysteroid hormone synthesis in crustacean Y-organs. Am Zool 39, 496-512.

Styrishave, B., Lund, T., Andersen, O., 2008. Ecdysteroids in female shore crabs

Carcinus maenas during the moulting cycle and oocyte development. J Mar Biol Assoc Uk 88, 575-581.

Subramoniam, T., 2000. Crustacean ecdysteriods in reproduction and embryogenesis.

Comp Biochem Physiol C Toxicol Pharmacol 125, 135-156.

24

Sumiya, E., Ogino, Y., Miyakawa, H., Hiruta, C., Toyota, K., Miyagawa, S., Iguchi, T., 2014. Roles of ecdysteroids for progression of reproductive cycle in the fresh water crustacean Daphnia magna. Front Zool 11.

Tom, M., Manfrin, C., Giulianini, P.G., Pallavicini, A., 2013. Crustacean oxi-

reductases protein sequences derived from a functional genomic project potentially involved in ecdysteroid hormones metabolism - A starting point for function examination. Gen Comp Endocr 194, 71-80.

Warren, J.T., Petryk, A., Marques, G., Jarcho, M., Parvy, J.P., Dauphin-Villemant, C.,

O'Connor, M.B., Gilbert, L.I., 2002. Molecular and biochemical characterization of two P450 enzymes in the ecdysteroidogenic pathway of Drosophila melanogaster. P Natl Acad Sci USA 99, 11043-11048.

Warren, J.T., Petryk, A., Marques, G., Parvy, J.P., Shinoda, T., Itoyama, K.,

Kobayashi, J., Jarcho, M., Li, Y., O'Connor, M.B., Dauphin-Villemant, C., Gilbert, L.I., 2004. Phantom encodes the 25-hydroxylase of Drosophila melanogaster and Bombyx mori: a P450 enzyme critical in ecdysone biosynthesis. Insect Biochem Mol Biol 34, 991-1010.

Yoshiyama, T., Namiki, T., Mita, K., Kataoka, H., Niwa, R., 2006. Neverland is an

evolutionally conserved Rieske-domain protein that is essential for ecdysone synthesis and insect growth. Development 133, 2565-2574.

Yoshiyama-Yanagawa, T., Enya, S., Shimada-Niwa, Y., Yaguchi, S., Haramoto, Y.,

Matsuya, T., Shiomi, K., Sasakura, Y., Takahashi, S., Asashima, M., Kataoka, H., Niwa, R., 2011. The Conserved Rieske Oxygenase DAF-36/Neverland Is a Novel Cholesterol-metabolizing Enzyme. J Biol Chem 286, 25756-25762.

25

Figure legends

Fig. 1. Alignment of deduced amino acid (aa) sequences of Lepeophtheirus salmonis

7,8-dehydrodenase/neverland (LsNvd) (A), disembodied (LsDib) (B), and shade

(LsShd) (C) with other species. A) The conserved Rieske motif and the non-heme

domain are market off in the alignment. The sequences used in the alignments were as

follows; L. salmonis nvd (LsNvd; accession no. submitted), Daphnia wassermani

(DwNvd; AFD97360.1), Crassostrea gigas cholesterol desaturase daf-36-like

(CgDaf-36; XP_011445555.1), Xenopus laevis neverland (XlNvd; BAK39959.1),

Clupea harengus cholesterol desaturase daf-36-like (ChDaf-36; XP_012685571.1). B)

Conserved P450 motifs are shown. The sequences used in the alignments were as

follows; L. salmonis disembodied (LsDib; Genbank accession no. XX), Daphnia

pulex disembodied (DpDib; EFX63066.1), Paracyclopina nana CYP 302A1 (PnDib;

AKH03533.1), Leptinotarsa decemlineata CYP 302A1 (LdDib; AGT57842.1),

Tigriopus japonicus CYP 302A1 (TjDib; AIL94171.1). C) Conserved P450 motifs

are shown. The sequences used in the alignments were as follows; L. salmonis shade

(LsShd; accession no submitted), Tigriopus japonicus shade (TjShd; AIL94172.1),

Daphnia magna (DmShd; BAF35770.1), Paracyclopina nana (PaShd; AKH03534.1)

and Sogatella furcifera (SfShd; AGI92296.1).

Fig. 2. Quantitative real-time PCR analysis of relative expression of A) LsNvd, B)

LsDib and C) LsShd in different developmental stages. Each point represents the

mean and confidence intervals (n= 5 parallels of approx. 150 animals for the nauplia

I/II and free-living copepodid (Free. Cop.) stages, 10 animals for the parasitic

copepodid (Par. Cop.), Chalimus I (Chal. I) and Chalimus II (Chal. II.) and one

animal for the pre-adult male (Pre-A. M.), pre-adult female (Pre-A. F.), adult male

(Adult M.) and adult female (Adult F.). The relative expression of the nauplia stage

was set to 1. Different letters denote significant differences between stages (ANOVA;

p < 0.05). Note different scaling of the axes.

Fig. 3. Localisation of Lepeophtheirus salmonis LsNvd, A) LsDib B-C) and LsShd D-

F) transcripts. In situ hybridisation was performed for each gene on sections from life

stages where transcript was highest expressed using specific anti-sense RNA. Negative

controls (sense RNA) applied to parallel sections are shown framed in corners.

26

Positive staining was seen for LsNvd in adult female ovaries (A). LsDib was detected

in the intestine of the adult female and weakly throughout the copepodid (B, C). A

strong positive signal was seen for LsShd in specific cells in the adult male

spermatophore, adult female intestine and throughout the copepodid (D, E, F).

Scalebar; A) = 300 μm; B, D, E) = 200 μm and C, F) = 100 μm.

Fig. 4. Functional assessment of Lepeophtheirus salmonis LsNvd, LsDib, LsShd and

LsOct3βR by RNA interference in copepodids. (A) Transcript level of LsNvd, LsDib,

LsShd and LsOct3βR after injection of dsRNA. (B) Transcript level of LsNvd, LsDib,

LsShd, LsEcR and LsRXR in LsOct3βR knock-down lice. The expression levels of the

respective genes in the control groups were set to 1. Mean ± Confidence Interval of

treated copepodid samples (n = 5) is shown in graphs. *Statistically significant (T-

test; P < 0.05). Phenotype of salmon louse larvae treated with dsRNA specific to

LsShd (Cb-d) and LsOct3βR (Dc-d). Controls are shown in Ca and Da. Scalebar = 300

μm.

Fig. 5. Functional assessment of Lepeophtheirus salmonis LsNvd using RNA

interference in adult female lice. The control lice produced normal egg strings (A) that

hatched and gave viable offspring. The first egg string of the LsNvd dsRNA treated

lice (D) hatched normally and did not show any macroscopic differences compared to

the control lice. Histological sections (stained with Toluidine) showed a less

organized packing of the oocytes in the LsNvdkd lice (E) compared to the control (C).

In addition, irregularities in the lining of the ovaries and oogonia were evident in the

LsNvdkd (F) animals compared to the control (C). Scalebar (A, D) = 1 mm, (B, E) =

100 μm and (C, F) = 500 μm.

Fig. 6. Detection of ecdysteroid level using tandem mass spectrometry. (A) Schematic

presentation of separation carried out in adult female lice; CT: cephalothorax and

Ab/G: abdomen/genital segment. The level of ecdysone E, 20-hydroxyecdysone (20E)

and ponasterone A (PonA) was investigated in the CT (B), Ab/G (C) and free-

swimming and parasitic larval stages (D). Different letters denote significant

differences between stages in B, C (ANOVA; p < 0.05). No statistical analysis was

performed on larval samples due to few samples. Ecdysteroid level is measured in

27

ng/g in adult females and ng/animal in larval stages. Note different scaling of the axes

and name of Y-axis.

28

a SYBR® Green assays were provided by Applied Biosystems, Branchurg, NJ, USA. b All general primers were purchased from Sigma-Aldrich, St Louis, MO, USA.

RACE, rapid amplification of cDNA ends; TOPO, DNA topoisomerase I; dsRNA, double-stranded RNA; RTq-

PCR, real-time quantitative PCR

Primer nameb Sequence (5`-3`) Method

LsNvd_5´RACE GTCACAGGTCTCCACAAGGGTCTTTCAG RACE

LsNvd_3´RACE CTGAAAGACCCTTGTGGAGACCTGTGAC RACE

LsDib_5´RACE TTCCACAGGGGAGGTCCATTGTCCG RACE

LsDib_3´RACE CTGTGCCGAAAGGGACTGTTCTTGTGAG RACE

LsShd_5´RACE GCGAGCAGTGTCCAAGGCAATAGTGTCG RACE

LsShd_3´RACE CCTTGCCGACACTATTGCCTTGGACACT RACE

LsShd_3´RACE_nested TCACAGCAGGAGTAGACACCATAGG RACE

M13_f GTAAAACGACGGCCAG Topo cloning

M13_r CAGGAAACAGCTATGAC Topo cloning

Cod_specific ATAGGGCGAATTGGGTACCG dsRNA

Cod_specific AAAGGGAACAAAAGCTGGAGC dsRNA

LsNvd_P1_F GACCCTTGTGGAGACCTGTG dsRNA/In situ

LsNvd_P1_R TTGGCAATGTGGGGATTGGT dsRNA/In situ

LsNvd_ P2_F GGAGAGGGTCGATACCACCA dsRNA

LsNvd_ P2_R GCATTCACCTACGGAAAGACTG dsRNA

LsDib_P1_F CCTCCCCTGTGGAAGGTTTTT dsRNA/In situ

LsDib_P1_R GAACAGTCCCTTTCGGCACA dsRNA/In situ

LsDib_P2_R AGCTGAAGCCCTTTTCATCCA dsRNA

LsDib_P2_R TCTGGGTCCAAATCCAAAGGG dsRNA

LsShd_P1_F ACCGTCATTTTCGCCCTTCT dsRNA/In situ

LsShd_P1_R AGGCTCTTGTTGTGTGCGTA dsRNA/In situ

LsShd_P2_F AAGGTACTCAAAGTGCTCATGCT dsRNA

LsShd_P2_R AAGGCTCTTGTTGTGTGCGT dsRNA

LsOct3βR_P1_F AGAGGATGAGAGCCTTCCCT dsRNA/In situ

LsOct3βR_P1_R TCAACCCTCGATCGAAACCAG dsRNA/In situ

LsOct3βR_P2_F AATCTGACACAGGAGGCAGC dsRNA

LsOct3βR_P2_R CCGTCCAGGCTACTCCAATC dsRNA

LsEF1α_F_ SYBR® CATCGCCTGCAAGTTTAACCAAATT RT-qPCR

LsEF1α_R_ SYBR® CCGGCATCACCAGACTTGA RT-qPCR

LsNvd F_SYBR® AACCAATCCCCACATTGCCA RT-qPCR

LsNvd R_SYBR® GGCCAGAAGCGATTTGTGTA RT-qPCR

LsDib_F_ SYBR® ACCGTCATTTTCGCCCTTCT RT-qPCR

LsDib_R_ SYBR® GCCTGGAGGAAGTGAGTGTC RT-qPCR

LsShd_F_ SYBR® GCTATGGGCCTGTAGTGAGG RT-qPCR

LsShd_R_ SYBR® AACATCCGCCTCATTCGGAG RT-qPCR

LsOct3βR_F_ SYBR® ATCCTCAGTTCCCTCGCAAC RT-qPCR

LsOct3βR_R_ SYBR® ATCCTCTGTTGAGGGGCTGA RT-qPCR

LsEcR_F_ SYBR® TCGCCCAACTCACGATTCAG RT-qPCR

LsEcR_R_ SYBR® GGGGAGTAAGGATGGGGTTC RT-qPCR

LsRXR_F_ SYBR® CCTAGTTGAACTCATCGCCAAAATG RT-qPCR

LsRXR_R_ SYBR® TGAAGAGTATGATGGCTCGTAGACA RT-qPCR

P i b S (5` 3`)Table. 1. Primer sequences and SYBR® Green assays used in this study.

29

Table. 2. Gradient for analysis of selected ecdysteroids of this study

Time (min) Flow (µL/min) A % B %

0 300 80 20

25 300 10 90

28 300 10 90

30 300 80 20

50 300 80 20

Table. 3. SRM conditions used in tandem mass spectrometry analysis.

Ecdysteroids Retention time (min) SRM transistion(m/z) CE (V)

E 19.7 465/429 20

20E 18 481/445 20

PonA 24.7 465/447 30 SRM: selected reaction monitoring, CE: collision energy, E: ecdysone, 20E:

20-hydroxyecdysone, PonA: ponasterone A.

30

Fig. 1.

31

Fig. 1.

32

Fig. 2.

33

F

FFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFFF

Fig. 3.

34

Fig. 4.

35

Fig. 5.

36

Fig. 6.

A

Ab/G CT

D

B