The lethal and sublethal effects of the anti-sea lice formulation Salmosan® on the Pacific spot prawn

(Pandalus platyceros)

by Kate Mill

B.Sc. (Hons., Environmental Chemistry), Queen’s University, 2014

Project Submitted in Partial Fulfillment of the

Requirements for the Degree of

Master of Environmental Toxicology

in the

Department of Biological Sciences

Faculty of Science

© Kate Mill 2019

SIMON FRASER UNIVERSITY

Spring 2019

Copyright in this work rests with the author. Please ensure that any reproduction or re-use is done in accordance with the relevant national copyright legislation.

ii

Approval

Name: Kate Mill

Degree: Master of Environmental Toxicology

Title: The lethal and sublethal effects of the anti-sea lice formulation Salmosan® on the Pacific spot prawn (Pandalus platyceros)

Examining Committee: Chair: Jonathan Moore Associate Professor

Chris Kennedy Senior Supervisor Professor

Vicki Marlatt Supervisor Assistant Professor

Curtis Eickhoff External Examiner Senior Environmental Toxicologist Nautilus Environmental

Date Defended/Approved: 25 April 2019

iii

Abstract

Sea lice outbreaks in salmonid aquaculture can impact both farmed and wild salmon. Anti-

sea lice chemotherapeutants used to treat these outbreaks are released directly into the

water column after treatment, potentially exposing non-target organisms. Salmosan®

(active ingredient: azamethiphos) has recently been approved for use in British Columbia

as a sea lice treatment. In the present study, the lethal and sublethal effects of Salmosan®

were examined in intermolt and post-molt Pacific spot prawns (Pandalus platyceros). Post-

molt prawns were found to be more sensitive than intermolt prawns, and this sensitivity

was exacerbated at higher exposure temperatures. Repeated (3 x) 1-h LC50 values for

post-molt prawns were 39.8, 27.1, and 17.1 µg/L at 5, 11 and 17 °C, respectively. All

intermolt prawns survived 3 x 1-h exposures up to 100 µg/L azamethiphos at 5, 11 and

17 °C. Intermolt prawns held at 17 °C molted 83 – 91% sooner and experienced 70 – 73%

greater mortality than those held at 5 or 11 °C; azamethiphos did not affect either of these

parameters. In a separate experiment, intermolt prawns displayed an 86 – 103% reduction

in antennule flicking, a chemoreception-mediated behavior, at 24 h following repeated

(3 x) 1-h exposures to 50 and 100 µg/L azamethiphos. These results may aid in the

development of regulatory protocols and guidelines for the use of anti-sea lice pesticides

in Canada.

Keywords: Aquaculture, sea lice, prawn, Salmosan®, azamethiphos, toxicity, pesticide, molt, behaviour, lethality

iv

Acknowledgements

I would first like to thank my supervisory committee for their support in this project.

Chris, thank you for allowing me the flexibility to pursue research questions within my

realm of interest, as well as to pursue interests outside the realm of my research. I leave

equipped with a jampacked toxicological toolbox and an ability to send thorough yet

concise emails. Vicki, thank you for your support in this project, and for piquing my curiosity

throughout this degree. There is a swarm of MET students looking for endocrine disruptors

wherever they go.

Many thanks to all the people who made the cold room a lot warmer. My army of

undergrad volunteers, thank you for your constant level of stoke about prawns: Meg

Ludlam, Cole Milton, Mariah Mund, Maryam Shakeri, Manpreet Jhutty, Harman Malhans,

Donya Divsalar, Jordan Wilson, Mi Mumuwin. Special thanks to Steven Barrett, Jill

Bennett and Kassie Rhodenizer for your prawn collaboration (mix tape coming soon).

Endless thanks to Bruce Leighton for never letting us be without solutions or seawater. BMSC humans – thank you for lending your brilliant minds and, equally cherished, your

ready distractions. A special shout out to Tam Hillhouse and Sian Mill for lending their

computers as mine made moves towards the electronic afterlife. Kennedy and Marlatt lab

inhabitants, thanks for keeping the bar high and the hilarity higher. Finally, and most

importantly, my family – thank you for the support in all shapes and forms, I promise I will

make the most of it.

This project was funded by the National Contaminants Advisory Group. For optimal

palatability, this thesis pairs best with a light-bodied pinot noir.

v

Table of Contents

Approval ............................................................................................................................ ii Abstract ............................................................................................................................ iii Acknowledgements .......................................................................................................... iv Table of Contents .............................................................................................................. v List of Tables ................................................................................................................... vii List of Figures ................................................................................................................. viii List of Acronyms ............................................................................................................... ix

Chapter 1. Introduction ................................................................................................ 1 1.1. Open net pen farming in Canada ............................................................................ 1 1.2. Sea lice ................................................................................................................... 2

1.2.1. Sea lice and salmon ........................................................................................ 2 1.2.2. Sea lice and open net pens ............................................................................. 3

1.3. Anti-sea lice chemotherapeutants in Canada ......................................................... 4 1.3.1. Regulation ....................................................................................................... 4 1.3.2. Current use treatments .................................................................................... 5

1.4. Risk of Salmosan® to non-target organisms ........................................................... 7 1.4.1. Environmental fate ........................................................................................... 7 1.4.2. Exposure ......................................................................................................... 7 1.4.3. Toxicity ............................................................................................................ 8

1.5. Study organism: Pacific spot prawn ...................................................................... 21 1.5.1. BC spot prawn fishery ................................................................................... 21 1.5.2. Life history and habitat .................................................................................. 22

1.6. Environmental temperature and toxicity ............................................................... 22 1.7. Study goal and objectives ..................................................................................... 25

Chapter 2. The lethal and sublethal effects of Salmosan® on molting in the Pacific spot prawn, Pandalus platyceros .......................................................... 26

2.1. Introduction ........................................................................................................... 26 2.2. Material and methods ........................................................................................... 28

2.2.1. Organism collection and holding ................................................................... 28 2.2.2. Chemicals ...................................................................................................... 28 2.2.3. The effect of Salmosan® exposures on intermolt stage prawns .................... 28 2.2.4. The effect of Salmosan® exposures on post-molt stage prawns ................... 30 2.2.5. Statistical analysis ......................................................................................... 32

2.3. Results .................................................................................................................. 33 2.3.1. Water quality .................................................................................................. 33 2.3.2. The effect of Salmosan® exposures on intermolt stage prawns .................... 34 2.3.3. The effect of Salmosan® exposures on post-molt stage prawns ................... 37

2.4. Discussion ............................................................................................................ 38 2.4.1. The effect of Salmosan® exposures on intermolt stage prawns .................... 38 2.4.2. The effect of Salmosan® exposures on post-molt stage prawns ................... 40

vi

Chapter 3. The effect of Salmosan® on chemosensory-mediated behaviours of the Pacific spot prawn, Pandalus platyceros .................................................... 46

3.1. Introduction ........................................................................................................... 46 3.2. Methods ................................................................................................................ 48

3.2.1. Organism collection and holding ................................................................... 48 3.2.2. Chemicals ...................................................................................................... 48 3.2.3. The effect of Salmosan® exposures on prawn behaviour during exposures . 48 3.2.4. The effect of Salmosan® exposures on behavioural responses to L-glutamate .................................................................................................................. 50 3.2.5. Statistical analysis ......................................................................................... 51

3.3. Results .................................................................................................................. 52 3.3.1. Water quality .................................................................................................. 52 3.3.2. The effect of Salmosan® exposures on prawn behaviour during exposures . 52 3.3.3. The effect of Salmosan® exposures on behavioural responses to L-glutamate .................................................................................................................. 54

3.4. Discussion ............................................................................................................ 57 3.4.1. The effect of Salmosan® exposures on prawn behaviour during exposures . 57 3.4.2. The effect of Salmosan® exposures on behavioural responses to L-glutamate .................................................................................................................. 60

Chapter 4. Conclusion ............................................................................................... 62

References ..................................................................................................................... 64

vii

List of Tables

Table 1. Summary of the lethal toxicity values for azamethiphos for various aquatic species ..................................................................................................... 12

Table 2. Summary of sublethal toxicity values for azamethiphos in various aquatic species ..................................................................................................... 17

Table 3. Estimated LC50 values (95% confidence intervals) and NOECs for intermolt and post-molt prawns ................................................................ 38

Table 4. Definitions for prawn behaviours examined in behavioural experiments (adapted from Lee and Meyers (1996) and Park (2013). ......................... 50

viii

List of Figures

Figure 1.. Schematic diagram of intermolt prawn exposure design. Intermolt prawns were acclimated at 5, 11 or 17 °C with 2 prawns per tank ....................... 29

Figure 2. Schematic diagram of post-molt prawn exposure design ........................ 32 Figure 3. Effect of Salmosan® and temperature on survival during molt of Pacific

spot prawns after 3 x 1-h exposures to Salmosan® (active ingredient: azamethiphos) at various temperatures ................................................... 35

Figure 4. Effect of Salmosan® and temperature on the time to molt of male Pacific spot prawns after 3 x 1-h exposures to Salmosan® (active ingredient: azamethiphos) at various temperatures ................................................... 35

Figure 5. Effect of Salmosan® and temperature on the relative change in carapace length (CL) of Pacific spot prawns ........................................................... 36

Figure 6. Effect of Salmosan® and temperature on the relative change in mass of Pacific spot prawns .................................................................................. 36

Figure 7. Effect of Salmosan® and temperature on Fulton’s K condition factor of male Pacific spot prawns ......................................................................... 37

Figure 8. Schematic diagram of behavioural experiment design ............................ 49 Figure 9. Total time spent moving during 3 x 1-h exposures of prawns to

Salmosan® ............................................................................................... 53 Figure 10. Feeding response of prawns measured by the change in number of

antennule flicks (∆ flicks) in response to L-glutamate at 24 h and 96 h following 3 x 1-h exposures to Salmosan® ............................................... 55

Figure 11. Feeding response of prawns measured by the change in time spent moving (∆ time spent moving) in response to L-glutamate at 24 h and 96 h following 3 x 1-h exposures to Salmosan® ....................................... 55

Figure 12. Feeding response of prawns measured by the change in total number of antennule and antennae wipes (∆ wipes) in response to L-glutamate at 24 h and 96 h following 3 x 1-h exposures to Salmosan® ........................ 56

Figure 13. Feeding response of prawns measured by the change in number of dactyl probes (∆ probes) in response to L-glutamate at 24 h and 96 h following 3 x 1-h exposures to Salmosan® .............................................................. 56

Figure 14. Kaplan-Meier curves showing the proportion of prawns that reacted to L-glutamate, a food stimulant, over time at t = 24 and 96 h after 3 x 1-h exposures to Salmosan® .......................................................................... 57

ix

List of Acronyms

AI Active ingredient ACh Acetylcholine AChE Acetylcholinesterase ANOVA Analysis of variance ANCOVA Analysis of covariance BC British Columbia

CL Carapace length CNS Central nervous system DDT Dichlorodiphenyltrichloroethane DFO Fisheries and Oceans Canada EC50 Median effective concentration EcR Ecdysteroid receptor GLMM Generalized linear mixed effects model LC50 Median lethal concentration LOEC Lowest observed effects concentration L-glu L-glutamate MIH Molt-inhibiting hormone MS222 Ethyl 3-aminobenzoate methanesulfonate NOEC No observed effects concentration OP Organophosphate RXR Retinoid X receptor

1

Chapter 1. Introduction

1.1. Open net pen farming in Canada

Declining fish stocks worldwide have prompted a rapid expansion of the

aquaculture industry to meet the growing global demand for seafood (FAO, 2018). Wild

fisheries capture rates have remained relatively static since the late 1980’s while global

population as well as per capita demand for seafood have continued to increase (FAO,

2018). In response, global aquaculture production has been steadily increasing and

approaching wild capture harvest rates (FAO, 2018). In 2016, aquaculture was estimated

to make up 47% of the world’s seafood production by tonnage and 64% of the total sale

value (FAO, 2018).

In Canada, aquaculture accounts for roughly 20% of total seafood production. This

industry generates $1.3 billion in revenue annually, creates 1400 full time equivalent jobs,

and generates a labour income of $618 million (DFO, 2018a). Domestically, an estimated

95% of jobs and wages created from aquaculture benefit rural and coastal communities

(DFO, 2018a). The largest contributor to Canadian aquaculture in both tonnages

harvested and economic value is the open net pen industry, accounting for just over half

of all aquaculture production (FAO, 2018; Statistics Canada, 2018).

Open net pens are used to farm fish by placing permeable enclosures directly in

near-shore coastal waters. The fish in these systems benefit from the open flow of

seawater, and the natural fluctuations in water conditions such as salinity, temperature,

and dissolved oxygen. Although Chinook salmon (Oncorhynchus tshawytscha) and coho

salmon (Oncorhynchus kisutch) are also farmed in Canada, Atlantic salmon (Salmo salar)

is the most common species farmed using the open net pen system (Statistics Canada,

2018). Atlantic salmon are an ideal species for this type of cultivation due to an efficient

feed to body conversion rate and an adaptability to the confines of net pens (DFO, 2018a).

Atlantic salmon farming operations were launched in the Bay of Fundy in Atlantic

Canada in the 1970s, chosen for the protected bays, deep coastal waters, and high tidal

2

heights (DFO, 2015). These operations grew steadily across Atlantic Provinces and were

introduced into the Pacific coast of British Columbia in the 1980s. Today, Canada is the

fourth largest producer of Atlantic salmon in the world, led by Norway, Chile and Scotland

(DFO, 2018a).

The economic benefits of open net pen farming of Atlantic salmon are contrasted

by potential ecological impacts. The immersion of these open net pens in coastal waters,

while beneficial to the cultured salmon, pose risks to proximal ecosystems through the

release of biological and chemical pollutants (Haya et al., 2001). Biological pollution

includes fecal waste, and pathogens such as viruses, bacteria and parasites. Chemical

pollution includes pesticides and antibiotics (collectively referred to as

chemotherapeutants) to treat these pathogens, as well as chemicals added to fish feed

(e.g. antioxidants, dyes) and those used in construction and maintenance (e.g. anti-

foulants, preservatives) (Burridge and Van Geest, 2014; Haya et al., 2001). Parasitic sea

lice represent a major concern surrounding the open net pen industry, as outbreaks pose

implications for both farmed and wild salmon, and chemical treatment options pose risks

to non-target organisms.

1.2. Sea lice

Sea lice, Lepeoptheirus salmonis and Caligus spp., occur naturally in wild salmon

populations. Under native conditions with no anthropogenic influences, these parasites

cause ecologically inconsequential impacts on wild salmonid populations (Costello, 2006).

However, the introduction of large-scale aquaculture has manifested pathogenic

conditions for this parasitic copepod (Costello, 2006).

1.2.1. Sea lice and salmon

Sea lice are naturally occurring ecto-parasites that cause damage to salmon by

attaching to and feeding on the host’s skin, mucus and underlying tissues (Costello, 2006).

Sea lice have rasping mouth parts that are used to graze over the skin of their host while

gripping with their antennae or frontal filaments. The damage at the site of feeding is

characterized by a loss of skin and blood, increased mucus discharge, and overall tissue

necrosis (Costello, 2006; Heuch et al., 2005). Heavy infestations may even lead to the

loss of fins (Heuch et al., 2005).

3

Damage to the host is typically characterized by sublethal impacts that may

indirectly lead to mortality. Sublethal impacts documented on salmon include reduced

appetite and food-conversion efficiency, reduced swimming fitness, poor osmoregulation,

and slower growth (Costello, 1993; Godwin et al., 2017; Wagner et al., 2004). Further, the

immunocompetence of the host is often compromised, resulting in higher susceptibility to

secondary infections (Johnson and Fast, 2004; Tully and Nolan, 2002). Sea lice infections

can also cause behavioural changes to the host. For example, infected salmon have been

observed to jump in an attempt to dislodge lice, which can attract predators and waste

energy (Atkinson et al., 2018; Costello, 1993). In Norway and Scotland, sea lice outbreaks

in sea trout have been reported to cause a premature return to fresh water as an act to

dislodge the attached sea lice (Birkeland, 1997).

1.2.2. Sea lice and open net pens

Open net pens are densely populated, facilitating the proliferation of pathogens

and parasites such as sea lice. Sea lice outbreaks represent the greatest source of both

mortality and financial losses to fish farms (Marine Harvest, 2017; Mustafa et al., 2001).

These outbreaks cost the industry $500 million annually worldwide in both direct losses

and indirect management costs, and account for 6% of product value (Costello, 2009;

Mustafa et al., 2001).

Open net pens are integrated directly into natural wild salmon habitat, and as such

these outbreaks are not isolated from wild populations. Sea lice travel from farms to wild

salmon through two primary routes of exposure. The first route is through escaped infected

fish transferring lice to wild fish. In Norway it has been estimated that escaped farmed

Atlantic salmon carry 10-fold more sea lice than wild Atlantic salmon (Heuch et al., 2005).

The second route of exposure is mediated by the location of net pens, in which wild fish

may swim near or through infected sites. Planktonic nauplii or free-living copepods have

been found up to 30 km away from the site of origin, suggesting that sea lice are capable

of travelling large distances and infecting wild populations within a 30 km radius of

aquaculture facilities (Heuch et al., 2005; Krkosek et al., 2005).

In BC, there is concern regarding out-migrating juvenile smolts whose migration

paths can cross densely packed net pen gauntlets, as smolts are more vulnerable to

infection and disease in this early life stage (Costello, 2009; Heuch et al., 2005; Krkošek

4

et al., 2007). Under non-farm conditions, sea lice are more common on relatively hearty

adults that pick up sea lice from the open ocean. Their return to natal streams is separated

from the outmigration of smolts by 1 – 2 months, providing a mechanism of protection

against sea lice transfer from adult to smolt. However, one cohort of Atlantic salmon will

occupy the net pens for approximately 18 months, providing a year-round reservoir of sea

lice in coastal environments and a mechanism for more vulnerable juvenile smolts to pick

up sea lice (Costello, 2006; Krkošek et al., 2007; Pike and Wadsworth, 1999). The collapse

of sea trout stocks in Norway, Scotland, and Ireland have implicated fish farms as an

important contributor (Heuch et al., 2005; Tully and Whelan, 1993). Declines in Pacific

salmon stocks have also been associated with sea lice outbreaks in BC coinciding with

outmigration of juvenile smolts (Krkosek et al., 2005; Krkošek et al., 2007).

Sea lice outbreaks have consequences both for farmed and wild populations.

However, as with any agricultural industry, these ecological impacts have trade-offs with

socio-economic benefit. Appropriate management is necessary for both the industry and

the marine ecosystem.

1.3. Anti-sea lice chemotherapeutants in Canada

Sea lice management includes both proactive and reactive strategies. Proactive

strategies include good animal husbandry, site fallowing, infrastructural modifications, and

regular monitoring. While reactive measures include some non-chemical strategies such

as warm water treatments and feeder fish, it is more common in Canada to use anti-sea

lice chemotherapeutants to control sea lice outbreaks.

1.3.1. Regulation

Anti-sea lice chemotherapeutants are regulated federally by Health Canada.

These are used either as in-feed treatments or in bath treatments. In-feed treatments are

considered antibiotics and regulated under the Food and Drugs Act, whereas bath

treatments are considered pesticides and are regulated through the Pest Management

Regulatory Agency (PMRA) under the Pest Control Products Act (Burridge and Van

Geest, 2014; Yossa and Dumas, 2016). The key role of Health Canada is to assess

environmental fate and toxicity data (often proprietary) submitted by registrants, and to

determine whether the level of risk to the environment or humans is acceptable (PMRA,

5

2016). In cases where the severity of an infestation outweighs the risk of bypassing this

full registration process, Health Canada may issue an emergency registration. Once a

chemotherapeutant is registered, fish farm managers can obtain a formulation through a

prescription from a licensed veterinarian.

The first series of anti-sea lice chemotherapeutants were issued in 1994 under

emergency registration in response to sea lice infestations in New Brunswick (Burridge

and Van Geest, 2014). These were the first sea lice infestations experienced in fish farms

in Canada. The products issued emergency registrations included natural pyrethrin

formulations, Salartect® (active ingredient (AI): hydrogen peroxide), ivermectin, and

Salmosan® (AI: azamethiphos). Exis® (AI: cypermethrin) was not issued an emergency

registration due to objections from Environment Canada and Fisheries and Oceans

Canada (DFO), however it was issued under a research permit instead (Burridge and Van

Geest, 2014). Salmosan® and Salartect® were fully registered in 1995; however, Salartect®

was less effective, leading to reliance on Salmosan®. Sea lice soon demonstrated

resistance to Salmosan® in Atlantic Canada. In response, SLICE® (AI: emamectin

benzoate) was issued an emergency registration in 1999 and was fully registered in 2009.

SLICE® was used as the first choice for treatment due to its ease of use as an in-feed

treatment, and its ability to target a full life cycle of the sea louse. Salartect® and

Salmosan® were not renewed for registration and SLICE® became the only product

registered for use in Canada. Shortly after, sea lice in Atlantic Canada began to

demonstrate resistance to SLICE® leading to major outbreaks in 2009 and 2010. Health

Canada issued emergency registrations for Salmosan®, Paramove®50 (AI: hydrogen

peroxide), and AlphaMax® (AI: delthamethrin). AlphaMax® was not renewed after 2010.

Until 2016, SLICE® was the only compound under full registration in Canada, and the only

compound used in BC. Salmosan® and Paramove®50 were used in Atlantic Canada under

emergency registration. Paramove®50 was fully registered in 2016 and Salmosan® was

fully registered in 2017 (PMRA, 2016; PMRA, 2017).

1.3.2. Current use treatments

There are currently 3 anti-sea lice formulations approved by Health Canada for

treatment against sea lice outbreaks: one in-feed treatment, SLICE®; and two water-

soluble bath treatments, Paramove®50 and Salmosan®. SLICE® is the most common

treatment method on the Atlantic coast as well as in BC, largely due to its efficacy and

6

ease of use. SLICE® is added to fish feed pellets as an in-feed treatment (Burridge et al.,

2010). Once consumed by the fish, the active ingredient, emamectin benzoate, is

absorbed by the gut and distributed to the skin and then ingested by attached and feeding

sea lice. Uneaten medicated fish feed will settle in the sediment beneath the net pen,

potentially impacting non-target organisms. There are concerns regarding its potential to

persist in the environment, due to its long half-life (174 – 427 d) in marine sediment and

tendency to adhere to sediment and organic matter (log KOW = 5) (Burridge et al., 2010;

SEPA, 2017).

Bath treatments are applied either directly into the net pen, or the fish are

transported into and treated in a specialized vessel called a well-boat. If applied directly

into the net pen, a skirt or tarp is first used to enclose the pen from the surrounding

environment. In the case of Paramove®50, an additional step is required to scoop the sea

lice from the net pen following treatment. The active ingredient of Paramove®50 is

hydrogen peroxide, which causes air bubbles to form in the hemolymph of the lice, often

resulting in paralysis but not death (Bruno and Raynard, 1994). The active ingredient in

Salmosan® is azamethiphos which causes death through the inhibition of

acetylcholinesterase and requires no additional removal (Burridge and Van Geest, 2014).

As with SLICE®, Paramove®50 and Salmosan® are not effective against all life stages of

the sea louse, and the application process may be repeated to target multiple generations.

Following bath treatments, the water from the net pens or well boats, containing

the active chemotherapeutants, is released directly into the water column. Upon release

into the water column, both Paramove®50 and Salmosan® are expected to remain in the

aqueous phase due to their high water solubility (1.1 g/L for Salmosan®; Paramove®50 is

completely miscible) and low hydrophobicity (log KOW = 1.05 for Salmosan®; log

KOW = -1.57 for Paramove®50) (Burridge and Van Geest, 2014; Solvay, 2015). Therefore,

they will be dispersed in the environment through dilution. Further, they are not expected

to persist in the environment, due to relatively short half-lives (6 – 9 d for Salmosan®; 12 –

120 h for Paramove®50) (Burridge and Van Geest, 2014; Solvay, 2015).

On the Pacific coast of Canada, SLICE® remains the most common anti-sea lice

chemotherapeutant in use. However, there is concern regarding reliance on SLICE® and

the potential for resistance to develop. Fish farm managers have already begun to include

Paramove®50 in their sea-lice management strategies and may look to incorporate

7

Salmosan® as well (PMRA, 2018). To date, Salmosan® has only been used on the east

coast of Canada. As such, the bulk of toxicity data for Salmosan® has focused on the

American lobster (Homarus americanus) an economically important species indigenous

to the east coast. In response to this data gap, the present study was conducted to

evaluate the risk of Salmosan® to a non-target crustacean indigenous to the Pacific coast,

the Pacific spot prawn (Pandalus platyceros).

1.4. Risk of Salmosan® to non-target organisms

The release of Salmosan® into the water column following sea lice treatment is of

concern for proximal non-target organisms (Burridge and Van Geest, 2014; Burridge et

al., 2010). The impact of these exposures will depend on the environmental fate, the

duration and concentration each organism is exposed to, and the organism’s sensitivity to

the chemical.

1.4.1. Environmental fate

Azamethiphos, the active ingredient in Salmosan®, has a low octanol-water

partition coefficient (log KOW = 1.05) and a high solubility in water (1.1 g/L) (Tomlin, 1997).

Azamethiphos is degraded through hydrolysis and photolysis with a half-life of 6 to 9 d

(Burridge et al., 2010; Helgesen and Horsberg, 2013). Due to its tendency to stay in the

aqueous phase, dilution will be the major factor determining its decrease in concentration

in the environment (Ernst et al., 2014).

1.4.2. Exposure

Organisms living close to net pens will be exposed to short term, low

concentrations of azamethiphos due to immediate dilution in the water column. Dilution of

azamethiphos following release is variable, depending on several factors such as tidal

amplitude, current, water depth and weather. A field study by Ernst et al., (2014) reported

azamethiphos concentrations within 10 m of a fish farm to be a maximum of 10 µg/L

following release of Salmosan® from the net pen. However, the authors did not report the

time after release that the samples were taken. Burridge et al., (2000) used a mathematical

approach to estimate concentrations surrounding net pens following release and

determined a 103 dilution factor 10 m from the site within 3 h. Based on the target

8

concentration of 100 µg/L of azamethiphos in the net pen, the range of concentrations

non-target organisms living within 0 – 10 m of net pens is predicted to range from 0.1 to

10 µg/L of azamethiphos within the first 3 h of release.

Organisms may also be subjected to repeated pulses of Salmosan® as farm

managers may apply consecutive treatments within a site to target all life stages of the

sea louse, or to treat multiple pens within one site (Burridge et al., 2008; Dounia et al.,

2016). Further, proximal farms may be treating outbreaks simultaneously, as outbreaks

can spread between sites. These multiple exposures would likely occur within 6 h and/or

repeated at intervals of approximately 12 h, to coincide with an outgoing tide as stated by

the label protocol (PMRA, 2016).

1.4.3. Toxicity

Azamethiphos, like all organophosphates (OPs), interferes with nervous system

functioning through the phosphorylation and consequent inhibition of acetylcholinesterase

(AChE) (Colovic et al., 2013; Weinbroum, 2004; Xuereb et al., 2009). AChE hydrolyses

acetylcholine (ACh) into choline and acetic acid. ACh is an excitatory neurotransmitter,

the hydrolysis of which allows neurons to return to their resting state after activation

(Colovic et al., 2013). Under AChE inhibition, acetylcholine builds up at neural synapses,

causing neurons to repeatedly fire. Eventually, the neuron becomes insensitive to new

acetylcholine and loses the ability to send and receive signals (Colovic et al., 2013;

Weinbroum, 2004). Symptoms of AChE inhibition include involuntary convulsions,

twitching, agitation, and eventual partial or complete paralysis (Couillard and Burridge,

2014; Fulton and Key, 2001; Xuereb et al., 2009).

AChE inhibition can lead to death both directly and indirectly. The most direct route

to death occurs through hypoxia, as an organism with paralysed muscles cannot actively

pump oxygenated water over its gills (in the case of aquatic organisms), leading to cardiac

impairment (Colovic et al., 2013; Dounia et al., 2016; Fulton and Key, 2001; Weinbroum,

2004). Paralysis can indirectly lead to death through the inability to forage or escape from

predators (Xuereb et al., 2009). AChE inhibition may also interrupt the response to

chemical cues as ACh is the primary sensory neurotransmitter in many crustaceans

(Florey, 1973). This can have consequences on aquatic organisms that rely on waterborne

chemical signals for functions such as foraging, mating, and detecting predators.

9

There is some evidence that OPs can exert effects through additional or alternative

modes of action to AChE inhibition. Malathion, an OP known to inhibit AChE, caused both

lethal and sublethal (reduced growth) effects in grass shrimp (Palaemonetes pugio) but

did not cause inhibition in AChE (Key et al., 1998). Conversely, no adverse effects were

observed in Daphnia magna exposed to the OP acephate, despite levels of AChE

inhibition that did cause mortality in the same species exposed to other AChE-inhibiting

OPs (Printes and Callaghan, 2004). A similar trend was observed in the amphipod

Gammarus fossarum, for which the OPs chlorpyrifos and methomyl caused comparable

levels of AChE inhibition (66-70%), but caused mortality only for chlorpyrifos (Xuereb et

al., 2009). These results led the authors in both studies to suggest AChE activity can be

altered through pathways other than direct inhibition of the enzyme, and therefore that

these OPs can bind sites other than AChE. While this has not been further investigated in

crustaceans, OPs in mammals can inhibit butyrylcholinesterase (BuChE) and

carboxylesterase (CaE), as well as bind to nicotinic and muscarinic acetylcholine

receptors (Mileson et al., 1999; Pope, 1999).

The majority of Salmosan® toxicity data describes lethal thresholds for crustaceans

as this chemotherapeutant is formulated to target a crustacean, the sea louse (Table 1).

In Canada, the American lobster has been studied frequently due to its economic

importance as well as its sensitivity to azamethiphos at concentrations below the target

bath concentration of 100 µg/L. Larval stages I, II, III, IV and adult lobsters were subjected

to toxicity tests with 48-h LC50s spanning 0.61 – 3.57 µg/L; 24-h LC50s spanning 2.8 –

8.9 µg/L; and, 1-h LC50s spanning 20.70 – 86.5 µg/l (Burridge et al., 1999; Burridge et al.,

2014; Pahl and Opitz, 1999). These toxicity tests demonstrate life stage sensitivity, with

the least sensitive life stage being larval stage I, compared to successive larval and adult

stages. American lobsters also show seasonal sensitivity to azamethiphos, with lobsters

being more sensitive during higher seasonal temperatures and during their post-molt

stage (Burridge et al., 2005). An economically important crustacean native to Chile, the

Southern rock crab (Metacarcinus edwardsii) has also shown environmentally relevant

sensitivity to azamethiphos, with a 30-min LC50 of 2.85 µg/L (Gebauer et al., 2017). Brine

shrimp (Artemia salina) and planktonic copepods appear to be the least sensitive

crustaceans for which toxicity values are reported, with no mortality or immobility observed

after 1-h exposures to 10,000 and 500 µg/L, respectively (Ernst et al., 2001; Van Geest et

al., 2014). In comparison, the 1-h LC50 of the target sea louse, L. salmonis, is 0.18 µg/L

10

(Roth et al., 1996). Longer duration exposures (i.e. 24 to 96 h) are less environmentally

relevant but do reveal LC50s below the target application concentration of 100 µg/L

azamethiphos for Daphnia magna; sand shrimp (Crangon septemspinosa), unidentified

Mysid sp., and, a copepod (Tisbe battagliai) (Burridge et al., 2014; Ernst et al., 2014;

Macken et al., 2015; Rose et al., 2016).

Lethal thresholds have also been reported for non-crustacean aquatic species,

however in far fewer numbers. After a 1-h exposure to 1000 µg/L azamethiphos (in

formulation as Alfacron 50WP®), 15% mortality was reported in Atlantic salmon (Sievers

et al., 1995). This is 10x the target concentration of azamethiphos. The 96-h LC50 for

stickleback (Gasterosteus aculeatus) was reported to be 190 µg/L (Ernst et al., 2001). The

european eel (Anguilla Anguilla) and the European seabass (Dicentrarchus labrax)

survived 240-min exposures at 100 µg/L while the rainbow trout (Onchorhynchus mykiss)

experienced 100% mortality under the same exposure scenario (Intorre et al., 2004).

Lethal thresholds were 20 – 100x greater than the target azamethiphos concentrations for

a rotifer (Branchionus plicatilis), polychaete (Polydora cornuta), sea urchin

(Strongylocentrotus droebachiensi), and bacterium (Vibrio fischeri) (Ernst et al., 2001).

Sublethal effects of azamethiphos have been documented at both the organismal

and biochemical levels (Table 2). At the organismal level, the most concerning sublethal

effect was a reported 48% decrease in spawning incidence in the American lobster after

4 x 1-h exposures of 10 µg/L repeated at 2-week intervals (Burridge et al., 2008).

Behavioural studies reported no effect of 1-h exposure at 500 µg/L on feeding rates of

copepods, or 3 x 30-min exposure at 1000 µg/L on shelter use of American lobsters

(Abgrall et al., 2000; Van Geest et al., 2014). However, anecdotal observations of

behaviours associated with neurotoxicity were reported, such as erratic flopping, loss of

claw control, decrease in activity, sporadic twitching, and rolling onto backs (Burridge et

al., 2000b; Burridge et al., 2014; Dounia et al., 2016). These behaviours were not

quantified but were used to determine a 24-h lowest observed effects concentrations

(LOEC) of 1.20 µg/L for the sand shrimp; 1-h LOECs of 3.70 and 11.1 µg/L for adult and

larval American lobsters, respectively; and, repeated exposure 1-h NOECs of 0.5 and 1.03

µg/L for adult and larval American lobsters, respectively (Burridge et al., 2000b; Burridge

et al., 2014; Dounia et al., 2016). At the biochemical level, azamethiphos exposure

decreased abdominal AChE activity and increased levels of calcium and lactate in the

11

hemolymph of the American lobster, symptoms associated with hypoxia (Dounia et al.,

2016).

Studies on the sublethal effects of azamethiphos in fish are scarce, with the only

effect reported being reduced AChE activity in the brain of eel, seabass, and trout (Intorre

et al., 2004). Intorre et al. (2004) anecdotally reported hyperactivity at the onset of the

exposure, inferring some level of agitation or avoidance. Sublethal effects on other

organisms include inhibition of fertilization in a sea urchin (Lytechnius pictus) and inhibition

of AChE in the gill and hemolymph of the blue mussel (Mytilus edulis) (Canty et al., 2007).

However, these effects occur at 7 – 70x the target concentration (Table 2).

12

Table 1. Summary of the lethal toxicity values for azamethiphos for various aquatic species. M = measured; N = nominal; F = formulation; AI = active ingredient; LC50 = median lethal concentration; EC50 = median effects concentration; CI = confidence interval.

Species Life stage Measured or

nominal

Formulation or active

ingredient

Exposure scenario Endpoint [Azamethiphos] (µg/L)

(95%CI)

Reference

Crustaceans Artemia salina F 24-h LC50 > 10 000 (Ernst et al., 2001) Copepod (various) M F 1-h LC50 > 500 (Van Geest et al., 2014) Crangon septemspinosa

M F 24-h LC50 191 (149 – 33)

(Burridge et al., 2014)

Crangon septemspinosa

M F 1-h LC50 > 85.5 (Burridge et al., 2014)

Crangon septemspinosa

M F 24-h

LC50 4.8 – 33.6 (Ernst et al., 2014)

Daphnia magna N AI 24-h EC50 0.167 (0.147 – 0.197)

(Rose et al., 2016)

Eohaustorius estuarius

F 48-h LC50 > 20 (Ernst et al., 2001)

Eohaustorius estuarius

F 48-h EC50 (mortality + immobility)

3.0 (2.1-4.4)

(Ernst et al., 2001)

Gammarus spp F 96-h LC50 < 5 (Ernst et al., 2001) Homarus americanus

Larvae M F 12-h LC50 0.90 – 1.33 (Pahl and Opitz, 1999)

13

Species Life stage Measured or

nominal

Formulation or active

ingredient

Exposure scenario Endpoint [Azamethiphos] (µg/L)

(95%CI)

Reference

Homarus americanus

Larvae M F 6-h LC50 3.50 – 5.40 (Pahl and Opitz, 1999)

Homarus americanus

Larvae M F 1-h LC50 20.70 – 26.50 (Pahl and Opitz, 1999)

Homarus americanus

Larvae M F 30-min LC50 27.01 – 37.70 (Pahl and Opitz, 1999)

Homarus americanus

Larvae M F 5-min LC50 33.90 – 50.40 (Pahl and Opitz, 1999)

Homarus americanus

Larval stage I M F 48-h LC50 3.57 (1.76 – 5.37)

(Burridge et al., 1999)

Homarus americanus

Larval stage I M F 24-h LC50 8.9 (Burridge et al., 2014)

Homarus americanus

Larval stage I M F 1-h LC50 > 86.5 (Burridge et al., 2014)

Homarus americanus

Larval stage II M F 48-h LC50 1.03 (0 – 4.28)

(Burridge et al., 1999)

Homarus americanus

Larval stage III M F 48-h LC50 2.29 (0.72 – 3.88)

(Burridge et al., 1999)

Homarus americanus

Larval stage IV

M F 48-h LC50 2.12 (1.06 – 3.18)

(Burridge et al., 1999)

Homarus americanus

Adult

M F 48-h LC50 0.61 – 3.24 (0.5 – 4.0)

(Burridge et al., 2005)

14

Species Life stage Measured or

nominal

Formulation or active

ingredient

Exposure scenario Endpoint [Azamethiphos] (µg/L)

(95%CI)

Reference

Homarus americanus

Adult M F 48-h LC50 1.39 (0.78 – 2.02)

(Burridge et al., 1999)

Homarus americanus

Adult M F 24-h LC50 2.8 (1.8 – 3.7)

(Burridge et al., 2014)

Homarus americanus

Adult M F 1-h LC50 24.8 (21.7 – 27.9)

(Burridge et al., 2014)

Homarus americanus

Male adult M F 5 x 1-h at t = 0, 4, 24, 28, 48-h

93% mortality 5 (Dounia et al., 2016)

Homarus americanus

Male adult M F 5 µg/L 3 x 1-h

at t = 0, 4, 24

LT50 26-h (Dounia et al., 2016)

Homarus americanus

Adult M F 10-d + simulated stress

33% mortality 0.061 (Couillard and Burridge, 2014)

Lepeoptheirus salmonis

Adult and pre-adult

N F 1-h

LC50 0.07 – 0.18 (0.06 – 0.21)

(Roth et al., 1996)

Metacarcinus edwardsii

Zoea I N F 30-min LC50 2.85 (2.46 – 3.24)

(Gebauer et al., 2017)

15

Species Life stage Measured or

nominal

Formulation or active

ingredient

Exposure scenario Endpoint [Azamethiphos] (µg/L)

(95%CI)

Reference

Metacarcinus edwardsii

Zoea I N F 30-min EC50 (mortality + immobility)

0.94 (0.79 – 1.09)

(Gebauer et al., 2017)

Mysid sp. M F 24-h LC50 12.5 (5.4 – 19.6)

(Burridge et al., 2014)

Mysid sp. M F 1-h LC50 > 85.5 (Burridge et al., 2014) Mysis stenolepsis M F 24-h LC50 4.8 – 23.5 (Ernst et al., 2014) Tisbe battagliai Copepodid M AI 48-h LC10 3.6

(2.6 – 4.5) (Macken et al., 2015)

Tisbe battagliai Copepodid M AI 48-h LC50 7.7 (6.8 – 8.5)

(Macken et al., 2015)

Tisbe battagliai Nauplius M AI 48-h LC10 3.4 (2.4 – 5.3)

(Macken et al., 2015)

Tisbe battagliai Nauplius M AI 48-h LC50 6.7 (5.9 – 7.3)

(Macken et al., 2015)

Fish Gasterosteus aculeatus

F 96-h LC50 190 (140 – 250)

(Ernst et al., 2001)

Salmo salar Adult N F 1-h 15% mortality 1000 (Sievers et al., 1995) Salmo salar Adult N F 1-h 100% mortality 5000 (Sievers et al., 1995)

16

Species Life stage Measured or

nominal

Formulation or active

ingredient

Exposure scenario Endpoint [Azamethiphos] (µg/L)

(95%CI)

Reference

Anguilla anguilla Adult N AI 240-min LC50 > 100 (Intorre et al., 2004) Dicentrarchus labrax

Adult N AI 240-min LC50 > 100 (Intorre et al., 2004)

Onchorhynchus mykiss

Adult N AI 120-min LC50 > 100 (Intorre et al., 2004)

Onchorhynchus mykiss

Adult N AI 240-min 100% mortality 100 (Intorre et al., 2004)

Other Branchionus plicatilis

F 24-h LC50 > 10 000 (Ernst et al., 2001)

Polydora cornuta Juvenile F 96-h LC50 2310 (650 – 590 000)

(Ernst et al., 2001)

Strongylocentrotus droebachiensis

Adult F 96-h LC50 > 1000 (Ernst et al., 2001)

Vibrio fischeri F 15-min EC50 (mortality + immobility)

11000 (1880 – 64500)

(Ernst et al., 2001)

17

Table 2. Summary of sublethal toxicity values for azamethiphos in various aquatic species. M = measured; N = nominal; F = formulation; AI = active ingredient; LC50 = median lethal concentration; EC50 = median effects concentration; CI = confidence interval.

Species Life stage Measured or

Nominal

Formulation or active ingredient

Exposure Endpoint [Azamethiphos] (µg/L)

(95% CI)

Reference

Crustaceans Copepod (various) various M F 1-h EC50

(Feeding inhibition) > 500 (Van Geest et al.,

2014) Crangon septemspinosa

M F 24-h LOEC 1.20 (Burridge et al., 2014)

Homarus americanus

Juvenile N AI 3 x 10-min 1-h intervals

EC50 (Shelter exit)

< 1000 (Abgrall et al., 2000)

Homarus americanus

Larval stage I M F 1-h LOEC 11.1 (Burridge et al., 2014)

Homarus americanus

Larval stage IV

M F 9 x 120-min over 3-d.

3-h intervals, repeated daily.

NOEC 1.03 (Burridge et al., 2000b)

Homarus americanus

Male adult M F 5 x 1-h at t = 0, 4, 24,

28, 48-h

Decrease in abdominal AChE

activity

0.5 (Dounia et al., 2016)

Homarus americanus

Male adult M F 5 x 1-h at t = 0, 4, 24,

28, 48-h

Increase in hemolymph

[lactate]

5 (Dounia et al., 2016)

18

Species Life stage Measured or

Nominal

Formulation or active ingredient

Exposure Endpoint [Azamethiphos] (µg/L)

(95% CI)

Reference

Homarus americanus

Male adult M F 5 x 1-h at t = 0, 4, 24,

28, 48-h

Increase in hemolymph

[calcium]

5 (Dounia et al., 2016)

Homarus americanus

Female adult N F 4 x 1-h 2-wk intervals

48% decrease in spawning incidence

10 (Burridge et al., 2008)

Homarus americanus

Adult M F 10-d + Subjected to

shipping stress

Effects on hepatosomatic and

gonadosomatic indices; lipid and water content in hepatopancreas

0.061 (Couillard and Burridge, 2014)

Homarus americanus

Adult M F 1-h LOEC 3.70 (Burridge et al., 2014)

Homarus americanus

Male adult M F 3 x 1-h at t = 0, 4, 24-h

NOEC 0.5 (Dounia et al., 2016)

Tisbe battagliai M AI 7-d EC50 Development

> 3.6 (Macken et al., 2015)

Fish Anguilla anguilla Adult N AI 240-min Decreased brain

AChE activity (with recovery)

100 (Intorre et al., 2004)

19

Species Life stage Measured or

Nominal

Formulation or active ingredient

Exposure Endpoint [Azamethiphos] (µg/L)

(95% CI)

Reference

Dicentrarchus labrax Adult N AI 240-min Decreased brain AChE activity (with

recovery)

100 (Intorre et al., 2004)

Onchorhynchus mykiss

Adult N AI 120-min Decreased brain AChE activity (with

recovery)

100 (Intorre et al., 2004)

Onchorhynchus mykiss

Adult N AI 240-min Decreased brain AChE activity (no

recovery)

100 (Intorre et al., 2004)

Other Lytechinus pictus F 20-min IC50

Fertilization 6840 (4880 – 8095)

(Ernst et al., 2001)

Lytechinus pictus F 20-min IC25 3340 (1150 – 4810)

(Ernst et al., 2001)

Mytilus edulis N AI 30-min in vitro

IC50 AChE activity (gill)

736 (Canty et al., 2007)

Mytilus edulis N AI 30-min in vitro

IC50 AChE activity (hemolympoh)

1300 (Canty et al., 2007)

Mytilus edulis N AI 1-h EC50 Feeding rate

> 100 (Canty et al., 2007)

Mytilus edulis N AI 24-h EC50 > 100 (Canty et al., 2007)

20

Species Life stage Measured or

Nominal

Formulation or active ingredient

Exposure Endpoint [Azamethiphos] (µg/L)

(95% CI)

Reference

Feeding rate Mytilus edulis N AI 1-h EC50

Phagocytic activity > 100 (Canty et al., 2007)

Mytilus edulis N AI 24-h 91% inhibition of phagocytic activity

> 100 (Canty et al., 2007)

21

1.5. Study organism: Pacific spot prawn

This study revolves around concern with the release of Salmosan® from open net

pens in BC and the potential adverse effects on proximal populations of Pacific spot

prawns (Pandalus platyceros). The Pacific spot prawn is an economically important

species whose habitat overlaps with areas occupied by open net pen farms.

Azamethiphos, the active ingredient in Salmosan®, is known to adversely affect non-target

crustaceans with studies showing both molt-related sensitivity and behavioural changes

(Abgrall et al., 2000; Burridge et al., 2005). Further, fishermen in Atlantic Canada have

reported an increase in dead shrimp and crabs near aquaculture sites following well-boat

treatments using Salmosan® (Wiber et al., 2012).

Decapod crustaceans (i.e. shrimps, crayfish, lobsters) are useful in toxicological

studies as they have a well understood molting cycle as well as a thoroughly described

hierarchy of behaviours associated with chemoreception (Hallberg and Skog, 2011; Lee

and Meyers, 1996). Therefore, the Pacific spot prawn was the focus of this study due to

its economic importance, its potential to be impacted by the release of Salmosan® from

open net pens, and the availability of accessible and quantifiable toxicological endpoints.

Further, protocols had been previously established in the Kennedy lab at Simon Fraser

University for obtaining, transporting, and holding spot prawns.

1.5.1. BC spot prawn fishery

The BC spot prawn fishery is considered one of the most sustainable fisheries in

the province (DFO, 2018b; Favaro et al., 2010). Although spot prawns are occasionally

caught in trawls, they are not the target species in the shrimp trawl fishery and are instead

targeted through traps (DFO, 2018b). Shrimp fishing by trap is associated with relatively

low bycatch and low habitat destruction (Favaro et al., 2010). This fishery is managed by

the DFO through in-season population surveys and enforced through seasonal closures

(DFO, 2018b). The commercial fishery is typically open for 4 to 8 weeks starting the 1st of

May. Recreational fishing can take place all year, with some regional closures during the

spawning season from January until March. The commercial prawn and shrimp by trap

fishery is one of the most valuable in BC, generating roughly $35 million annually in landed

value and $3 million in direct wages (DFO, 2018b).

22

1.5.2. Life history and habitat

The Pacific spot prawn is the largest of 7 commercially harvested shrimp species

in BC and is the only one colloquially referred to as a ‘prawn,’ a term usually reserved for

Penaeid shrimp (Butler, 1964; DFO, 2018b). The spot prawn is native to the West coast

of North America from California to Alaska, and the Sea of Japan to the Korea Straight.

As adults, their habitat ranges from the intertidal to 400 m depth, and they are typically

found around 50 to 90 m (DFO, 2018b).

The life cycle of a spot prawn begins in March or April, hatching from eggs into a

planktonic life stage until settling into the benthic habitat by May or June. As they transition

through their final larval stages into their juvenile stage, they migrate into shallower waters.

Adult prawns migrate back into deeper waters after their first year (Butler, 1964). Typically,

adult prawns will remain in deeper waters during the day and migrate to shallower waters

to forage at night, where they feed opportunistically on varied food sources including

smaller shrimp, amphipods, worms, limpets, plankton and dead material (Butler, 1964).

Prawns are protandric hermaphrodites, meaning that they are born male and transition to

female at some point in their life. Spot prawns enter this transitional phase around year

two and are female by year four. Prawns mate through September and October, after

which the females carry 2000 – 4000 eggs until they are ready to hatch in early spring

(Butler, 1964; Kelly et al., 1977).

The near shore environment that is optimal for open net pens is also optimal habitat

for the Pacific spot prawn. This environment provides easy migration from deep waters to

shallow intertidal for nocturnal foraging. Prawns may also congregate around net pens,

attracted to the fecal matter and detritus falling from the pens above, as has been reported

in American lobsters in Atlantic Canada (Findlay et al., 1995). Further, adults do not

migrate to a significant extent, and are estimated to remain within a 10 km radius (Bower

et al., 1996). Therefore, there is potential for significant overlap between spot prawn

habitat and net pen sites.

1.6. Environmental temperature and toxicity

Crustaceans are ectothermic organisms; therefore, their internal temperature is

regulated by the external environment. Unlike endothermic organisms such as mammals

23

and birds that will actively maintain a relatively constant internal temperature regardless

of their environment, aquatic crustaceans exhibit body temperatures close to the ambient

temperature of the water column they occupy. Body temperature plays a critical and

pervasive role at all levels of biological organization (Hochachka and Somero, 1984).

Therefore, a changing body temperature has the potential to impact an organism’s

response to environmental stressors such as chemical contamination. As such, a brief

summary on how temperature affects biological systems of ectotherms, and consequently

toxicity, is warranted.

Temperature influences physiological processes by directly affecting reaction rates

and reaction equilibria (Hochachka and Somero, 1984). Increased temperatures result in

more molecules possessing energy equal to or greater than the activation energy required

by a given reaction, causing reaction rates to increase (Hochachka and Somero, 1984).

As such, the rate of metabolic processes increases with temperature, influencing

processes related to the breakdown of molecules for energy release (i.e. glycolysis,

cellular respiration) and the biotransformation and excretion of both endogenous and

exogenous compounds (Hochachka and Somero, 1984; Kennedy and Walsh, 1997). As

many of these metabolic processes require oxygen, oxygen consumption also typically

increases with temperature (Claësson et al., 2016).

Temperature also directly effects equilibrium constants associated with the

formation of weak, non-covalent bonds (Hochachka and Somero, 1984). Unlike strong

covalent bonds, these relatively weaker bonds are readily disrupted by small temperature

changes. If bond formation is an exothermic reaction (i.e. enthalpy is negative), an

increase in temperature will act to destabilize the bonds. Likewise, if the reaction is

endothermic (i.e. enthalpy is positive) then an increase in temperature will stabilize the

bond. Hydrogen bonds, van der Waals forces, and ionic bonds are destabilized by

increased temperatures, while hydrophobic bonds are stabilized by increased

temperatures. An important biological structure vulnerable to thermal perturbation is the

phospholipid bilayer in cellular membranes, typically held together by van der Waals

forces and hydrogen bonds. Increased temperatures destabilize this bilayer, leading to

increased membrane fluidity and permeability, and potentially altering the functioning of

membrane-associated proteins (Guschina and Harwood, 2006). Further, enzyme-ligand

complexes, protein structures, and nucleic acid structures often involve the formation of

24

weak non-covalent bonds and thus can be influenced by temperature changes

(Hochachka and Somero, 1984).

To withstand long-term seasonal shifts in temperature, ectotherms must be able

to compensate for the effects of temperature on the structure and function of biological

molecules. This process of temperature acclimation, typically occurring on a scale of days

to weeks, primarily involves transcriptional changes (Hochachka and Somero, 1984). To

maintain an optimal rate of metabolic functioning, ectotherms can adjust catalytic

efficiency through changing the quantity of enzymes transcribed, or through transcribing

specific isoenzymes that perform optimally at given temperatures (Hochachka and

Somero, 1984). To account for the increased oxygen demand at higher metabolic rates,

mitochondrial densities may increase at higher temperatures (Portner and Knust, 2007).

Another important function of acclimation is to restore optimal membrane fluidity through

homeoviscous adaptation, in which the composition of saturated (less fluid) and

unsaturated fatty acids (more fluid) in the phospholipid bilayer is altered (Hochachka and

Somero, 1984). As well, higher temperatures may trigger the transcription of heat shock

proteins that act to protect proteins and ensure proper folding and structure is maintained

(Portner and Knust, 2007). The ability to acclimate to new temperatures means that

ectotherms will experience less pronounced temperature effects after an acclimation

period compared to after an acute temperature change (Kennedy and Walsh, 1997;

Kennedy et al., 1989).

By causing acute or long-term shifts in physiological processes, temperature has

the potential to act as an indirect modifier of toxicity (Cairns et al., 1975; Heugens et al.,

2001). Temperature can modify toxicity by altering the internal exposure (i.e. dose), or the

susceptibility to the toxicant. As oxygen demand increases in response to an increased

metabolic rate, ventilation typically increases, thereby increasing the uptake of

contaminants from the water column (Abdel-Lateif et al., 1998; Kennedy et al., 1989).

Oxygen solubility is also lower in warm water, often exacerbating the need for increased

ventilation. Temperature is also a factor determining chemical solubility, potentially

influencing the amount of a chemical available for uptake and transport. Further, if the

degree or duration of the temperature change surpasses a threshold, the organism can

experience a mismatch between oxygen availability and supply (Heugens et al., 2001;

Portner and Knust, 2007). The consequence of this on toxicity may be a reduced energy

supply or capacity to direct towards metabolism of the toxicant. Because such a variety of

25

complex processes are affected by temperature, no common rule exists for how

temperature alters toxicity of a chemical. However, temperature dependent toxicity often

follows one of two models: (1) increasing toxicity with temperature or (2) increasing toxicity

in either direction away from an optimal temperature (Cairns et al., 1975; Li et al., 2014;

Zhou et al., 2014).

The typical benthic temperature is roughly 6 to 12 °C for the Strait of Georgia

(Butler, 1964; DFO, 2009). However, prawns migrate nightly to forage and may therefore

interact with warmer surface waters, where seasonal maximums in BC typically reach

18 °C, however, may reach up to 20 °C (DFO, 2018c). The upper lethal temperature limit

of Pacific spot prawns has been reported as 22.9 °C, and the lower limit to be 3 °C (Kelly

et al., 1977; Whyte and Carswell, 1982).

1.7. Study goal and objectives

The goal of this study was to characterize the lethal and sublethal effects of

azamethiphos, in formulation as Salmosan®, on the Pacific spot prawn under

environmentally relevant exposure scenarios. The lethal and sublethal effects were

investigated through two experiments. The objective of the first experiment (Chapter 2)

was to determine the effect of Salmosan® on the ability of prawns to molt, the time to molt,

and growth during molt. Further, sensitivity to Salmosan® was determined at two molt

stages: the intermolt stage and the post-molt stage. The objective of the second

experiment (Chapter 3) was to determine the effect of Salmosan® exposure on locomotory,

olfactory and grooming behaviours during and following exposure.

In both experiments the prawns were subjected to short term (1-h) exposures to

simulate the immediate dilution of Salmosan® upon release into the water column; and,

repeated exposures (3 x) to simulate repeated sea lice treatments on one site or nearby

sites treating outbreaks simultaneously (Abgrall et al., 2000; Burridge et al., 2000b; Dounia

et al., 2016; Van Geest et al., 2014). Further, all exposures were run under three

temperatures regimes (5, 11 and 17 °C) to investigate temperature as a potential modifier

of Salmosan® toxicity.

26

Chapter 2. The lethal and sublethal effects of Salmosan® on molting in the Pacific spot prawn, Pandalus platyceros

2.1. Introduction

Like all crustaceans, Pacific spot prawns (Pandalus platyceros) molt periodically

throughout their life. Prawns molt in order to grow, as well as to pass through successive

larval, juvenile and adult life stages. The shedding of the exoskeleton is referred to as

ecdysis while the entire process is referred to as molting. For spot prawns, ecdysis occurs

every 3 to 4 months. Roughly 5 to 7 days before ecdysis, prawns begin to grow a new

exoskeleton and produce enzymes to degrade the old exoskeleton (Hosamani et al.,

2017). Water uptake increases to create pressure to push the old exoskeleton off. The

actual shedding of the exoskeleton – which once shed is referred to as an exuvia – takes

only minutes. Growth occurs in the 5 to 7 days following ecdysis, during which time the

prawn continues to take in water to expand its malleable new exoskeleton.

The molting process is regulated through the endocrine system, specifically the Y-

organ/ecdysteroid receptor (EcR) axis. Ecdysteroids are constantly released by paired Y-

organs located in the anterior cephalothorax in between the gills (Nakatsuji et al., 2009).

These ecdysteroids are transported through the hemolymph to various target tissues,

where they interact with EcRs and subsequently heterodimerize with the crustacean

retinoid X receptor (RXR) (Hosamani et al., 2017; Zou, 2005). It is this EcR/RXR dimer

that binds to DNA response elements in certain genes, initiating transcription of proteins

responsible for regulating the molting process. One such protein induced by this pathway

is chitobiase, an enzyme responsible for degrading the exoskeleton (Zou and Fingerman,

1999). The levels of ecdysteroids in the hemolymph remains relatively low due to inhibition

of the Y-gland by a polypeptide neurohormone, the molt inhibiting hormone (MIH),

produced in and released from the X-organ-sinus gland complex located in the eyestalk

(Nakatsuji et al., 2009). When MIH levels drop, ecdysteroidogenesis intensifies, and

molting shortly follows.

27

An environmental chemical that interferes directly or indirectly with the Y-

organ/EcR axis has the potential to affect the molting cycle. The most common molt-

related effect of toxicant exposure appears to be the inhibition of molting as reported by:

Schimmel et al. (1979) in blue crabs (Callinectus sapidus) exposed to kepone; Cantelmo

et al. (1981) in blue crabs exposed to polycyclic aromatic hydrocarbons; Zou and

Fingerman (1997a) in Daphnia magna exposed to diethylstilbestrol; Zou and Fingerman

(1997b) in D. magna exposed to diethyl phthalate; Baer and Owens (1999) in D. magna

exposed to methoxyclor; Snyder and Mulder (2001) in American lobster (Homarus

americanus) exposed to heptachlor; Rodríguez Moreno et al. (2003) in estuarine crab

(Chasmagnathus granulate) exposed to cadmium; and Dounia et al. (2018) in American

lobsters exposed to emamectin benzoate. The induction of molting is less common but

has been shown by Weis and Mantel (1976) and Weis et al. (1992) in fiddler crabs (Uca

pugilator) exposed to dichlorodiphenyltrichloroethane (DDT); and, by Waddy et al. (2002)

in American lobsters exposed to emamectin benzoate.

The broad range of environmental toxicants that affects molting in crustaceans

alludes to the presence of several modes of action on the Y-organ/EcR axis. However, the

specific mechanisms of action on induction or inhibition are largely unknown. The

induction of molting due to the emamectin benzoate and DDT was hypothesized by Weis

and Mantel (1976) to be due to interference in the release of MIH, which is under the

control of the nervous system. If azamethiphos exposure interrupts normal functioning of

the nervous system, this may induce molting in the spot prawns.

Molting is energetically exhaustive, and crustaceans that have just molted (i.e.

post-molt stage) have been reported to be more susceptible to the effects of environmental

pollutants compared to those that are in between ecdysis events (i.e. intermolt stage)

(Bechmann et al., 2018; Lee and Buikema, 1979; Price and Uglow, 1979). Further,

crustaceans continue to intake water during in the days following ecdysis, therefore post-

molt individuals are likely exposed to higher internal concentrations of an environmental

contaminant that is present in the water column (Hosamani et al., 2017). Consequently, it

is expected that post-molt prawns will be more susceptible to azamethiphos toxicity than

intermolt prawns.

The objectives of this experiment were to compare the sensitivity of Pacific spot

prawns to azamethiphos (in formulation as Salmosan®) between the intermolt and post-

28

molt stages; and, to determine if any effects occurred on the time to molt, ability to molt

and growth during molt. This was achieved by subjecting intermolt and post-molt prawns

to identical exposure scenarios of Salmosan®, and then monitoring the intermolt prawns

throughout their molting cycle.

2.2. Material and methods

2.2.1. Organism collection and holding

Adult Pacific spot prawns (Pandalus platyceros) were purchased from T&T

Supermarket in Vancouver, BC in May of 2017 and June of 2018. The prawns were

transported to Simon Fraser University in source seawater in a cooler at approximately

10 °C under constant aeration. Prawns were held communally in 500 L holding tanks at

11 ± 1 °C under constant aeration and a 12:12 photoperiod. They were fed a mixed diet

of frozen brine shrimp, fish, and squid ad libitum 3 to 4 times per week. All prawns were

acclimated to laboratory conditions for at least 2 weeks prior to exposures. Prawns

weighed an average of 27.4 g, ranging from 17.5 to 48.5 g. The maximum loading density

was 1 prawn per 5 L seawater.

2.2.2. Chemicals

Salmosan® (Fish Vet Group®, Inverness, Scotland) is a wettable powder

formulation containing 49.8% azamethiphos and was obtained from Fisheries and Oceans

Canada (DFO). All exposures were conducted in formulation as Salmosan® and prepared

at target concentrations of azamethiphos by thoroughly mixing in seawater for 1 h. All

animal euthanizations were performed in seawater using 1 g/L ethyl 3-aminobenzoate

methanesulfonate (MS222, Sigma Aldrich, ON) buffered with 1 g/L sodium bicarbonate

(Sigma Aldrich, ON).

2.2.3. The effect of Salmosan® exposures on intermolt stage prawns

The objectives of the intermolt stage experiments were to determine the 3 x 1-hr

median lethal concentrations (LC50) for intermolt prawns; and, to determine the effects of

3 x 1-hr Salmosan® exposures on survival during molt, time to molt, and growth during

molting. The experimental set up for the intermolt exposures consisted of 6 waterbaths

29

each holding 10 x 9 L tanks and maintained at a target temperature of either 5, 11 or

17 °C, with 2 waterbaths per temperature (Figure 1). Prawns were acclimated for 24 h in

a tank with 2 prawns per tank. Each tank was randomly assigned to a nominal treatment

concentration of 0, 10, 50 or 100 µg/L azamethiphos and subjected to 3 x 1-h exposures

at t = 0, 6, and 24 h. Azamethiphos concentrations were selected based on previous range

finding tests. The final concentrations of azamethiphos were achieved by preparing a

stock solution of Salmosan® in seawater and adding an appropriate volume to the 9 L tank

with a syringe. The prawns were held in clean seawater in between exposures. A total of

120 prawns were exposed, with 10 prawns in 5 tanks per treatment. Prawns were fed

EWOS® (Bergen, Norway) Micro fish feed pellets ad libitum 3 times a week. Approximately

1 hr after feeding, any remaining food and feces were removed along with 50 – 60% of

the water, which was replaced with clean seawater.

Figure 1.. Schematic diagram of intermolt prawn exposure design. Intermolt

prawns were acclimated at 5, 11 or 17 °C with 2 prawns per tank. Prawns in each tank were subjected to 3 x 1-h exposures to azamethiphos at 0, 10, 50, or 100 µg/L in formulation as Salmosan®. Following exposure, prawns were monitored for mortality, time to molt, molting success, and growth.

Following exposure, the prawns were monitored in clean water for an additional 72

hr and mortality was then recorded for estimation of LC50 values at each temperature. To

ensure no prawns were in a pre-molt phase instead of an intermolt phase during exposure,

30

any prawns that molted within 1 week following exposure were euthanized and removed

from further analysis. Mass was recorded 1 d after exposure. Temperature, dissolved

oxygen, pH and conductivity were recorded during exposures and 3 times a week during

the monitoring period.

The prawns were then further monitored until they passed through ecdysis.

Observations were recorded at time of ecdysis, and 7 d following ecdysis after

euthanization with MS222 (Figure 1). The time to molt was recorded as the number of

days between the final exposure and ecdysis. Molting success was recorded, defined as

survival within the 7 d following ecdysis. The carapace length (CL) of the shed exuvia was

measured (CLexuvia), as well as the CL of the euthanized prawn (CL7d). Relative CL growth

was then calculated for each prawn as:

!"#$%&'")*+,-.%ℎ = 1)*23 − )*56789:

)*56789:; <100%

Sex was determined at euthanization by the presence of the appendix masculina (male)

on the second pleopod. Mass was also recorded at the time of euthanization. The relative

change in mass between the time of exposure (massexp) and time of euthanization

(Mass7d) was calculated for each prawn as:

!"#$%&'"@ℎ$A+"&AB$CC = DB$CC23 − B$CC56E

B$CC56EF <100%

The mass7d and CL7d were used to calculate Fulton’s K condition factor for each prawn as:

GH#%-AICJ@-AK&%&-AL$@%-, = DB$CC23

)*23M F <100

2.2.4. The effect of Salmosan® exposures on post-molt stage prawns

To compare sensitivity to Salmosan® between molt stages, post-molt prawns were

subjected to identical exposures as the intermolt prawns. The experimental set up for the

post-molt exposures consisted of 6 waterbaths each containing 9 x 9 L tanks and

maintained at a target temperature of either 5, 11 or 17 °C, with 2 waterbaths per

temperature (Figure 2). Prawns were acclimated for 24 h in the individual tanks containing

1 prawn per tank and monitored daily for molting throughout July and August, a molting

31

period for Pacific spot prawns previously observed in the lab. Prawns were fed frozen

mysid shrimp (Piscine Energetics, Vernon, Canada) ad libitum 3 times a week. An hour

after feeding, any remaining food and feces as well as 50 – 60% of the water were

siphoned out and replaced with clean seawater. Prawns were checked for molting each

day at 5:00 pm and exposed by 9:00 am.

Once a prawn molted, it was individually transferred into a separate tank

(individually housed) within the same waterbath system and randomly assigned a nominal

treatment concentration of Salmosan® at 0, 10, 50 or 100 µg/L azamethiphos. Prawns

were then subjected to 3 x 1-h exposures of Salmosan® at t = 0, 6 and 24 h and held in

clean water in between exposures. The final concentrations of azamethiphos were

achieved by preparing a stock solution of Salmosan® in seawater and adding an

appropriate volume to the 9 L tank with a syringe. Following the exposure period, the

prawn was monitored for an additional 72 h in clean seawater. Mortality was then recorded

for calculation of the LC50 values of Salmosan® at each temperature. At 5 and 11 °C,

n = 6 prawns were subjected to each combination of Salmosan® and temperature. Pre-

exposure mid-ecdysis mortalities at 17 °C resulted in n = 4 – 6 prawns subjected to each

treatment combination at this temperature. Surviving prawns were euthanized with MS222

at the end of an experiment. Temperature, dissolved oxygen, pH and conductivity were

recorded during exposures and 3 times a week during the monitoring period.

32

Figure 2. Schematic diagram of post-molt prawn exposure design. Pacific spot

prawns (Pandalus platyceros) were housed individually at 5, 11 or 17 °C and monitored until they molted. After molting, each prawn was individually subjected to 3 x 1-h exposures of 0, 10, 50 or 100 µg/L azamethiphos in formulation as Salmosan® and monitored an additional 72 h.

2.2.5. Statistical analysis

LC50 values and 95% confidence intervals for the post-molt and intermolt repeated

1-h exposures at each temperature were estimated using the Speaman Karber method in

the ecotoxicology package in R (Gama, 2015; R Development Core Team, 2017). LC50

values were compared between each temperature treatment group using the ratio test

according to Wheeler et al. (2006). For the exposures in which no LC50 could be

determined, the highest concentration without a significant difference in mortality

compared to the control groups was reported, representing the no observed effects

concentration (NOEC). Comparisons between proportion dead after the third pulse in each

treatment group were made using a contingency analysis in JMP® Version 13.1.0 (SAS

Institute Inc., 2016). A p-value of < 0.05 was used to infer statistical significance.

A generalized linear mixed effects model (GLMM) was used to determine the

effects of temperature and concentration on survival during molt using the lme4 package

in R version 1.1.4 (Bates et al., 2015; R Development Core Team, 2017). Separate

two-way analyses of variance (ANOVAs) were used to determine the effects of

temperature and concentration on time to molt, relative CL change, relative change in

mass and Fulton’s K in JMP® Version 13.1.0 (SAS Institute Inc., 2016). All analyses

33

included waterbaths and tanks as random effects. A p-value of < 0.05 was used to infer

statistical significance.

Because sex was determined at euthanization, sex was not considered in the

experimental design and as a result there was an unbalanced number of males and

females in each treatment group. Further, there were fewer females (n = 25) than males

(n = 95), and not enough (< 2) females in each treatment group to run separate analyses

on each sex. Therefore, the effect of sex was investigated in each endpoint. Where sex

had a significant effect on the endpoint measured, the females were removed from the

analysis. A generalized linear mixed effects model (GLMM) was used to determine the

effects of temperature, concentration and sex on survival during molt using the lme4

package in R version 1.1.4 (Bates et al., 2015; R Development Core Team, 2017).

Separate three-way analyses of variance (ANOVAs) were used to determine the effects

of temperature, concentration and sex on time to molt, relative CL change, relative change

in mass and Fulton’s K in JMP® Version 13.1.0 (SAS Institute Inc., 2016). All analyses

included waterbaths and tanks as random effects. A p-value of < 0.05 was used to infer

statistical significance. Consequentially, females were removed from analysis on time to

molt and Fulton’s K condition factor. Females were included in the analyses for all other

endpoints.

2.3. Results

2.3.1. Water quality

Temperature, dissolved oxygen, pH and conductivity were recorded 3 times a

week during the intermolt exposures and monitoring period. The water quality parameters

ranged within 2 °C of the target temperature. For the 5 °C target treatment, the average

temperature was 5.1 °C (range 5 – 7 °C). For the 11 °C target treatment, the average

temperature was 10.8 °C (range 10 – 12 °C). For the 17 °C target treatment, the average

temperature was16.9 °C (range 15 – 18 °C). The average pH was 8.41 (range 7.74 – 8.87.

The average conductivity was 47.6 µS/cm2 (range 43.2 – 52.3 µS/cm2). The average

dissolved oxygen was 8.12 mg/L (range 7.0 – 8.9 mg/L).

Temperature, dissolved oxygen, pH and conductivity were recorded 3 times a

week during the post-molt exposures and monitoring period. The water temperature

34

remained within 1.5 °C of the target temperature for all temperatures. For the 5 °C target

treatment, the average temperature was 5.5 °C (range 5 – 6.5 °C). For the 11 °C target

treatment, the average temperature was 11.3 °C (range 10 – 12.5 °C). For the 17 °C target

treatment, the average temperature was 16.9 °C (range 16 – 18 °C). The average pH was

8.10 (range 7.06 – 8.62). The average conductivity was 48.1 µS/cm2 (range 43.9 –

59.0 µS/cm2). The average dissolved oxygen was 7.8 mg/L (range 6.9 – 8.9 mg/L).

2.3.2. The effect of Salmosan® exposures on intermolt stage prawns

Prawns were subjected to repeated short-term exposures of Salmosan® during

their intermolt phase and subsequently monitored throughout their molting cycle to

determine the effect of these exposures on the time to molt, the ability to molt, and growth

during molt. All exposures were conducted at 5, 11 and 17 °C to determine whether

temperature modified any observed effects.

There was no evidence of an effect of Salmosan® on any of the endpoints

measured (molting success, time to molt, relative CL or mass growth, or Fulton’s K

condition factor) (p > 0.05, Figures 3 – 7). However, there was an effect of temperature

on molting success; there was significantly lower survival in the prawns held at the highest

temperature of 17 °C than at 5 °C (p = 0.014) or 11 °C (p = 0.008) in the control group as

well as the 10 and 100 µg/L groups (Figure 3). Overall, the 17 °C treatment group had

56 ± 9% survival, compared to 97 ± 3% at 11° and 95 ± 4% at 5 °C. All mortalities occurred

during ecdysis. Temperature also affected the mean time to molt, with prawns held at 5 °C

taking longer to molt than at 11 °C or 17 °C (p < 0.05; Figure 4). The mean time to molt

was 44 ± 3 d for prawns held at 5 °C, compared to 25 ± 2 d at 11 °C and 23 ± 2 d at 17 °C.

There was no evidence of an effect of either Salmosan® or temperature on relative

CL growth (p > 0.05; Figure 5), relative change in mass (p > 0.05; Figure 6), or Fulton’s K

condition factor (p > 0.05; Figure 7). The mean change in mass was negative for all

treatment groups (Figure 6).

35

Figure 3. Effect of Salmosan® and temperature on survival during molt of

Pacific spot prawns after 3 x 1-h exposures to Salmosan® (active ingredient: azamethiphos) at various temperatures. Prawns at 17 °C ( ) had lower survival than at 5 °C ( , p = 0.014) or 11 °C ( , p = 0.008). There was no effect of Salmosan® (p = 0.48). Bars indicate mean values; whiskers indicate standard error.

Figure 4. Effect of Salmosan® and temperature on the time to molt of male

Pacific spot prawns after 3 x 1-h exposures to Salmosan® (active ingredient: azamethiphos) at various temperatures. Prawns held at 5 °C ( ) took longer to molt than at 11 °C ( ) and 17 °C ( ) (p < 0.05). There was no effect of Salmosan® (p = 0.93). Lines indicate the medians; boxes indicate the first and third quartiles; whiskers indicate the minima and maxima.

36

Figure 5. Effect of Salmosan® and temperature on the relative change in

carapace length (CL) of Pacific spot prawns after 3 x 1-h exposures to Salmosan® (active ingredient: azamethiphos) at 5 °C ( ), 11 °C ( ), or 17 °C ( ). There was no evidence of an effect of Salmosan® (p = 0.78) or temperature (p = 0.15) on relative CL change. Lines indicate the medians; boxes indicate the first and third quartiles; whiskers indicate the minima and maxima.

Figure 6. Effect of Salmosan® and temperature on the relative change in mass

of Pacific spot prawns after 3 x 1-h exposures to Salmosan® (active ingredient: azamethiphos) at 5 °C ( ), 11 °C ( ), or 17 °C ( ). There was no evidence of an effect of Salmosan® (p = 0.69) or temperature (p = 0.98) on mean change in mass. Lines indicate the medians; boxes indic