Acute Toxicity of Β-N-Methylamino-L-Alanine (BMAA

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5-2015 Acute Toxicity of β-N-Methylamino-L-Alanine (BMAA) to Fathead Minnow (Pimephales promelas) and Zebrafish (Danio rerio) Jiayi Wang University of Nebraska-Lincoln, [email protected]

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Wang, Jiayi, "Acute Toxicity of β-N-Methylamino-L-Alanine (BMAA) to Fathead Minnow (Pimephales promelas) and Zebrafish (Danio rerio)" (2015). Dissertations & Theses in Natural Resources. 116. http://digitalcommons.unl.edu/natresdiss/116

This Article is brought to you for free and open access by the Natural Resources, School of at DigitalCommons@University of Nebraska - Lincoln. It has been accepted for inclusion in Dissertations & Theses in Natural Resources by an authorized administrator of DigitalCommons@University of Nebraska - Lincoln. ACUTE TOXICITY OF β-N-METHYLAMINO-L-ALANINE (BMAA) TO

FATHEAD MINNOW (PIMEPHALES PROMELAS) AND ZEBRAFISH (DANIO

RERIO)

Jiayi Wang

A THESIS

Presented to the Faculty of

The Graduate College at the University of Nebraska

In Partial Fulfillment of Requirements

For the Degree of Master of Science

Major: Natural Resource Sciences

Under the Supervision of Professors Kyle Hoagland and Daniel Snow

Lincoln, Nebraska

May, 2015

ACUTE TOXICITY OF β-N-METHYLAMINO-L-ALANINE (BMAA) TO

FATHEAD MINNOW (PIMEPHALES PROMELAS) AND ZEBRAFISH (DANIO

RERIO)

Jiayi Wang, M.S.

University of Nebraska, 2014

Advisors: Kyle Hoagland and Daniel Snow

β-N-methylamino-L-alanine (BMAA) is a neurotoxic amino acid produced by most species of cyanobacteria. Exposure to BMAA has been hypothesized as a cause of

ALS and possibly Parkinson’s and Alzheimer’s diseases for several decades. Both in vitro and in vivo experiments revealed that exposure to elevated concentrations of

BMAA can damage motor neurons and cause motor dysfunctions. However, the exact mechanism of BMAA-induced neurotoxicity has not been well understood.

Based on the available literature and in spite of its water-soluble and non-protein nature, BMAA appears to be able to bioaccumulate in organisms. The ubiquity of cyanobacteria and the potential for bioaccumulation of BMAA, arouse wide human health concern. Previous investigations into toxicity of BMAA mainly focused on various mammalian test models, including rats, mice and primates. However, the toxic effect of BMAA on aquatic organisms has attracted limited attention. Due to the potential for widespread BMAA production in aquatic ecosystems, it is important to understand the toxicity of BMAA in aquatic organisms, such as fish, from an ecotoxicological perspective

Because limited information exists about how exposure to BMAA influences fish, we

investigated the acute toxic effect of β-N-methylamino-L-alanine (BMAA) on two widely adopted model fish: fathead minnow (Pimephales promelas) and zebrafish

(Danio rerio). The 96 h toxicity tests revealed that BMAA is not acutely lethal to fathead minnow (juveniles: 96 h LC50 > 10 mg/L; larvae: 96 h LC50 > 10 mg/L) or zebrafish (larvae: 96 h LC50 > 10 mg/L; embryos: 10-day LC50 > 100 mg/L) at concentrations well above those reported in the literature. However, exposure to

BMAA at 100 mg/L significantly affected the heart rate of zebrafish embryos.

Additionally, in the zebrafish embryo-larvae behavioral assay, a range of locomotor function anomalies in zebrafish embryo-larvae exposed to BMAA were observed. On one hand, BMAA was found to accelerate the onset of spontaneous contractions and enhanced touch-evoked contractions. On the other hand, BMAA reduced performance in touch-evoked escape, free swimming and startle response.

To our knowledge, we are the first to investigate the acute toxicity of BMAA in fathead minnow and its behavioral effects on fish. The finding that BMAA influenced locomotor behaviors of fish is of both neurotoxicological and ecological importance.

iii

ACKNOWLEDGEMENTS

First, I would like to thank my advisors Dr. Kyle Hoagland and Dr. Daniel Snow. I am so grateful that I could be a student of Dr. Hoagland, a nice and responsible advisor.

Without him, I could not have been able to work on this project and pursue my M.S. degree in this country. Thank you to Dr. Daniel Snow for his valuable advice on my research design and academic writing.

I would like to thank Dr. Blair Siegfried and Dr. Ayse Kilic, for being willing to be my committee members. Thank you both for your support and suggestions on my thesis.

I would also like to thank all staff in the Water Science Lab, especially Dr. David

Cassada. His patient and professional guidance helped me to have a better understanding of HPLC analysis.

Besides, I want to thank Vicky Samek and Sara Findrick in the Life Sciences Annex, for their assistance in establishment and maintenance of the lab. I want to thank Drs.

Mark Pegg, Steve Thomas and Amy Burgin, for their help in providing equipment needed for my experiment. I also want to thank Kelly Heath and Kathy Pinkerton for their useful suggestions on animal care. Especially, please allow me to thank Dr. John

Carroll, Dr. Ayse Kilic, and Dr. John Holz and Mr. Tadd Barrow

(HABAquaticSolutions) for their generous financial help when I had trouble to continue my study. Without your support, I could never have had the chance to finish my research.

Finally, I want to say thank you to my family and my girlfriend Mary Wang. It was they who gave me the strength to finish my degree. I could not imagine how I could iv

be here without their care and encouragement.

TABLE OF CONTENTS

Abstract ...... i

ACKNOWLEDGEMENTS ...... iii

TABLE OF CONTENTS ...... v

LIST OF FIGURES ...... ix

ABBREVIATIONS ...... x

CHAPTER ONE. LITERATURE REVIEW

1.1 β-N-methylamino-L-alanine (BMAA)...... 1

1.1.1 ALS/PDC and the discovery of BMAA ...... 1

1.1.2 Structure and biochemistry of BMAA ...... 2

1.2 Neurotoxicity of BMAA ...... 3

1.2.1 BMAA ALS/PDC hypothesis ...... 3

1.2.2 Bioaccumulation of BMAA ...... 4

1.2.3 Experimental evidences of BMAA-induced neurotoxicity ...... 5

1.2.4 Mechanisms of BMAA-induced neurotoxicity ...... 7

1.3 BMAA in ecosystems ...... 9

1.3.1 Source of BMAA ...... 9

1.3.2 Occurrence of BMAA in diverse environments...... 11

1.3.3 Bioaccumulation of BMAA in wild organisms ...... 11

1.4 Ecotoxicity of BMAA ...... 13

1.4.1 Toxic effects of BMAA on terrestrial organisms ...... 13

1.4.2 Toxic effects of BMAA on aquatic organisms ...... 15

CHAPTER TWO: MATERIALS AND METHODS

2.1 Introduction…...... 18

2.2 Chemicals…………………………………………………………………………...... 22

2.3 Test organisms and maintenance ...... 23

2.4 Acute fathead minnow juvenile test ...... 24

2.4.1 Lab conditions ...... 24

2.4.2 Preparation before exposure ...... 24

2.4.2.1 Handling of fathead minnow juveniles ...... 24

2.4.2.2 Acclimation of fathead minnow juveniles ...... 25

2.4.3 Range-finding test ...... 25

2.5 Acute fathead minnow larvae test ...... 26

2.5.1 Lab conditions ...... 26

2.5.2 Preparation before exposure ...... 26

2.5.2.1 Handling of fathead minnow larvae ...... 26

2.5.2.2 Acclimation of fathead minnow larvae ...... 27

2.5.3 Range-finding test ...... 27

vii

2.6 Acute zebrafish larvae test ...... 28

2.6.1 Lab conditions ...... 28

2.6.2 Preparation before exposure ...... 29

2.6.2.1 Handling of zebrafish eggs ...... 29

2.6.2.2 Hatching of zebrafish eggs ...... 29

2.6.3 Range-finding test ...... 29

2.7 Zebrafish embryo-larvae test ...... 30

2.7.1 Lab conditions ...... 30

2.7.2 Preparation before exposure ...... 31

2.7.2.1 Handling of zebrafish embryos ...... 31

2.7.2.2 Handling of zebrafish adults ...... 31

2.7.2.3 Accumulation of zebrafish adults ...... 31

2.7.2.4 Preparation for spawning ...... 32

2.7.2.5 Spawning...... 32

2.7.2.6 Collection of eggs ...... 32

2.7.3 Range-finding test ...... 33

2.7.4 10-day LC50 test ...... 35

2.7.5 Zebrafish embryo-larvae behavioral assay ...... 37

2.7.5.1 Spontaneous contractions...... 39

viii

2.7.5.2 Touch-evoked contractions ...... 39

2.7.5.3 Touch-evoked escape ...... 40

2.7.5.4 Free swimming...... 41

2.7.5.5 Startle response ...... 42

2.7.5.6 Anxiety ...... 43

2.8 Statistical analysis ...... 45

CHAPTER THREE: RESULTS AND DISCUSSION

3.1 Results ...... 46

3.1.1 Acute toxicity of BMAA to fathead minnow and larval zebrafish ...... 46

3.1.2 Acute toxic effects of BMAA to zebrafish embryo-larvae ...... 46

3.1.2 Behavioral effects of BMAA to zebrafish embryo-larvae ...... 48

3.2 Discussion ...... 53

3.2.1 Conclusions ...... 57

3.2.2 Importance and future work ...... 57

LIST OF FIGURES

Figure 1.1...... 95

Figure 1.2...... 95

Figure 2.1...... 96

Figure 3.1...... 96

Figure 3.2...... 97

Figure 3.3...... 97

Figure 3.4...... 98

Figure 3.5...... 98

Figure 3.6...... 99

Figure 3.7...... 100

Figure 3.8...... 101

Figure 3.9...... 103

ABBREVIATIONS

AD: Alzheimer’s disease

AFT: acute fish test

ALS: amyotrophic lateral sclerosis

AMPA: α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid

ANOVA: analysis of variance

BBB: blood-brain barrier

BMAA: β-N-methylamino-L-alanine

BOAA: β-N-oxalylamino-L-alanine bpm: beats per minute cpm: contractions per minute

DI: deionized dpf: days post fertilization

DO: dissolved oxygen dph: days post hatch

EAA: excitatory amino acid

EC50: median effective concentration

EPA: Environmental Protection Agency

ER: endoplasmic reticulum

FET: fish embryo test hpf: hours post fertilization hph: hours post hatch

IC: internal plate control

IC50: median inhibitory concentration i.p.: intraperitoneal injection

LC50: median lethal concentration

LD50: median lethal dose

LOEC: Lowest-Observed-Effect Concentration

MS222: tricaine methanesulfonate mGluR5: metabotropic glutamate receptor 5

NEC: No Effect Concentration

NMDA: N-methyl-D-aspartate

NOEC: No-Observed-Effect Concentration

NPAA: non-protein amino acid

OECD: Organisation for Economic Cooperation and Development

PFOS: perfluorooctane sulphonic acid

ROS: reactive oxygen species

GSK3β: β-type glycogen synthase kinase-3

NOEC: No-Observed-Effect Concentration

PD: Parkinson’s disease

PDC: Parkinson dementia complex

SEM: standard error of the mean

CHAPTER ONE. LITERATURE REVIEW

1.1 β-N-methylamino-L-alanine (BMAA)

1.1.1 ALS/PDC and the discovery of BMAA

Amyotrophic lateral sclerosis (ALS), also known as Lou Gehrig’s disease, is a progressively debilitating, fatal disease caused by the degeneration of the motor neurons (Rowland and Shneider, 2001). Although evenly distributed in most areas of the world (Bradley and Mash, 2009), ALS has three high-incidence epicenters in

Guam, the Kii Peninsula of Japan, and west New Guinea, where it is often accompanied with Parkinson’s disease (PD) and dementia (Vyas and Weiss, 2009).

The incidence of ALS/Parkinson - dementia complex (ALS/PDC) on Guam was unnaturally higher (approximately 100 times) than elsewhere at its peak in 1950’s

(Arnold et al., 1953; Garruto and Yase, 1986; Prasad and Kurland, 1997), leading to a worldwide concern and a high interest in finding the etiology of the disease on this island (Arnold et al., 1953; Koerner, 1952; Mulder et al., 1954).

After ruling out genetic factors (Kurland and Mulder, 1954), the investigators turned to link ALS/PDC to special environmental conditions on Guam or unique habits of indigenous Chamorro people (Reed et al., 1987). Soon, an unusual traditional diet of Chamorros to eat food such as tortillas, soups and dumplings made from native cycad (Cycas micronesica Hill) seed flour became a popular research subject until the early 1970’s (Lythgoe and Riggs, 1949; Nishida et al., 1955; Whiting, 2

1964; Whiting et al., 1966). Although no successful lab animal models of ALS were established in cycad material toxicity tests (Laqueur et al., 1963; Levene et al., 1964;

Mickelsen et al., 1964), a new amino acid of potential neurotoxicity,

β-N-methylamino-L-alanine (BMAA), was isolated from cycad seeds in 1967 (Vega and Bell, 1967; Whiting, 1988).

The discovery of BMAA benefited from the original attempt to find

β-N-oxalylaminol-L-alanine (BOAA), a known neurotoxin responsible for paralytic disease lathyrism with symptoms similar to ALS/PDC (Bell, 2009; Rao et al., 1964).

Instead of BOAA, BMAA was incidentally found in cycad seeds and gradually became a widely studied potential cause of ALS/PDC (Bradley et al., 2009; Chiu et al.,

2011; Pablo et al., 2009; Spencer et al., 1987).

1.1.2 Structure and biochemistry of BMAA

β-N-methylamino-L-alanine (BMAA) is a small non-protein amino acid

(NPAA), with a chemical formula of C4H10N2O2 and a molecular weight of 118.13

(Dossaji and Bell, 1973; Jonasson et al., 2008). BMAA is structurally an alanine with a methylated amino group on β-carbon (Figure 1.1), which is similar to another neurotoxic amino acid, β-N-oxalylaminol-L-alanine (BOAA), only with a methyl group instead of an oxalyl group (Banack et al., 2007).

BMAA is a polar base and can be easily dissolved in water. Despite of its non-lipophilicity nature, BMAA is capable of bioaccumulating in organisms probably by binding to certain proteins in a protein-associated form (Cox et al., 2003; Murch et

al., 2004a). Free BMAA can be transformed into protein-associated BMAA during protein synthesis, and it can also be slowly released back to free form through acid hydrolysis (Murch et al., 2004a; Lopičić et al., 2011).

1.2 Neurotoxicity of BMAA

1.2.1 BMAA ALS/PDC hypothesis

The hypothesis that BMAA caused ALS/PDC was supported by toxicity test on chicks soon after its isolation (Vega and Bell, 1967). Both natural and synthetic

BMAA (i.p., 0.2-0.8 g/kg) caused weakness, inability and discoordination in young

(1-3 days) chicks (Whiting, 1988).

Similar tests on rats and mice also confirmed the acute toxicity of BMAA, showing symptoms including weakness, convulsion and discoordination (Vega et al.,

1968). However, because only acute and non-progressive toxic effects were observed, the hypothesis of BMAA as the causal agent of ALS/PDC was questioned and almost excluded (Polsky et al., 1972).

The “BMAA-ALS/PDC” hypothesis had been seldom mentioned until 1987, when a landmark paper reporting the neurotoxicity of BMAA to primates was published (Spencer et al., 1987). Based on observing a transitory hyperexcitable state with a following long whole-body shake/wobble in mice treated with BMAA (Ross and Spencer, 1987), Spencer and colleagues moved on to monkeys and recorded many ALS/PDC-like neurodegenerative symptoms, such as shaking and paralysis, in male cynomolgus macaques (1 year) fed by BMAA (oral, 0.1-0.3 g/kg per day) for up

to 12 weeks. This study presented clinical, neurophysiological and neuropathological evidence to support that BMAA could induce a primate motor system disorder involving the upper and lower motor neurons and the extrapyramidal system (Spencer et al., 1987).

Although Spencer’s observation was a great breakthrough, the high BMAA dose used in his work was criticized by Duncan et al. (1988) and Garruto et al. (1988), who at that time believed it was too high for Chamorros to reach the neurotoxicity threshold by consuming cycad seeds.

1.2.2 Bioaccumulation of BMAA

Due to BMAA’s non-lipophilicity and non-protein nature (Krüger et al., 2010),

BMAA was initially expected to be readily excreted from the human body after ingestion (Strong, 2003). However in 2002, unlike other polar substances, BMAA was surprisingly found to bioaccumulate in flying fox bats that fed on cycad seeds on

Guam (Cox and Sacks, 2002a, 2002b). Murch et al. (2004a) then suggested a bioaccumulation mechanism that dissolved BMAA (free BMAA) can be bound to proteins in an associated form (protein-associated BMAA), which was proved by

Cheng and Banack (2009) who detected a high content of protein-associated BMAA in cycad seeds. The bioaccumulation capacity of BMAA, together with the fact that only free BMAA was measured in early studies (Cheng and Banack, 2009), and the fact that flying fox bats were a traditional food of Chamorros (Cox et al., 2003), indicated a much lower previous estimation of the BMAA consumption by people on

Guam (Chiu et al., 2011).

The bioaccumulation of BMAA became an important base of the

BMAA-ALS/PDC hypothesis, because protein-associated BMAA possibly functions as an endogenous neurotoxic reservoir that slowly releases free BMAA in human bodies. Given that BMAA passes the blood-brain barrier (BBB) (Duncan et al., 1991;

Kisby et al., 1988; Smith et al., 1992), the gradual release of free BMAA was suggested to cause a chronic BMAA exposure in human brains (Murch et al., 2004a).

This suggestion was corroborated by the discovery of BMAA in brain tissues from

Chamorro ALS/PDC patients and Canadian Alzheimer’s disease (AD) patients, but not in healthy controls (Cox et al., 2003; Murch et al., 2004a, 2004b). The work of

Pablo et al. (2009) also supported the bioaccumulation of BMAA by finding high levels of BMAA in almost all (49 out of 50) brain samples from ALS and AD patients in North America.

It was noteworthy that two other investigations failed to detect any free or protein-associated BMAA in Chamorro ALS/PDC victim or US AD victim brains

(Montine et al., 2005; Snyder et al., 2009). However, the insensitivity of their methods was suggested as the cause for a lack of detection (Bradley and Mash, 2009;

Karamyan and Speth, 2008).

1.2.3 Experimental evidences of BMAA-induced neurotoxicity

Based on the inspiring observation of Spencer et al. (1987), as well as the finding of BMAA bioaccumulation, a number of BMAA neurotoxicity investigations

have subsequently been done in the past three decades. Most investigations, both in vivo and in vitro, supported the BMAA-ALS/PDC hypothesis (Banack et al., 2010;

Chiu, 2011; Karamyan and Speth, 2008; Karlsson et al., 2014; Lopičić et al., 2011;

Papapetropoulos, 2007; Rodgers, 2014).

BMAA exposure in vivo caused motor function anomalies such as ataxia, rigidity, weakness and convulsion (Chang et al., 1993; Dawson et al., 1998; Matsuoka et al., 1993; Rakonczay et al., 1991; Seawright et al., 1990), vacuolization and mitochondria dysfunction of spinal cord motor neurons (de Munck et al., 2013b), degeneration of hippocampal neurons followed by long-term cognitive function deficits (Karlsson et al., 2009c, 2011, 2012), selective damage of cerebellum

(Seawright et al., 1990), probable damage of dopaminergic terminals (Santiago et al.,

2006), and perturbation of serum metabolism (Engskog et al., 2013) to rats. In vivo mice experiments showed motoric impacts of BMAA similar to those in rats (Smith and Meldrum, 1990). Al-Sammak (2012), in a 14 day-BMAA exposure (i.p.) test on mice, observed a median lethal dose (LD50) of 3 mg/g and symptoms including dyspnea, urinary and fecal incontinence, and uncoordinated limb and body movement.

Other neurotoxic effects of BMAA on mice included acute locomotive activity impairments and hyperactivity (Karlsson et al., 2009b), neurodegeneration of hippocampal neurons (Buenz and Howe, 2007), and retinal neuron death (Santucci et al., 2009).

Additionally, various in vitro studies also proved the neurotoxicity of BMAA by reporting BMAA-induced death of spinal cord (Rao et al., 2006) and cortical

neurons in rodents (Lobner et al., 2007; Ross et al., 1987; Weiss and Choi, 1988;

Weiss et al., 1989).

It was notable that not all investigations supported the BMAA-ALS/PDC hypothesis. Perry et al. (1989) fed 2-month-old female mice with a total dose of 15.5 g/kg BMAA over 11 weeks; Cruz-Aguado et al. (2006) fed 6 month old male mice with daily 28 mg/kg BMAA for 30 days. Although high doses of BMAA were used, no behavioral and neuropathological impacts of BMAA were observed in either study.

However, the lack of toxic effects of BMAA in both works was explained as inadequate behavior observation methods and improper dose selection, respectively

(Karamyan and Speth, 2008). More recently, Lee and McGeer (2012) investigated the toxicity of BMAA to three different neuron-derived human cancer cell lines. The researchers failed to observe any strong cell damage caused by BMAA exposure and concluded that the BMAA-ALS/PDC hypothesis became untenable. However, Chiu et al. (2011) questioned their conclusion because the cell lines they used were highly proliferative immortalized cells, which are significantly different in physiological characteristics from normal neurons in vivo.

1.2.4 Mechanisms of BMAA-induced neurotoxicity

Despite the various studies implicating BMAA in ALS/PDC, the exact mechanism of its neurotoxicity has not been well understood. The first mechanistic

BMAA paper was published in 1988 by Weiss and Choi (1988). The authors found

− that the presence of a physiological concentration (≥ 10 mM) of bicarbonate (HCO3 )

ions greatly enhanced the neurotoxicity of BMAA in vitro. The bicarbonate dependency of BMAA might contribute to the formation of a BMAA-carbamate

(Figure 1.2) that had a structure similar to glutamic acid (glutamate), a well-known excitatory amino acid (EAA) (Nunn and O’Brien, 1989; Myers and Nelson, 1990).

A number of studies have provided evidence that BMAA interacted with different EAA receptors and induced excitotoxicity (Chiu et al., 2012): BMAA acted as an agonist for NMDA receptors, AMPA receptors and metabotropic glutamate receptor 5 (mGluR5), finally promoting neural cell damage (Chiu et al., 2012; Goto et al., 2012; Liu et al., 2009; Lobner et al., 2007; Nedeljkov et al., 2005; Rakonczay et al., 1991; Weiss and Choi, 1988); BMAA activated non-NMDA ionotropic glutamate receptors, causing a decrease of input membrane and an imbalance between intracellular [Na+] and [K+] activity (Lopicic et al., 2009). BMAA was also found to produce an in vitro elevation of [Ca2+] level (Brownson et al., 2002; Chiu et al., 2012;

Cucchiaroni et al., 2010; Okle et al., 2013a; Rao et al., 2006), which in brains plays an important role in EAA-induced neurotoxicity and cell death (Choi, 1987, 1992;

Randall and Thayer, 1992; Tymianski et al., 1993).

In addition, BMAA inhibited the cysteine/glutamate antiporter system Xc- mediated cystine uptake, leading to decreased cellular glutathione and increased oxidative stress in cells (Liu et al., 2009). BMAA-involved oxidative stress was supported by various other studies recording a rise of reactive oxygen species (ROS) level both in vitro and in vivo (Chiu et al., 2012; Cucchiaroni et al., 2010; de Munck et al., 2013a; Okle et al., 2013b; Lobner et al., 2007; Rao et al., 2006; Santucci et al.,

2009).

In 2009, Karlsson et al. (2009a) found that BMAA interacted with melanin and increasingly accumulated in melanin and neuromelanin containing cells, suggesting a possible link between ALS/PDC and its common symptom of pigmentary retinopathy

(Cox et al., 1989). Nunn and Ponnusamy (2009) also proposed a biochemical mechanism of BMAA toxicity in which BMAA reacted non-enzymatically with pyridoxal-50-phosphate, releasing methylamine. The formation and release of methylamine might be important because chronic exposure to methylamine in rats caused oxidative stress (Deng et al., 1998).

Some more recent findings include: BMAA induced human protein misfolding and aggregation by misincorporated into proteins in place of L-Serine, indicating a trigger for neurological diseases associated with protein-misfolding including ALS

(Dunlop et al., 2013; Rodgers and Dunlop, 2011); BMAA enhanced the level of

β-type glycogen synthase kinase-3 (GSK3β) and TAR-DNA-binding protein 43

(TDP-43), which were both involved in neurodegenerative diseases (de Munck et al.,

2013b); BMAA caused dysregulation of the cellular protein homeostasis followed by endoplasmic reticulum (ER) stress and increasing Caspase 12 cleavage in human neuroblastoma cells (Okle et al., 2013a).

1.3 BMAA in ecosystems

1.3.1 Sources of BMAA

Although BMAA has been isolated from endemic Guam cycads (Cycas

micronesica Hill) in 1967, its ultimate source in cycads remained unknown for decades. In 2003, Cox et al. (2003) first discovered that BMAA was produced by nitrogen-fixing cyanobacteria of the genus Nostoc that lived in endosymbiosis with specialized coralloid roots of the cycads.

Cyanobacteria have been well known for their ability to produce a wide range of secondary metabolites (Carmichael 1992; Moore, 1996; Namikoshi and Rinehart,

1996; Welker and Von Döhren, 2006). Many of these by-products are dermal irritants and known to affect gastrointestinal tract, liver, or neural system of both animals and humans, and known collectively as cyanotoxins (Chorus and Bartram, 2003; Dow and

Swoboda, 2000; Mohamed, 2008; Stewart et al., 2008). Generally, each of these cyanotoxins seems to be produced by only a few species of cyanobacteria (Rippka et al., 1979). However, BMAA was reported to be produced by most species of cyanobacteria, including symbionts and free-living groups from both terrestrial and aquatic environments (Banack et al., 2007; Bidigare et al., 2009; Cox et al., 2005;

Esterhuizen-Londt et al., 2011a; Roney et al., 2009).

BMAA belongs to a class of over 900 non-protein amino acids (NPAAs) found in nature produced by plants, fungi, and microorganisms (Rubinstein, 2000). Although

NPAAs have many unknown functions (Vranova et al., 2011), a majority of them appear to be nitrogen-storage molecules and potential anti-predation toxins (Karlsson,

2010). However, limited information exists on the nature and role of BMAA production in cyanobacteria (Downing et al., 2011; Berntzon et al., 2013). Recently, planktonic diatoms (Jiang et al., 2014) and dinoflagellate Gymnodinium catenatu

(Lage et al., 2014) were also suggested to be potential BMAA producers.

1.3.2 Occurrence of BMAA in diverse environments

Cox et al. (2005) pointed out that the occurrence of BMAA might not be limited to Guam, but be globally distributed due to the ubiquity of cyanobacteria.

Although there is not enough evidence to prove the ubiquity of BMAA (Faassen, 2014) so far, many studies have confirmed the wide extent of BMAA in diverse ecosystems.

BMAA was found in cyanobacteria samples collected from fresh and brackish waterbodies in the British Isles (Metcalf et al., 2008), coastal and open-water areas of the Baltic Sea (Jonasson et al., 2010), urban waters in the Netherlands (Faassen et al.,

2009), freshwater impoundments in South Africa (Esterhuizen and Downing, 2008;

Esterhuizen-Londt and Downing, 2011), arid inland areas (Roney et al., 2009) and freshwater lakes (Li et al., 2010; Jiao et al., 2014) in China, and highland lakes in

Peru (Johnson et al., 2008). Water or planktonic samples from shallow springs of the

Gobi Desert in Mongolia (Craighead et al., 2009), dams in Brazil (Moura et al., 2009), reservoirs in the Midwest United States (Al-Sammak et al., 2013, 2014), west coastal marine environments of Sweden (Jiang et al., 2014) and two coastal waterbodies of

Portugal (Lage et al., 2014) also contained BMAA. In addition to aquatic environments, BMAA was detected in dried cyanobacterial crusts and mats from the deserts of Gulf region, particularly Qatar (Cox et al., 2009).

1.3.3 Bioaccumulation of BMAA in wild organisms

BMAA was previously demonstrated to bioaccumulate in Guam’s cycad seeds

and flying fox bats through the food chain from endosymbiotic cyanobacteria (Banack and Cox, 2003; Cox and Sacks, 2002a; 2002b; Cox et al., 2003). Therefore, the ubiquity of cyanobacteria also suggested that similar situation might occur in other environments containing cyanobacteria, especially aquatic ecosystems.

To investigate the potential for bioaccumulation of cyanotoxins in Nebraska,

Al-Sammak et al. (2014) examined 72 plant samples and 248 fish samples collected from local reservoirs between 2009 and 2010; 15 plant samples and 31 fish samples contained BMAA (Al-Sammak et al., 2014). Bioaccumulation of BMAA was also detected in zooplankton, mussels, oysters and fish in the Baltic Sea (Jonasson et al.,

2010), shrimp, lobsters, crabs, oysters and fish in South Florida coastal waters

(Banack et al., 2014; Brand et al., 2010), fish and crocodiles in a South African natural freshwater body (Esterhuizen-Londt, 2010), and mollusks, crustaceans and fish in Chinese lake Taihu (Jiao et al., 2014).

In Sweden, BMAA was found in oysters and fish from local fish market

(Spáčil et al., 2010), as well as mussels on Swedish west coast (Salomonsson et al.,

2013). BMAA was also detected in organs of sharks in South Florida (Mondo et al.,

2012) and commercial shark cartilage (fin) food supplements (Mondo et al., 2014).

The wide spread occurrence of cyanobacteria-produced BMAA, together with its potential for bioaccumulation within different organisms, indicate multiple potential paths of human exposure such as direct contact during recreational activities, consumption of water and foods, and even inhalation of aerosols and dusts (Banack et al., 2010; Stommel et al., 2013). Besides the human health concern, the ubiquity of

cyanobacteria also suggests an ecotoxicological concern about the fate and impact of

BMAA in diverse ecosystems, especially aquatic ecosystems.

1.4 Ecotoxicity of BMAA

In order to establish a causative association between BMAA exposure and human ALS/PDC, most toxicological tests of BMAA focused on proving its neurotoxicity in several mammalian test models such as rats, mice and primates (Chiu et al., 2011; Karamyan and Speth, 2008; Lopičić et al., 2011; Rodgers, 2014).

Compared to these human health risk investigations, quite limited attention has been drawn to the ecological risk assessment of BMAA. Bidigare et al. (2009) reported possible implications of cyanobacteria and BMAA in avian vacuolar myelinopathy

(AVM), a neurological disease that produces uncoordinated behavior in affected birds in wetland ecosystems of the southeastern United States. Unfortunately, the authors provided no toxicological evidences to support such a link (Bidigare et al., 2009). In fact, only a few studies involving the ecotoxicity of BMAA exist to date.

1.4.1 Toxic effects of BMAA on terrestrial organisms

In 2009, Zhou et al. (2009) reported the toxic effects of BMAA on fruit fly

(Drosophila melanogaster). Adult fruit flies (0-3 days) were raised on sucrose-agar medium mixed with water-dissolved BMAA at a final concentration of 4 mM. Dietary intake of BMAA not only reduced lifespan of both male (< 35 days) and female (< 25 days) fruit flies compared to that of control (> 50 days), but also weakened the

climbing performance of both males and females. The authors also found that the adverse effects of BMAA could be partly restored by cofeeding with leucine or lysine, both of which might share the same transporter system with BMAA on blood-brain barrier of fruit flies (Zhou et al., 2009). Zhou and colleagues (2010) continued their research on fruit flies but used sucrose-agar media containing different concentrations of BMAA (0, 2, 4, 6, 8 or 10 mM). The result again showed that the BMAA-induced reduction of lifespan and climbing ability were dose-dependent. Some other effects of

BMAA on fruit flies were also observed, including BMAA accumulation in brain, female fertility decrease and learning and memory ability impairment (Zhou et al.,

2010).

In work of Goto et al. (2012) using the same Drosophila model, virgin female fruit flies (1-3 days) were fed with food pellets containing BMAA (25 mM) for 6 days.

Starting from 2 days of feeding, fruit flies have gradually exhibited motor dysfunctions in standing, walking, climbing, flying, and wing beat. Physiological effects of BMAA ingestion were also reported in this study. Postsynaptic response abnormalities, such as more rise time, reduced amplitude, and longer duration, of an identified glutamatergic cell were detected in fruit flies fed with BMAA, indicating that they suffered from intracellular excitotoxicity (Goto et al., 2012).

Okle et al. (2013a) investigated the toxicity of BMAA on summer bees of the

European honeybee (Apis mellifera). BMAA (0.625 or 1.25 mM) bath application on the bee brains for 30 min lead to the generation of ROS and an increase of antennal lobe activity with higher Ca2+ level in brain neurons. Feeding bees with sugar water

mixed with bee saline containing BMAA (final concentration 5 mM) significantly increased the mortality after 24 h and impaired their odor learning ability after 5 days.

By feeding 14C-BMAA, BMAA was found to occur in head and thorax, and accumulate in abdomen, of bees with little or no excretion. 14C-BMAA was also transferred from one bee to another via trophallaxis, resulting in an exposure of the whole beehive (Okle et al., 2013a).

It should be noted that Drosophila and honeybees are both important insect models of neuroscience research (Bilen and Bonini, 2005; Menzel and Muller, 1996).

On one hand, the observations of motor dysfunctions in fruit flies and odor learning impairment in honeybees are important proofs of BMAA-induced neurotoxicity. On the other hand, the finding that BMAA reduced lifespan and female fertility in fruit flies, and that BMAA was transferred among bees via trophallaxis are of ecological importance.

1.4.2 Toxic effects of BMAA on aquatic organisms

BMAA has been detected in diverse ecosystems, especially aquatic ecosystems (Al-Sammak et al., 2014; Faassen, 2014). Given the worldwide increasing frequency and magnitude of cyanobacteria blooms (Carmichael, 2001), cyanobacteria-produced BMAA might increasingly occur in aquatic ecosystems. Thus, there is a need to understand the outcome of BMAA exposure in a variety of aquatic organisms. However, using aquatic organisms as the test models is relatively novel in

BMAA toxicity research.

In 2011, Esterhuizen et al. (2011) reported a rapid uptake and accumulation of

BMAA by lab cultured aquatic macrophyte Ceratophyllum demersum. The same group also investigated the effect of BMAA on the oxidative stress responses of C. demersum (Esterhuizen-Londt et al., 2011b). In vivo exposure test revealed that

BMAA (0.5-100 mg/L) had a significant inhibitory effect on all tested oxidative stress response enzymes, including superoxide dismutase, catalase, guaiacol peroxidase, glutathione peroxidase and glutathione reductase. In contrast, enzymes not related to oxidative stress response, such as alcohol dehydrogenase, alkaline phosphatase and acetylcholinesterase, were not affected by BMAA (5-500 mg/L) in in vitro binding experiment. The authors suggested that one important mechanism of BMAA is to induce oxidative stress by inhibiting antioxidant enzymes (Esterhuizen-Londt et al.,

2011b).

Lürling et al. (2011) explored both acute and chronic toxicity of BMAA to

Daphnia magna. Although BMAA was not acutely lethal (48 h LC50 > 10,000 μg/L) to neonate (< 24 h) Daphnia, it reduced mobility (48 h IC50 40 μg/L) and affected life history characteristics such as reproduction and population growth rate.

Bioconcentration of BMAA was also detected after a 96 h exposure to 100-10,000

μg/L BMAA in juvenile (> 48 h and < 72 h) Daphnia, and after a 15 days exposure to

50 and 100 μg/L BMAA in neonates (< 24 h) (Lürling et al., 2011).

The first paper focusing on the effects of BMAA on aquatic vertebrates was published in 2009. Purdie et al. (2009b) showed physiological and morphological toxicity of BMAA to the early-life stage development of zebrafish (Danio rerio).

BMAA exposure (5-50,000 μg/L) in zebrafish embryo at 6 h post fertilization (hpf) for 5 days caused a range of neuromuscular and developmental abnormalities, including reduced heart rate, pericardial edema, abnormal spinal axis formation, clonus-like convulsions and significant premature point of hatch. According to this study, Danio rerio might be a useful model of BMAA toxicity research because of its sensitivity to BMAA (Purdie et al., 2009b).

Purdie’s group then expanded their study on BMAA to two other aquatic species besides zebrafish: brine shrimp (Artemia salina) and the protozoan Nassula sorex (Purdie et al., 2009a). Responses to BMAA exposure included loss of phototaxis for brine shrimp and increased mortality for all species: Danio rerio larvae (10-day

LC50 10 μg/L), Artemia salina nauplii (24 to 72 h LC50 3,600 to 1,500 μg/L) and

Nassula sorex (72 h LC50 5,000 μg/L). Artemia salina and Nassula sorex were also suggested to be potential models for BMAA investigations because both of them are consumers of cyanobacteria (Purdie et al., 2009a).

In short, BMAA is toxic to several organisms from both terrestrial and aquatic environments. Due to the potential global occurrence of BMAA produced by ubiquitous cyanobacteria (Cox et al., 2005), extensive ecotoxicity of BMAA might exist, especially in aquatic ecosystems. More investigations are needed to evaluate the exposure level, biological fate and toxic effects of BMAA on different aquatic organisms.

CHAPTER TWO: MATERIALS AND METHODS

2.1 Introduction

A toxicity test is usually conducted to investigate the toxicity of a substance on living organisms, such as primates, rodents, fish, or Daphnia, for either a human safety evaluation or an ecological impact assessment (Abbott, 2005; Ecobichon, 1997;

Guilhermino et al., 2000). In a toxicity test, the toxicity of the test substance is often evaluated by measuring its quantifiable effects on exposed organisms at certain endpoints such as survival, growth, reproduction, behavior, and metabolism, within a certain period of time (Stokes, 2002; Suter, 1990). According to different exposure times of test substances, toxicity tests are divided into three types: acute, subacute, and chronic (Gupta and Bhardwaj, 2012). Due to its relatively short time and low cost, an acute toxicity test is the first test conducted in most cases. An acute toxicity test provides a preliminary understanding of the toxic nature of the test substance and serves as an information base for further subacute and chronic tests (Akhila et al.,

2007). In an acute toxicity test, the test organisms are usually treated with high doses of test substance, in order to cause changes in at least one measurable endpoint within a short period of time.

Mortality rate is one of the most easily observable endpoints in a toxicity test.

Therefore, the lethality of a substance to test organisms has become an important aim of an acute toxicity test (Toth, 1997, 2000). In a toxicity test using aquatic organisms, the acute lethality of a test substance is often expressed as the Median Lethal

Concentration (LC50), which causes death of 50% of the test organisms. A typical aquatic toxicity test for LC50 determination starts with a range-finding test, followed by a LC50 test. The range-finding test is aimed at finding a proper range of toxin concentrations as a reference for the LC50 test (Carpenter et al., 1974).

Generally in the range finding test, test organisms are exposed to a test substance at 5-6 gradient concentrations (≤ 100 mg/L), spaced by a large factor, such as 10. After the exposure, a range from the highest concentration that causes no mortality to the lowest substance concentration that causes absolute mortality of the test organisms is recorded. In the LC50 test, the test organisms are exposed to 5-6 gradient concentrations bordered by the range recorded in the range-finding test. After the same exposure time, the LC50 value can be calculated from the

Concentration-Mortality curve for the test substance.

Besides mortality rate, sub-lethal anomalies of test organisms in development, behavior, physiology, and genetics are also widely adopted endpoints in toxicity tests.

No Effect Concentration (NEC), the highest substance concentration that causes no anomalies in the test organism, reflects the threshold concentration of a substance before initiating significant toxic effects (van der Hoeven, 1997). Because the actual

NEC is almost impossible to measure, it is usually estimated from the

No-Observed-Effect Concentration (NOEC) - Lowest-Observed-Effect Concentration

(LOEC) interval (Laskowski, 1995). Median effective concentration (EC50) and median inhibitory concentration (IC50) are two other common sub-lethal endpoints, referring to the substance concentration that induces effects and inhibition,

respectively, in 50% of the test organisms (Sebaugh, 2011). NOEC - LOEC, EC50 and

IC50 endpoints can be investigated along with the LC50 value as important supplements to the toxicity of a test substance.

In aquatic toxicity tests, fish are crucial test models (Harshbarger and Clark,

1990), not only due to their indispensable roles in aquatic food chains, but also because they are an important aquatic food source for humans (Lammer et al., 2009).

Fish have been used in toxicity tests for more than 150 years (Penny and Adams, 1863, as cited in Hunn, 1989). Due to an increasing need for toxicant assessment and ecological study in recent decades, a number of valuable fish models, such as fathead minnow (Pimephales promelas), zebrafish (Danio rerio), and Japanese medaka

(Oryzias latipes), have been established and widely used in toxicity or ecotoxicity investigations (Ankley and Johnson, 2004; Ankley and Villeneuve, 2006; T.

Braunbeck et al., 2005; Hatanaka et al., 1982; Hill et al., 2005; McGrath and Li, 2008;

Örn et al., 2003). Advantages of these fish models include small body size, well characterized development/growth process, short maturation/reproduction cycle, high fecundity, and high tolerance of different environmental stresses.

Fathead minnow, a member of the demersal cyprinid family, is broadly distributed in temperate waters across North America (Divine, 1968; Eddy and

Underhill, 1974; Held and Peterka, 1974; Isaak, 1961; Page and Burr, 1991; Zimmer et al., 2001), and plays a key role in the regulation of structure and function of aquatic ecosystems (Gingras and Paszkowski, 1999; Paszkowski et al., 2004; Scott and

Crossman, 1973; Zimmer et al., 2002). As a native fish species, fathead minnow has

become the most widely studied toxicological fish model in North America (Ankley and Villeneuve, 2006), owning one of the biggest aquatic toxicology databases

(Ankley et al., 2001; Gray et al., 2002; Keddy et al., 1995; Miracle et al., 2003; Sinks and Schultz, 2001).

Zebrafish (Danio rerio), another demersal cyprinid species, is originated from the Ganges River system, Burma, Malakka and Sumatra (Eaton and Farley, 1974;

Engeszer et al., 2007; Talwar and Jhingran, 1991). Zebrafish has been proved to be a valuable fish model and used for decades in a variety of toxicity investigations, including chemical screening, water quality control, ecotoxicological assays, and neurotoxicology studies (Coverdale et al., 2004; Goldsmith, 2004; Goolish et al., 1999;

Hill, et al., 2005; Hisaoka and Battle, 1958; Laale, 1977; Lele and Krone, 1996; Nagel,

2002; Parng et al., 2002).

In the traditional acute fish test (AFT) with an endpoint of lethality, such as the LC50 test, exposure to toxic substances is hypothesized to bring fish severe distress and suffering (Chandroo et al., 2004; Nagel, 2002). Under the push of the animal rights legislations and animal protection organizations, a few alternative test approaches have been established, such as the in vitro fish cell assay and in vivo fish embryo test (FET) (Babich and Borenfreund, 1991; Bichara et al., 2014; Embry et al.,

2010; Henn and Braunbeck, 2011; Nagel, 2002; Ryan and Hightower, 1994; Tollefsen et al., 2006; van den Brandhof and Montforts, 2010). Numbers of in vitro cytotoxicity assays using fish cells have been developed in last 30 years (Castaño et al., 2003), and they showed a good correlation with in vivo fish test in chemical toxicity level ranking

(Kilemade and Quinn, 2003). However, in vitro tests are limited by their inability to actually represent natural situations in the whole organism (Fleming, 2007), and

Segner (2004) also concluded that the in vitro cytotoxicity assay was less sensitive than in vivo toxicity test for fish by reviewing previous works. By contrast, in vivo

FET, especially the zebrafish embryo test, has been proposed to be a promising alternative to the traditional AFT (Belanger et al., 2013; Lammer, et al., 2009; Scholz et al., 2008; Strähle et al., 2012). McKim (1977) has found that toxic effects on early life-stage are able to predict chronic toxicity of a substance, and embryo-larvae and early juvenile are the most sensitive stages of fish development. The Zebrafish embryo model is ideal for FET because zebrafish develop fast and normally hatch within 4 days post fertilization (dpf) at 26 °C (Nagel, 2002). In addition, the embryonic development of zebrafish is phenotypically identifiable and has been well-described in detail (Berry et al., 2007; Hinton et al., 2005; Kimmel et al., 1995).

The observation of zebrafish embryonic development can be easily achieved through the transparent chorion. The increasing popularity of zebrafish embryo tests also benefits from a reliable correlation (R2 = 0.854; p = 0.05; n = 56) between the zebrafish embryo tests and the traditional AFT, in spite of a prediction weakness

(Ratte and Hammers-Wirtz, 2003).

2.2 Chemicals

β-N-methylamino-L-alanine (BMAA), tricaine methanesulfonate (MS222),

3,4-dichloroaniline (3,4-DCA) and sodium bicarbonate (NaHCO3) were purchased

from Sigma-Aldrich (St. Louis, MO). All other chemicals used were of analytical grade quality. BMAA, NaHCO3 and 3,4-DCA stock solutions were all prepared in deionized (DI) water (Milli-Q, Millipore, MA) one day before the onset of corresponding test and stored at 4 °C. MS222 solutions were freshly prepared before use.

2.3 Test organisms and maintenance

All test organisms, including fathead minnows (Pimephales promelas) juveniles and larvae, and zebrafish (Danio rerio) adults and eggs, were obtained from

Aquatic Research Organisms (Hampton, NH). Maintenance and testing of organisms were all carried out in large growth incubators (Percival model I-35LLVL, Percival

Scientific, Boone, IA) kept in the Life Science Annex (LScA) at the University of

Nebraska-Lincoln (UNL), with room temperature, humidity and pressure controlled.

All glassware used for fish holding were pre-acid-washed with acetic acid soaking for at least 24 h and thoroughly rinsed with corresponding dilution water. Dilution water was aerated for at least 12 h in growth incubator before use, in order to reach a proper temperature and a dissolved oxygen (DO) level of near saturation. After aeration, the final pH of dilution water was adjusted to 7.5 (for fathead minnow) or 7.3 (for zebrafish) with the addition of 1 M HCl or 1 M NaOH. Water parameters such as DO, pH, temperature, hardness and alkalinity were measured every 24 h. If necessary, a semi-static water renewal method was applied to prevent DO deficiency and ammonia accumulation. The water renewal was handled in a very careful manner to remove

most food residues and fish feces without causing much stress in fish. All test organisms were observed regularly. Dead fish were removed as soon as observed.

Abnormal behaviors and the mortality rate of fish were recorded every 24 h.

2.4 Acute fathead minnow juvenile test

2.4.1 Lab conditions

The temperature in the growth incubators was maintained at 20 ± 1 °C and the light/dark cycle was set at 16 h/8 h. Both the acclimation and the toxicity test chambers were 3.8 L sphere glass fish bowls (Anchor Hocking #25178, Lancaster,

Ohio). The dilution water was the dechlorinated tap water obtained by use of two activated carbon treatment tanks (Culligan, Rosemont, IL). The final pH of the dilution water and all other test solutions was adjusted to 7.5. Other lab conditions were controlled following the US EPA guidelines (US EPA, 2002).

2.4.2 Preparation before exposure

2.4.2.1 Handling of fathead minnow juveniles

One month old fathead minnow juveniles were shipped by air in a plastic container. After arrival, the container was put in the growth incubator for 1-2 h until it had a water temperature close to 20 °C. Then the juveniles were randomly selected and transferred to acclimation chambers containing 3 L dilution water (< 30 fish/fish bowl) using a nylon fish net.

2.4.2.2 Acclimation of fathead minnow juveniles

Before the toxicity test, fish were acclimated under lab conditions for 7 days, regularly fed with commercial fish food flakes (TetraMin, Tetra, Blacksburg, VA) twice a day. All of the water in each acclimation chamber was renewed every 24 h with the help of another acclimation chamber as the “water-change” bowl, in a method described as follows: first, a hemispheric fish net was used to cover the mouth of the empty “water change” bowl; second, fresh dilution water was added into the

“water-change” bowl until the fish net bottom was submerged 1-2 cm into the water; third, approximate 3/4 of the water in the acclimation chamber was poured out in a very gentle and careful manner to prevent the fish from escaping; fourth, the remaining water containing the fish in the acclimation chamber was gently poured into the “water-change” bowl through the fish net, which had holes big enough for feces to pass but small enough to trap the fish safely; then, the acclimation chamber was cleaned and refilled with 3 L fresh dilution water; finally, the fish trapped in the fishnet were transferred quickly but carefully back to the acclimation chamber.

2.4.3 Range-finding test

Fathead minnow juveniles were treated with either the dilution water as the control or BMAA. BMAA solutions were freshly diluted from 10 g/L stock solution before use.

Acclimated fathead minnow juveniles (1 month) were randomly selected and carefully transferred into test chambers using a fish net. As a result, fathead minnow

juveniles were divided into five test groups, exposed to dilution water (control) and four gradient concentrations of BMAA (300 μg/L, 1,000 μg/L, 3,000 μg/L and 10,000

μg/L), respectively. Each test group had one test chamber. Each test chamber contained 1 L test solution and 10 fathead minnow juveniles (10 juveniles/1 L test solution/test chamber). The total exposure time was 96 h and no feeding was applied during the test. The test solution of each test chamber was renewed every 24 h with the same renewal method adopted during acclimation; A 10 ml solution sample from each test group was taken every 24 h and stored at -20 °C.

2.5 Acute fathead minnow larvae test

2.5.1 Lab conditions

The temperature in growth incubators was maintained at 20 ± 1 °C and the light/dark cycle was 16 h/8 h. Both the acclimation and the toxicity test chambers were 3.8 L glass fish bowls. The dilution water was the dechlorinated tap water. The final pH of dilution water and all other test solutions was adjusted to 7.5. Other lab conditions were controlled following the US EPA guidelines (US EPA, 2002).

2.5.2 Preparation before exposure

2.5.2.1 Handling of fathead minnow larvae

Fathead minnow larvae at the age of 5 days post hatch (dph) were shipped by air in a plastic container. After arrival, the container was put in the growth incubator

for 1-2 h until it had a water temperature close to close to 20 °C. Then, larvae were randomly selected and transferred to acclimation chambers containing 3 L dilution water (< 30 fish/fish bowl) using a small siphon hose. The siphon was handled in a very gentle and careful manner to avoid causing any physical damage to the larvae.

2.5.2.2 Acclimation of fathead minnow larvae

Before the toxicity test, fish were acclimated under lab conditions for 7 days, regularly fed with commercial larvae diet (Zeigler Larvae AP100 #LD250, Pentair

Aquatic Habitats, Apopka, FL). Given that most fathead minnow larvae stayed around the bottom of the acclimation chamber, a mixture of larvae diet and 1 ml fresh dilution water was added into each acclimation chamber so that the larvae diet microcapsules could sink slowly to the water bottom. Approximate 3/4 of the water in each acclimation chamber was renewed every 24 h using a small siphon hose. The renewal method using a fishnet was not adopted in this case because the larvae were too small and too vulnerable to netting.

2.5.3 Range-finding test

Fathead minnow larvae (12 dph) were treated with either the dilution water as the control or BMAA. Before use, the BMAA solutions were freshly diluted from 10 g/L stock solution.

After acclimation, fathead minnow larvae (12 dph) were randomly selected and transferred into test chambers using a small siphon hose. To prevent the larvae

from staying in the siphon hose, the siphon hose had a size that allowed the larvae to swim forward in one direction but hardly turn around. As a result, fathead minnow larvae were divided into three test groups, exposed to the dilution water (control) and two concentrations of BMAA (1,000 μg/L and 10,000 μg/L), respectively. Each test group had one test chamber. Each test chamber contained 500 ml test solution and 10 fathead minnow larvae (10 larvae/500 ml test solution/test chamber). The total exposure time was 96 h without feeding. 3/5 of the test solution (300 ml) in each test chamber was renewed every 24 h; A 10 ml solution sample from each test group was taken every 24 h and stored at -20 °C.

2.6 Acute zebrafish larvae test

2.6.1 Lab conditions

The temperature in growth incubators was maintained at 26 ± 1 °C with the light/dark cycle at 16 h/8 h. Both the hatching and the toxicity test chambers were 3.8

L glass fish bowls. The dilution water was commercial bottled water (Ice Mountain

100% Natural Spring Water, Nestlé, Stamford, CT) diluted by a factor of two using distilled water. Dechlorinated tap water with a high hardness of around 200 mg/L was not used in this test because zebrafish are hardness sensitive and prefers soft water with hardness less than 50 mg/L. The dilution water had a final pH of 7.3 after aeration. The pH of all other test solutions prepared using the dilution water was also adjusted to 7.3. Other lab conditions were controlled following the OECD guidelines

(OECD, 2012).

2.6.2 Preparation before exposure

2.6.2.1 Handling of zebrafish eggs

Zebrafish eggs were shipped by air in a plastic container soon after fertilization, and had an age of less than 24 hpf upon arrival. The container was put in the growth incubator for 1-2 h until it had a water temperature close to 26 °C. Then, the eggs were transferred into a hatching chamber using an opening-widened 5 ml pipette (Pipetman, Gilson, Middleton, WI).

2.6.2.2 Hatching of zebrafish eggs

Zebrafish eggs were kept in the hatching chamber for 48 h when more than 90% of the eggs hatched. No water renewal was applied during hatching. In order to start the BMAA exposure at the early life-stage of zebrafish, the larvae less than 24 h post hatch (hph) old were directly used in the toxicity test without accumulation.

2.6.3 Range-finding test

Zebrafish larvae were treated with either the dilution water as the control or

BMAA. Before use, the BMAA solutions were freshly diluted from 10 g/L stock solution.

Newly hatched zebrafish larvae (< 24 hph) were randomly selected and carefully transferred into the test chambers using a 25 ml beaker, with as little water as possible. A siphon hose seemed less applicable to transfer zebrafish larvae because they reacted and escaped much faster than fathead minnow larvae. As a result,

zebrafish larvae were divided into three test groups, exposed to the dilution water

(control) and two concentrations of BMAA (1,000 μg/L and 10,000 μg/L), respectively. Each test group had one test chamber. Each test chamber contained 500 ml test solution and 10 zebrafish larvae (10 larvae/500 ml test solution/test chamber).

The total exposure time was 96 h. No feeding was applied because zebrafish larvae were able to absorb nutrition through yolk sac till 5 dpf (Jardine and Litvak,

2003; Rubinstein, 2003); 3/5 of the test solution (300 ml) in each test chamber was renewed every 24 h using a small siphon hose; A 10 ml solution sample from each test group was taken every 24 h and stored at -20 °C.

2.7 Zebrafish embryo-larvae test

2.7.1 Lab conditions

The temperature in growth incubators was maintained at 26 ± 1 °C, with the light/dark cycle at 16 h/8 h. The fish care chambers used for acclimation and spawning were 20 L (40 cm × 20 cm × 25 cm) glass fish tanks (Aqueon #10005,

Franklin, WI). The toxicity test chambers were 3.5 ml 24-well tissue culture plates

(Falcon #353226, Corning, Corning, NY). All plates were pre-saturated with corresponding test solutions for at least 24 h to avoid unexpected absorption of

BMAA. The dilution water was Ice Mountain bottled water diluted by a factor of 2 using distilled water. The dilution water and all other test solutions had a final pH of

7.3. Other lab conditions were controlled following the OECD guidelines (OECD,

2012).

2.7.2 Preparation before exposure

2.7.2.1 Handling of zebrafish embryos

Zebrafish embryos were shipped by air in a plastic container soon after fertilization, and had an age of less than 24 hpf upon arrival. After arrival, the container was put in the growth incubator for 1-2 h until it had a water temperature

~26 °C. Due to the nature of FET, zebrafish embryos (< 24 hpf) were directly used in the rang-finding test without accumulation.

2.7.2.2 Handling of zebrafish adults

Adult zebrafish (3 males and 6 females) were shipped by air in a plastic bag.

After arrival, the bag was put in the growth incubator for 1-2 h until its water had a temperature close to 26 °C. Then, the adults were transferred into a 20 L fish tank containing 16 L dilution water. A sponge filtration-aeration system was applied in the tank to maintain a sufficient DO level and to avoid toxic ammonia accumulation. To aid bacteria growth in the sponge filter, the sponge filtration-aeration system started running three days in water before zebrafish were introduced.

2.7.2.3 Accumulation of zebrafish adults

Zebrafish males and females were acclimated together under lab conditions for

7 days, regularly fed with fish food flakes twice a day. Approximate 3/4 of the water in the tank was renewed every 24 h using a mid-size siphon hose (Aqueon #06227,

Franklin, WI).

2.7.2.4 Preparation for spawning

After the acclimation, zebrafish males were transferred using a fish net into another 20 L fish tank also containing 16 L dilution water and a well-functioning sponge filtration-aeration system. The males and females were kept separately under lab conditions for 7 days, regularly fed with fish food flakes twice a day and frozen brine shrimp (Sally’s, San Francisco Bay, Newark, CA) once a day. 1/4 of the water in the tank was renewed every 24 h using a mid-size siphon hose.

2.7.2.5 Spawning

The fish tank keeping the zebrafish males was selected as the spawning tank.

On the day before the spawning, zebrafish females were transferred into the spawning tank using a fish net 2 h prior to the onset the darkness. After being held together overnight, the males and the females started mating and spawning in 2 h after the onset of light. To prevent eggs from being consumed, the adults were confined close to the surface of water by a pre-installed horizontal nylon mesh submerged 3 cm into the water. After the onset of light, a plastic green plant was placed into the tank as spawning stimulus (DIN, 2001). The aeration-filtration speed was properly reduced to avoiding heavy perturbation of the dilution water.

2.7.2.6 Collection of eggs

During the spawning, most eggs freely passed through the mesh and fell onto the bottom of the spawning tank. 2.5 h after the onset of light, the zebrafish adults and

the mesh were removed from the spawning tank. Then, eggs were siphoned out using a small siphon hose and collected in a 1,000 ml beaker. Given that most eggs collected were spawned between 1.5 h and 2.5 h after the onset of light, 0 hpf was defined here as 2 h after the onset of light. For convenience, zebrafish prior to 96 hpf were defined as embryos and fish from 96 hpf to the end of test were defined as larvae.

Due to the nature of FET, zebrafish embryos (< 1 hpf) were directly used in the LC50 test and the behavioral assay.

2.7.3 Range-finding test

Zebrafish embryos were treated with the dilution water as a negative control

(nC) or 3,4-DCA as a positive control (pC), or BMAA. Before use, the pC solution and the BMAA solutions were freshly diluted from 4 g/L and 10 g/L stock solutions, respectively.

Immediately after arrival, zebrafish embryos (< 24 hpf) were transferred into a

120 mm Petri dish (Pyrex, Corning, Corning, NY) using an opening-widened 5 ml pipette. The embryos in the Petri dish were washed for 30 s using 50 ml dilution water.

The healthy zebrafish embryos (< 24 hpf) were randomly selected and placed onto 24-well plates using an opening-widened 200 μl pipette, with as little water as possible. As a result, zebrafish embryos were divided into four test groups, exposed to four test solutions, respectively: the dilution water (nC), 4 mg/L 3,4-DCA (pC), and two concentrations of BMAA (1 mg/L and 10 mg/L). Each test group had one 24-well

plate with 10 embryos in 10 wells. These 10 wells occupied the same locations on each plate (A1-A5 and B1-B5) and were pre-loaded with 2 ml corresponding test solution (1 embryo/2 ml test solution/well). Each plate was covered by a low-evaporation lid to attenuate the change of solution volume and concentration due to evaporation in the growth incubator.

In order to start the exposure in all test groups at the same time with minimum delay, a pre-exposure method was applied before zebrafish embryos were placed onto the 24-well plates. Right after washing, more than double embryos (< 24 hpf) required for the test were randomly selected and transferred from the 120 mm Petri dish into four 90 mm Petri dishes (> 20 embryos/Petri dish) using an opening-widened 1,000 μl pipette. Each of the four Petri dishes was pre-loaded with 20 ml fresh test solution corresponding to one of the four plates. After a careful examination under a dissecting microscope (Nikon model SMZ-1 ESD, Nikon, Japan) at 35× magnification, abnormal embryos were removed and 10 healthy embryos from each Petri dish were randomly selected and placed onto the corresponding plate.

The total exposure time was 10 days with no feeding applied. Half of the test solution (1 ml) in each well was renewed every 24 h using a 1,000 μl pipette. A 10 ml sample of test solution removed from each test group was taken every 24 h and stored at -20 °C.

All zebrafish were regularly observed under a dissecting microscope. Dead zebrafish were removed from the plates as soon as observed. Mortality of a zebrafish embryo was determined by coagulation, no tail detachment, no somite formation or no

heart beat.

2.7.4 10-day LC50 test

In the 10-day LC50 test, zebrafish embryos were treated with the dilution water as a negative control (nC) or NaHCO3 as a solvent control (sC), or BMAA + NaHCO3.

Before use, the sC solution was freshly diluted by the dilution water from 20 M

NaHCO3 stock solution and the BMAA solutions were diluted by the fresh sC solution from 10 g/L stock solution.

Newly collected zebrafish eggs (< 1 hpf) were transferred into a 120 mm Petri dish using an opening-widened 5 ml pipette. The eggs in the Petri dish were washed for 30 s using 50 ml dilution water.

At 3 hpf, the healthy zebrafish embryos were randomly selected and placed onto 24-well plates using an opening-widened 200 μl pipette. As a result, zebrafish embryos were divided into seven test groups, exposed to seven test solutions, respectively: the dilution water (nC), 20 mM NaHCO3 (sC), and five concentrations of BMAA (1 mg/L, 3 mg/L, 10 mg/L, 30 mg/L and 100 mg/L). Each test group had one 24-well plate with 20 embryos in 20 wells. These 20 wells on each plate occupied the same locations (A1-A5, B1-B5, C1-C5 and D1-D5) and were pre-loaded with 1 ml corresponding test solution (1 embryo/1 ml test solution/well). Additional 4 embryos were placed in the remaining 4 wells (A6, B6, C6 and D6) on each plate pre-loaded with 1 ml dilution water (1 embryo/1 ml dilution water/well) as an internal plate control (iC). In the end, all test groups including iC were distributed on plates as

shown in Figure 2.1. Each plate was covered by a low-evaporation lid to attenuate the change of solution volume and concentration due to evaporation in the growth incubator.

In order to start the exposure in all test groups at the same time with minimum delay, a pre-exposure method similar to that described in the range-finding test was applied. Briefly, washed zebrafish eggs (< 1 hpf) were randomly selected and transferred into seven 90 mm Petri dishes (> 100 eggs/Petri dish for nC and > 40 eggs/Petri dish for other groups) pre-loaded with 20 ml respective test solutions. At 3 hpf, unfertilized eggs were removed and 20 healthy embryos from each Petri dish as well as 4 embryos (iC) from the Petri dish for nC were randomly selected and placed onto the corresponding plate.

The total exposure time was 10 days without feeding. Half of the test solution

(500 μl) in each well was renewed every 24 h using a 1,000 μl pipette. 10 ml sample of test solution removed from each test group was taken every 24 h and stored at

-20 °C.

All zebrafish were regularly observed under a dissecting microscope. Dead zebrafish were removed from the plates as soon as observed. Abnormal growth and the mortality rate of zebrafish in each test group were recorded every 24 h. Hatching time was measured starting at 48 hpf until all embryos hatched. All embryos were observed every 3 h under the microscope and the hatching time of each embryo was recorded. Heart rate was measured at 72 hpf, 144 hpf and 10 dpf. 10 embryos from each test group were randomly selected and individually observed under the

microscope. The heart rate of each embryo was calculated by converting the time taken for 30 beats to beats per minute (bpm). Body length was measured at 144 hpf and 10 dpf. 10 embryos from each test group were randomly selected and the body length of each embryo was individually measured with a ruler under the dissecting microscope. Sub-lethal endpoints including anomalies in eyes, pigmentation, blood circulation, pectoral fins, swimming bladder, yolk sac and pericardia were also recorded. For convenience, zebrafish larvae older than 96 hpf were individually anesthetized by 10 ml 60 mg/L MS222 in a 55 mm Petri dish before observation.

After observation, each larva was allowed to recover in 100 ml fresh dilution water for 5 s and then transferred back to its corresponding well.

2.7.5 Zebrafish embryo-larvae behavioral assay

In the behavioral assay, zebrafish embryo-larvae were treated with the dilution water as a negative control (nC) or NaHCO3 as a solvent control (sC), or BMAA +

NaHCO3. All test solutions were freshly prepared before use from stock solutions with the same method described in the 10-day LC50 test.

Newly collected zebrafish eggs (< 1 hpf) were transferred into a 120 mm Petri dish using an opening-widened 5 ml pipette. The eggs in the Petri dish were washed for 30 s using 50 ml dilution water.

the dilution water (nC), 20 mM NaHCO3 (sC), and four concentrations of BMAA (0.1 mg/L, 1 mg/L, 10 mg/L and 100 mg/L). Each test group had one 24-well plate with 20 embryos in 20 wells which occupied the same locations (A1-A5, B1-B5, C1-C5 and

D1-D5) on each plate and were pre-loaded with 1 ml corresponding test solution (1 embryo/1 ml test solution/well). Each plate was covered by a low-evaporation lid to attenuate the evaporation in the growth incubator.

In order to start the exposure in all test groups at the same time with minimum delay, a pre-exposure method similar to that described in the 10-day LC50 test was applied. Briefly, washed zebrafish eggs (< 1 hpf) were randomly selected and transferred into six 90 mm Petri dishes (> 40 eggs/Petri dish) pre-loaded with 20 ml respective test solutions. At 3 hpf, unfertilized eggs were removed and 20 healthy embryos from each Petri dish were randomly selected and placed onto the corresponding plate.

The total exposure time was 10 days and no feeding was applied during the test. Half of the test solution (500 μl) in each well was renewed every 24 h with the same renewal method adopted in the 10-day LC50 test. 10 ml sample of test solution removed from each test group was taken every 24 h and stored at -20 °C.

All zebrafish embryos were regularly observed under a dissecting microscope.

Dead zebrafish were removed from the plates as soon as observed. Only normally developed zebrafish were used in the behavior assay.

2.7.5.1 Spontaneous contractions

The first locomotor behavior of zebrafish embryos is spontaneous contractions, a series of slow and alternating tail coils occurring at 17-18 hpf (Colwill and Creton,

2011). After reaching the peak of frequency around 21 hpf, spontaneous contractions gradually slow down and almost disappears after 27 hpf (Saint-Amant and Drapeau,

1998).

Starting at 16 hpf, embryos were observed under a dissecting microscope until all embryos showed spontaneous contractions. The observation was done every hour and the onset time of spontaneous contractions for each embryo was recorded. Onset of spontaneous contractions for an embryo was determined here as occurrence of at least 1 contraction during an observation time of 20 s.

At 22 hpf, all embryos from each test group were placed in a 55 mm Petri dish containing 20 ml fresh dilution water using an opening-widened 200 μl pipette. After

2 min acclimation, the movements of embryos in 3 min were recorded by a digital camera (PowerShot SX260 HS, Canon, Japan). The frequency of spontaneous contractions for each embryo was calculated by converting the time taken for 20 contractions to contractions per minute (cpm). After recording, all embryos from each test group were transferred back onto a new 24-well plate pre-loaded with 1 ml corresponding fresh test solution and continued the exposure.

2.7.5.2 Touch-evoked contractions

Stereotypically, zebrafish embryos show response to mechanical stimuli

around 21 hpf, right after the peak of spontaneous contractions (Downes and Granato,

2006). Touching the head, tail or yolk sac of an embryo triggers 1-3 in situ body contractions each containing a rapid tail coil and a following slower relaxation

(Saint-Amant and Drapeau, 1998).

In order to measure the touch-evoked contractions of zebrafish embryos more easily, all embryos were dechorionated with 2 pairs of very fine forceps at 24 hpf. At

27 hpf, 10 embryos from each test group were randomly selected and individually placed at the center of a 55 mm Petri dish containing 20 ml fresh dilution water using an opening-widened 200 μl pipette. After 2 min acclimation, each embryo was stimulated by continually touching the head using a soft painting brush hair. The total contractions of the embryo in response to 10 touches with at least 5 s intervals were counted. Although the spontaneous contractions of zebrafish embryos had a very low frequency at 27 hpf (McKeown et al., 2009), the timing of each touch was carefully selected to avoid miscounting due to a high similarity between the two behaviors.

After observation, each embryo was transferred back to the corresponding well and continued the exposure.

2.7.5.3 Touch-evoked escape

By the end of 2 dpf, dechorionated zebrafish embryos still rested most of time with little movement. However, they were able to show escape response to mechanical stimuli via high frequency tail contractions. Normally, an embryo touched on the head rapidly re-orientates its body direction and swims away from the stimulus

(Ahmad et al., 2012).

At 48 hpf, 10 embryos from each test group were randomly selected and individually placed at the center of a 120 mm Petri dish containing 20 ml fresh dilution water using an opening-widened 200 μl pipette. The Petri dish was placed over an A4 size grid paper covered with 4.5 mm squares. After 2 min acclimation, each embryo was stimulated by continually touching the head using a painting brush hair. The incidence of the embryo showing escape behaviors in response to 10 touches with at least 5 s intervals was counted. The distance that the embryo swam during each escape was also measured. The escape distance was estimated from the number of squares away from the touching location multiplied by the width of a square (4.5 mm). To ensure an enough swimming space, the embryo was re-placed to the center of the Petri dish after each escape. After observation, each embryo was transferred back to the corresponding well and continued the exposure.

2.7.5.4 Free swimming

After 4 dpf, zebrafish larvae gradually started spontaneous free swimming in a pattern of “beat and glide” after inflation of the swim bladder (Airhart et al., 2007;

Buss and Drapeau, 2001). At 120 hpf, most larvae were able to free swimming, although still in a relatively slow and low-frequency manner.

At 144 hpf, 12 zebrafish larvae from each test group were randomly selected and individually placed in a 55 mm Petri dish containing 10 ml fresh dilution water using an opening-widened 1,000 μl pipette. The Petri dish was placed over an A4 size

grid paper covered with 4.5 mm squares. After 5 min acclimation, the swimming activity of each larva in 3 min was recorded by a digital camera. The total swimming distance, the time spent on swimming, the swimming speed, and the incidences of freezing bouts and erratic swimming of the larva during the recording were measured.

Total swimming distance was estimated from the number of squares that the larva swan multiplied by 4.5 mm. Swimming speed was calculated from the total swimming distance divided by the time spent on swimming. A freezing bout was defined here as an entire absence of spontaneous body motion for at least 2 s between two swimming movements. Erratic swimming was defined here as a sudden transverse shift of body position or a rapid change of swimming velocity (direction and speed). The free swimming recording was performed in group of 6 larvae from the same test group in 6 respective Petri dishes. After recording, each larva was transferred back into the corresponding well and continued the exposure.

2.7.5.5 Startle response

Startle response of animals is an unconditioned behavior which reflects the integration of sensory and motor functions (Koch, 1999). A startle response, usually representing as rapid contractions of body muscles, can be triggered by visual, acoustic or tactile stimuli (Bang et al., 2002; J. Best et al., 2010; J. D. Best et al.,

2008). Zebrafish larvae showed no startle response until 5 dpf, when taping the plates caused a sudden acceleration of swimming speed.

At 7 dpf, 10 zebrafish larvae from each test group were randomly selected and

placed together in a 55 mm Petri dish containing 20 ml fresh dilution water using an opening-widened 1,000 μl pipette. The Petri dish was placed over an A4 size grid paper covered with 4.5 mm squares. After 5 min acclimation, the swimming activity of larvae in 1.5 min was recorded by a digital camera as the baselines. Subsequently, larvae were startled by continually tapping the rim of Petri dish using the head of a falling 200 μl pipette tip. The tail of pipette tip was impaled by a horizontal thumbtack fixed on top of a flat base, so that the pipette tip could freely rotate in the vertical plane. The responses of larvae to 20 taps with 5 s intervals were also recorded by camera. The number of squares that each larva swam in 2 s after each tap was counted. To ensure the strength consistency of each tap, the pipette tip was dropped by the observer from the same horizontal angle of 45°. After recording, all larvae from each test group were transferred back onto a new 24-well plate pre-loaded with 1 ml corresponding fresh test solution and continued the exposure.

2.7.5.6 Anxiety

Anxiety of animals can be caused by a number of factors, such as predation, environmental changes and drug exposure (Blaser and Gerlai, 2006; Egan et al., 2009;

Wong et al., 2010). Anxiety of zebrafish can be reliably evaluated by measuring thigmotaxis (Peitsaro et al., 2003; Richendrfer et al., 2012; Schnörr et al., 2012).

Thigmotaxis describes a “centrophobic” behavior in which animals strongly prefer staying or moving in close to the wall rather than the center of an apparatus (C.

Maximino et al., 2010a).

The zebrafish larvae used in the anxiety test were from the same batch of collected eggs as those used in other behavioral tests, but were reared in different apparatus. Right after the exposure started at 3 hpf, the remaining washed eggs in the

120 mm Petri dish were carefully examined under a dissecting microscope.

Unfertilized eggs were removed and healthy embryos were transferred to a new 120 mm Petri dish containing 40 ml fresh dilution water using an opening-widened 200 μl pipette. 3/4 of water (30 ml) in the Petri dish was renewed every 24 h using a 5 ml pipette and no feeding was applied before the anxiety test. Unhealthy and dead zebrafish were removed from the plates as soon as observed.

At 7 dpf, 42 zebrafish larvae were randomly selected and divided into six test groups (7 larvae/ groups), exposed to six test solutions, respectively: the dilution water (nC), 20 mM NaHCO3 (sC) and four concentrations of BMAA (0.1 mg/L, 1 mg/L, 10 mg/L and 100 mg/L). Larvae from each test group were individually placed in a 55 mm Petri dish containing 10 ml corresponding fresh test solution using an opening-widened 1,000 μl pipette. A 38 mm diameter circle was marked on the bottom of Petri dish to divide it into the inner zone and the outer zone (area ratio 1:1).

The Petri dish was placed over an A4 size grid paper covered with 4.5 mm squares.

After 10 min exposure, the swimming activity of larvae in 5 min was recorded by a digital camera. The swimming performance of each larva was evaluated with the same method adopted in the free swimming test. In addition, thigmotaxis of each larva in distance and in time was also measured during the recording. Thigmotaxis in distance and thigmotaxis in time were defined here as the percentage (%) of the number of

squares and the percentage (%) of the time, respectively, that the larva swam in the outer zone. The recording was performed in group of 6 larvae from 6 test groups in 6 respective Petri dishes. After recording, larvae were removed from their test groups to avoid recounting.

2.8 Statistical analysis

The lethality of BMAA in zebrafish embryo-larvae was analyzed using the US

EPA Probit Analysis Program_1.5 (Environmental Monitoring Systems Laboratory,

OH). The 10-day LC50 and its 95% confidence interval were both calculated.

Developmental and behavioral endpoints analyses were carried out using

Excel_2010 (Microsoft, Bellevue, WA) and JMP_10.0.2 (SAS, Cary, NC). For each endpoint, data were input into Excel to calculate mean of observations and the standard error of the mean (SEM) for each test group. One-way analysis of variance (ANOVA) and post hoc Dunnett’s test (*p < 0.05; **p < 0.01; ***p < 0.001) were performed in JMP to determine significant differences between control and other test groups. All data are presented as mean ± SEM.

CHAPTER THREE: RESULTS AND DISCUSSION

3.1 Results

3.1.1 Acute toxicity of BMAA to fathead minnow and larval zebrafish

Short term (96-hr) BMAA exposure (up to 10,000 μg/L) neither influenced mortality nor caused obvious adverse effects in juvenile and larval fathead minnow

(Pimephales promelas) or larval zebrafish (Danio rerio). Because none of the three test organisms died during respective range-finding tests, final LC50 tests were not conducted.

3.1.2 Acute toxic effects of BMAA to zebrafish embryo-larvae

3.1.2.1 10-day LC50

Although no lethality was observed in the range-finding test, one larva exposed to 10 mg/L BMAA exhibited weakness, abnormal spine, and entire body albinism and edema. Therefore, a final 10-day LC50 test was conducted using BMAA with higher concentrations (1-100 mg/L). The result confirmed that BMAA had little influence on mortality of zebrafish embryos. By the end of 10-day LC50 test, no embryos died in nC group; one embryo died in sC, the 1 mg/L and 100 mg/L BMAA groups; 2 embryos died in the 10 mg/L and 30 mg/L BMAA groups; 3 embryos died in the 3 mg/L BMAA group. Because only one iC embryo died in the 100 mg/L

BMAA group, all test groups were accepted and used for LC50 calculation (OECD,

2012).

3.1.2.2 Hatching time

Embryos started hatching after 48 hpf and most of them hatched by 66 hpf.

BMAA had no significant influence on the average hatching time of zebrafish embryos. The hatching seemed to be slightly accelerated in the BMAA groups which had a higher average hatching rate compared to both control groups at all observation points before reaching 100%. The maximum difference (30%) between the BMAA groups and nC occurred at 57 hpf, when the hatching rates of them were 75% and

45%, respectively. However, the acceleration was not obvious between the BMAA groups and sC, with the maximum hatching rate difference of only 8% at 63 hpf

(Figure 3.1).

3.1.2.3 Heart rate

Zebrafish embryos exposed to BMAA showed a significantly accelerated heart rate compared to sC at 72 hpf (Figure 3.2A). The heart rate of embryos from 30 mg/L

BMAA and 100 mg/L groups were 18% (***p < 0.001; n = 10) and 23% (***p <

0.001; n = 10) faster than sC (197.28 ± 4.10 bpm), respectively.

Interestingly, BMAA exposure significantly reduced the heart rate of embryos from all five BMAA groups at 144 hpf (Figure 3.2B). The reductions in heart rate of embryos compared to sC (228.37 ± 3.47 bpm) were: 10% (**p < 0.01; n = 10) for the

1 mg/L BMAA group, 9% (**p < 0.01; n = 10) for the 3 mg/L BMAA group, 10%

(**p < 0.01; n = 10) for the 10 mg/L BMAA group, 11% (***p < 0.001; n = 10) for the 30 mg/L BMAA group, and 20% (***p < 0.001; n = 10) for the 100 mg/L BMAA group.

At 10 dpf, BMAA had no significant effects on heart rate of embryos exposed to 10 mg/L and 100 mg/L BMAA. However, heart rates of embryos from the 1 mg/L,

3 mg/L and 100 mg/L BMAA groups were significantly slower than sC (170.57 ±

1.70 bpm), with reductions of 14% (***p < 0.001; n = 10), 12% (***p < 0.001; n =

10) and 8% (*p < 0.05; n = 10), respectively (Figure 3.2C).

No significant differences were observed between nC and sC groups at all observation points (Figure 3.2).

3.1.2.4 Body length

Zebrafish larvae had little difference in body length between the BMAA groups and controls (Figure 3.3). The average body length of larvae for all test groups

(n = 7) was 4.21 ± 0.01 cm at 144 hpf and 4.11 ± 0.01 cm at 10 dpf.

3.1.2 Behavioral effects of BMAA to zebrafish embryo-larvae

For behavioral endpoints monitored during the embryo life stage, there were no significant differences between zebrafish from nC and sC. However, the two control groups gradually presented significant differences at the larvae life stage (data not shown). Given that nC was not the direct control, only sC was used as the control group in all ANOVA and post hoc Dunnett’s tests.

3.1.2.1 Spontaneous contractions

Spontaneous contractions first occurred at 17 hpf, and all embryos showed movements by 21 hpf. Exposure to BMAA caused an early onset of spontaneous contractions (Figure 3.4A). Embryos from the 100 mg/L BMAA group started to show spontaneous contractions at 19.00 ± 0.16 hpf, nearly 1 h earlier (**p < 0.01; n =

20) than sC (19.88 ± 0.15 hpf).

At 22 hpf, the average spontaneous contraction rate of zebrafish embryos was

9.23 ± 0.37 cpm for all test groups (n = 6; except nC). Embryos from the BMAA groups had no significant difference in the frequency of spontaneous contractions compared to sC (Figure 3.4B).

3.1.2.2 Touch-evoked contractions

By 27 hpf, all zebrafish embryos were able to exhibit at least one contraction in response to 10 continual touches on the head. The responses of the embryos were almost the same between nC (4.00 ± 0.93 contractions/10 touches) and sC (3.90 ±

0.75 contractions/10 touches). Generally, BMAA enhanced the touch-evoked contractions of embryos (Figure 3.5). Approximately twice (*p < 0.05; n = 10) as many contractions as sC were observed in the 10 mg/L BMAA group (7.90 ± 1.28 contractions/10 touches). Although the difference was not significant, touch-evoked contractions of embryos in the 0.1 mg/L BMAA group (6.50 ± 1.24 contractions/10 touches) was 67% higher than sC. Additionally, embryos from the 0.1 mg/L and 100 mg/L BMAA groups responded 28% and 41% more times than controls, respectively,

without significant differences.

3.1.2.3 Touch-evoked escape

BMAA exposure affected the touch-evoked escaping behavior of zebrafish embryos (Figure 3.6). Embryos from all BMAA groups exhibited an approximately

30-50% lower incidences of touch-evoked escape movements compared to sC (5.40 ±

0.85 escapes/12 touches) (Figure 3.6A). Especially, this reduction was significant in embryos from the 10 mg/L BMAA group (*p < 0.05; n = 10) and the 100 mg/L

BMAA groups (**p < 0.01; n = 10), with escape incidences of 2.80 ± 0.47 and 2.67 ±

0.58 escapes/12 touches, respectively.

There were no significant differences between the BMAA groups and sC in the embryos’ average swimming distance during each escape. However, the average escape distances of embryos from four BMAA groups were all approximately 40% shorter than sC (2.15 ± 0.38 cm) (Figure 3.6B).

3.1.2.4 Free swimming

BMAA exposure resulted in obvious changes in the free swimming activity of zebrafish larvae at 144 hpf (Figure 3.7). The total swimming distance of larvae from the 1 mg/L BMAA group in 3 min (20.96 ± 4.18 cm) and their swimming speed during movements (7.58 ± 1.40 cm/min) were both significantly lower (*p < 0.05 for total distance and **p < 0.01 for speed; n = 12) than sC (total distance: 39.86 ± 6.07 cm; speed: 14.97 ± 1.85 cm/min). Similarly, the swimming performance of larvae

from the 100 mg/L BMAA group (total distance: 12.44 ± 2.80 cm; speed: 4.63 ± 0.91 cm/min) was also significantly reduced (**p < 0.01 for total distance and ***p <

0.001 for speed; n = 12) (Figure 3.7A). In addition, larvae from the 10 mg/L BMAA group had a significant reduction (*p < 0.05; n = 12) in swimming speed (9.09 ± 1.20 cm/min) and a non-significant 38% reduction in total swimming distance (24.86 ±

4.39 cm) in comparison with sC (Figure 3.7B). It was notable that the larvae’s swimming activity of the 0.1 mg/L BMAA group seemed to be enhanced in spite of no statistical significance. Larvae from this group swam 36% longer in distance

(54.34 ± 6.23 cm) and 25% faster in speed (18.48 ± 2.04 cm/min) compared to sC

(Figures 3.7A-B).

During the 3 min swimming, larvae from sC had 2.18 ± 0.63 freezing points,

28.00 ± 9.87 s freezing time and 2.33 ± 0.47 erratic movements. Larvae from the

BMAA groups did not significantly differ from each other or sC in performances at these three endpoints. However, larvae showed a dose dependent increase from 0.82 ±

0.30 to 4.58 ± 1.52 in the incidence of freezing points among BMAA groups (Figure

3.7C). It was also noteworthy that larvae from the 0.1 mg/L BMAA group spent only

3.18 ± 1.48 s in freezing which was much lower than other BMAA groups and sC

(Figure 3.7D). Besides, larvae from the 100 mg/L BMAA group had 55-75% less erratic movements (0.75 ± 0.30) in comparison with other test groups (Figure 3.7E).

3.1.2.5 Startle response

BMAA reduced the startle response of zebrafish larvae at 7 dpf (Figure 3.8).

The average distance that larvae from sC swam in 2 s in response to 20 taps was 0.91

± 0.10 cm. In contrast, the startle responses of larvae from the 10 mg/L BMAA group

(0.66 ± 0.05 cm/startle) and the 100 mg/L BMAA group (0.71 ± 0.03 cm/startle) were both significantly reduced (*p < 0.05, n = 20) (Figure 3.8E). For each BMAA group, the swimming distances of larvae in 2 s after each tap are shown in Figures 3.8A-D and the curve of 20 responses was compared with sC. No taps generated a significantly longer swimming distance of larvae for all BMAA groups. On the other hand, 10-18 out of 20 taps resulted in reduced movements for larvae from the BMAA groups, in which 1-2 reductions were significantly different. A general trend of decreasing swimming distances along continual taps was found in all test groups (n =

6; except nC) by observing a negative slope of the linear regression line across 20 responses, indicating larvae’s habituation to startles (Figures 3.8A-D). The startle habituation of each larva was defined here as a process that its responses declined close to the baseline (normally 0-0.5 cm/2 s). The response curve of each larva was first smoothed with a moving average of three consecutive values. Then, the habituation process was determined as the series of responses before reaching the bottom of the moving average curve. No significant differences existed for larvae from different test groups in the number of taps required for startle habituation

(Figure 3.8F).

3.1.2.6 Anxiety

10 min exposure to 0.1-100 mg/L BMAA did not significantly influence the

swimming activity and thigmotaxis of larvae at the age of 7 dpf (Figure 3.9) The longest swimming distance (25.33 ± 6.01 cm) and the highest swimming speed (9.67

± 1.55 cm/min) were both observed in larvae exposed to 0.1 mg/L BMAA (Figures

3.9C-D). Larvae exposed to 20 mM NaHCO3 in dilution water (sC) swam 66.98 ±

13.90% of distance and spent 69.29 ± 13.72% of time in the outer zone of Petri dish.

Larvae exposed to 1 mg/L BMAA exhibited the highest thigmotaxis in both distance

(95.56 ± 2.91%) and time (96.11 ± 2.46%) (Figures 3.9A-B).

3.2 Discussion

The 96 h toxicity tests revealed that BMAA was not acutely lethal to fathead minnow (juveniles: 96 h LC50 > 10,000 μg/L; larvae: 96 h LC50 > 10,000 μg/L) or zebrafish (larvae: 96 h LC50 > 10,000 μg/L). The 10-day zebrafish embryo-larvae toxicity test further proves the low lethality of BMAA by showing a 10-day LC50 >

100 mg/L. The zebrafish embryo-larvae test also showed that BMAA non-significantly accelerated the hatching of embryos. BMAA did significantly affect heart rate but had little effect in the body length of zebrafish larvae.

The high 10-day LC50 (> 100 mg/L) for BMAA found in this study is contrary to previous work reporting a 10-day LC50 of BMAA as low as 10 μg/L in the same zebrafish embryo model (Purdie et al., 2009a). Purdie and colleagues in their investigation used different dilution water (dechlorinated water: hardness 10.6 mg/L, alkalinity 31.3 mg/L and pH 8.2) and different bicarbonate donor (Na2CO3) compared to this study (Purdie et al., 2009b). However, the huge difference (> 10,000 times)

between two LC50 values acquired is still surprising. Besides, the onset time of

BMAA exposure is earlier in this study (< 1 hpf) than the previous work (24 hpf), which should normally result in a higher lethality in zebrafish embryos (T.

Braunbeck and Lammer, 2006). Purdie et al. (2009b) also reported a reduction in heart rate of embryos at 72 hpf. However, several other sub-lethal effects reported by them, including pericardial edema, abnormal spinal axis formation, clonus-like convulsions, and significant premature point of hatch, were not observed in this study.

Despite a low acute lethality in zebrafish, BMAA influenced embryo-larvae behaviors, especially motor function related behaviors. Spontaneous contractions are the first behavior of zebrafish embryos during their development. This movement is driven by simple electrical coupling of motor neurons in the spinal cord, independent of supraspinal inputs (Saint-Amant and Drapeau, 1998). That BMAA caused an earlier onset of spontaneous contractions, indicates a BMAA-induced earlier development of motor nervous system in zebrafish embryos. The frequency of embryonic spontaneous contractions has been shown to be affected by toxicants such as perfluorooctanesulphonicacid (PFOS) and chlorpyrifos (Huang et al., 2010;

Selderslaghs et al., 2010). In contrast, BMAA as a neurotoxin had little influence on the frequency of spontaneous contractions at 22 hpf. However, at 27 hpf, BMAA showed a significant enhancement in the embryos’ touch-evoked contractions. Unlike the spontaneous movement stage when the formation of glutamate receptors just starts

(Cox et al., 2005; Hoppmann et al., 2008), touch response of zebrafish embryos appears only after the glutamatergic synapses gradually take over the

neurotransmission between spinal motor network and sensory neurons as well as interneurons (Pietri et al., 2009). A significant amount of evidence supports the hypotheiss that BMAA is an agonist for many glutamate receptors such as NMDA,

AMPA and mGluR5 (Chiu et al., 2012; Goto et al., 2012; Liu et al., 2009; Lobner et al., 2007). This indicates that the increase in touch response might result from the

BMAA-induced hyperactivity of motor network. The involvement of glutamate receptors in the neurotoxicity of BMAA might also explain why touch-evoked contractions but not spontaneous contractions were influenced by BMAA.

The adverse behavioral effect of BMAA was first observed in touch-evoked swimming activity of zebrafish embryos at 48 hpf. The significantly fewer responses and shorter (though not significant) escaping distances indicated that BMAA caused impairment of locomotor functions in embryos. Declined locomotor activity was also found in the free swimming of larvae at 144 hpf. The reduction in swimming distance and speed of larvae exposed to 1-100 mg/L BMAA agrees with the increase in the incidence of freezing points and the freezing time. Although larvae from higher

BMAA concentration groups exhibited fewer erratic movements, it did not mean that they swam better in this case. This is because if a larva swims slowly all the time, its body might actually not be strong enough to perform continuous fast “beat and glide” movements. Thus, a lack of erratic swimming might refer to the weakness of motor function. It should be noted that larvae exposed to BMAA at the lower concentration

(0.1 mg/L) seemed to have a higher (though not significant) swimming activity compared to sC. Interestingly, in the anxiety test, the longest swimming distance and

the fastest speed (though not significant) also belonged to larvae exposed to 0.1 mg/L

BMAA. The pattern that BMAA enhances the swimming activity at a lower concentration but depresses the locomotion at a higher concentration is similar to what has been found in zebrafish treated with alcohol (Gerlai et al., 2000).

The NMDA glutamatergic mediation appears to play an important role in the startle response of zebrafish (Caio Maximino et al., 2010b; Roberts et al., 2011). A previous study found that the NMDA receptor antagonist memantine increases the startle response (J. D. Best, et al., 2008). The reduction in startle response of larvae exposed to the NMDA receptor agonist BMAA is supported by previous results, consistent with a neurotoxic effect of BMAA exposure on zebrafish. No obvious change in the startle habituation of larvae indicates that the sensorimotor dysfunction rather than a faster habituation was mainly responsible for the reduction in the startle response.

Due to the addition of 20 mM NaHCO3 as the bicarbonate donor for BMAA in the zebrafish embryo-larvae test, a NaHCO3 control group (sC) was necessary besides nC. However, from 48 hpf, embryos from the two control groups began to present obvious differences in locomotor activity. This suggests that the additional 20 mM

NaHCO3 accelerates the development of embryos. Because no feeding was applied during the exposure, all nutrition acquired by embryo-larvae were via yolk sac absorption (Jardine and Litvak, 2003; Rubinstein, 2003). Therefore, the elevation of larvae’s locomotor activity in sC compared to nC might be attributed to sufficient minerals in test solution.

3.2.1 Conclusions

(1) BMAA had a 96 h LC50 of over 10,000 μg/L to three test organisms: 1

month juvenile fathead minnow, 12 dph larval fathead minnow and 72 hpf

larval zebrafish.

(2) BMAA showed a 10-day LC50 of higher than 100 mg/L to embryonic

zebrafish (< 1 hpf).

(3) BMAA exposure accelerated the heart rate of zebrafish embryos at 72 hpf,

but reduced it at 144 hpf and 10 dpf.

(4) BMAA exposure did not influence the body length of zebrafish larvae.

(5) BMAA exposure induced an early onset of spontaneous contractions in

zebrafish embryos.

(6) BMAA exposure enhanced touch-evoked contractions of zebrafish

embryos at 27 hpf.

(7) BMAA depressed several locomotor activities of zebrafish embryo-larvae

from 48 hpf to 7 dpf, including touch-evoked escape, free swimming and

startle response.

(8) BMAA did not cause anxiety in 7 dpf larval zebrafish.

3.2.2 Importance and future work

This is to our knowledge the first investigation into the behavioral effects of

BMAA on fish. This study links motor dysfunctions in zebrafish embryo-larvae with

BMAA, providing extra evidence of its neurotoxicity. The zebrafish embryo-larvae

model seems to be an ideal model of BMAA research because it provides rapid and diverse behavioral endpoints for neurotoxicity and neurodegeneration assessments (de

Esch et al., 2012; Tierney, 2011). More investigations into the neurotoxicity of BMAA using this model are guaranteed in the future. This is also the first to study the acute toxicity of BMAA on fathead minnow. Although no behavioral endpoints were monitored in fathead minnow, its locomotor performance might also be influenced by

BMAA. The finding that BMAA depresses touch-evoked escape, free swimming and startle response of fish is also of ecological importance, because these are all related to feeding and predator-related behaviors. Fish exposed to BMAA might have greater difficulty with respect to prey capture and predator avoidance. Due to the nature of an acute toxicity test, the BMAA concentrations selected in this study are much higher than the level detected in real aquatic environment (Al-Sammak et al., 2014; Faassen,

2014). However, given the potential ubiquity of BMAA and its ability to bioaccumulate (Esterhuizen and Downing, 2008; Murch et al., 2004a), even BMAA at low concentrations in aquatic ecosystems might result in chronic toxic effects on fish and other aquatic organism. Future study is required to understand the fate of BMAA in aquatic ecosystems and its chronic effects on different aquatic organisms.

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Figure 1.1. Chemical structure of BMAA.

Figure 1.2 Comparison of the structure of L-BMAA-carbamate and L-glutamate

Figure 2.1. Distribution of test groups on 24-well plates in zebrafish embryo-larvae

10-day LC50 test. 1 reperents 1 mg/L BMAA; 2 reperents 3 mg/L BMAA; 3 reperents 10 mg/L BMAA; 4 reperents 30 mg/L BMAA; 5 reperents 100 mg/L BMAA; nC reperents negative control (dilution water); sC reperents solvent control

(20 mM NaHCO3); and iC reperents internal plate control (dilution water).

54 Hatching Hatching time(hpf)

51 nC sC 1 3 10 30 100 BMAA concentration (mg/L)

Figure 3.1. Hatching time of embryos exposed to nC (dilution water), sC (20 mM

NaHCO3) or BMAA (1-100 mg/L) in sC. Bars represent the average value ± SEM per test group (n = 12).

A B 260 *** 240 +++ *** ** ** +++ ** *** 240 220 ++ + ++ ++

220 200 *** +++

Heart rate (bpm) Heartrate 200 180 Heart rate (bpm) Heartrate

180 160 nC sC 1 3 10 30 100 nC sC 1 3 10 30 100 BMAA concentration (mg/L) BMAA concentration (mg/L)

C 180

* *** + 160 *** +++ +++

140 Heart rate (bpm) Heartrate

120 nC sC 1 3 10 30 100 BMAA concentration (mg/L)

Figure 3.2. Heart rate of embryos exposed to nC (dilution water), sC (20 mM

NaHCO3) or BMAA (1-100 mg/L) in sC at (A) 72 hpf, (B) 144 hpf, and (C) 10 dpf. Bars represent the average value ± SEM per test group (+p < 0.05, ++p < 0.01, +++p < 0.001 vs nC; *p < 0.05, **p < 0.01, ***p < 0.001 vs sC; n = 10).

A B

4.4 4.3

4.3 4.2

4.2 4.1

4.1 4 Body length (cm) length Body 4.0 (cm) length Body 3.9

3.9 3.8 nC sC 1 3 10 30 100 nC sC 1 3 10 30 100 BMAA concentration (mg/L) BMAA concentration (mg/L)

Figure 3.3. Body length of embryos exposed to nC (dilution water), sC (20 mM

NaHCO3) or BMAA (1-100 mg/L) in sC at (A) 144 hpf, and (B) 10 dpf. Bars represent the average value ± SEM per test group (n = 10).

A 21 B 12

20 10

** 8 19 6

18 (cpm) rate

contractions (hpf) contractions 4 Onset time of spontanous spontanous of Onsettime 17 contraction Spontaneous 2 sC 0.1 1 10 100 sC 0.1 1 10 100 BMAA concentration (mg/L) BMAA concentration (mg/L) Figure 3.4. (A) Onset time and (B) frequency of spontaneous contractions at 22 hpf for embryos exposed to sC (20 mM NaHCO3) or BMAA (1-100 mg/L) in sC. Bars represent the average value ± SEM per test group (**p < 0.01 vs sC; n = 10).

10 * 8

2 Contractions/10touches

0 sC 0.1 1 10 100 BMAA concentration (mg/L)

Figure 3.5. Incidence of contractions in response to 10 touches for embryos exposed to sC (20 mM NaHCO3) or BMAA (1-100 mg/L) in sC at 27 hpf. Bars represent the average value ± SEM per test group (*p < 0.05 vs sC; n = 10).

A B

7 2.5

6 2.0 5 4 1.5 * ** 3 1.0 2 0.5

Escapes/12 touches Escapes/12 1 Escapedistance (cm) 0 0.0 sC 0.1 1 10 100 sC 0.1 1 10 100 BMAA concentration (mg/L) BMAA concentration (mg/L) Figure 3.6. (A) Incidence of escapes in response to 12 touches and (B) swimming distance during each escape for embryos exposed to sC (20 mM NaHCO3) or BMAA (1-100 mg/L) in sC at 48 hpf. Bars represent the average value ± SEM per test group (*p < 0.05, **p < 0.01 vs sC; n = 10).

100

A B

70 25

50 40 15 * ** *

30 10 (cm/min)

distance (cm) distance 20 ** *** Total swimming Total Swimming speedSwimming 5 10 0 0 sC 0.1 1 10 100 sC 0.1 1 10 100 BMAA concentration (mg/L) BMAA concentration (mg/L)

C D 7 50

40 5 4 30

3 20 2 Freezing points Freezing 10 1 (s) in freezing Time 0 0 sC 0.1 1 10 100 sC 0.1 1 10 100 BMAA concentration (mg/L) BMAA concentration (mg/L)

1 Erratic swimming Erratic

0 sC 0.1 1 10 100 BMAA concentration (mg/L)

Figure 3.7. (A) Total swimming distance, (B) swimming speed during movement, (C) incidence of freezing, (D) time spent in freezing and (E) incidence of erratic swimming for larvae exposed to sC (20 mM NaHCO3) or BMAA (1-100 mg/L) in sC at 144 hpf. Bars represent the average value ± SEM per test group (*p < 0.05, **p < 0.01, ***p < 0.001 vs sC; n = 12).

101

A 2 sC

0.1 mg/L BMAA 1.5

0.5

* * Swimming Swimming distance(cm) 0 0 5 10 15 20 Startles

B 2 sC

1 mg/L BMAA 1.5

0.5

* Swimming Swimming distance(cm) 0 ** 0 5 10 15 20 Startles

C 2 sC

10 mg/L BMAA 1.5

* 0.5 *** Swimming Swimming distance(cm) 0 0 5 10 15 20 Startles

D 2 sC

100 mg/L BMAA 1.5

0.5

Swimming Swimming distance(cm) * 0 0 5 10 15 20 Startles

102

E F

1.0 14

12 0.8 * * 10 8 0.6 6

0.4 habituation 4

Startles needed for needed Startles 2 Swimming distance (cm) (cm) distance Swimming 0.2 0 sC 0.1 1 10 100 sC 0.1 1 10 100 BMAA concentration (mg/L) BMAA concentration (mg/L) Figure 3.8. Startle response to continual 20 taps for larvae exposed to sC (20 mM

NaHCO3) or BMAA (1-100 mg/L) in sC at 168 hpf. The average swimming distance of larvae in 2 s after each tap was compared between sC and the (A) 0.1 mg/L, (B) 1 mg/L, (C) 10 mg/L and (D) 100 mg/L BMAA groups, respectively. “ ” marks the beginning of taps. Data points represent the average value ± SEM per test group (*p < 0.05, **p < 0.01, ***p < 0.001 vs sC; n = 10). (E) Average swimming distance over 20 tap and (F) number of taps needed for startle habituation for larvae from different test groups. Bars represent the average value ± SEM per test group (*p < 0.05 vs sC; n = 10).

103

A B

100 100

80 80

60 60

40 40

20 20 Thigmotaxis (%) in time Thigmotaxis

Thigmotaxis (%)in distance Thigmotaxis 0 0 sC 0.1 1 10 100 sC 0.1 1 10 100 BMAA concentration (mg/L) BMAA concentation (mg/L)

C D

35 12

30 10

25 8 20 6

15 (cm/min)

distance (cm) distance 4

Total swimming Total 10 Swimming speedSwimming 5 2 0 0 sC 0.1 1 10 100 sC 0.1 1 10 100 BMAA concentration (mg/L) BMAA concentration (mg/L) Figure 3.9. Thigmotaxis and swimming activity of larvae at the age of 7 dpf after 10 min exposure to sC (20 mM NaHCO3) or BMAA (1-100 mg/L) in sC. (A) Percentage of distance larvae swam and (B) percentage of time larvae spent in outer zone of Petri dish. (C) Total swimming distance and (D) swimming speed during movements for larvae from different test groups. Bars represent the average value ± SEM per test group (n = 7).