Physiological and Molecular Developmental Effects of a PPAR-agonist, GW7647, on zebrafish (Danio rerio)

Wenwan Dong

Projektrapport från utbildningen i

EKOTOXIKOLOGI Ekotoxikologiska avdelningen

Nr 139

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Contents

Acknowledgments…………………………………………………………………… 3 Abstract…………………………………………………………………………4

1. Introduction…………………………………………………………... . .5 2. Materials and Methods………………………………………………..9

2.1 Experimental solution…………………………………………………………….9

2.2 Zebrafish and egg production………………..………………………………9

2.3 Exposure………………………………………………......……………………….10

2.4 Determination of mortality…………………………………......…………….10

2.5 Examination of heartbeat rate……………………………………………………10

2.6 RNA isolation and cDNA synthesis…………………………………………………..11

2.7 Quantitative real-time PCR…………………………………………………………11

2.8 Data analysis……………………………………………………………………12

2.9 Statistics…………………………………………………………………………………12

3. Resu l t s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14 3.1 Morta l i ty. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .14

3.2 Heartbeat rate.... . .. . .. .. . .. . .. . .. . .. . .. . .. . .. . .. . .. . .. .. . .. . .. . .. . .. . .. . .. . .. . .. . .. . .. .14

3.3 mRNA expression.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .15

4. D i s c u s s i o n . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 1 4.1 M o r t a l i t y. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 1

4.2 Heartbeat rate..... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .21

4.3 mRNA expression.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .23

5. Conclusion... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .27

References.....................................................................................................28

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Acknowledgments

I thank Prof. Björn Brunström for teaching and supervising of whole project. I also

thank Dr. Jan Olsson for his technical advice, teaching of qPCR assays and analysis.

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Abstract

Pharmaceuticals and their metabolites are emerging environmental contaminants.

Among these pollutants of concern, agonists to peroxisome proliferator-activated

receptors (PPARs) are among those most frequently reported to be present in the

environment. In contrast to knowledge about their environmental occurrence, little is

known about their effects on organisms in the environment, especially on aquatic

species. In this project, the aim was to analyze physiological and molecular effects of

GW7647 in fish. GW7647 is known to be a selective PPARα agonist in humans.

Zebrafish (Danio rerio) embryos/larvae were exposed in 96-well plates to

concentrations of 0.1 μM, 0.3 μM, 1 μM and 3 μM GW7647 for 8 days. After 3 days

of exposure, the heartbeat rates of newly hatched larvae were determined. After 8

days of exposure, the fish larvae were analyzed for expressions of PPARs and three

related genes by real-time quantitative polymerase chain reaction (real-time qPCR).

GW7647 caused a significant decline in heartbeat rate from 0.3 μM to 3 μM. There

was no significant change in mRNA expression for any of the tested genes. The

heartbeat decrease may be linked to activation of PPARs but the mRNA expression

results did not indicate PPAR activation.

Keyword: zebrafish Danio rerio, GW7647, developmental stage, heartbeat rate,

PPARs, acyl-CoA oxidase, liver fatty acid-binding protein

Abbreviation:

Peroxisome proliferator-activated receptors: PPARs;

Acyl-CoA oxidase: ACOX;

Liver fatty acid-binding protein: L-Fabp;

Enoyl-CoA hydratase/ L-3-hydroxyacyl-CoA dehydrogenase: Ehhdh;

Real-time quantitative polymerase chain reaction: real-time qPCR.;

Retinoid X Receptor: RXR

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1. Introduction

Pharmaceuticals and their metabolites are emerging environmental contaminants

and there is a growing use of pharmaceuticals in human and veterinary medicine.

Due to disposal of unused drugs, elimination from patient’s body, and discharge from

the pharmaceutical industry, they are present in sewage treatment plant effluents

and enter into the environment (Daughton and Ternes, 1999). Fibrates are among the

most frequently reported pharmaceuticals contaminating aquatic environments. The

concentrations of fibrates are up to μg/L levels in surface water and they are found in

drinking water and some of them become persistent toxicants (Daughton and Ternes,

1999; Fent et al., 2005; Togola and Budzinski, 2007). Although the individual

concentration of a drug may not cause acute toxicity in the environment, the

combined effects of drugs and the sensitivities of nontarget organisms are unknown

(Daughton and Ternes, 1999).

A variety of environmental contaminants including fibrates are classified as agonists

of the peroxisome proliferator-activated receptors (PPARs). PPARs are nuclear

receptors and fibrates exert their effects by activating these ligand-dependent

transcription factors. Activated PPARs form heterodimers with the Retinoid X

Receptor (RXR) and bind to specific regions of target DNA sequences to regulate the

transcription of RNA. Therefore, activated PPARs induce transcription of peroxisomal

enzymes, especially acyl-CoA oxidase (ACOX), and cause corresponding effects on

lipid metabolism (Berger and Moller, 2002).

Since the PPAR (PPARα) was discovered in 1990 (Issemann and Green, 1990), the

roles of PPARs in the regulation of metabolism, development and tumorigenesis have

been studied. In this early study on PPARs, they were referred to increase the size

and numbers of liver peroxisomes in rodent liver tissue (Issemann and Green, 1990).

Then PPARs were identified as receptors to induce peroxisome proliferation in

Xenopus frog cells (Dreyer et al., 1992). In the paper by Dreyer and co-workers, the

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presence and functions of PPARs during development were firstly studied in Xenopus

frog embryos.

Three subtypes of PPARs have been identified and named as PPARα, PPARβ (also

called PPARδ) and PPARγ. The subtypes have different tissue distributions (Berger

and Moller, 2002). It was demonstrated that all three distinct PPAR subtypes are

present in zebrafish (Danio rerio). PPARα was expressed mainly in liver, proximal

tubules of kidney, enterocytes, and pancreas. PPARβ showed a widespread

distribution, e.g. in kidney, pancreas, intestine, skin epithelium, lymphocytes, and

gonads. PPARγ was expressed weakly in pancreas, intestine, and gonads (Ibabe et al.,

2002). Additionally, the three subtypes of PPARs are also expressed in early

developmental stages of frog (Xenopus) and zebrafish (Dreyer et al., 1992, Ibabe et

al., 2005). The expression levels of the PPAR subtypes differ in different

developmental stages in zebrafish. In adult females, PPARα and PPARγ were reported

to be strongly expressed in the early stage oocytes, and moderately in late stage

oocytes. PPARβ expression was generally more intense in juveniles than in other

stages. For the different developmental stages, PPARβ was distributed similarly to

PPARα but showed much weaker expression than PPARα. The expression of PPARγ

was higher in the early stages than in adults (Ibabe et al., 2005).

Since contaminants that modulate PPAR-mediated activities were found in the

aquatic environment, the potential risk of PPARs regulators to aquatic organisms are

of particular concern. Opposing to animals that at least spend some time in

terrestrial settings, aquatic organisms are subject to continual lifecycle exposure

(Daughton and Ternes, 1999). There are many papers showing that PPARs agonists

induce peroxisome proliferation and peroxisomal enzymes in both fish cell culture

and exposed fish. In an earlier study, an environmentally relevant waterborne

concentration of gemfibrozil (1.5 μg/L) induced oxidative stress in goldfish liver and a

higher concentration (1500 μg/L) of exposure for 28 days reduced PPARβ mRNA

levels (Mimeault et al., 2006). Micromolar concentrations of clofibrate or gemfibrozil

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induced an embryonic malabsorption syndrome in zebrafish, resulting in little yolk

consumption and small-sized larvae (Raldua et al., 2008). Clofibric acid induced the

PPAR-regulated enzyme ACOX activity in male fathead minnows after 21 days of

exposure to 108.9 mg/L (Weston et al., 2009). Injection of ciprofibrate in rainbow

trout caused dose-related increases in peroxisomal ACOX activity (Yang et al., 1990).

Fenofibrate was also reported to increase the peroxisome-related activities such like

catalase and ACOX in rainbow trout (Du et al., 2004).

Knowing the regulation mechanism by PPAR-agonists and the effects of some

PPAR-modulators in fish led us to investigate the physiological and molecular effects

of a PPAR-agonist, GW7647, on a common experimental aquatic vertebrate species,

zebrafish.

GW7647 is a potent and subtype-selective human PPARα agonist. It is a man-made

urea-substituted thioisobutyric acid, which activates the human PPARα, PPARγ and

PPARδ with EC50 values of 0.006, 1.1 and 6.2 μM, respectively (Brown et al., 2001).

In a similar assay on murine PPARα, PPARγ and PPARδ receptors, GW7647 was also

highly selective having EC50s of 0.001, 1.3 and 2.9 μM, respectively (Brown et al.,

2001). In the present study, the tropical freshwater species zebrafish was chosen as

the experimental fish. Because zebrafish are small, easy and inexpensive to care for,

and produce large numbers of transparent embryos that develop outside of the

mothers, they are common and important model organisms for studies of

development. In last few years, numbers of investigations of toxic side effects of

pharmaceuticals used the zebrafish (Rubinstein, 2006). Due to the sensitivity during

developmental stages, zebrafish embryos and larvae are also used to investigate the

toxicity of environmental contaminants (Hill et al., 2005). For example, the impact of

ibuprofen (an over-the-counter drug) on the development of zebrafish implicated the

drug’s embryotoxicity at a high (>10 μg/L) dose level (Anuradha and Pancharatna,

2009).

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In this report, GW7647 was used to expose early stages of zebrafish and physiological

effects and effects on mRNA expression of the PPARs and the target genes ACOX,

Liver fatty acid-binding protein (L-Fabp) and enoyl-CoA hydratase/

L-3-hydroxyacyl-CoA dehydrogenase (Ehhdh) were studied.

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2. Materials and Methods

2.1 Experimental solution

GW7647 and DMSO (dimethylsulfoxide) were purchased from Sigma-Aldrich (St.

Louis, MO, USA). A stock solution of 15 mM GW7647 was prepared in DMSO. Groups

of embryos were exposed to four concentrations of GW7647 (3 μM, 1 μM, 0.3 μM,

0.1 μM). All the exposure waters and the control water were prepared to the same

final concentration of DMSO (0.02%). The water used for zebrafish embryo exposure

was treated under the conditions as described in the OECD Guideline 210 (1992).

2.2 Zebrafish and egg production

Zebrafish were obtained from a local pet store (Kalhälls Akvarium AB, Stockholm,

Sweden) and kept in the aquarium facility at Department of Environmental Toxicology,

Uppsala University. Fish were raised and maintained under 14-hour light: 8-hour dark

cycle in a tank with 40% water changed daily. Water quality and environmental

conditions were according to The Zebrafish Book (Westerfield, 2000). Fish were fed

Aquarium Nature Tropical S Fishfood once a day.

A stainless steel cage with a mesh size of 1-2 mm was hanged into an aquarium as a

breeding chamber. The cage was placed about 10 cm above to the bottom of the

tank, thus allowing the eggs to be collected on the bottom and to be protected from

predation by the adults. Artificial plants were put into the cage as breeding stimulant

and substrate and the water temperature was about 28oC. Twenty-two adult fish (the

ratio of male: female was about 1:1) were placed into the cage before the onset of

darkness. Within 30 min after the onset of light in the next morning, all adults were

removed with the cage and the eggs were collected.

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2.3 Exposure

In order to start exposure with minimum delay, the collected eggs were divided

evenly in five groups and transferred immediately into 100 mm crystallization dishes

containing different test concentrations (3 μM, 1 μM, 0.3 μM, 0.1 μM), and control

solution, respectively. An inverted microscope with magnification of 160× was used

to distinguish fertilized eggs. The fertilized eggs were transferred into a 96-well plate

with the same concentration of GW7647 that they were exposed to before. One

fertilized egg was placed in each well. The embryos were reared in a dark incubator

at 27oC. Half the volume of solution in each well was renewed every day. The whole

exposure period was 8 days (from the spawning day to the fifth day after hatching),

under food deprivation condition.

2.4 Determination of mortality

After selection of fertilized eggs, the embryos in each treatment were counted and

the number noted. All the fertilized eggs per treatment were used to assess mortality

in the different groups. The lethal endpoint included coagulation, tail-not-detached,

no-somite-formation and no-heart-beat. Eggs were checked every 24 hours and the

deaths were noted as “died before hatching” or “died after hatching”. The mortalities

were expressed as a percentage.

2.5 Examination of heartbeat rate

After three days of exposure, the surviving hatched larvae were examined for

heartbeat rate. Sixty individuals in each treatment or control were collected

randomly to examine the heart rate. Twenty heartbeats for each larva was timed by

stopwatch and noted. The data were used to calculate the number of heartbeats per

minute. The final results for the different treatments were expressed as average

heartbeat rate in one minute (beats/min).

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2.6 RNA isolation and cDNA synthesis

After 8 days of exposure, 25 surviving larvae per concentration were considered as

one sample and transferred into a micro tube; five samples were prepared for each

concentration. All the samples of larvae were killed by instant freezing in liquid

nitrogen.

RNA was isolated from the samples using the Aurum Total RNA Fatty and Fibrous

Tissue kit according to the manufacturer’s instruction (Catalog #732-6830, Bio-Rad,

Laboratories, CA, US). Once the RNA was bound and purified on the RNA binding

column, it was eluted by nuclease-free water instead of the elusion solution in the kit.

The concentrations and purities of the RNA samples were assessed using a NanoDrop

ND-1000 Spectrophotometer (NanoDrop Technologies Inc., DE, US). A standard

volume of RNA (1 µg per sample) was reverse transcribed to cDNA using iScript cDNA

Synthesis kit (Catalog #170-8891, Bio-Rad). cDNA was then diluted 1:100 in

nuclease-free water.

2.7 Quantitative real-time PCR

Quantitative PCR reactions were conducted on a Rotor-Gene 6000 (Corbett Research,

Sydney, Australia) using SYBR green technology. The reagents were acquired from

Bio-Rad. The reactions were carried out using the iQ SYBR Green Supermix kit

(Catalog #170-8885, Bio-Rad). The thermal profile for the SYBR green reactions was 3

min at 95 oC followed by 45 cycles of 15s at 95 oC, 15s at annealing temperature (51oC,

54 oC or 57 oC, different for different primers), and 20s at 68 oC. Data were collected

during the 68 oC extension period.

Primers for each gene were published before or they were designed with Primer3 (v.

0.4.0) and evaluated with NetPrimer from PREMIER Biosoft (CA, USA). The primer

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sequences are shown in Table 1.

Table 1: Nucleotide sequences for the primers used in the quantitative real-time PCR reactions. Name Sequence (5’-3’) Accession number

DR* Forward TCTGGAGGACTGTAAGAGGTATGC NM 212784

Rpl13α Reverse AGACGCACAATCTTGAGAGCAG

DR Forward AAGGAAGCCGCTGAGATG ENSDART

EF1α Reverse AGCACAGCACAGTCAGCC 00000023156

(Lin et al., 2009)

DR Forward CTTCTTGGGTATGGAATCTTGC BC154531

β-actinB2 Reverse GTACCACCAGACAATACAGTG (Drew et al., 2008)

DR Forward CATCTGCTGTGGAGACCGTC DQ 017612

PPARα Reverse CTTCTGTCTTGTTGATCTCCTGC

DR Forward GTCGCCGCAATCATCC XM 694808

PPARβ1 Reverse GCGTTCTCCGTCACCAG

DR Forward ATGACGCAATAAGGTACGG NM 131468

PPARδb Reverse CAGGTAGGCTGTGTTGACG

DR Forward CGCATACACAAGAAGAGCC NM 131467

PPARγ Reverse CGGTGACTTCGCTGATGG

DR Forward GAGGAGTTTCTCAGAGCCATCTC AF254642

L-Fabp Reverse TCCATAGTGGTGATTTCAGCCT

DR Forward GATTCTGTGGAGGTGCTGAC NC 007120

Ehhdh Reverse CAATGCGGTAATGACAGACTA

DR Forward AATAGAAGGAGAGAAATAGAGTC NM 001005933

ACOX Reverse CAACAGTCTTGTAGGAGTAGAT

* DR = Danio rerio (zebrafish)

2.8 Data analysis

Quantitative PCR data were analyzed using the Rotor-Gene 6000 application software.

The difference in target gene mRNA abundance between dosed samples and the

DMSO control was calculated using the 2-ΔΔCt equation (Schefe et al., 2006).

2.9 Statistics

Statistical data analysis was performed by Microsoft Excel (Microsoft Corporation,

Washington, USA) and GraphPad Prism 5 Demo (Ver. 5.3, GraphPad Software Inc., CA,

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USA). Statistical differences between exposed groups and the DMSO control were

calculated by a one-way analysis of variance (ANOVA) followed by Dunnett’s T-test.

Differences were considered significant if P<0.05.

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3. Results

3.1 Mortality

After the selection of fertilized eggs, the exposed embryos were 174, 144, 160, 144

and 180 in concentrations of 3 µM, 1 µM, 0.3 µM, 0.1 µM and control, respectively.

Before hatching, there were 6, 2, 3, 3 and 4 dead embryos in these treatments. After

hatching to the end of exposure, the numbers of dead larvae were 5, 2, 1, 2 and 2,

respectively. Therefore, the total mortalities during the whole exposure period were

6.3%, 2.8%, 2.5%, 3.5% and 3.3% from the group exposed to the highest

concentration to the control.

Besides the exposure from 0.1 µM to 3 µM of GW7647, an initial experiment was

done using the same conditions but different exposure concentrations. Mortality

among embryos exposed to 10 µM GW7647 was significantly higher (64.6%) than in

the control group (3.1%). This concentration was excluded in the later experiment.

3.2 Heartbeat rate

After 72h of exposure, the heartbeat frequency of hatched larvae was examined. In

each group, data from 60 larvae was collected randomly to calculate the mean value

of heartbeats in 1 minute (See Fig 1). Generally, the control larvae had the fastest

heartbeat rate (161 per min in average), while the heartbeat rate in the group

exposed to 0.3 µM GW7647 was lowest (132 per min).

The heartbeat rate of larvae in 0.1 µM was faster (154 per min) than in other

exposed groups, but the highest exposure concentration (3 µM) did not lead to the

slowest heartbeat rate (140 per min).

Comparing with the control, there were significant differences for the groups

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exposed to 0.3 µM, 1 µM and 3 µM. The P value was less than 0.0001 for the

One-way Analysis of Variance and less than 0.0001 for Dunnett’s Multiple

Comparison Test.

Fig1: Mean heartbeat numbers in 1 minute after 72 hours (from fertilization to hatching) of exposure to different concentrations of

GW7647. Means were calculated from 60 individuals in one

treatment. Variation is shown as standard error.

***P<0.0001 based on one-way ANOVA and Dunnett’s T-test.

3.3 mRNA expression

Ct values were obtained for three reference genes and seven test genes. Gene

Rpl13α, EF1α and β-actinB2 were considered as reference genes and their

expressions in the different groups were examined. Exposure of zebrafish early stages

to GW7647 in different concentrations did not alter the Rp13α mRNA expression

level compared to the control (data not shown). Therefore, the Rp13α was chosen

for use in the later result analysis because it was expressed more stably than the

other two genes.

For all genes examined (four PPARs genes and three PPARs-regulated genes) no

significant difference in expression was observed between the undosed larvae and

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the larvae exposed to GW7647 (see Fig 2).

The relative mRNA expressions of all the PPARs genes in the treatment groups were

higher than in the control but the differences were not significant (See Fig. 2a-d).

Without considering the variations, the mean expressions in the treated groups for

PPARα were approximately 2.5- fold, 2- fold, 1.7- fold and 1.5- fold the control value,

respectively (Fig. 2a). For PPARβ1, the mean mRNA expression increased with

increasing concentration, from 2 times higher (0.1 µM) to 2.5 times (3 µM) higher

than the mean expression in the control (Fig. 2b). For PPARδb, mean values of mRNA

expressions in the treatment groups were from approximately 3- fold (0.1 µM and 1

µM) to 2- fold (3 µM) the control value (Fig. 2c). For PPARγ, the relative mRNA

expressions were approximately 4- to 5- fold in dosed larvae compared with the

average value of the controls (Fig. 2d).

The variances were high and therefore differences were not significant in spite of

large differences in mean values.

In GW7647-exposed groups, ACOX was nominally higher than the control value but

no significant differences were found. When comparing the averages of all exposed

groups to control, they were approximately 2.2- fold, 1.8- fold and 2.0- fold higher,

respectively (Fig. 2e). Embryos exposed to 0.1 µM and 3 µM of GW7647 showed a

slightly higher Ehhdh transcript abundance than the mean of the control (Fig. 2f),

while embryos exposed to 0.3 µM and 3 µM showed a slightly higher L-Fabp

transcript abundance than the control (Fig. 2g).

For all genes studied, the lowest variance always occurred in the control (see Fig. 2).

It indicates that control larvae may grow and develop more stably than dosed larvae.

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Fig. 2(a): Mean (+S. E.) mRNA expression of PPARα after 8 days (from embryo to larvae stage) of exposure to GW7647. Means

were calculated using data from five independent larva replicates

and each replicate included 25 individuals.

Fig. 2(b): Mean (+S. E.) mRNA expression of PPARβ1 after 8 days (from embryo to larvae stage) of exposure to GW7647. Means

were calculated using data from five independent larva replicates

and each replicate included 25 individuals.

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Fig. 2(c): Mean (+S. E.) mRNA expression of PPARδb after 8 days (from embryo to larvae stage) of exposure to GW7647. Means

were calculated using data from five independent larva replicates

and each replicate included 25 individuals.

Fig. 2(d): Mean (+S. E.) mRNA expression of PPARγ after 8 days

(from embryo to larvae stage) of exposure to GW7647. Means

were calculated using data from five independent larva replicates

and each replicate included 25 individuals.

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Fig. 2(e): Mean (+S. E.) mRNA expression of ACOX after 8 days (from embryo to larvae stage) of exposure to GW7647. Means

were calculated using data from five independent larva replicates

and each replicate included 25 individuals.

Fig. 2(f): Mean (+S. E.) mRNA expression of Ehhdh after 8 days

(from embryo to larvae stage) of exposure to GW7647. Means

were calculated using data from five independent larva replicates

and each replicate included 25 individuals.

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Fig. 2(g): Mean (+S. E.) mRNA expression of L-Fabp after 8 days

(from embryo to larvae stage) of exposure to GW7647. Means

were calculated using data from five independent larva replicates

and each replicate included 25 individuals.

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4. Discussion

Developmental toxicity is a very important area of toxicology, since the development

of embryos involves many organs and physiological processes. Effects can include low

birth weight, malformations, behavioral deficits or even death of the newborn (Bailey,

2008). Studies in developmental toxicology show the potential adverse effects of

chemicals during sensitive life stages and may be useful to predict the hazard of

chemicals.

4.1 Mortality

Exposure to GW7647 in early stages might interfere with survival of zebrafish larvae.

Combining the results of mortality data from the two experiments, the mortalities in

controls were stable and acceptable, only about 3% (3.33% and 3.13%). The mortality

in the group exposed to 3 µM of GW7647 was about twice the mortality in the

control (6.32%) and mortality when the embryos were exposed to 10 µM was 64.62%.

This result suggests that the LC50 (50% lethal concentration) of GW7647 is between

3 µM and 10 µM.

The purpose of counting number of dead embryos/larvae during the exposure was to

find a suitable range of doses. Therefore, 0.1 µM, 0.3 µM, 1 µM and 3 µM were

chosen as exposure concentrations since most individuals (more than 90%) survived

and there were no marked malformations at these doses.

4.2 Heartbeat rate

The heartbeat rate of dosed larvae significantly declined from 0.1 µM to 3 µM

compared with the control. This effect may involve activation of PPARs because all

PPAR subtypes play an important role in controlling transcriptional expression of

enzymes, for instance those involved in glucose metabolism (Marx et al., 2004; Xiao

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et al., 2009). The regulation of cardiac PPARα by GW7647 was proved in a study of

Yue et al. (2003). In this study, GW7647 was reported to attenuate the

downregulation of PPARα and fatty acid oxidation enzymes in mice when myocardial

injury from ischemia/reperfusion was induced. In PPARα-null mice, GW7647 did not

protect the heart (Yue et al., 2003). Down-regulation of PPARα signaling could

preserve heart function against pressure overload, while cardiac PPARα

overexpression led to lipid accumulation and might cause a diabetic heart (Marx et

al., 2004).

All three subtypes of PPARs are expressed in vascular smooth muscle cells in heart.

As a highly selective and potent PPARα agonist, GW7647 was demonstrated to

stimulate glucose uptake in cardiac muscles. In isolated papillary muscles cells

exposed to 5 μM, 10 μM and 20 μM of GW7647 for 30 min a significant increase in

glucose uptake was found (Xiao et al., 2009). In addition, the glucose uptake was

demonstrated to lead to hemodynamic change. In rabbits treated with glucose, their

heartbeat rate was remarkably decreased (Farias et al., 1986).

Heartbeat of zebrafish could be affected by factors like exposure to chemicals, or

special living conditions (Padilla and Roth, 2001; Langheinrich et al., 2003). During

the study, all zebrafish embryos and larvae were raised under the same living

condition without contamination with other chemicals than GW7647. Therefore, I

suggest that zebrafish exposure to GW7647 activated the PPARα mainly in cardiac

muscle cells. By the transcriptional expression, activated PPARα induced cardiac

metabolism involved in glucose uptake. An unusual cardiac metabolism might cause

hemodynamic change and lead to significant decrease of the heartbeat rate.

However, there was no evidence to demonstrate directly that the slow heartbeat

rates in exposed zebrafish larvae were caused by GW7647 via regulation of PPARs.

Further studies are needed to prove the mechanisms of GW7647 interaction with the

cardiovascular system of zebrafish larvae.

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Studies in animal models suggest that PPARα agonists may be used in clinical

modulations to protect from cardiovascular disease in the future (Yue et al., 2003;

Xiao et al., 2009). Therefore, understanding of their function has important clinical

implications.

4.3 mRNA expression

The main objective of the study was to determine the effect of GW7647 exposure in

zebrafish larvae by focusing on proteins involved in lipid metabolism. As a PPARs

activator, GW7647 is expected to affect mRNA levels of PPARs and their related

target genes.

Our study indicates that GW7647 had no significant effect on these mRNA

expressions even at the highest concentration of 3 µM. There are some evidences

that a high concentration of GW7647 (3 mg/kg/d) regulated mRNA expression of

PPARα and RXRα in rodents (Yue et al., 2003). Furthermore, after incubation for 24

hours with 1 µM of GW7647, all three subtypes of PPARs were activated in human

cell lines (Seimandi et al., 2005). No data was reported on effects of GW7647 in

zebrafish.

GW7647 is a potent and selective activator of PPARα in humans and rodents and it

shows >100-fold and >1000-fold selectivity for PPARα over PPARβ and PPARγ in

humans and mice, respectively (Brown et al., 2001). The relative PPARγ mRNA

expression in the present study was slightly higher than that of the other PPAR

subtypes. This is supported by former studies on PPARs expression in different

developmental stages of zebrafish showing that PPARγ expression is higher in the

early stages than in adults (Ibabe et al., 2005).

As a PPARα-selective agonist, GW7647 should affect fatty acid metabolism as do

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other PPARα agonists such as fenofibrate and bezafibrate. The relative gene

expression changes in common may indicate the genes regulated by PPARα

stimulation. From the expression data obtained from rodents and dogs, it was

reported that PPAR agonists induced the peroxisomal enzymes ACOX and Ehhdh (Guo

et al., 2006; Guo et al., 2007).

Acyl-CoA oxidase (ACOX) is a target gene regulated by PPARs. In this study, relative

mRNA expression of the ACOX gene was tested to evaluate its regulation by

activation of PPARα. The ACOX gene did not change significantly by treatment with

GW7647. In many previous studies, ACOX regulation by peroxisome proliferators was

shown by measuring ACOX activity. The activity was assessed by using a method that

measures the H2O2 production upon oxidation of leucodichlorofluorescein catalysed

by ACOX (Small et al., 1985).

Exposure of fish to peroxisome proliferators sometimes led to significant effects on

ACOX, but some papers also reported lack of effect on ACOX after exposure. It was

demonstrated that 1 mM clofibric acid or 1 mM bezafibrate administered to salmon

(Salmo salar) hepatocytes in culture, resulted in a significant increase of PPARs mRNA

expression and of ACOX activity (Small et al., 1985). However, when fathead minnows

(Pimephales promelas) were exposed to bezafibrate at concentrations up to 106.7

mM, no effect was observed. When exposed to clofibric acid, only the fathead

minnows at the highest tested concentration (108.9 mM) showed induced activity of

ACOX and there was no increase in expression of PPARs mRNA (Weston et al., 2009).

In addition, in rainbow trout (Onchorynchus mykiss) exposure to another peroxisome

proliferator, gemfibrazil, led to activity change of ACOX after injection daily for 14

days with the doses 46, 87 or 152 mg/kg/day (Scarano et al., 1994). However, when

goldfish (Carassius auratus) were exposed to gemfibrazil (up to 1.5 mM) for the same

duration (14 days), there was no significant change of ACOX activity (Mimeault et al.,

2006). It thus seems that even after exposure to the same lipid regulators, the

experimental results could be different. One explanation to this may be differences in

25

sensitivity between fish species, but the exposure pathway is also important for the

results. Exposure in cell culture may give a sensitive response and injection is a more

direct pathway than to expose fish to the chemical via the water. In the studies

referred to above, most peroxisome proliferators tested did not activate ACOX or

regulate PPARs mRNA expression by exposure via the water, which is in agreement

with my result.

After exposure to PPARα agonists, the activation of PPARα firstly results in induction

of the enzyme ACOX and then Ehhdh is induced as the next enzyme in the cascade

(Guo et al., 2006). Ehhdh was not significantly increased in my project, but Ehhdh can

be activated by a novel PPARα and γ coagonist, LY465608, in rodent and dog cells

(Guo et al., 2007). Also, treatment of mouse primary hepatocytes with bezafibrate

and fenofibrate can elevate the Ehhdh mRNA expression level (Guo et al., 2006).

Human and rodent data provide evidence that L-Fabp interacts with PPARα and

PPARγ but not with PPARβ, by protein–protein contacts. During activation, L-Fabp is

considered as a co-activator in PPAR-mediated gene control (Wolfrum et al., 2001).

There was no effect on L-Fabp in my study, but in rat hepatocytes cultured in

presence of bezafibrate, the L-Fabp mRNA expression was clearly higher than in

non-drug-treated cells (Besnard et al., 1993). Following exposure of PPARα-deficient

mouse hepatocytes to the classical peroxisome proliferators, clofibrate and

Wy-14,643, the L-Fabp level did not change and there was no fatty acid metabolism

(Lee et al., 1995). These studies show that PPARα mediates the induction of L-Fabp. It

was reported that in most cases the expression pattern is similar in all mammals but

may be different in fish (Haunerland and Spener, 2004). In a recent study on fish, it

was demonstrated that under complex environmental stresses, the PPARβ and

L-Fabp mRNA expressions in livers of goldfish (Carassius auratus) were significantly

higher than in control (Wang et al., 2008). So L-Fabp may be linked to all three PPARs

subtypes in fish.

26

In this study, it seems there was no evidence that GW7647 activated the PPARs or

affected the expression of these receptors. There were very few studies on exposure

of fish to GW7647 before. Many studies where fish were exposed to other PPARs

agonists showed a lack of effects on mRNA expression (Mimeault et al., 2006; Wang

et al., 2008; Weston et al., 2009). Therefore, my and other studies may imply that

activation of PPARs is not very easy and has high individual or species differences.

The high variation values for the mRNA expressions of the exposed fish in the

present study may confirm the individual differences.

To investigate PPAR-regulated genes, besides studying the mRNA levels of PPARs,

ACOX, L-Fabp and Ehhdh, the relative mRNA expression of Retinoid X Receptor (RXR)

should be examined since all PPARs need to heterodimerize with it to bind to the

response elements on DNA. So RXR expression and activation are important in PPARs

regulated processes and need to be studied.

27

5. Conclusion

Determining the physiological effects and molecular actions of PPARs agonists in fish

species is important for environmental risk assessment. This study shows that

exposure to a selective PPARα agonist in humans, GW7647, is capable of regulating

the heart function and leads to the slow heartbeat as response. This effect may be

caused by GW7647 regulation of PPARs but the mRNA expression results cannot

support this hypothesis. However, abnormal heartbeat rates are induced by GW7647

and the mechanism needs further study. As in previous studies, exposure of fish to

PPARs modulators in the aquatic environment seems not to activate the PPARs.

Future studies are needed to show whether PPARα-dependent expression can be

induced in fish larvae.

Besides studying the PPARα agonist GW7647, future studies can also be expanded to

include PPARs agonists found in the environment, such as fibrates, and their effects

on fish.

28

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