Drug disposition in the zebrafish embryo and larva: focus on … · 2020. 1. 7. · Universiteit...

Faculty of Pharmaceutical, Biomedical and Veterinary Sciences

Department of Veterinary Sciences

Drug disposition in the zebrafish embryo

and larva: focus on cytochrome P450

activity

Dissertation for the degree of doctor in Veterinary Sciences (PhD) at

the University of Antwerp to be defended by

EVY VERBUEKEN

Supervisors:

Prof. Dr. Steven J. Van Cruchten

Prof. Dr. Chris J. Van Ginneken Antwerp, 2019

Faculteit Farmaceutische, Biomedische en

Diergeneeskundige Wetenschappen

Departement Diergeneeskundige Wetenschappen

Geneesmiddelendispositie in het

zebravisembryo en -larve met de focus op

cytochroom P450 activiteit

Proefschrift voorgelegd tot het behalen van de graad van doctor in

de Diergeneeskundige Wetenschappen aan de Universiteit

Antwerpen te verdedigen door

EVY VERBUEKEN

Promotoren:

Prof. Dr. Steven J. Van Cruchten

Prof. Dr. Chris J. Van Ginneken Antwerpen, 2019

Doctoral committee

Supervisors

Prof. Dr. S.J. Van Cruchten Faculteit Farmaceutische, Biomedische en


Universiteit Antwerpen

Prof. Dr. C.J. Van Ginneken

Faculteit Farmaceutische, Biomedische en



Members of the PhD Steering Committee

Prof. Dr. D. Knapen Faculteit Farmaceutische, Biomedische en



Dr. Luc Van Nassauw

Faculteit Geneeskunde en Gezondheids-

wetenschappen


Members of the PhD Examination Committee

Prof. Dr. P. Annaert Faculteit Farmaceutische Wetenschappen

Katholieke Universiteit Leuven

Prof. Dr. J. Legler

Faculteit Diergeneeskunde

Universiteit Utrecht

ISBN: 9789057286391

Depot number: D/2019/12.293/25

Cover created by Ronny Verbueken – Zebrafish images © 2018 Zebrafishlab

Cover design by Natacha Hoevenaegel, Nieuwe Media Dienst , UA

1

Table of contents

List of abbreviations………………………………………………………..…7

Chapter 1: General introduction……………………………………………11

1. Preface…………………………………………………………………………13

2. Developmental toxicity studies: an overview……………………………..14

2.1 Terminology……………………………………………………………...14

2.2 Historical and legal context……………………………………………..14

2.3 Traditional and alternative test systems in developmental toxicity..18

3. The Zebrafish…………………………………………………………………27

3.1 Zebrafish in their natural habitat………………………………………27

3.2 Zebrafish in a laboratory setting……………………………………….28

3.3 Reproduction and breeding…………………………………………….32

3.4 Embryonic and larval development…………………………………...34

4. Disposition of xenobiotics in zebrafish……………………………..……...49

4.1 ADME in mammals and zebrafish……………………………………..49

4.1.1 Absorption……………………………………………………….50

4.1.2 Distribution……………………………………………………...51

4.1.3 Metabolism………………………………………………………51

4.1.4 Excretion…………………………………………………………53

4.2 Cytochrome P450 enzymes in humans and zebrafish………………..55

4.3 Phase II enzymes in humans and zebrafish…………………………...62

4.4 Transport proteins in humans and zebrafish…………………………47

5. References…………………………………………………………………......74

Chapter 2: Aims of the doctoral project…………………………………...85

Chapter 3: In Vitro Biotransformation of Two Human CYP3A Probe

Substrates and Their Inhibition during Early Zebrafish

Development……………………………………………………………........89

1. Abstract………………………………………………………………………..91

2

2. Introduction…………………………………………………………………..91

3. Materials and Methods………………………………………………………95

3.1 Fish maintenance and breeding………………………………………...95

3.2 Tissue sampling…………………………………………………………..96

3.2.1 Adult zebrafish………………………………………………….96

3.2.2 Zebrafish embryos………………………………………………96

3.3 Isolation of microsomes…………………………………………………97

3.3.1 Adult zebrafish………………………………………………….97

3.3.2 Zebrafish embryos……………………………………………....98

3.4 Benzyloxy-methyl-resorufin assay in adult zebrafish liver

microsomes……………………………………………………………….98

3.5 Benzyloxy-methyl-resorufin assay in microsomes from whole

zebrafish embryo homogenates……………………………………….100

3.6 Inhibition studies with adult zebrafish liver microsomes……….....100

3.6.1 Ketoconazole…………………..……………………………….100

3.6.2 CYP3cide……..…………………………………………………101

3.6.3 Preliminary study with 1–aminobenzotriazole………..……102

3.7 Benzyloxy-methyl-resorufin assay in CYP Baculosomes®……...…102

3.8 Benzyloxy-methyl-resorufin assay in recombinant

zebrafish CYPs………………………………………………………….103

3.9 Luciferin-IPA assay with adult zebrafish liver microsomes...……..104

3.10 Mathematical and statistical analyses……………………………105

4. Results……………..…………………………………………………………106


microsomes and in microsomes from whole zebrafish embryo

homogenates ……………………………………………………………106

4.2 Inhibition studies with adult zebrafish benzyloxy-methyl-resorufin

assay in cytochrome P450 (CYP) Baculosomes® and in recombinant

zebrafish CYPs liver microsomes…………………………….………..108

4.2.1 Ketoconazole and CYP3cide……………………………….....108

4.2.2 Preliminary study with 1–aminobenzotriazole………..……110

3

4.3 Benzyloxy-methyl-resorufin assay in cytochrome P450 (CYP)

Baculosomes® and in recombinant zebrafish CYPs……………...….110

4.4 Luciferin-IPA assay with adult zebrafish liver microsomes……..….111

5. Discussion…………………………………………………………………....112

6. Conclusions…………………………………………………………...……..116

7. References…..………………………………………………………………..118

Chapter 4: From mRNA Expression of Drug Disposition Genes to In

Vivo Assessment of CYP-Mediated Biotransformation during

Zebrafish Embryonic and Larval Development……………………...…123

1. Abstract………………………………………………………………………125

2. Introduction…………………………………………………………………125

3. Materials and Methods……………………………………………………..131

3.1 In vitro study on cytochrome P450 activity in zebrafish embryos,

larvae and adults………………………………………………………..131

3.1.1 Fish maintenance and breeding……………………………...131

3.1.2 Tissue collection and isolation of microsomes……..….……133

Benzyloxy-methyl-resorufin assay in microsomes prepared

from whole zebrafish embryos, larvae and adults………….134

3.1.3 Mathematical and statistical analyses…...………..…………136

3.2 In vivo study on cytochrome P450 activity in zebrafish embryos and

larvae………………………………………………………...…………...137

3.2.1 Fish maintenance and breeding……………...……………….137

3.2.2 Benzyloxy-methyl-resorufin assay in zebrafish embryos and

larvae…………………………………………………………....139

3.2.3 Preliminary inhibition study in zebrafish embryos of 98 and

122 hpf……………………………………………………….….142

3.2.4 Mathematical and statistical analyses…………….………….142

3.3 mRNA Expression of Phase I and Phase II enzymes and P–

glycoprotein……………………………………………………………..143

3.3.1 Fish Maintenance and Breeding……………………….....…..143

3.3.2 Quantification of mRNA levels by means of qPCR……..….145

4

3.3.3 Mathematical and statistical analyses…………………….…148

4. Results………………………………………………………………………148


larvae and adults…………………………………………………….….148


larvae……………………………………………………………………..151

4.2.1 Quantitative analysis of resorufin formation…………….…151

4.2.2 Qualitative analysis of resorufin formation….……….……..152

4.2.3 Preliminary inhibition study in zebrafish embryos of 98 and

122 hpf…………………………………………………………..158


glycoprotein……………………………………………………………..159

5. Discussion…………………………………………..………………………167

5.1 Ontogeny of in vitro and in vivo cytochrome P450 activity in

zebrafish embryos, larvae and adults…………………………...……167

5.1.1 In vitro versus in vivo……………………...……………….…167

5.1.2 Benzyloxy-methyl-resorufin versus 7–ethoxyresorufin…...169

5.1.3 Literature versus current study…………………………..…..170

5.2 Ontogeny of cytochrome P450 mRNA expression in zebrafish

embryos and larvae…………………………………………………….171

5.2.1 Cytochrome P450 mRNA expression during zebrafish

organogenesis…………………………………………………..172

5.2.2 Cytochrome P450 mRNA expression during zebrafish larval

development…………………………………………………....173

5.3 Ontogeny of mRNA expression of two Phase II enzymes and a P-

glycoprotein in zebrafish embryos and larvae………..……….…….175

6. Conclusions…….……………………………………………………………177

7. References………………………………………………………..…………..178

Chapter 5: General discussion…………………………………………….187

1. Implications of the current findings for developmental toxicity studies

using zebrafish embryos……………………………..…………………….191

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2. Possible solutions to prevent false negative results in alternative

developmental toxicity testing…………………………………………….197

3. Xenobiotic metabolism in zebrafish: an intra– and interspecies

comparison…………………………………………………………………..206

3.1 Xenobiotic metabolite formation during zebrafish development....206

3.2 The ontogeny of drug disposition enzymes and transporters in

zebrafish versus humans……………………………………………....213

3.3 Xenobiotic metabolite formation in zebrafish versus mammals…..217

4. General conclusion and recommendations………………………………220

5. References…………………………………………………………………....222

Summary…………………………………………………………………..…231

Samenvatting………….……………………………………………………..235

Dankwoord…….……….……………………………………..………..……239

7

List of abbreviations

ABC

ABZ

ADME

AhR

AOP

ATP

BAC

BBB

BOMR

CAR

CRO

CYP

Dpf

E2

EDTA

EMA

ATP–binding cassette

Albendazole

Absorption, Distribution, Metabolism and Excretion

Aryl hydrocarbon receptor

Adverse Outcome Pathway

Adenosine triphosphate

Baculosomes®

Blood–brain barrier

Benzyloxy–methyl–resorufin

Constitutive androstane receptor

Contract Research Organization

Cytochrome P450

Days post–fertilization

17β–estradiol

Ethylenediaminetetraacetic acid

European Medicines Agency

ER

ER

EROD

EST

FDA

FET

FETAX

FMO

GI

Endoplasmic reticulum

7–Ethoxyresorufin

Ethoxyresorufin–o–deethylase

Embryonic Stem cell Test

Food and Drug Administration

Fish Embryo acute toxicity Test

Frog Embryo Teratogenicity Assay–Xenopus

Flavin containing monooxygenase

Gastrointestinal

8

HLM

Hpf

IPA

LC–MS

LLOD

LLOQ

MAS

MDR

MP

MXR

NBD

NCE

OECD

P–gp

PPAR

PXR

qPCR

3Rs

RA

REACH

S.B.

SLC

SULT

TCDD

TMD

TST

Human liver microsomes

Hours post–fertilization

Isopropyl acetal

Liquid chromatography–mass spectrometry

Lower limit of detection

Lower limit of quantification

Metabolic Activating System

Multidrug resistance

Microsomal protein

Multixenobiotic resistance

Nucleotide–binding domain

New chemical entity

Organization for Economic Co–operation and Development

P–glycoprotein

Peroxisome proliferator–activated receptor

Pregnane X receptor

Quantitative polymerase chain reaction

Replacement, Reduction and Refinement

Retinoic acid

Registration, Evaluation, Authorization and Restriction of

Chemicals

Swim bladder

Solute carrier

Sulfotransferase

2,3,7,8–tetrachlorodibenzo–p–dioxin

Transmembrane domain

Testosterone

9

UGT

WEC

ZEDTA

ZEM

ZLaM

ZLM

ZM

Uridine 5'–difosfo–glucuronosyltransferase

Whole Embryo Culture assay

Zebrafish Embryo Developmental Toxicity Assay

Microsomes prepared from whole zebrafish embryo homogenates

Microsomes prepared from whole zebrafish larva homogenates

Zebrafish liver microsomes

Microsomes prepared from whole adult zebrafish homogenates

11

Chapter 1: General introduction

13

1 Preface

The thalidomide tragedy in the late fifties and early sixties lead to the

obligatory use of at least two animal species, i.e. a rodent and a non–rodent

model, in the in vivo developmental toxicity studies for safety testing of drugs

prior to exposing women of childbearing potential. In view of the 3Rs concept

within laboratory animal sciences—Replacement, Reduction and Refinement—

the zebrafish embryo gained interest as an alternative model for developmental

toxicity studies since the zebrafish is not considered to be a test animal until it

reaches the stage of independent feeding, i.e. 120 h post–fertilization (EU

Directive 2010/63/EU, p. 39) [1]. The zebrafish embryo developmental toxicity

assay is currently being explored for regulatory acceptance, as the exposure

window covers largely zebrafish organogenesis. However, the externally

developing zebrafish embryos depend on their intrinsic biotransformation

capacity for the detoxification or bioactivation of xenobiotics, which is in contrast

to mammalian embryos relying on maternal metabolism. This difference is

particularly relevant in view of proteratogens, which need to be bioactivated to

exert their teratogenic potential. Therefore, the xenobiotic–metabolizing capacity

of zebrafish embryos and larvae has been investigated in several studies over the

last two decades. Since the overall results from these studies are contradictory,

the main aim of this thesis is to contribute to a better understanding of the

ontogeny of drug disposition in zebrafish. To this end, the thesis investigates the

ontogeny of cytochrome P450 (CYP) enzymes on mRNA as well as on activity

level, and to a lesser extent also the expression levels of Phase II enzymes and a

transporter protein, i.e. abcb4, at different time–points during zebrafish

development.

In order to correctly interpret the findings of the doctoral project, the current

chapter provides relevant background information on the zebrafish (embryo)

model as well as on the processes and enzymes that are involved in drug

disposition. To start with, this chapter describes the relevance of the use of

zebrafish embryos as an alternative animal model in developmental toxicity

studies. Subsequently, the general characteristics of the zebrafish model are

described as well as its embryonic and larval development with special emphasis

14

on the ontogeny of pivotal drug–metabolizing organs such as the digestive

system. The latter information is pivotal with regards to the localization of CYP

activity during zebrafish development. The last part of the introduction

elaborates on drug disposition, also called ADME (Absorption, Distribution,

Metabolism and Excretion), with special emphasis on Metabolism since the

ontogeny of zebrafish biotransformation capacity underlies the subject of the

doctoral project. In this section on drug disposition, a comparison is made

between the traditional mammalian model and the zebrafish.

2 Developmental toxicity studies: an overview

2.1 Terminology

The ICH S5 guideline on reproductive toxicology aims to provide key

considerations for developing a testing strategy to identify hazards and

characterize reproductive and developmental risk for human pharmaceuticals [2].

According to the test guideline TG 414 of the Organization for Economic Co–

operation and Development (OECD) [3], developmental toxicity is defined as

‘the study of adverse effects on the developing organism that may result from

exposure prior to conception, during prenatal development, or postnatally to the

time of sexual maturation. The major manifestations of developmental toxicity

include 1) death of the organism, 2) structural abnormality, 3) altered growth, and

4) functional deficiency’. The term developmental toxicity is often used

interchangeably with teratology (< teratos, meaning monster), which represents

the study of abnormal development in an embryo or fetus due to exposure to a

particular agent, a so–called teratogen, during gestation [4]. Developmental

toxicity/teratology is part of a more general term, i.e. reproductive toxicity, which

includes ‘harmful effects on the offspring as well as an impairment of male and

female reproductive functions or capacity’ [5].

2.2 Historical and legal context

Already in the second half of the nineteenth century, Camille Dareste was

able to induce congenital malformations in chicken embryos by exposing them to

15

environmental factors such as hyperthermia, hypothermia and anoxia. This study

made the French zoologist one of the pioneers of developmental toxicity studies

(reviewed by [6]). During the first half of the twentieth century, more studies were

performed to assess the influence of environmental factors such as x–radiation

and dietary deficiency on mammalian development (reviewed by [6]), [7].

However, in spite of the alarming results of several developmental toxicity

studies, the contemporary community remained convinced that the placental

barrier protected human embryos from all external influences (reviewed by [6]).

This conviction was ended by the thalidomide tragedy in the late 1950s and early

1960s, which proved that human embryos are indeed susceptible to

environmental insult. Although high doses of thalidomide did not have toxic

effects in adult mammals and studies in pregnant rats were found to be safe

(reviewed by [8,9]), the drug caused phocomelia, i.e. malformations of arms and

legs, and craniofacial malformations in human newborns when it was taken

during the first trimester of pregnancy as a treatment of morning sickness (Figure

1). The first known case of the thalidomide disaster was a girl who was born

without ears in 1956 in Stolberg, Germany. Her father, who worked at the

Grünenthal company which synthesized thalidomide, received samples of the

new drug for his pregnant wife [10]. A few years later, an Australian obstetrician

and a German pediatrician began to notice birth defects in babies whose mothers

had used thalidomide during pregnancy and alerted the medical community to

the teratogenic effects of the compound [11]. Nevertheless, the drug remained on

the market with devastating consequences worldwide since a retrospective study

showed that around 10,000 to 12,000 babies were born with serious birth defects

during the late 1950s and early 1960s (reviewed by [12,13]). In order to prevent

such a tragedy from happening in the United States, U.S. drug laws were

reformed by the Kefauver–Harris Drug Amendments in 1962 [14] which gave the

U.S. Food and Drug Administration (FDA) new authorities, e.g.: prove

effectiveness of drug products by well–controlled clinical studies before they go

on the market and report any serious side effects; set good manufacturing

practices for industry, including regular inspections of production facilities; and

requirement of FDA approval before marketing of the drug in the U.S. (website:

16

https://www.fda.gov). In general, the U.S. FDA, which was founded in 1906,

regulates the safety, efficacy, and security of human and veterinary

pharmaceuticals as well as the safety of national food supply and cosmetics

(website: https://www.fda.gov).

Besides a reformation of the drug laws, the thalidomide disaster also led to

the introduction of 3–Segment Studies by the FDA, which implied 1) study of

fertility and reproductive performance, 2) study of in utero development (in vivo

teratology study) and 3) perinatal and postnatal study (reviewed by [6]). The in

vivo teratology study required at least two species, i.e. in most cases a rodent

(mouse, rat) and a non–rodent (rabbit) model since significant interspecies

differences were found in the types of effects after thalidomide exposure

(reviewed by [6]). The choice of the rabbit as a non–rodent model is not surprising

since it showed to be sensitive to thalidomide, giving the same type of

malformations (limb defects) as humans [15,16], whereas the rat did not show

malformations with this human teratogen (reviewed by [8]). Furthermore, in

teratology studies, treatment/exposure should cover the period of organogenesis

for the respective species, i.e. days 6–17 of gestation for rats and days 6–19 of

gestation for rabbits, since it represents the sensitive period for teratogenic effects

(reviewed by [6]).

Figure 1. A picture of so–called ‘Thalidomide Babies’: children with thalidomide–

induced phocomelia. Source: http://www.chm.bris.ac.uk/motm/thalidomide/first.html.

https://www.fda.gov/

https://www.fda.gov/

17

Pharmaceuticals for human use need to undergo reproductive toxicity testing

in order to evaluate the risk to women of childbearing potential who may be

treated with the compound (reviewed by [17]). In the United States as well as in

Europe, developmental toxicity studies are currently regulated by federal

agencies, i.e. the FDA in the U.S. (see above) and the European Medicines Agency

(EMA) in Europe. The EMA was founded in 1995 by the European Union with

the aim of harmonizing the work of existing national regulatory bodies. The

agency is responsible for the scientific evaluation, supervision and safety

monitoring of pharmaceuticals in the EU (website: http://www.ema.europa.eu).

Since 1970, the U.S. Environmental Protection Agency (EPA) has been

involved in the assessment of chemicals and environmental pollutants for

adverse health outcomes, including developmental toxicity (website:

https://www.epa.gov). The European Chemicals Agency (ECHA), founded by

the EU in 2007, is the driving force among regulatory authorities in implementing

the EU's chemicals legislation called Registration, Evaluation, Authorization and

Restriction of Chemicals (REACH) (website: https://echa.europa.eu). The REACH

regulation requires companies to register all chemicals that are manufactured in

or imported into the EU by generating a dossier containing data on

physicochemical characteristics as well as (eco)toxicological properties, hazards

and risks of chemical substances. Furthermore, REACH encourages the

identification of chemicals that may pose unacceptable hazards to human health

and/or the environment in order to reduce or restrict their usage [18,19]. Besides

REACH, the Organisation for Economic Co–Operation and Development

(OECD), an intergovernmental economic organisation founded in 1961, plays a

major role in ecotoxicology. The organisation, which consists of representatives

from 36 countries, sets international standards on a wide range of topics, from

agriculture over education and employment to tax and trade. One of its

accomplishments is the set–up of OECD Test Guidelines (TG) for the testing of

chemicals which represents a collection of internationally agreed testing methods

used by governments, industry and independent laboratories to assess the safety

of chemical products with attention to animal welfare (website:

http://www.oecd.org).

http://www.ema.europa.eu/

https://www.epa.gov/

https://echa.europa.eu/

http://www.oecd.org/

18

2.3 Traditional and alternative test systems in developmental toxicity

studies

The traditional developmental toxicity studies are performed in vivo in

pregnant mammals by exposing them to the test compound during the period of

organogenesis after which the near–term fetus is examined for skeletal, visceral

and external malformations (reviewed by [20]). In these studies, rat, and to a lesser

extent mouse, are the most commonly used rodent species in the in vivo

developmental toxicity studies, because of their well–understood biology and

pharmacokinetics and the large amount of historical background data about these

species [2]. As mentioned above, the thalidomide tragedy emphasized the

necessity to use of a second, non–rodent, species in developmental toxicity

studies since substantial interspecies differences in effects were detected after

exposure to this human teratogen (reviewed by [6]). However, the actual reason

behind the dissimilarities in toxic effects is unknown. Since the rabbit fetus

showed the same type of malformations as humans after exposure to thalidomide

and since this species is suitable for artificial insemination due to the easiness of

semen collection, it is often the non–rodent species of choice [15,16]. However, in

vivo developmental toxicity studies have some major drawbacks: they are time–

consuming, labour–intensive, expensive and they have numerous interfering

factors such as nutritional state of the dam, placental function and variability in

developmental age of embryos within or between litters (reviewed by [20]).

Besides these drawbacks, the increasing awareness of animal welfare after the

introduction of the 3Rs by Russell and Burch [21] led to the gradual

implementation of alternative developmental toxicity test systems in the 1970s

and 1980s (reviewed by [20]). The alternative assays are characterized by their

high–throughput capabilities, time– and cost–efficiency and the reduction of

interfering factors. The alternative test systems have the potential to be used as

screening assays in the early phase of the drug development process to select

compounds for further in vivo testing in a mammalian model. With regards to

screening assays, a positive response in an alternative test system would indicate

a potential human hazard while a negative response would not indicate the

absence of a hazard. Hence, a negative response in a screening assay would be

19

followed by in vivo mammalian testing, while a positive response would require

no further testing unless the investigator is concerned about a potential false

positive response. Therefore, the alternative assays cannot replace the testing of

compounds in the more complex mammalian model, but they reduce in vivo

mammalian studies [2]. Besides the pharmaceutical industry, regulations and

organizations that are involved in the surveillance of environmental safety such

as REACH and OECD also encourage the use of alternative test methods in

developmental toxicity studies [18]. The following test systems are considered as

alternatives for the traditional in vivo developmental toxicity assays:

In silico test systems

In silico test systems are computer simulated models such as

(Quantitative) Structure–Activity Relationship ((Q)SAR) models or

physiologically based pharmacokinetic (PBPK) modelling. PBPK

modelling can predict the absorption, distribution, metabolism and

excretion (ADME) properties of compounds, whereas (Q)SAR models

can predict the biological activity of a compound based on its

physico–chemical properties. Because of their predictive value, both

approaches have been used in the drug discovery process since the

early 2000s (reviewed by [12]), [22]. More recently, the use of (Q)SAR

models in predicting in vivo responses in developmental and

reproductive toxicity studies is being explored, i.e. the so–called

Adverse Outcome Pathways (AOPs) [23]. An AOP is an analytical

construct that describes a sequential chain of causally linked events

(key events (KE)) at different levels of biological organisation that

lead to a toxicological effect (adverse outcome (AO)) (Figure 2). In

silico models may be used to investigate the molecular initiating event

(MIE) of an AOP (Figure 2). In this respect, the role of AOPs is twofold

since they reduce in vivo mammalian studies and they provide

information on the mechanism that causes a toxicological effect. A

guidance document of the OECD provides a detailed description of

how AOPs are to be developed, reviewed and published [24].

20

Figure 2. Schematic representation of Adverse Outcome Pathways (AOPs).

In vitro test systems with mammalian–derived tissue or cells

o Limb bud micromass culture

This technique, which has been used since the 1970s, involves

culturing of limb buds which are dissected from fore– or

hindlimbs of rat, mouse, rabbit or chicken embryos. After

culturing for a designated period, morphological criteria such

as the overall appearance, size and shape of the limb bud are

used to assess the degree of normal development of the

explant (Figure 3). Chondrogenesis is considered as an

important parameter to evaluate teratogen–induced

malformations of the limb buds such as phocomelia and

syndactyly. Cartilage development can be microscopically

visualized by staining of the limb bud with alcian blue or

toluidine blue (reviewed by [25]).

o Embryonic Stem cell Test (EST)

The EST includes a culture of embryonic stem cells that were

isolated from blastomeres of the early mouse embryo. These

cultured pluripotent cells develop into differentiated cells of

all three primary germ layers, which has made the EST a

widely used system to study gene expression patterns and

cellular developmental processes during early embryogenesis

(reviewed by [20]). The EST is based on the assessment of three

toxicological endpoints after 10 d of chemical exposure: 1) the

inhibition of differentiation into beating cardiomyocytes, 2)

the cytotoxic effects on differentiating embryonic stem cells

and 3) the cytotoxic effects on 3T3 (“3–day transfer, inoculum

Molecular level

In silico / in vitro

Organellelevel

In vitro

Cellular level

In vitro

Tissue level

In vitro

Organlevel

In vivo

Organismlevel

In vivo

Population

Field studies

MolecularInitiatingEvent (MIE)

Adverse Outcome (AO)Key Events (KE)

21

3x105 cells”) fibroblasts [26]. The EST has been used as an

alternative for in vivo developmental toxicity testing since the

1990s [27].

o Whole Embryo Culture (WEC)

The post–implantation WEC typically uses rodent embryos

which are usually explanted with their visceral yolk sac at

presomite or early somite stages (day 9 or 10 of gestation). The

embryos are cultured on a rotating platform in an oxygenated

mixture of serum and culture medium for 24–48 h during the

sensitive stage of organogenesis. Since the 1980s, the WEC has

been used as a test system for developmental toxicants. In the

WEC, the following endpoints are evaluated: 1) embryonic

death, 2) growth retardation, 3) structural and functional

abnormalities to a.o. neural tube closure, heart development

and the formation of branchial arches. In comparison with the

limb bud micromass culture and the EST, the WEC more

closely resembles the in vivo developmental toxicity assay

(reviewed by [28,29]. However, two days of culturing only

represents a small part of the organogenesis period (6–17 days

for rats and 6–19 days for rabbits, in function of strain [3]),

which results in nondetection of insults to developmental

events outside this period such as palate closure and the

formation of digits [29].

22

Figure 3. Limb bud micromass culture of rat (A and B) and rabbit (C and D) after

treatment with thalidomide. Pictures on the left represent 6–carboxy–2',7'

dichlorofluorescin diacetate (DCF) staining for oxidative stress and pictures on the right

represent Nomarski images. The arrow indicates the nucleus. The figure is reproduced

from Hansen et al. (2002) [30].

Non–mammalian test systems

o Alternatives with invertebrates

Simultaneously with the WEC and EST, invertebrates such as

Drosophila melanogaster or fruit fly and Hydra attenuata or

freshwater polyp came into the picture as alternative animal

models in developmental toxicity studies. Drosophila

embryo(s) (cell cultures) have been used to assess the

teratogenic effect of compounds on muscle and/or neuron

differentiation, heat shock proteins, neurotransmitter levels as

well as morphological development (reviewed by [31]).

Developmental toxicity assays with Hydra attenuata use

artificial embryos which consist of dissociated terminally

differentiated and pluripotent cells of adult Hydra. The assay

is based on the adult/developmental (A/D) ratio that expresses

the relationship of toxic doses in the adult and offspring

(reviewed by [31]) [32]. Although the genetic elements of

23

development between invertebrates and vertebrates are

highly conserved, most developmental toxicants seem to act

at the level of cytoplasmic processes, which seemingly vary

between species. Moreover, vertebrates and invertebrates

differ with regards to embryogenesis. Hence, the substitution

of vertebrates by invertebrates in developmental toxicity

studies has been reduced to a minimum (reviewed by [31]).

o Alternatives with vertebrates

Alternative test systems with non–mammalian vertebrates

most commonly involve amphibians such as aquatic frogs

(Xenopus spp.) and fish such as zebrafish (Danio rerio), Japanese

medaka (Oryzias latipes) and fathead minnow (Pimephales

promelas) (Table 1). These non–mammalian vertebrates share

some characteristics which make them suitable as alternative

test systems: 1) fish and amphibians are evolutionary closer to

humans compared to invertebrates, 2) they produce eggs in

large quantities, 3) the embryos develop externally and grow

rapidly which makes them suitable for high–throughput

screening in a multiwell–format (Figure 4), 4) the embryos and

their genes are easy to manipulate, 5) the cost per embryo is

low [33,34] and 6) non–mammalian vertebrates are not subject

to Directive 2010/63/EU on the protection of animals used for

scientific purposes until they reach the stage of independent

feeding (EU Directive 2010/63/EU, p. 39) [1]. Xenopus, medaka

and fathead minnow embryos are mainly used in

developmental toxicity studies with regards to environmental

contaminants, whereas screening assays with zebrafish

embryos are used in environmental toxicology, i.e. the fish

embryo acute toxicity test (FET) [35], as well as in the drug

discovery/development process, i.e. the zebrafish embryo

developmental toxicity assay (ZEDTA) [36,37] (Table 1).

24

Regulatory acceptance of the FET is under consideration as an

alternative for the fish acute toxicity test (TG 203 OECD, 1992)

[38]. Since the zebrafish has a short generation time, a rapid

embryonic development, a highly transparent chorion and

produces a high number of eggs per spawning act, this fish

species has been the first choice for alternative embryo toxicity

testing. Hence, the zebrafish (embryo) plays a key role in this

thesis. In part 2 of the general introduction, we will further

elaborate on the husbandry, anatomy, physiology and

embryonic development of this animal species.

A major disadvantage of the in vitro test systems such as WEC and EST and

of the alternative test systems with ex utero developing vertebrates is the lack of

maternal metabolism [12,28,39]. Hence, the alternative vertebrate models such as

the zebrafish depend on their intrinsic biotransformation capacity for the

detoxification and/or bioactivation of xenobiotics. With regards to their use in

developmental toxicity studies, knowledge of the ontogeny of the

biotransformation capacity of these alternative models is pivotal, which underlies

the subject of the doctoral project. Part 3 of the general introduction will focus on

ADME (Absorption, Distribution, Metabolism and Excretion) in zebrafish with

special emphasis on metabolism, but first the general characteristics of the

zebrafish model will be discussed.

25

Table 1. A comparison between non–mammalian vertebrates that are most commonly used as alternative test systems for

developmental toxicity.

FET: Fish Embryo Acute Toxicity Test; FETAX: Frog Embryo Teratogenesis Assay–Xenopus; hpf: h post–fertilization; LAGDA: Larval Amphibian Growth and Development Assay;

ZEDTA: Zebrafish Embryo Developmental Toxicity Assay.

AFRICAN CLAWED FROG

(XENOPUS LAEVIS)

JAPANESE MEDAKA

(ORYZIAS LATIPES)

FATHEAD MINNOW

(PIMEPHALES PROMELAS)

ZEBRAFISH

(DANIO RERIO)

REF.

ORIGIN Sub–Saharan Africa Japan, China, South Korea Central North America India, Burma, Malakka,

Sumatra

[34,39,40]

ENVIRONMENT Aquatic Aquatic Aquatic Aquatic

WATER TEMP. IN

LABORATORY

24 ± 2 °C 26 ± 1 °C 26 ± 1 °C 27 ± 1 °C [34,39,41,42]

# EGGS PER MATING 500–3000+ 20–40 100–250 100–200 [34,40,43]

GENERATION TIME 6–10 months 2–3 months 4–5 months 3–4 months [43,44]

EXTERNAL

EMBRYONIC

DEVELOPMENT

Yes

Yes Yes Yes

APPEARANCE OF

EGGS/EMBRYOS

3–layered jelly envelope

Developing larvae and

tadpoles become

gradually transparent

Stable chorion with spiny

hooks -> adhesion to anal fin

of female

Moderately transparent

Chorion only hardens in

multicellular stage

Sticks to surfaces

Transparent

Stable chorion

Non–sticky

Highly transparent

[33,34,43,44]

EMBRYONIC

DEVELOPMENT AT 26

°C

± 19 hpf: somite dev.

± 44 hpf: heart–beat visible

± 50 hpf: hatching



± 160 hpf: hatching



± 120 hpf: hatching

±18 hpf: somite dev.


± 72 hpf: hatching

[33,34]

DEVELOPMENTAL

TOXICITY TEST

TYPES: DURATION +

TOXICOLOGICAL

ENDPOINTS

FETAX: 96 h; mortality,

growth inhibition and

malformations

LAGDA: 16 w; mortality,

growth, metamorphosis,

reproductive maturation

FET: 232 h; mortality

(coagulation), tail

detachment, somite

development, heart beat/

blood circulation, hatching

FET: 120 h; mortality

(coagulation), tail

detachment, somite

development, heart beat/

blood circulation, hatching

FET: 96 h; mortality, tail

detachment, somite

development, heartbeat,

hatching

ZEDTA: 120 h; viability,

morphology of somites,

neural tube, notochord, tail,

fins, heart, facial structures

[34,35,37,39,45]

26

Figure 4. A comparison between the life cycles of Xenopus laevis (A), Danio rerio (B) and Mus musculus (C).

Source: http://www.mun.ca/biology/desmid/brian/BIOL3530/DB_03/DBNVert1.html.

http://www.mun.ca/biology/desmid/brian/BIOL3530/DB_03/DBNVert1.html

27

3 The Zebrafish

3.1 Zebrafish in their natural habitat

Zebrafish ((Brachy)danio rerio) are small freshwater fish, which belong to the

Cyprinidae family (reviewed by [40]). The fish are indigenous to South Asia and

are broadly distributed across parts of India, Bangladesh, Nepal, Myanmar and

Pakistan (reviewed by [46]). Zebrafish inhabit still or slow–moving waters and

shallow ponds, which are often connected to rice cultivation (reviewed by [40,46]).

Indeed, the name Danio derives from the Bengali name ‘dhani’, which means ‘of

the rice field’ [47]. The natural range of temperatures in which zebrafish live is

from as low as 6 °C in winter to over 38 °C in summer, which classifies them as

eurythermal, i.e. having a high temperature tolerance. Adult zebrafish are

characterized by five to seven blue longitudinal stripes extending from behind

the operculum into the caudal fin (reviewed by [40]). Although sexual

dimorphism in zebrafish is rather subtle, males have a more streamlined body

shape while gravid females have a more rounded shape as well as a small

urogenital papilla or ovipositor in front of the anal fin origin [48] (Figure 5). The

Standard Length (SL; from the tip of the snout to the origin of the caudal fin) of

adult zebrafish rarely exceeds 40 mm (Figure 5). The natural diet of the

omnivorous zebrafish consists primarily of zooplankton and insects. Although

the mean lifespan of domesticated zebrafish is around 42 months, the lifespan of

zebrafish in the wild is not well documented [49].

28

Figure 5. Representation of a female (upper picture) and a male (lower picture)

zebrafish. The Standard Length (SL), i.e. the length from the tip of the snout to the

origin of the caudal fin, is represented in the upper picture and has an average value

of 30 mm . The SL is used sometimes as an indicator of developmental progress [50].

Both pictures are obtained from the Zebrafishlab website: © 2018 Zebrafishlab.

3.2 Zebrafish in a laboratory setting

In a laboratory setting, adult zebrafish are maintained in aquaria with a

filtering system or in a recirculating aquaculture system at a 14/10 h light/dark

cycle with an average maximum fish density of seven fish per liter [51]. However,

the maximum fish density may vary since it depends on the housing system, the

age of the fish and the amount of feeding. Although zebrafish exhibit a tolerance

for a wide range of environmental conditions, it is pivotal to maintain optimal

water quality parameters for zebrafish in a laboratory setting. Indeed, adult

zebrafish maintained under sub–optimal conditions need more energy to

maintain their homeostasis, rather than for growth, reproduction and immune

function [46]. The following gives an overview of the optimal water quality

parameters for laboratory zebrafish.

29

Temperature

A temperature range of 24–30 °C is considered to be the preferred

range for laboratory zebrafish [52]. However, in most laboratories,

temperatures of 26–28.5 °C are being used. Moreover, 28.5 °C is the

most commonly used temperature for breeding and embryonic

development since it is thought to be close to the optimal temperature

for zebrafish growth [51].

pH

Most zebrafish facilities use a pH of between 7.0 and 8.0, since this is

the optimal pH range for the bacterial flora in the biological filter (see

below) and it is within the general range recommended for freshwater

fish [46].

Hardness

Water hardness is a measure of the quantity of divalent ions,

primarily calcium and magnesium [46]. Besides their role in

osmoregulation, zebrafish require these ions for a number of

physiological processes. Since tap water has varying degrees of

hardness, zebrafish facilities commonly use distilled or reverse

osmosis water to which calcium and magnesium salts are added to

bring hardness values to the recommended range for freshwater fish,

i.e. 75–200 mg/L CaCO3.

Conductivity/Salinity

The conductivity is a measure of water’s capability to conduct an

electrical current. Conductivity is directly related to the concentration

of dissolved ions in the water, i.e. salinity [53]. In most zebrafish

facilities, salinity is not measured directly, but is instead derived from

the conductivity measurement. Freshwater fish in general are

hyperosmotic to the media in which they live and thus tend to gain

water and lose salts by diffusion across the gills and skin [46]. In order

to minimize the energetic cost to maintain their internal water and salt

balance, zebrafish exhibit a preferred range of conductivity levels, i.e.

300–1000 µS [51].

30

Nitrogenous wastes

Zebrafish are ammonotelic organisms since they excrete nitrogen as

ammonia (NH3), which is highly toxic for the fish. Besides the

excretion of ammonia across the branchial epithelium and in feces of

the zebrafish, it is also produced by decomposing organic matter

(Figure 6). The highly toxic ammonia (NH3) and the non–toxic

ammonium (NH4+) exist at equilibrium in artificial aquatic systems. In

zebrafish housing systems with a biological filter,

ammonia/ammonium is eliminated by Nitrosomonas bacteria, which

oxidize the highly toxic compound into nitrite (NO2-) (Figure 6). Since

nitrite is as well toxic to zebrafish, this intermediate product must be

eliminated by Nitrobacter bacteria in the biological filter via oxidation

into the non–toxic nitrate (NO3-) (Figure 6). Although nitrates are

generally not toxic to zebrafish, prolonged exposure of the fish to high

nitrate levels may adversely affect their health [46]. In the presence of

water plants, nitrates are incorporated into the plant proteins.

However, most housing systems in zebrafish laboratories do not

contain plants because of the dirt they produce. Hence, the water in

aquaria or recirculating systems should be renewed to remove excess

of nitrate (Figure 6). The recommended values for nitrogenous wastes

in a zebrafish housing system are as follows: NH3 0 mg/L, NO2- < 0.3

mg/L and NO3- ≤ 12.5 mg/L.

31

Figure 6. Nitrogen cycle in a fish aquarium. Nitrosomonas and Nitrobacter bacteria are

maintained in the biological filter of the zebrafish housing system.

Zebrafish in captivity are preferably fed live diets or frozen food,

supplemented with artificial diets. Live diets such as Artemia and Paramecium or

frozen food such as Artemia, bloodworms (Chironomid larvae) and Daphnia are

visually attractive to zebrafish and are highly digestible. The artificial diets can

be used to deliver specific nutrients that may not be present in live diets or frozen

food [46]. Although the specific nutritional requirements of zebrafish are not well

understood, several studies showed that live diets or frozen diets are essential to

maintain good growth and survival rates (reviewed by [46]). With regards to

feeding procedures, there are two general approaches that are used in a zebrafish

housing system, i.e. feeding according to the “five–minute rule” and body weight

feeding. The five–minute rule implies presenting the fish the amount of food

which they can fully consume within five minutes. However, this less accurate

method may lead to under- or over feeding of the fish. In case of feeding by body

weight, a fixed percentage of zebrafish body weight is provided each day. The

amount of food depends on the age of the fish, i.e. 50–300% for larval fish and 1–

10% for adults [46].

Decomposing organic matter (i.e., dead fish, uneaten food, plant fragments)

Ammonia/Ammonium(NH3/NH4

+)

Waste products

Food

Oxidation byNitrosomonas bacteria

Nitrites(NO2

-)

Oxidation byNitrobacter bacteria

Nitrates(NO3

-)

Incorporatedinto plant protein

Water change

32

For detailed information regarding zebrafish maintenance as well as feeding

and diet of larval and adult fish, we refer to the Materials and Methods section of

Chapter 3 and 4 of the thesis.

3.3 Reproduction and breeding

During early life, both male and female zebrafish have undifferentiated

ovary–like gonads, which start developing at approximately 12–14 days post–

fertilization (dpf). Later during development, around 70 dpf, the ovary–like

gonads transform into well–differentiated ovaries or testes according to a 1:1 sex

ratio [54,55]. The exact time period when female and male zebrafish begin to

develop differentiated ovaries and testes respectively may vary. For instance,

body growth, which depends on environmental factors such as feeding

conditions, stocking density and water temperature, may cause inter–laboratory

differences in the timing of gonadal differentiation in zebrafish. Besides the body

growth, the use of different zebrafish strains may also give rise to variations in

the timing of sexual differentiation [54]. Testosterone and estrogen are considered

natural inducers of sex in fish [56]. Cytochrome P450 aromatase, which

aromatizes testosterone into estrogen, shows a sexually dimorphic expression

pattern in zebrafish [57,58]. Consequently, the aromatase enzyme may play an

important role in the transformation of undifferentiated gonads into testes or

ovaries [57,59].

Although zebrafish reach sexual maturity at 10–12 weeks, the optimal

breeding age is between 7 and 18 months [51]. In laboratory conditions, zebrafish

breed all year round, whereas in nature spawning mainly occurs during the

monsoon season, i.e. from June to August. Moreover, zebrafish are photoperiodic

in their breeding since they commonly spawn within the first few hours of

daylight. In a zebrafish housing system with a 14/10 h light/dark cycle spawning

starts within the first minute of exposure to light following darkness, continuing

for about an hour [60]. However, spawning does not seem to be strictly limited to

this time period [46].

33

Ovulation in female zebrafish is induced by exposure to male gonadal

pheromones. After ovulation, females release pheromones which in turn trigger

mating behavior in males [40]. This mating behavior immediately elicits

oviposition by females and external fertilization of the eggs [46]. A single female

may produce clutches of several hundred eggs in a single spawning [40]. Fish of

12 months old have an inter–spawning interval of 1.9 days, which increases with

age [61]. However, with the aim of producing a maximal number of embryos in a

laboratory setting, an inter–spawning interval of about six or seven days is

recommended [51].

Most zebrafish breeding facilities currently use a small breeding tank with a

mesh or grill bottom, which is placed inside a larger tank filled with water (Figure

7). Other breeding facilities use spawning nets, which are put inside the aquaria.

Fish are added in pairs or in small groups to the spawning tank or net the evening

before mating. When the fish spawn the next morning, the eggs, which have a

diameter of approximately 0.7 mm, fall through the bottom of the breeding tank

or net and are thereby protected from being eaten by the adults [40]. The eggs are

collected by siphoning them up from the bottom of the tank 30–40 min after the

lights turned on in the zebrafish housing system[51]. For a detailed description of

the collection and raising of zebrafish embryos, we refer to the Materials and

Methods section of Chapter 3 and 4 of the thesis.

34

Figure 7: Zebrafish breeding tank consisting of a solid external tank, an internal

tank with perforated bottom, a divider to separate male and female fish and a lid.

Source: https://www.tecniplast.it/uk/product/07l-tank.html.

3.4 Embryonic and larval development

Zebrafish embryos can be raised within a temperature range of 26°C to

28.5°C. As a result, the time–points (expressed in hours or days post–fertilization

(hpf or dpf)) at which developmental events occur, may vary between the

different laboratories. This section describes zebrafish embryonic and larval

development at 28.5 °C, which is the best temperature for growth [51], according

to the classification of Kimmel et al. (1995) [62] and Nüsslein–Volhard et al. (2002)

[63], respectively (Table 2).

The period of organogenesis, i.e. development of cardiovascular,

gastrointestinal, urogenital and central nervous system, coincides with

embryonic development (Table 2). Zebrafish embryos depend on the yolk for

nutrition until the onset of exogenous feeding, which occurs during the larval

period. Larval development officially begins at 72 hpf (Table 2), which is

approximately one day after hatching [63]. However, embryo–larval transition

occurs gradually and implies the period between the onset of exogenous feeding

and complete yolk absorption. Although yolk absorption is complete around 7

dpf, most research facilities already start feeding the zebrafish around 96 hpf in

order to stimulate their food–seeking behavior [64]. During the larval period,

35

zebrafish grow approximately 2 mm/week and most vital organs, i.e. brain, heart

and gastro–intestinal system have already been fully developed and are

functional at the beginning of this period (Table 2). Nevertheless, zebrafish

maturation is highly asynchronous due to environmental fluctuations as well as

individual differences. The larval–juvenile transition reflects the period of

metamorphosis in which larval morphology is transformed into that of a juvenile,

which is characterized by scale development, ossification of the skull bones and

maturation of fins and fin rays. The juvenile period begins around 30 dpf and

includes the development of the definite kidney, i.e. mesonephros, and the

gonads (Table 2). However, zebrafish only reach sexual maturity around 90 dpf,

when they are considered adults [63,64].

In view of endpoint evaluation in developmental toxicity studies and

considering the localization of biotransformation activity in zebrafish embryos

and larvae (Chapter 2: aims of doctoral project), knowing the normal

organogenesis in zebrafish is a prerequisite. Hence, in the following section we

provide a concise summary of the critical periods in the development of some

major zebrafish organs (Table 2). However, in view of this project, we will further

elaborate on the organs that have a pivotal role in drug metabolism. Moreover,

important differences between zebrafish and mammalian organ development are

briefly outlined.

36

Table 2: An overview of key events during embryonic, larval and juvenile development of the zebrafish.

DEV. PERIOD TIMING DEVELOPMENTAL EVENTS ORGAN DEVELOPMENT REFERENCES

ZYGOTE PERIOD 0 – 0.75 hpf Segregation of cytoplasm towards the

animal pole

None [62]

CLEAVAGE

PERIOD

0.75 – 2.25

hpf

Cleavage of cytoplasm (mitosis)

to form blastomeres

None [62]

BLASTULA

PERIOD

2.25 – 5.25

hpf

-Further cleavage of cytoplasm

-Development of yolk syncytial layer

-Beginning of epiboly (i.e., cell movements)

None [62]

GASTRULA

PERIOD

5.25 – 10.33

hpf

-Epiboly continues

-Development of epiblast and hypoblast

-Notochord rudiment

-Brain rudiment (neural plate)

[62]

SEGMENTATION

PERIOD

10.33 – 24 hpf -Development of somites

-Elongation of embryonic tail

-Development of brain neuromeres

-Bilateral pronephros rudiment

-Development of otic vesicle + otoliths

-Development of optic vesicle + lens

-Development of linear heart tube

[62,65,66]

PHARYNGULA

PERIOD

24 – 48 hpf -Straightening of head

-Development of pharyngeal arches

-Pigmentation of the skin (melanophores)

-Paired pectoral fin rudiments

-Development of heart atrium and ventricle

-Development of vascular system

-Development of gut tube

-Liver primordium

-Pancreas primordium

-Swim bladder rudiment

[62,66-71]

Developmental periods are based on the classification by Kimmel et al. (1995) [62] and Nüsslein–Volhard et al. (2002) [63].

Hpf: hours post–fertilization. In this thesis, developmental stages of the organogenesis period are represented as hours post-fertilization

(hpf). Later developmental stages are shown as days post-fertilization (dpf).

37

Table 2: Continued.

DEV. PERIOD TIMING DEVELOPMENTAL EVENTS ORGAN DEVELOPMENT REFERENCES

HATCHING

PERIOD

48 –72 hpf -Embryo out of chorion

-Pectoral fin development

-Glomerular blood filtration in pronephros

-Development of esophageal lumen

-Development of endocrine pancreas

-Liver growth and vascularization

-Formation of posterior chamber swim bladder

[62,67-72]

LARVAL

PERIOD

72 hpf – 30

dpf

-Active feeding

-Free swimming

-Startle response

-Growth

-Opening mouth and anus

-Development of pharyngeal lumen

-Functional digestive tract and liver

-Development of exocrine pancreas

-Development pronephric nephron complete

-Onset of mesonephric nephron development

-Air inflation of both swim bladder chambers

-Yolk absorption

[63,67,68,70,71,73-

75]

JUVENILE

PERIOD

30 – 90 dpf -Beginning of scale development

-Growth -Mesonephros (adult kidney) replaces

pronephros

-Development of well–differentiated ovaries

and testes

[54,75]

Developmental periods are based on the classification by Kimmel et al. (1995) [62] and Nüsslein–Volhard et al. (2002) [63].

Hpf: hours post–fertilization; dpf: days post–fertilization. In this thesis, developmental stages of the organogenesis period are represented

as hours post-fertilization (hpf). Later developmental stages are shown as days post-fertilization (dpf).

38

The digestive system

The development of the zebrafish digestive system shows some differences

with the development of the mammalian gastrointestinal system. The

mammalian gut comprises the pharynx, esophagus, stomach, intestine and colon

and the mammalian liver and pancreas develop from the gut as endodermal

buds. However, in zebrafish the digestive system is formed from individual

organ anlagen rather than a common endodermal tube. In zebrafish, the intestinal

primordium or gut develops first from the endodermal tube. After completion of

the intestinal anlage, the rostral digestive tract (pharynx and esophagus) begins

to form from endoderm rostral to the zebrafish gut. Moreover, the stomach and

caecum are absent in the zebrafish digestive system [70].

o The digestive tract: the pharynx, esophagus and intestine

According to the nomenclature of the zebrafish digestive system, the

alimentary canal is divided into the pharynx, esophagus, intestinal bulb, mid-

intestine and posterior intestine (Figure 8) [67,70,73].

In the early zebrafish embryo, endodermal progenitor cells are situated at the

blastoderm margin. During gastrulation, the endodermal cells start to migrate

medially and reach the embryonic midline during the mid-segmentation period,

around 18 hpf [76]. The endoderm shows no obvious histological organization

until 21 hpf when the endoderm between the fin buds and the anterior end of the

yolk extension becomes radially organized [70].

During the pharyngula period, the radial organization of endoderm extends to

the region dorsal to the posterior end of the yolk extension [70](Wallace et al.,

2003). This radial organization also implies that the endodermal cells, which

initially adopt a bilayer configuration, become separated by the formation of

several small cavities, which eventually coalesce to create a central lumen. As a

result, the lining endodermal cells are arranged into a monolayer configuration

[73]. However, zebrafish gut formation is different from mammals, since the

mammalian gut lumen develops by folding of the endoderm instead of by a

cavitation process (reviewed by [77]). By 34 hpf, an elliptical tube, i.e. the zebrafish

39

“gut”, has been formed between the fin buds and the region of the future anus.

However, the endoderm rostral to the gut shows no obvious histological

organization. In other words, the pharynx and esophagus are undeveloped at this

stage [70].

At the start of the hatching period, at 50 hpf, the esophageal primordium has

become visible [70]. By 52 hpf, cells from the lateral plate mesoderm have started

to encircle the zebrafish gut. These cells will proliferate and differentiate into

connective tissue, which will give rise to the muscle layers. Moreover, at around

52 hpf, enteroendocrine cells, which produce hormones such as somatostatin and

glucagon, appear in the posterior end of the intestinal primordium [73]. The

lumen of the anterior pharynx is formed between 54 and 58 hpf, whereas the

lumen of the posterior pharynx is still not obvious at this point in time. The

esophageal lumen is now clearly patent and contiguous with the rostral part of

the gut [70].

By the start of the larval period, at 74–76 hpf, the zebrafish mouth has opened

and the lumen of the posterior pharynx has become visible. Furthermore, the

endodermal cells throughout the length of the gut have polarized into columnar

epithelium and enteroendocrine cells are now scattered throughout the intestinal

primordium [70,73]. By 98–102 hpf, the opening of the anus has become visible.

Hence, the zebrafish intestinal primordium is now a completely open–ended

tube. At this time, the development of the digestive tract is also characterized by

the compartmentalization of the intestinal tube into the intestinal bulb, i.e. an

expansion of the lumen of the intestinal tube, mid-intestine and posterior

intestine (Figure 8) [73]. At approximately 120 hpf, the zebrafish has a functional

but still immature digestive tract [70,73,78].

By 28 dpf, the intestinal tract has adopted a coiled configuration. At this point

of the juvenile period, the digestive tract is essentially in its adult shape and

consists of three anatomical segments: an anterior segment, which runs

rearwards from the intestinal bulb; a middle segment, which is directed forwards;

and a posterior segment, which runs rearwards to the anus (Figure 8) [78]. Both

anterior and middle anatomical segments are supposed to be involved in lipid

40

absorption, whereas most of the posterior segment and the caudal extremity of

the posterior segment are supposed to be involved in protein absorption and

water/ion transport, respectively [78,79].

o The pancreas

Similar to mammals, the development of the zebrafish pancreas is

characterized by two distinct pancreatic anlagen, namely a dorsal posterior and

a ventral anterior anlage, which join to form the definitive pancreas [68]. The

zebrafish pancreas develops from endoderm rostral to the gut, whereas the

mammalian pancreas develops from dorsal and ventral protrusions of the gut,

which later join together [70,80]. In zebrafish, the dorsal posterior anlage

comprises the endocrine cells of the Islet of Langerhans, whereas the ventral

anterior anlage comprises the exocrine cells, the pancreatic duct and a small

number of endocrine cells [68].

During the segmentation period, at 16 hpf, two bilateral populations of pdx1

(pancreatic duodenal homeobox 1)–expressing cells start to converge to the

midline and eventually fuse at 18 hpf in order to form the dorsal posterior

pancreatic anlage at the level of the fourth somite around 24 hpf [68,81]. The

movement of the two pdx1-expressing populations to the midline is supposed

not to be specific to pancreas morphogenesis, but it is postulated to be part of the

early endoderm movement [68]. During the pharyngula period, by 30 hpf, leftward

looping of the zebrafish gut has dislocated the dorsal posterior anlage on the right

side of the gut [82,83].

There are two different hypotheses about the development of the exocrine

pancreatic anlage. According to a classical model, the ventral anterior anlage

begins to form as a ridge on the ventral side of the intestinal bulb primordium at

around 34 hpf (Figure 8). Because of the ventral ridge’s close proximity to the

lateral plate mesoderm and the yolk syncytial layer, these latter structures may

be considered potential sources for signaling molecules to stimulate anterior

pancreatic bud morphogenesis. By 40 hpf, the ventral anterior bud has extended

ventrally from the intestinal bulb primordium to the embryo’s right. The anterior

bud grows out towards the posterior anlage and by 44 hpf, the two anlagen have

41

come into contact (Figure 8) [68]. This hypothesis contrasts with an alternative

theory where the exocrine anlage is supposed to develop from endoderm rostral

to the zebrafish gut. According to this theory, the fusion of the exocrine anlage

with the gut and the subsequent anlage growth might resemble the budding of

the ventral anterior bud as described in the classical model [70,74].

The hatching period is characterized by the fusion of the two pancreatic

anlagen at 52 hpf. The posterior anlage becomes surrounded by increasing

numbers of exocrine cells from the anterior anlage to form the pancreatic islet. By

this time, the posterior anlage is no longer in direct contact with the intestine,

whereas the anterior anlage maintains its connection with the intestine to form

the pancreatic duct, which is located between the esophagus and the intestinal

bulb (Figure 8) [68].

When the zebrafish embryo becomes a larva, the exocrine cells caudal to the

pancreatic islet proliferate to form the tail of the larval pancreas, which may be

seen at 76 hpf (Figure 8). During the larval period, there is a continuous expansion

and differentiation of exocrine cells surrounding the pancreatic islet [68,74]. By 5.5

dpf, the pancreatic islet has been totally engulfed by exocrine tissue, which now

extends from the first to the sixth or seventh somite, and the endocrine islet is

located at the level of the third and fourth somite. The zebrafish pancreas may

now be divided into a “head” region, which contains the single Islet of

Langerhans, and a “tail” region. The existence of a single endocrine islet in the

head region of the zebrafish pancreas contrasts with the mammalian islets of

Langerhans, which are distributed throughout the pancreas [81].

42

Figure 8: Schematic representation of the development and anatomy of the zebrafish

gut. A: Anterior segment; M: Middle segment (intestinal bulb) and P: posterior

segment. The figure is reproduced from Crosnier et al. (2005) [78].

Figure 9: Schematic representation of the development of the zebrafish pancreas,

liver and swim bladder. The images represent the anterior (arrow) and posterior

(arrowhead) pancreatic buds. All images show ventral views of the endoderm,

anterior to the top. S: swim bladder; gb: gall bladder; L: Liver. The figure is

reproduced from Field et al. (2003b) [68].

43

o The liver

There are two different hypotheses regarding the onset of zebrafish liver

development [67,84,85].

The first hypothesis postulates that the liver–specific marker ceruloplasmin

(cp) may be detected on the left–hand side of the endoderm at 16 hpf. According

to this model, the progenitor cells migrate and aggregate in order to form the liver

bud, which becomes detectable after 32 hpf. However, there is no concrete

evidence that the early cp–positive cells really contribute to the formation of the

liver bud at a later stage of zebrafish development [84,85].

The second hypothesis is based upon two transcription factors hhex

(hematopoietically-expressed homeobox) and prox1 (prospero homeobox 1),

which are two important hepatoblast markers. The expression of hhex and prox1

may first be detected within the endoderm rostral to the zebrafish gut at

approximately 24 hpf, which is concurrent with the onset of liver development.

Hhex and prox1 genes are expressed in an endodermal cluster anterior to the

pdx1-expressing cells, which are important in pancreatic development [69,86].

According to the second hypothesis, liver morphogenesis may be divided into

two phases, namely budding and growth. The budding process occurs from 24 to

50 hpf and may be subdivided into three stages, based upon distinct liver

morphology. The subsequent growth phase takes place between 50 and 96 hpf

and is characterized by an impressive change in liver size, shape and placement.

The second hypothesis is the most likely one, because it is supported by data from

anatomic [67] and genetic studies [69,86]. Therefore, in this thesis, the description

of the liver development is based upon the second theory.

The onset of the pharyngula period indicates the beginning of the budding

phase, which may be divided into three stages. Stage I begins around 24 hpf and

is characterized by an aggregation of prehepatic cells on the ventral surface of the

endoderm rostral to the zebrafish gut. As a result, by 28 hpf, the liver primordium

has become a ventral thickening, which is positioned slightly left of the midline

at the level of the first somite. The liver primordium is situated anterior to the

dorsal pancreatic anlage, which may be detected at the level of the fourth somite

44

(Figure 9) [67-69]. Stage II of the budding process takes place between 28 and 34

hpf. During this stage, the prehepatic thickening increases in size and the

intestinal bulb primordium undergoes a leftward bend at the level of the

developing liver [67,69,82]. Stage III of liver budding begins at approximately 34

hpf, when a furrow starts to form between the anterior edge of the liver and the

future esophagus. During this stage, the furrow expands posteriorly in order to

separate the liver from the intestinal bulb primordium (Figure 9) [67].

During the hatching period, stage III of the budding phase continues.

Therefore, by 50 hpf, the tissue that connects the liver to the intestinal bulb

primordium has formed the hepatic duct, which consists of columnar epithelial

cells (Figure 9). The formation of the hepatic duct indicates the end of stage III

and thus the end of the budding phase [67,69]. Liver budding in zebrafish

contrasts with mammalian liver development, which is characterized by

hepatocytes that appear to dissociate from one another and migrate into the

mesenchyme of the adjacent septum transversum [87]. The budding process is

followed by the growth phase, which is typified by changes in liver size, shape

and placement. Furthermore, during this phase, the liver becomes vascularized

and it is supposed to exert its physiological functions for the first time. At the

beginning of the growth phase, the liver size remains unchanged. Moreover,

around 50 hpf, endothelial cells are closely associated with the liver periphery

until approximately 60 hpf, when they begin to invade the outer layers of the liver

[67,69]. By 72 hpf, the endothelial cells, which appear to derive from subintestinal

vessel, have permeated the entire liver [67,69,88]. This process of endothelial

invasion differs from liver vascularization in mammals, where hepatocytes

invade the adjacent mesenchyme and arrange themselves around the vascular

network, which is already present [87]. By the end of the hatching period, the liver

size has moderately increased, but the liver shape has not altered (Figure 9) [67].

During the larval period, around 96 hpf, the zebrafish liver is essentially in its

adult configuration. At this point in time, the liver overlaps with the anterior

portion of the remaining yolk and its anterior edge is in contact with the

pericardial cavity. The liver may now be seen as a medial expansion, which

45

extends from the left side of the larva across the midline ventral to the esophagus

[67].

The kidney

In the course of vertebrate evolution, three distinct forms of kidneys of

increasing complexity have been generated, namely the pronephros,

mesonephros and metanephros. In zebrafish, the pronephros is the functional

kidney during early larval live [89]. The pronephros consists of two glomeruli that

fuse at the embryonic midline and are connected to the pronephric ducts by two

pronephric tubules [65]. When the zebrafish larva becomes a juvenile, a

mesonephros forms along the length of the pronephros and will later serve as the

final adult kidney. The metanephric kidney is formed exclusively in amniotes

such as mammals, in which it will become the definitive kidney [72,90].

The nomenclature of the zebrafish pronephros has changed over the years

[72,91,92]. Historically, only a short stretch of tubule was supposed to exist in the

pronephros, connecting the glomerulus to a long pronephric duct (Figure 10a)

[91]. However, based upon new molecular data, there is now a consensus that the

long stretch of tubular epithelium, which has previously been considered the

pronephric duct, is actually subdivided into two proximal tubule segments

(proximal convoluted tubule or PCT and proximal straight tubule or PST), two

distal tubule segments (distal early or DE and distal late or DL) and a short duct.

Furthermore, what has traditionally been considered “tubule” is now believed to

represent a “neck” segment (Figure 10b) [93]. These subdivisions of the

pronephric tubular epithelium in zebrafish are in many ways homologous to the

segments of the metanephric tubules in mammals [91]. However, the zebrafish

pronephric duct is a short segment, which connects the tubules to the cloaca,

whereas the mammalian metanephros possesses a complex collecting system in

order to receive the waste from thousands of nephrons. Between the DE and DL

segments, the zebrafish pronephros comprises the Corpuscles of Stannius, which

represent clusters of endocrine glands and which are situated at the junction

(Figure 10b) [93]. The Corpuscles of Stannius are responsible for maintaining

calcium and phosphate homeostasis [94].

46

Figure 10: Schematic representation of the pronephros in zebrafish larvae according to

the historical nomenclature (a) [92] and the recent nomenclature (b) [91].

Figure (a): g: glomerulus; pt: pronephric tubules; pd: pronephric ducts. Figure (b): P:

podocytes of renal corpuscle; N: neck; PCT: proximal convoluted tubule; PST: proximal

straight tubule; DE: distal early; CS: Corpuscle of Stannius; DL: distal late; PD: pronephric

duct; C: cloaca; T: tubule. Figure 11a and 11b are reproduced from Hostetter et al. (2003)

[92] and Wingert et al. (2008) [91], respectively.

The development of the zebrafish pronephros starts in the early segmentation

period, at 13 hpf, when the rudiment of the pronephros is first evident as a paired

mass of intermediate mesoderm, which lies under the third somite [62,65].

At the beginning of the pharyngula period, around 24 hpf, the tissue that will

later form the glomerulus and tubular neck segments exists as paired, disk–

shaped nephron primordia situated ventral to the third somite. Each nephron

primordium appears as an invagination of the coelomic lining and is still

connected to the coelom by a nephrostome. Furthermore, by this time

epithelialization of the pronephric tubules has been completed. As a result, the

epithelial cells of the pronephric tubules are polarized with apical and basolateral

domains containing ion transport proteins [65,72]. At the same time, the bilateral

pronephric ducts have fused at their posterior end and exit the embryo at the

position of the cloaca [65,90,95]. Around 32-33 hpf, the separation of the nephron

primordia from the coelom is complete and each of the two primordia appears as

(a) (b)

47

a separate group of cells with a central lumen but with no connection to the

coelom. Moreover, the first signs of pronephric nephron morphogenesis may

now be seen. This morphogenesis becomes obvious at 40 hpf, when each nephron

primordium is partitioned into a medial glomerular domain and a lateral domain

representing the tubular neck segment. The developing glomerulus consists of

podocytes that form extensive foot processes, which interact with the capillaries

growing in from the overlying dorsal aorta between 40 and 48 hpf [65]. As a result,

glomerular blood filtration begins at around 48 hpf but is leaky at this time,

allowing large molecules to pass into the tubules [90].

By the beginning of the hatching period, by 50 hpf, the medial surfaces of the

two pronephric glomeruli have fused at the midline. At this time, the tubular neck

segments bend over laterally, where they are connected to the proximal tubule

segments (Figure 10b). Ten hours later, at around 60 hpf, a direct connection

between the pronephric glomeruli and tubular neck segments is established [65].

During the larval period, by 84 hpf, the development of the pronephric

nephron is essentially complete and the well–developed glomerular filtration

barrier leads to size–selective blood filtration [65,96]. The larval period also

indicates the start of mesonephric nephrogenesis since the first mesonephric

nephron appears around 12 dpf on top of the pronephric distal early (DE)

segment (Figure 10b). Around 14 dpf, the first mesonephric nephron becomes

functional due to fusion of the nephron with the lumen of the underlying

pronephros. Subsequently, additional mesonephric nephrons are progressively

added first caudal and then rostral to the first–forming nephron [75,97].

When the zebrafish larva reaches the juvenile period, around 30 dpf, the young

mesonephros morphologically resembles the fully mature adult mesonephros,

consisting of the “head”, “trunk” and “tail” regions (Figure 11). In contrast to the

pronephros, the mesonephros possesses a significantly higher degree of

structural and functional complexity. Moreover, mesonephric nephrogenesis

continues throughout the life of the zebrafish, with a rapid growth phase during

the juvenile period and a slower growth phase during adulthood [75,97].

48

Figure 11: The mesonephros of an adult zebrafish (90 dpf) with head, trunk

and tail regions. The figure is reproduced from Diep et al. (2015) [97].

49

4 Disposition of xenobiotics in zebrafish

4.1 ADME in mammals and zebrafish

Since maternal metabolism is lacking in ex utero developing organisms,

zebrafish embryos depend on their intrinsic biotransformation capacity for the

detoxification and/or bioactivation of xenobiotics. The biotransformation or

metabolism of a compound is one of the ADME–processes, i.e. Absorption,

Distribution, Metabolism and Excretion, that describe the disposition and fate of

xenobiotics within an organism (Figure 12) [98]. The metabolism and/or excretion

are sometimes referred to as ‘elimination’ of a drug [99]. The current section

describes the different ADME–processes, also referred to as pharmacokinetics,

whereas 4.2, 4.3 and 4.4 focuses on the drug–metabolizing capacity in zebrafish

since the latter underlies the subject of the doctoral project.

Figure 12: Schematic representation of ADME (Absorption, Distribution, Metabolism and

Excretion) in mammals for different routes of administration. Moreover, a comparison

between mammals and zebrafish has been made with regards to the different routes of

absorption: the light brown colored boxes represent routes of absorption which zebrafish

and mammals have in common, whereas the dark brown box represents the route of

absorption which is typical for aqueous organisms. SC: Subcutaneous; IM: intramuscular;

IP: intraperitoneal; IV: intravenous.

Oral administration

Gastrointestinal tract

Liver

Blood stream

Kidney BrainOtherorgans

A

D

M

E

Skin/gills:passivediffusion

SC, IM, IP, IV injectionsOcular, nasal, rectal and intratracheal administration

Faeces Urine

50

4.1.1 Absorption

Absorption refers to the process of how drugs go through the organs of the

body to reach the systemic circulation [98]. The different routes of administration

that are used in mammals are represented in figure 12. With regards to zebrafish,

the compounds of interest are usually dissolved in the incubation medium. The

route of absorption in case of aqueous exposure depends on the age of the

zebrafish embryo or larva. Until hatching, the zebrafish embryo is surrounded by

a chorion and the aqueous perivitelline space between the chorion and the

embryo (Figure 13). The chorion is an acellular permeable membrane of 1.5–10

µm thickness with circular pores with a diameter of 0.5–1.5 µm, which is

considered to be an effective barrier only for large molecules of a size of >3 kDa.

Until hatching of the zebrafish embryo, compounds <3 kDa pass through the

chorion into the perivitelline space to reach the skin of the embryo by which they

are taken up by passive diffusion (Figure 12) [100,101]. From the hatching period

onwards, the developing gills of the zebrafish embryo also contribute to the

passive diffusion of xenobiotics [62]. Due to opening of the mouth around 74 hpf,

exogenous compounds may be orally ingested by the zebrafish larva, which

makes the gastrointestinal tract another key site for absorption [73,101].

Alternatively, xenobiotics can be injected into the yolk sac or the vasculature,

from where they get distributed throughout the body.

The ADME–concept, as described originally, does not include the uptake of

compounds into the metabolizing cell. Around the turn of the century, the

cellular drug uptake processes were added to the ADME–concept and introduced

as phase 0 transport, which includes carrier–mediated uptake of drugs from, e.g.

the blood or gut lumen, into the metabolizing cell. In the mammalian liver and

kidney, phase 0 transporters are situated at the blood–facing basolateral

membrane of the metabolizing cell, whereas intestinal phase 0 transporters are

situated at the apical membrane (Figure 14) [reviewed by [102]). Section 4.4

further elaborates on transport proteins with special emphasis on what is known

about the transporters in zebrafish.

51

Figure 13: Zebrafish embryo of 27 hpf surrounded by a chorion a perivitelline space.

4.1.2 Distribution

Distribution is defined as the transportation of xenobiotics from one

tissue/organ to another tissue/organ, which is mainly performed via the blood

circulation (Figure 12). Drugs interact with plasma proteins in the blood and

unbound drugs interact with the membranes of the tissues. An equilibrium exists

between the free drug in blood plasma and drugs bound to plasma proteins [98].

In zebrafish, the distribution of xenobiotics has not yet been extensively studied,

except for the blood–brain barrier (BBB). In most vertebrates, the BBB consists of

tight junctions between adjacent endothelial cells that protect the brain by

allowing the absorption of only those drugs that are necessary for brain

metabolism [98]. The BBB in zebrafish is functionally and molecularly similar to

that of higher vertebrates and starts developing as early as 72 hpf [103].

4.1.3 Metabolism

Metabolism or biotransformation refers to the chemical modification of

exogenous compounds to increase their hydrophilicity and water solubility to

facilitate their excretion. Metabolism is classified into two main phases, i.e. phase

I and phase II [98].

52

4.1.3.1 Phase I

Phase I metabolism is characterized by the addition or unmasking of a

functional, polar moiety and includes primarily oxidation reactions, although

reduction and hydrolysis reactions are also possible. Phase I reactions are

governed mainly by cytochrome P450 (CYP) enzymes, which are responsible for

the oxidation of the majority (>60 %) of marketed drugs (Figure 15) [98,104]. Other

oxidative enzymes that are commonly involved in mammalian phase I

metabolism include flavin containing monooxygenase (FMO), aldehyde oxidase

(AO), monoamine oxidase (MAO), alcohol dehydrogenase (ADH) and aldehyde

dehydrogenase (ALDH). CYP and FMO enzymes are both located in the

endoplasmic reticulum and require NADPH (nicotinamide adenine dinucleotide

phosphate) and O2 as co–factors in order to become active. AO and MAO

enzymes have no co–factor requirements and are located in the cytosol and

mitochondria, respectively. The cytosolic ADH enzymes facilitate the reversible

oxidation of alcohols to aldehydes or ketones using NAD+/NADPH as a cofactor,

whereas ALDHs are NAD(P)+–dependent and catalyze the oxidation of a wide

range of aldehydes. The phase I enzymes that are involved in the

biotransformation of xenobiotics are differentially expressed in the mammalian

liver, intestine, kidney, lung and brain ([104]. Section 4.2 further elaborates on

CYP enzymes with special emphasis on what is known about CYPs in zebrafish.

4.1.3.2 Phase II

Phase II metabolism consists mainly of conjugation reactions, i.e.

glucuronidation, sulfonation, methylation, acetylation, amino acids and

glutathione (GSH) conjugation. The cofactors of these reactions react with

functional groups that are either present on the parent compound or are

introduced during phase I biotransformation. Indeed, phase I does not

necessarily precede phase II. The conjugated metabolites are relatively more

polar, which facilitates their excretion from the body via urine or faeces (Figure

14). The main conjugative enzymes that are involved in mammalian phase II

reactions are uridine 5’–diphospho (UDP)–glucuronosyltransferase (UGT),

sulfotransferase (SULT), glutathione S–transferase (GST), N–acetyltransferase

53

(NAT) and methyltransferase (MT). UGTs are located in the endoplasmic

reticulum, whereas NATs can be found in the cytosol. The location of SULTs,

GSTs and MTs depends on the class of enzymes since all three enzyme groups

include membrane–bound (e.g. endoplasmic reticulum or Golgi apparatus) as

well as cytosolic enzymes. Phase II enzymes are differentially expressed in a

broad range of mammalian tissues, in particular liver, kidney, intestine, lung and

brain [104]. Section 4.3 further elaborates on phase II enzymes with special

emphasis on what is known about conjugative enzymes in zebrafish.

4.1.4 Excretion

Excretion is the removal of xenobiotics from the body, either as a metabolite

or as unchanged drug. In mammals, there are many different routes of excretion,

including urine, bile, sweat, saliva, tears and milk. The liver and kidney are by

far the most important excretory organs. In the kidney, excretion of xenobiotics

depends on glomerular filtration, active tubular secretion and passive tubular

absorption. Drugs excreted by the liver appear in the bile and enter the

duodenum where they may be reabsorbed resulting in enterohepatic circulation.

Exogenous compounds with a high molecular weight (>300 Da) and lipophilic

groups are more likely to be excreted in bile, whereas drugs with a low molecular

weight and hydrophilic groups are mainly excreted in urine.

In pharmacokinetic studies, clearance is a measure of drug elimination (i.e.

metabolism and excretion) from the body [99]. Drug clearance refers to the

volume of plasma fluid that is cleared of drug per unit time. Moreover, clearance

may also be considered the fraction of drug removed per unit time [99]. Regarding

zebrafish, van Wijk et al. (2019) [101] showed an increase in clearance of

paracetamol in zebrafish larvae between 72 and 120 hpf, which is expected to

result from the continuous growth of eliminating organs such as the liver and

kidneys (Table 2).

Around the turn of the century, phase III transport was added to the ADME–

concept, which includes the active transport (efflux) of drugs out of the

metabolizing cell into excretion fluids by means of ATP–binding cassette (ABC)

transporters. The latter are mainly situated at the apical (luminal) membrane, e.g.

54

the bile–facing canalicular membrane in the liver or the urine–facing tubule brush

border membrane in the kidney (Figure 14). The ABC carriers consume ATP,

which is needed to transport the metabolite against a concentration gradient. In

some circumstances, the vectorial transcellular drug transfer through the

metabolizing cell is interrupted and the metabolites are returned back into the

blood by ABC transporters that are situated at the basolateral membrane (Figure

14). The prototype of phase III ABC carriers is P–glycoprotein, also called

multidrug resistance protein 1 (MDR1) (gene code ABCB1), which is the earliest

cloned multidrug carrier [102]. Section 4.4 further elaborates on transport proteins

with special emphasis on what is known about the transporters in zebrafish.

Besides the kidney and liver, phase 0 and phase III transporters are also

expressed in other mammalian organs such as lung, heart, intestine, pancreas,

brain and placenta. Indeed, in the mammalian placenta, P–glycoprotein (P–gp) is

expressed at the apical membrane of syncytiotrophoblast cells facing the maternal

blood where it seems to protect the fetus against xenobiotics by extruding them

into the maternal blood. Another important location of P–gp is the apical

membrane of brain endothelial cells, i.e. blood–brain barrier (BBB), where the

transporter protects the central nervous system from high concentrations of

drugs [102].

Within the metabolizing cell, compounds and/or metabolites are transferred

between the metabolism sites and the membrane transporters by cytosolic

binding proteins and cytoskeletal structures. This intracellular transport

mechanism is sometimes referred to as phase III transport (in this case, the efflux

of drugs is referred to as phase IV transport) [102,105]. However, in the current

thesis we will maintain the general accepted nomenclature as shown in Figure

14.

55

Figure 14: The figure represents de different steps of drug metabolism (Phase I and II)

and membrane transport (Phase 0 and III) in a hepatocyte. SLC*: solute carrier; ABC*:

ATP–binding cassette. The figure is adapted from Döring et al. (2014) [102].

4.2 Cytochrome P450 enzymes in humans and zebrafish

The name “cytochrome P450” (CYP) is derived from the characteristic

absorption band at 450 nm which was initially observed upon binding of the

enzyme to carbon monoxide [106]. CYP enzymes are a group of heme–containing

monooxygenases that are evolutionarily conserved and exist in all living

organisms from bacteria to humans. CYPs catalyse the oxidation of a wide variety

of substrates such as drugs, carcinogens, steroids, pesticides and other chemicals

with the aim of increasing the substrate’s polarity. The generally accepted

catalytic cycle for CYP enzymes is shown in Figure 15 and consists of a complex

multistep process: after binding of the substrate (RH), one electron is transferred

from NADPH–P450 reductase to the enzyme, O2 binds to the CYP ferrous ion

(Fe2+) and another electron is transferred to the Fe2+-O2 complex (steps 1–4). After

step 4, the successive events are rather unclear. According to the generalized CYP

reaction mechanism as show in Figure 15, step 4 is followed by the addition of

two protons (H+), the release of H2O, the removal of a proton from the substrate

Phase 0: Uptake

Drug

SLC*transporter

Phase III: Efflux

ATP ADP

ABC*transporter

Cofactor

-O-charged groupConjugate

TransferasesPhase II: Conjugation

-OHOxidated drug

Phase I: OxidationO2

Cytochromes P450RH + 2H+ + O2 + 2e- -> ROH + H2O

ATPADP

HEPATOCYTECanalicular/apicalmembrane

Basolateralmembrane

Phase III: Efflux

56

(RH) to FeO3+, the transfer of the hydroxyl group from FeOH3+ to the substrate

radical and finally the release of the oxidized substrate (ROH) (steps 5–9)

[107,108]. The following reaction mechanism is a simplified representation of the

CYP catalytic cycle [108]:

RH + 2H+ + O2 + 2e- -> ROH + H2O

The CYP superfamily is divided in families and subfamilies according to their

amino acid sequence homology: e.g. CYP1 refers to the family in which the

enzymes share at least 40% amino acid identity and CYP1A refers to the

subfamily in which the members share at least 55% amino acid identity. The final

Arabic number, e.g. CYP1A1, represents the individual enzyme or isoform. To

date, around 57 human CYP genes have been identified of which approximately

one quarter are considered to be involved in the biotransformation of xenobiotics,

i.e. CYP families 1, 2 and 3 (Table 3). The major CYP isoforms that are responsible

for the metabolism of drugs in man are shown in Figure 16. Although CYPs can

be found in almost all organs, the liver and intestinal epithelia are the

predominant sites for CYP–mediated biotransformation (Table 3). Besides their

role in drug metabolism, CYP families 1, 2 and 3 are also involved in the metabolic

conversion of a variety of endogenous compounds such as vitamins, bile acids

and hormones. The CYP isoforms from the other families (CYP families 4–51) are

generally involved in endogenous processes such as the synthesis, activation or

inactivation of endogenous regulatory molecules [109,110].

In zebrafish, Goldstone et al. (2010) [111] uncovered a total of 94 CYP genes

that fall into 18 CYP families, which are also found in humans. Similar to the

human situation, the zebrafish CYPs can also be divided into two major

functional groups, i.e. CYP families 1–3 that are primarily involved in the

metabolism of xenobiotics and CYP families 5–51 that are involved in

endogenous functions. Whereas most CYP genes in families 5–51 are direct

orthologues of human CYPs, complexity is much greater in the families that are

involved in xenobiotic metabolism since these CYP genes are much more diverse

(Table 3) [111]. In view of their use in developmental toxicity studies, the doctoral

project focuses on the ontogeny of some CYPs from families 1–3 that are supposed

to have a role in drug metabolism in zebrafish (Table 3: CYPs underlined and in

57

italics). Besides the above–mentioned functions, CYP enzymes in humans and

zebrafish also have a role in embryonic development such as CYP26 (retinoic acid

hydroxylase) which was first discovered in zebrafish [111-113]. Indeed, CYP26

enzymes have been shown to metabolize retinoic acid (RA) which affects

numerous developmental events such as the regulation of germ layer and body

axis formation, neurogenesis, cardiogenesis and the development of pancreas,

lung, and eye [114].

Figure 15: Generally accepted catalytic cycle for cytochrome P450 (CYP) enzymes. The

ferric (Fe3+) and ferrous (Fe2+) ions represent the heme group of the CYP enzyme and RH

and ROH represent the substrate and metabolite, respectively. The figure is reproduced

from Guengerich (2007) [107].

58

Figure 16: The contribution of individual cytochrome P450 (CYP) enzymes in the

metabolism of drugs in humans. The figure is reproduced from Guengerich (2006) [109].

Many of the human CYPs are induced by a diverse array of xenobiotics,

which are at the same time substrates of the corresponding CYPs. The induction

of CYP enzymes occurs by ligand activation of key receptor transcription factors

followed by de novo RNA and protein synthesis, which leads to increased CYP

expression and CYP activity levels (Figure 17). In humans, transcription factors

primarily involve the nuclear pregnane X receptor (PXR), the cytosolic

constitutive androstane receptor (CAR) and the cytosolic aryl hydrocarbon

receptor (AhR) [115]. The mechanism of CYP induction is highly conserved as it

is also found in many other species including zebrafish. However, zebrafish only

exhibit two of the above–mentioned regulatory mechanisms, i.e. AhR and PXR,

with CAR being absent in this species [116-118].

In mammals as well as in zebrafish, CYP enzymes are located in the

membrane of the endoplasmic reticulum (ER) (Figure 17). The ER can be

fragmented by differential centrifugation of the tissue homogenate at 10,000 and

100,000× g leading to small sealed vesicles referred to as microsomes. Hence,

59

microsomes are artificial structures containing CYP enzymes, which can be used

in in vitro drug metabolism studies to assess CYP activity [119].

Figure 17: Simplified schematic representation of cytochrome P450 (CYP) induction in

humans, which involves the activation of key transcription factors such as aryl

hydrocarbon receptor (AhR), constitutive androstane receptor (CAR) or pregnane X

receptor (PXR). The induction mechanisms result in the synthesis of CYP proteins, which

are located in the membrane of the endoplasmic reticulum (ER).

Nucleus

Cytoplasm

Xenobiotics

CAR / AhR

Gene transcription

mRNA

PXR

mRNATranslation

Increased drug metabolism / pro-

drug activation

CYP protein

ER

60

Table 3: Zebrafish and human CYP homologues which are involved in the metabolism of xenobiotics, including their tissue

distribution in adults.

CYP FAMILY ZEBRAFISH

CYPS

TISSUE DISTRIBUTION HUMAN CYPS TISSUE DISTRIBUTION REFERENCES

CYP1

CYP1A

CYP1B1

CYP1C(1,2)

CYP1D1

Liver, GI, kidney, heart, gills, eye

Brain, eye, heart, kidney, gills, liver, GI,

gonads

Heart, eye, gills, brain, liver, kidney, GI,

testes

Brain, liver, gills, GI, kidney

CYP1A1

CYP1A2

CYP1B1

—

CYP1D1P*

GI, lung, heart, brain, lymphocytes, liver

Liver

Skin, brain, heart, lung, liver, kidney, GI,

placenta

—

—

[98,110,111,120-124]

CYP2

CYP2AA(1–12)

CYP2AE(1,2)

—

—

—

—

—

CYP2K6

CYP2K(8,16-22, 31)

GI, kidney, liver, heart, brain, eye,

gonads

Unknown

—

—

—

—

—

Liver, ovary

Unknown

—

—

CYP2C(8,9)

CYP2C18

CYP2C19

CYP2D6

CYP2E1

CYP2W1

—

—

Liver, kidney, adrenal glands, GI, ovary

Epidermis

Liver, GI

Liver, kidney, placenta, lung, GI, brain

Liver, nose, oropharynx, lung, brain

Tumor–specific

[98,110,111,120,124-

127]

Bold text in column of zebrafish: tissues where the CYP enzymes are highly expressed. Underlined zebrafish CYPs: enzymes that were

investigated in this PhD project. Human CYP enzymes in bold: major CYPs that are involved in the biotransformation of xenobiotics.

CYP: cytochrome P450; GI: gastrointestinal; P*: pseudogenes.

61

Table 3: Continued.

CYP FAMILY ZEBRAFISH

CYPS

TISSUE DISTRIBUTION HUMAN CYPS TISSUE DISTRIBUTION REFERENCES

CYP2

CYP2AD(2,3,6)

CYP2J20

CYP2N13

CYP2P(6,7,10)

CYP2V1

CYP2U1

CYP2R1

CYP2X(6–10)

CYP2Y3

Unknown

Unknown

Unknown

Brain, gonads, liver, heart, kidney

Unknown

Unknown

Unknown

Unknown

Unknown

CYP2J2

CYP2U1

CYP2R1

—

CYP2A(6,13)

CYP2B6

CYP2F1

CYP2S1

Heart, kidney, lung, liver, GI

Brain, thymus

Liver

—

Liver

Liver, heart

Lung, testis

GI

[98,110,111,120,124-

127]

CYP3

CYP3A65

CYP3C(1–4)

Liver, intestine, brain, gills, eye

Liver, GI, kidney, brain, gills, gonads,

eye, heart

CYP3A-se1,-se2*

CYP3A(3,4)

CYP3A7

—

Liver, GI, kidney, lung, brain, placenta

Fetus, placenta, liver

[111,124,128,129]

Bold text in column of zebrafish: tissues where the CYP enzymes are highly expressed. Underlined zebrafish CYPs: enzymes that were

investigated in this PhD project. Human CYP enzymes in bold: major CYPs that are involved in the biotransformation of xenobiotics.

CYP: cytochrome P450; GI: gastrointestinal; se*: (single exon) pseudogenes.

62

4.3 Phase II enzymes in humans and zebrafish

Since the doctoral project includes the assessment of the developmental

mRNA expression of UGT and SULT in zebrafish (Chapter 2: aims of doctoral

project), the current section will focus on these two phase II enzymes.

The UGTs are a superfamily of enzymes that catalyze the covalent addition

of sugars from UDP–sugar donors to functional groups (most frequently

hydroxyl, carboxyl, or amine) of a broad range of lipophilic molecules. These

lipophilic molecules may be from exogenous sources, i.e. xenobiotics, as well as

from endogenous sources, e.g. bile acids, vitamins and hormones. The resulting

glucuronides are generally inactive and water soluble, thus facilitating their

excretion from the body through urine or feces (Figure 18) [130]. In humans, the

UGT superfamily comprises four families, i.e. UGT1, UGT2, UGT3 and UGT8,

from which UGT3 and UGT8 are distinguished from the first two families

because of their different UDP–sugar cofactor utilization. UGT1 and UGT2,

which use UDP–glucuronic acid as cofactor, have an important role in the

conjugation of pharmacological agents, whereas UGT3 and UGT8 may conjugate

mainly endogenous compounds. However, the functional properties of the UGT3

and UGT8 families remain to be fully elucidated [130]. Naturally, UGT1 and

UGT2 are highly expressed in organs of detoxification, while the expression of

UGT3 and UGT8 is relatively low in the major drug–metabolizing organs. The

UGT genes have been shown to be induced by a wide variety of ligand–activated

transcription factors, i.a. peroxisome proliferator–activated receptor (PPAR),

PXR, CAR and AhR [130,131].

Huang and Wu (2010) [132] identified 45 UGT genes in zebrafish that can be

divided into three families: UGT1, UGT2 and UGT5. The zebrafish UGT1 and

UGT2 genes are closely related to the human UGT1 and UGT2 genes, respectively,

whereas the zebrafish UGT5 genes are a novel UGT subfamily that does not exist

in humans (Table 4). It has been hypothesized that the zebrafish UGT1 and UGT2

families are duplicated into two unlinked gene clusters (a and b), i.e. UGT1a,

UGT1b and UGT2a, UGT2b [132]. Although the expression of the different UGT

genes in zebrafish was shown to be sex–specific, UGT1a and UGT1b were found

63

to be highly expressed in the liver and intestine of both females and males.

Besides the liver and intestine, UGT genes were also found to be expressed in

brain and gonads of adult zebrafish. The gene expression of UGT1a and of some

members of the UGT5 family is regulated by the AhR pathway. However, further

research is needed regarding the involvement of additional transcription factors

such as PXR in the regulation of UGT expression in zebrafish [104,133].

Similar to CYP enzymes, UGTs are located in the membrane of the

endoplasmic reticulum (ER). Hence, microsomes can be used to assess the

glucuronidation of xenobiotics in an in vitro setting, provided the addition of a

cofactor [104,134,135].

Figure 18: : In humans as well as in zebrafish, the nonsteroidal anti–inflammatory

drug diclofenac is glucuronidated by members of the UGT superfamily [134,135].

The current figure represents the glucuronidation of diclofenac by a member of the

human UGT2 family. The figure is reproduced from Knöspel et al. (2016) [135].

The SULTs catalyze the transfer of a sulfonate group (SO3-) from the sulfate

donor 3’–phosphoadenosine–5’–phosphosulfate (PAPS) to substrate compounds

containing hydroxyl or amino group(s) (Figure 19). The sulfation results in the

inactivation or increased water–solubility of the substrate compounds, thereby

facilitating their excretion from the body [104,136]. There are two classes of SULTs:

1) cytosolic SULTs which are involved in hormone regulation and in the

metabolism of numerous xenobiotics and 2) Golgi membrane–bound SULTs,

64

which are central players in a number of molecular recognition events and

biochemical signalling pathways such as T–cell response and cell

adhesion/proliferation [104,137]. Humans contain 13 cytosolic SULTs and 37 Golgi

membrane–bound SULTs. Since the Golgi membrane–bound SULTs are less

involved in drug metabolism, these enzymes will not be further discussed. The

human cytosolic SULTs fall into four distinct gene families, i.e. the two major

families SULT1 or phenol SULTs, and SULT2 or hydroxysteroid SULTs and the

lesser–known SULT4 and SULT6 (Table 4) [136,137]. The cytosolic SULTs are

primarily expressed in the human liver, lung, brain, skin, platelets, breast, kidney

and gastrointestinal tissue [138]. Similar to UGT, human SULT genes have been

shown to be induced by a wide variety of ligand–activated transcription factors,

i.a. PPAR, PXR, CAR and AhR [131,139].

In zebrafish, 20 cytosolic SULT genes have been identified, which can be

divided into six families, i.e. SULT1–6 (Table 4). Similar to human cytosolic

SULTs, a good proportion (nine of twenty) of zebrafish SULTs belong to the

SULT1 gene family. Zebrafish SULT1s were shown to be capable of sulfating a

wide spectrum of xenobiotics, including environmental pollutants and drug

compounds, as well as endogenous substrates such as thyroid hormones,

estrogens and dopamine. Zebrafish SULT2 enzymes are mainly involved in the

sulfation of endogenous compounds, hydroxysteroids in particular, whereas

members of the SULT3 family are capable of catalyzing the sulfation of

hydroxysteroids and xenobiotics. The SULT6 ST1 enzyme was shown to be

capable of sulfating dopamine and thyroid hormones as well as a number of

xenobiotics [140]. The functional relevance of the remaining SULT4 and SULT5

enzymes still needs to be elucidated. Besides the cytosolic SULTs, zebrafish also

contain Golgi membrane–bound SULTs such as heparin sulfotransferases and

tyrosylprotein sulfotransferases, which will not be discussed in this thesis [136].

To date, information regarding the regulation of SULT expression in zebrafish is

lacking.

Since the xenobiotic–sulfating SULT enzymes are located in the cytosol, the

activity of these enzymes can be assessed in the S9 fraction—the post–

65

mitochondrial supernatant fraction that contains a mixture of microsomes and

cytosol—provided the addition of a cofactor [141].

Figure 19: In humans as well as in zebrafish, the analgesic and anti–pyretic drug

acetaminophen (paracetamol) is sulfated by members of the SULT superfamily. PAP: 3’–

phosphoadenosine–5’–phosphate; PAPS: 3’–phosphoadenosine–5’–phosphosulfate;

SULT: sulfotransferase. The figure is reproduced from Liu et al. (2010) [140].

66

Table 4: Zebrafish and human UGT and SULT homologues.

ENZYME

FAMILY

ZEBRAFISH

ENZYME

HUMAN ENZYME REFERENCES

UGT

UGT1

UGT2

—

UGT5

—

UGT1

UGT2

UGT3

—

UGT8

[130,132]

SULT

SULT1 ST1

SULT1 ST2

SULT1 ST3

SULT1 ST4

SULT1 ST5

SULT1 ST6

SULT1 ST7

SULT1 ST8

SULT1 ST9

SULT2 ST1

SULT2 ST2

SULT2 ST3

SULT3 ST1

SULT3 ST2

SULT3 ST3

SULT3 ST4

SULT3 ST5

SULT4 A1

SULT5 A1

SULT6 B1

SULT1A(1,2,4)

SULT1A(1,2,4)

SULT1A1

SULT1B2

SULT1A(1,2,4)

SULT1C4

SULT1C2

SULT1A(1,2,4)

SULT1A(1,2,4)

—

SULT2B1

SULT2A1

SULT2B1

SULT2B1

—

—

—

—

—

SULT4A1

—

SULT6B1

[136]; (https://zfin.org)

Underlined zebrafish UGTs and SULTs: enzymes that were investigated in the doctoral project.

UGT: UDP–glucuronosyltransferase; SULT: sulfotransferase.

https://zfin.org/

67

4.4 Transport proteins in humans and zebrafish

In the doctoral project, we assessed the mRNA expression of a

multixenobiotic transporter in the developing zebrafish, i.e. abcb4, which is

functionally similar to P–glycoprotein in humans (Chapter 2: aims of doctoral

project). This section gives a brief overview of what is known about transporters

in humans and zebrafish. However, knowledge of transporter function as well as

the list of transport proteins that have been identified is still growing.

Transport proteins guide drugs and their metabolites in and out the cells and,

in particular, enable water soluble or charged drugs and metabolites to pass the

phospholipid membrane barrier. As mentioned in 4.1, two main clusters of

transporter families have been described, i.e. ATP–binding cassette transporters

(ABC) and solute carrier (SLC) transporters [102]. These transport proteins are

situated in the apical and/or basolateral membrane of epithelial cells that separate

nearly all body fluid compartments such as kidneys, liver and intestine, as well

as in brain endothelium, circulating blood cells, gonads, retina, placenta,

olfactory epithelium, mammary glands, etc. [142].

The SLC transporters are mainly responsible for the uptake of compounds

through the basolateral (e.g. in hepatocytes) or apical (e.g. in the intestine) cell

membrane. The SLC uptake of compounds occurs down the concentration

gradient via facilitated diffusion which does not rely directly on ATP hydrolysis

(uniporter in Figure 20). However, uptake may also occur against the

concentration gradient via a secondary active transport mechanism together with

a co–substrate (symporter and antiporter in Figure 20) [102]. In humans, the SLC

superfamily comprises 52 gene families which are expressed in SLC carriers that

only transport endogenous substrates such as signaling molecules and

physiological metabolites, carriers that transport xenobiotics in addition to

physiological compounds and SLC carriers that mainly transport xenobiotics.

Figure 22 gives an overview of SLC gene families that are involved in the

transport of xenobiotics (transporters with a negligible role in xenobiotic

transport were excluded from the scheme). SLC transporters are multispecific

68

since they allow permeation of a spectrum of compounds with variable chemical

structures as shown by the trivial names of transport proteins in Figure 22 [105].

The ABC carriers are efflux transporters which are mainly located at the

apical cell membrane, e.g. near the intestinal lumen, the proximal tubule lumen

in the kidney and the bile canaliculus in the liver. Hence, ABC transporters are

involved in the excretion of xenobiotics from body. However, these efflux

transporters may also be expressed at the basolateral membrane of e.g.

enterocytes and hepatocytes, where they pump the parent compound or its

metabolite back into the blood [105]. Unlike the SLC carriers, the efflux of

compounds or their metabolites occurs against the concentration gradient via a

primary active transport mechanism which relies on the direct expense of energy

(Figure 20) [102]. The energy is released by the hydrolysis of ATP at the

cytoplasmic nucleotide–binding domain (NBD), which is connected to a

polypeptide transmembrane domain (TMD) (Figure 21). Based on the

organization of NBDs and TMDs, the ABC transporters are classified into seven

different families, some of which are shown in Figure 22 [102,143]. Similar to SLC

transporters, ABC carriers may transport xenobiotics as well as endogenous

compounds such as physiological metabolites (e.g. bile acids, bilirubin

conjugates, uric acid), neurotransmitters, hormones and signaling molecules.

Moreover, individual SLC and ABC carriers have overlapping substrate

specificities within the respective superfamily as well as among these two

transporter groups. For instance, movement of an organic anion drug from the

blood side to the urinary side is thought to involve basolateral uptake (SLC22)

transporters and apical efflux (ABCC) transporters [142]. As mentioned in 4.2, P–

glycoprotein (P–gp), also called multidrug resistance protein 1 (MDR1) (gene

code ABCB1), is the most studied and well understood transporter in the ABC

superfamily (Figures 21 and 22) [143]. P–gp can transport structurally unrelated

hydrophobic, amphipathic compounds and plays an important role in limiting

the entry of various xenobiotics in the central nervous system (BBB) [143].

The tissue expression of transport proteins in humans is regulated by ligand–

activated nuclear receptors such as PXR, CAR, farnesoid X receptor (FXR) and

vitamin D receptor (VDR) [144].

69

Aquatic organisms including zebrafish are continuously exposed to toxicants

dissolved in water. Detoxification systems, such as phase I and phase II enzymes,

as well as transport proteins are therefore essential for survival of these

organisms. Table 5 gives an overview of the SLC and ABC transporters that were

shown to be expressed in zebrafish. Similar to the human situation, zebrafish SLC

and ABC carriers are responsible for the uptake and efflux of xenobiotics as well

as endogenous compounds. Moreover, zebrafish ABC transporters also contain

TMDs and NBDs as described for humans (Figure 21) [145]. With regards to the

widely studied P–gp, a direct orthologue of human ABCB1 is lacking in zebrafish.

However, these aquatic organisms possess two gene paralogues, i.e. abcb4 and

abcb5 (Table 5), with common ancestry to human ABCB4 and ABCB5,

respectively. Nevertheless, zebrafish abcb4 is functionally similar to human

ABCB1 (and not ABCB4) since it was shown to act as a multixenobiotic transporter

whereas human ABCB4 is mainly involved in the export of phosphatidylcholine

from hepatocytes into the bile [143,146,147]. In contrast to abcb4, zebrafish abcb5

appears not to be involved in the efflux of xenobiotics [146]. Although not much

is known about the regulation of transporter gene expression in zebrafish,

Jackson and Kennedy [147] showed that PXR may play a role in the transcriptional

regulation of abcb4.

Despite the similarities between human and zebrafish transporters, a large

number of carrier proteins are yet to be characterized in zebrafish. Furthermore,

knowledge of transporter function in this species is still scarce and needs to be

further elucidated.

70

Figure 20: A schematic representation of the two main clusters of transporter

families: the solute carrier (SLC) and ATP–binding cassette (ABC) transporters. The

SLC transporters do not rely directly on ATP hydrolysis although secondary active

transport occurs in the case of antiporters and symporters. On the contrary, ATP

transporters use the direct expense of energy by splitting of ATP for substrate

transport against the concentration gradient. The figure is reproduced from Döring

et al. (2014) [102].

Figure 21: Topology of P–glycoprotein (P–gp): P–gp is a single polypeptide which

consists of two homologous halves that arose from gene duplication. Each half

comprises six transmembrane helices (TMHs) and one nucleotide–binding domain

(NBD) located on the cytoplasmic side of the membrane. The figure is reproduced

from Sharom (2011) [143].

71

SLC

SLC 15 Proton-coupled oligopeptide transporters (POTs)

Monocarboxylate transporters (MCTs) SLC 16

SLC 21/SLCO Organic anion transporting polypeptides (OATPs)

SLC 22 Organic anion/cation transporters (OATs/OCTs)

SLC 28 Sodium-coupled concentrative nucleoside transporters (CNTs)

SLC 29 Equilibrative nucleoside transporters (ENTs)

SLC 46

SLC 47

Proton-coupled folate transporters (PCFTs)

Multidrug and toxin extrusion transporters (MATEs)

SLC 51 Organic solute transporters (OSTs)

ABC

ABCA

ABCB

ABCB1 Multidrug resistance protein 1 (MDR1)

P-glycoprotein (Pgp)

ABCC

ABCG

ABCA2

ABCB4 Multidrug resistance protein 2/3 (MDR2/3)

ABCB11 Bile salt export pump (BSEP)

Sister of P-glycoprotein (sPgp)

ABCC1

ABCC2

ABCC3

ABCC4

ABCC5

ABCC6

ABCC10

ABCC11

Multidrug resistance-associated protein 1(MRP1) MRP2 MRP3 MRP4 MRP5 MRP6 MRP7 MRP8

ABCG2 Breast cancer resistance protein (BCRP) Mitoxantrone resistance protein (MXR)

Placenta-specific ABC protein (ABCP)

Figure 22: Overview of human uptake transporters, i.e. solute carrier (SLC), and human efflux

transporters, i.e. ATP–binding cassette (ABC), that are involved in the transport of xenobiotics.

The first column in the scheme represents the SLC and ABC gene families; the second column

of the ABC transporters represents some members of the ABC gene families; and the column

on the right shows the trivial names of the transport proteins. SLCO: solute carrier organic

anion [102,142].

72

Table 5: Zebrafish and human transporter homologues, including their tissue distribution in adult zebrafish.

TRANSPORTER

SUPERFAMILY

ZEBRAFISH

TRANSPORTER

GENE FAMILY

ZEBRAFISH TRANSPORT

PROTEINS/GENES

TISSUE DISTRIBUTION

(ADULTS)

HUMAN HOMOLOGUE

(+ % AMINO ACID SEQUENCE

IDENTITY)

REF.

SLC

Slc2/Slco

Slc22

Slc47

Oatp1(c1, d1, e1, f1, f2, f3, f4)

Oatp2(a1, b1)

Oatp3(a1, a2)

Oatp4(a1)

Oatp5(a1, a2)

Oat(1, 3)

Oat2(a, b, c, d, e)

Oct(1, 2, 6)

Octn(1, 2)

Orctl(3, 4)

Mate(3, 4, 5, 6, 7, 8)

Liver, intestine, kidney,

brain, gills, skeletal muscle

Kidney, intestine, gills,

liver, brain, skeletal muscle

Brain, intestine, skeletal

muscle, kidney, gills

Unknown

Brain, kidney

Kidney, intestine, gills,

brain, eye, testis

Kidney, intestine, brain,

eye, gonads

Kidney, liver, intestine,

gills, brain, heart, skeletal

muscle, eye, gonads

Kidney, intestine, eye,

brain, testis

Kidney, intestine, brain

Kidney, testes, intestine,

eye, liver, brain, gills, ovary

Oatp1c1: 54% with OATP1C

Oatp2a1: 50-52% with OATP2A

Oatp2b1: 42-49% with OATP2B

Oatp3a: 71-75% with OATP3

OATP4A1

Oatp5a1: 45-54% with OATP5A

Oatp5a2: 68% with OATP5

Oat1: OAT1

Oat3: OAT3

Oat2 (a-e): OAT2

Oct1: OCT2

Oct2: OCT3

Oct6: OCT6

Octn1: OCTN1

Octn2: OCTN2

Orctl13: ORCTL13

Orctl14: ORCTL14

Mates (3-8): 40-52 % with

MATE1 and MATE2

[148,149];

(https://zfin.org)

[150]

[151]

The solute carrier (SLC) transporters shown in this table may be involved in the transport of xenobiotics as well as endogenous compounds. In the third

column, zebrafish transporters are shown as proteins or genes depending on the nomenclature that has been used in literature. Mate: multidrug and toxin

extrusion; Oat: Organic anion transporter; Oatp: Organic anion transporting polypeptide; Oct: Organic cation transporter; Octn: Organic cation/carnitine

transporter; Orctl: Organic cation transporter-like; Slco: solute carrier organic anion.

https://zfin.org/

73

Table 5: Continued.

TRANSPORTER

SUPERFAMILY

ZEBRAFISH

TRANSPORTER

GENE FAMILY

ZEBRAFISH TRANSPORT

PROTEINS/GENES TISSUE DISTRIBUTION

(ADULTS) HUMAN HOMOLOGUE (+ %

AMINO ACID SEQUENCE

IDENTITY)

REF.

ABC

Abcb

Abcc

Abcb4

Abcb5

Abcc1

Abcc4

Abcc5

Liver, intestine, muscle, gill,

eye, ovary, heart, testis

Liver, epidermis

Gonads, eye, intestine,

kidney, brain, muscle, gill

Gonads, intestine, kidney,

eye, brain, gills, heart,

muscle, liver

Gonads, brain, eye,

intestine, kidney, gills,

heart, muscle, liver

50-64% with ABCB1/ABCB4,

50-64% with ABCB5

70% with ABCC1

69% with ABCC4

73% with ABCC5

[145,146,152,153]

The ABC–binding cassette (ABC) transporters shown in this table may be involved in the transport of xenobiotics as well as endogenous compounds. In the

third column, zebrafish transporters are shown as proteins or genes depending on the nomenclature that has been used in literature. With regards to tissue

distribution, organs with negligible expression of the respective transporter are excluded from the table. Zebrafish Abcb4 has been investigated in the doctoral

project.

74

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152. Long, Y.; Li, Q.; Li, J.; Cui, Z. Molecular analysis, developmental function and

heavy metal-induced expression of ABCC5 in zebrafish. Comparative biochemistry

and physiology. Part B, Biochemistry & molecular biology 2011, 158, 46-55.

153. Lu, X.; Long, Y.; Sun, R.; Zhou, B.; Lin, L.; Zhong, S.; Cui, Z. Zebrafish Abcb4 is a

potential efflux transporter of microcystin-LR. Comparative biochemistry and


85

Chapter 2: Aims of the doctoral

project

87

The zebrafish embryo developmental toxicity assay (ZEDTA) is currently being

explored for regulatory acceptance as alternative assay in the drug development

process. Indeed, the ZEDTA bridges the gap between in vitro and in vivo

developmental toxicity testing due to the many advantages of the zebrafish

embryo model. In contrast to in vivo mammalian models, the zebrafish embryo

cannot rely on maternal metabolism, so it depends on its own drug–metabolizing

capacity for the detoxification or bioactivation of a compound. However, there is

some discrepancy in literature regarding the xenobiotic–metabolizing capacity of

zebrafish embryos during organogenesis, which is the exposure window for

developmental toxicity. Since this knowledge is pivotal with regards to the

predictivity of the ZEDTA for human risk assessment, the main goal of the

current doctoral project was to characterize drug disposition in zebrafish

during organogenesis with a main focus on CYP–mediated metabolism.

Since drug metabolism in human embryos and fetuses is immature, the drug–

metabolizing capacity of zebrafish embryos during early development is

expected to be negligible as well. Hence, we hypothesize that zebrafish lack the

intrinsic biotransformation capacity to detoxify or bioactivate xenobiotics

during organogenesis.

To test this hypothesis the following research objectives were addressed:

1. Obtain an overall view of CYP–mediated metabolism during zebrafish

organogenesis by performing an in vitro study in which microsomes

prepared from whole zebrafish embryo homogenates at several

developmental time–points were exposed to a fluorogenic non–specific

CYP substrate, i.e. benzyloxy–methyl–resorufin (BOMR). As a reference

for the embryos, CYP activity was also assessed in adult zebrafish liver

microsomes (ZLM) and in microsomes prepared from whole adults, using

the same substrate (Chapter 3 and 4). In addition, we investigated

whether ZLM are able to metabolize a human CYP3A4–specific substrate,

i.e. Luciferin–IPA, considering the predominant role of CYP3A4 in human

drug metabolism (Chapter 3).

88

2. Localization of CYP–mediated biotransformation in intact zebrafish

embryos during organogenesis by exposing them to BOMR. Zebrafish

embryos exposed to the CYP1–specific 7–ethoxyresorufin (ER) were

included as a positive control (Chapter 4). As such, the spatio–temporal

pattern of CYP–mediated metabolism in zebrafish embryos during

organogenesis can be assessed.

3. Get a more complete view of drug disposition in the zebrafish embryo by

performing a gene ontogeny experiment in which the mRNA expression

levels of several CYP (phase I) enzymes as well as phase II enzymes and

a drug transporter were assessed at different time–points during

organogenesis (Chapter 4).

4. Investigate drug disposition in zebrafish larvae beyond organogenesis by

performing the same CYP activity assays and drug disposition gene

analysis as in objectives 1, 2 and 3 at different time points during larval

development (Chapter 4). This way, we aimed to get a view on (full)

maturation of the CYP enzymes and the impact of key events that happen

during the larval period such as the onset of exogenous feeding and

complete yolk absorption.

89

Chapter 3: In Vitro Biotransformation

of Two Human CYP3A Probe

Substrates and Their Inhibition

during Early Zebrafish Development

Adapted from:

In Vitro Biotransformation of Two Human CYP3A Probe Substrates and Their

Inhibition during Early Zebrafish Development.

International Journal of Molecular Sciences. 2017; 18 (1): 217.

DOI 10.3390/ijms18010217

Evy Verbueken, Derek Alsop, Moayad A. Saad, Casper Pype, Els M. Van Peer,

Christophe R. Casteleyn, Chris J. Van Ginneken, Joanna Wilson and Steven J.

Van Cruchten

91

1 Abstract

At present, the zebrafish embryo is increasingly used as an alternative animal

model to screen for developmental toxicity after exposure to xenobiotics. Since

zebrafish embryos depend on their own drug-metabolizing capacity, knowledge

of their intrinsic biotransformation is pivotal in order to correctly interpret the

outcome of teratogenicity assays. Therefore, the aim of this in vitro study was to

assess the activity of cytochrome P450 (CYP)—a group of drug-metabolizing

enzymes—in microsomes from whole zebrafish embryos (ZEM) of 5, 24, 48, 72,

96 and 120 h post-fertilization (hpf) by means of a mammalian CYP substrate, i.e.

benzyloxy-methyl-resorufin (BOMR). The same CYP activity assays were

performed in adult zebrafish liver microsomes (ZLM) to serve as a reference for

the embryos. In addition, activity assays with the human CYP3A4-specific

Luciferin isopropyl acetal (Luciferin-IPA) as well as inhibition studies with

ketoconazole and CYP3cide were carried out to identify CYP activity in ZLM. In

the present study, biotransformation of BOMR was detected at 72 and 96 hpf;

however, metabolite formation was low compared with ZLM. Furthermore,

Luciferin-IPA was not metabolized by the zebrafish. In conclusion, the capacity

of intrinsic biotransformation in zebrafish embryos appears to be lacking during

a major part of organogenesis.

2 Introduction

The zebrafish (Danio rerio) embryo has emerged as an alternative animal

model for developmental toxicity—also called teratogenicity—screening of new

drugs and environmental pollutants (reviewed by [1-4]). The widespread use of

the zebrafish is mainly due to its many advantages such as its short generation

time and high fecundity resulting in 100–200 eggs per mating, which makes the

use of zebrafish embryos less time-consuming in comparison with in vivo

mammalian developmental toxicity studies [5]. Moreover, zebrafish embryos and

larvae can be used in medium—or high—throughput screening because of their

92

small size (0.5–4 mm) [6] and, as the embryos can be kept in small volumes (100

µL), only a small amount of compound is required (reviewed by [2]). The latter is

very useful during early drug development when the availability of a new

chemical entity (NCE)—as defined by the Food and Drug Administration (FDA)

[7]—is still very low. Additionally, the legislation of the European Union

concerning animal experimentation does not consider it to be a test animal until

120 h post-fertilization (hpf), i.e. the stage of independently feeding [8,9].

Zebrafish embryos develop ex utero and the embryos as well as their chorion are

optically transparent, which makes them suitable for microscopic observation of

(internal organ) malformations at different developmental time points [5,6]. The

lack of a maternal barrier during zebrafish development implies direct exposure

of the embryo to the parent compound in teratogenicity assays, while mammalian

embryos/fetuses are exposed to the parent compound and its metabolites due to

drug metabolism by, predominantly, the dam’s liver. Hence, zebrafish embryos

depend on their own drug-metabolizing capacity for detoxification and/or

bioactivation of xenobiotics. The latter is particularly important for compounds

that require bioactivation to exert their teratogenic potential, i.e. so-called

proteratogens. A lack of intrinsic biotransformation in the zebrafish embryo can

lead to false negative results in teratogenicity assays as proteratogens will be

missed. Since drug metabolism, i.e. phase I (mainly oxidation) (reviewed by [10])

and phase II (conjugation) metabolism (reviewed by [11]), in human embryos and

fetuses was shown to be immature, the drug-metabolizing capacity of zebrafish

embryos during early development is expected to be negligible as well. In

addition, the zebrafish liver and intestine—two important drug-metabolizing

organs—develop late in organogenesis, i.e. between 72 and 96 hpf, which

supports our hypothesis concerning the lack of intrinsic biotransformation by

zebrafish embryos. This hypothesis cannot be tested by just exposing zebrafish

embryos to known mammalian proteratogens, as has been done previously [12],

because this in vivo approach does not distinguish between teratogenic effects

caused by the parent compound or by its metabolite. For in vivo studies, also

other pharmacokinetic factors besides metabolism, such as absorption,

distribution and excretion, may determine the exposure in the zebrafish embryo

93

and thus also the teratogenic outcome. In the present in vitro study, these

confounding factors were excluded by focusing only on the intrinsic metabolizing

capacity of zebrafish (embryos). For this purpose, we used microsomes—

subcellular fractions of endoplasmic reticulum containing cytochrome P450

(CYP) isoenzymes—from whole embryo homogenates and from adult zebrafish

livers.

CYP enzymes represent a superfamily of hemoproteins from which the

CYP1, CYP2 and CYP3 families are mainly involved in the oxidative metabolism

of xenobiotics in man (reviewed by [13,14]). Furthermore, the human CYP3A

subfamily metabolizes approximately 50% of drugs that undergo oxidative

biotransformation (Table 1) (reviewed by [15]). CYP-mediated drug-metabolism

predominantly occurs in the liver, whereas other tissues such as the intestine,

brain, lung, kidney, skin, gonads, etc., contribute to a smaller extent (reviewed by

[15,16]). Goldstone and colleagues (2010) [17] were able to identify the full suite of

CYP genes in zebrafish and suggested that also in adult zebrafish the CYP

families 1–3 and, to a lesser extent, CYP4s are involved in the biotransformation

of xenobiotics. Nevertheless, zebrafish CYP3A genes do not phylogenetically

cluster with mammalian CYP3A genes, so differences in CYP3A activity between

zebrafish and mammals can be expected [18,19]. Besides the identification of CYPs

in adult zebrafish, Goldstone et al. also demonstrated distinct temporal patterns

of CYP expression over the course of zebrafish development [17]. In addition to

the research of Goldstone et al. (2010), other in vitro and in vivo studies have

already been performed on the expression and activity of CYP1 and, to a lesser

extent, CYP3 enzymes in adult and developing zebrafish [20-35]. However, results

from these studies are inconclusive and in some cases even contradictory. The

latter is most likely due to differences in study design, such as in vitro versus in

vivo, using mRNA versus protein versus activity level (induced versus basal CYP

activity), other developmental time points, quantitative versus qualitative

measurements, other substrates/substrate concentrations, etc. Therefore, the

drug-metabolizing capacity of zebrafish embryos still remains a point of debate

and requires further investigation.

94

The aim of the present in vitro drug metabolism study was to assess intrinsic

CYP activity in zebrafish embryos of 5–120 hpf and, as a reference for the

embryos, in the adult zebrafish liver. The activity assays were performed by

means of two mammalian CYP substrates, i.e. benzyloxy-methyl-resorufin

(BOMR) (Vivid® CYP450 Screening Kits User Guide 2012) and Luciferin isopropyl

acetal (Luciferin-IPA) [36,37], which are supposed to be metabolized by the

pharmacologically important CYP3A enzyme. Since the zebrafish liver develops

late in organogenesis, microsomes were prepared from the whole embryonic

body so as to take all the organs of the developing zebrafish into account. In

addition to the activity assays, inhibition studies with CYP inhibitors were

performed in adult zebrafish liver microsomes to distinguish between CYP–

mediated metabolism and non–CYP–mediated metabolism. To this end, the non-

specific and concentration-dependent CYP inhibitor ketoconazole [38], the pan–

CYP inhibitor 1–aminobenzotriazole [39], and the CYP3A4-specific inhibitor

CYP3cide [40] were used as inhibitors. The inhibition studies with CYP3cide as

well as the activity assays with Luciferin-IPA showed differences between

zebrafish and mammalian CYP3A activity, which is in concordance with the

phylogenetic difference in CYP3A gene expression. Furthermore, the results of

the present study support our hypothesis regarding the lack of intrinsic

biotransformation by zebrafish embryos as the latter were not able to metabolize

BOMR during a major part of organogenesis.

Table 1. Most important drug-metabolizing cytochrome P450 (CYP) enzymes in man:

relative abundance in human liver and contribution to oxidative biotransformation

of drugs (reviewed by [15,41]).

CYP Isoform Content in Liver (% of

Total CYP)

% of Drugs Metabolized by

CYP

CYP3A4/5 ±30 ±50

CYP2D6 ±4 ±30

CYP2B6 2–10 ±25

CYP2C8, -2C9, -2C19 ±20 ±16

CYP1A2 ±13 ±4

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3 Materials and Methods

3.1 Fish maintenance and breeding

Adult zebrafish (Danio rerio, in house wild-type AB zebrafish line) were

housed in glass aquaria of 60 L with filtration system at a density of <1 fish/L.

Fish were kept in reverse osmosis water to which commercial sea salts (Instant

Ocean® Sea Salt, Blacksburg, VA, USA) and sodium bicarbonate (VWR, Leuven,

Belgium) were added in order to obtain pH and conductivity values of 7.5 ± 0.3

and 500 ± 40 µS/cm, respectively. The water temperature was set to 28 ± 1 °C and

the fish were subjected to an automated light-dark cycle of 14/10 h. Water

parameters and fish health were checked daily and water was renewed once in a

fortnight to keep the levels of ammonia (NH3), nitrite (NO2−) and nitrate (NO3−)

below the detection limits, i.e. NH3 < 0.02 mg/L, NO2− < 0.3 mg/L and NO3− ≤ 12.5

mg/L. Fish were fed twice daily with thawed food—alternating Artemia nauplii,

Daphnia and Chironomidae larvae (Aqua Mila, Deurne-Diest, Belgium)—and

once daily with granulated food (sturgeon food Duvo+, Laroy Group™,

Wondelgem, Belgium) [5].

For the collection of zebrafish embryos, adult fish were transferred to a

spawning tank the day before mating. The next morning, eggs were collected 45

min after the light was turned on. Subsequently, feces and coagulated eggs were

removed by washing the embryos in freshly prepared egg water—same

composition as the adult fish medium—with pH and conductivity set to 7.5 and

480 µS/cm, respectively [5]. The zebrafish embryos were kept in egg water using

a density of 1 embryo/mL and under the same environmental conditions of light

and temperature as for the adults. Dead embryos were removed daily and egg

water was renewed every 48 h. The zebrafish embryos were raised until they

reached the desired developmental stage.

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3.2 Tissue sampling

3.2.1 Adult zebrafish

Only one sex was used for liver collection as hepatic CYP1A activity was

shown to be independent of gender in zebrafish [30]. Six batches of adult female

zebrafish between six months and one year of age were utilized for the

preparation of zebrafish liver microsomes. Batch 1 and Batch 2 were used for the

optimization of the activity assays as well as for the inhibition studies, for which

an adequate amount of microsomal protein is needed. Hence, Batch 1 and 2

consisted of 65 and 100 individuals per batch, respectively, whereas the

remaining batches (3–6) consisted of 10 animals per batch. After a food

deprivation period of 48 h, the fish were euthanized by rapid destruction of the

brain and decapitation [42]. The gastrointestinal system was carefully removed

from the zebrafish body, followed by identification and isolation of the liver. In

order to prevent bile contamination, the gall bladder was carefully discarded.

During the dissection process, livers were rinsed with pre-cooled washing buffer

(10 mM potassium phosphate (KPO4) buffer (BD Gentest™, Woburn, MA, USA)

containing 1.15% potassium chloride (KCl) (Analar Normapur®, VWR, Leuven,

Belgium) at pH 7.4). Liver samples were immediately snap-frozen in liquid

nitrogen and stored at −80 °C until the isolation of ZLM. The animal protocols

applied in this study were evaluated and approved by the Ethical Committee of

Animal Experimentation from the University of Antwerp (Antwerp, Belgium)

(ECD 2015-49; 18 September 2015).

3.2.2 Zebrafish embryos

For each developmental stage—i.e. 5, 24, 48, 72, 96 and 120 hpf—three

batches of zebrafish embryos were used and each batch consisted of

approximately 2500 embryos. When they reached the desired developmental

stage, the embryos were snap-frozen in liquid nitrogen and stored at −80 °C to be

used for microsomal protein preparation.

97

3.3 Isolation of microsomes

3.3.1 Adult zebrafish

The protocol for the isolation of ZLM is based on the one described by Hill

[43] regarding the preparation of rat liver microsomes. All homogenization steps

were performed on ice. Prior to homogenization, liver samples were thawed on

ice, weighed and washed with pre-cooled homogenization buffer (10 mM KPO4

buffer containing 1.15% KCl, 1 mM ethylenediaminetetraacetic acid (EDTA) and

one unit of Halt™ Protease Inhibitor Single-Use Cocktail per 10 mL buffer (the

latter two were purchased from Thermo Fisher Scientific, Waltham, MA, USA) at

pH 7.4) in order to remove possible remnants of hemoglobin. Subsequently, for

each gram of liver tissue, a twofold volume in a milliliter of homogenization

buffer was added. The tissue was then homogenized manually in a glass tube by

means of a Potter-Elvehjem PTFE pestle. As a final homogenization step, samples

were subjected to ultrasonication for (5 × 5) s with intervals of 10 s and an

amplitude of 75% using an Ultrasonic Processor VCX 130 (Sonics & Materials Inc.,

Newton, CT, USA). The homogenate was centrifuged at 12,000× g for 20 min at 4

°C, using a Heraeus™ Multifuge™ X3R Centrifuge (Thermo Fisher Scientific). In

order to remove the fat layer that had been accumulated on the surface of the

resulting supernatant, an additional centrifugation step was performed at 12,000×

g for 10 min at 4 °C. The purified supernatant—containing the S9-fraction—was

then subjected to ultracentrifugation at 100,000× g for 60 min at 4 °C, using an

Optima™ MAX-XP ultracentrifuge (Beckman Coulter, Indianapolis, IN, USA).

The resulting pellet was resuspended in homogenization buffer followed by a

second ultracentrifugation step at 100,000× g for 40 min at 4 °C. Finally, the

resulting microsomal pellet was resuspended in storage buffer (100 mM KPO4

buffer containing 250 mM sucrose (Sigma–Aldrich, St. Louis, MO, USA), 1 mM

EDTA and 1 unit of Halt™ Protease Inhibitor Single-Use Cocktail per 10 mL

buffer), aliquoted and stored at −80 °C until further use. The microsomal protein

concentration of the ZLM was determined by means of the microplate procedure

of the Pierce™ BCA Protein Assay Kit with bovine serum albumin as a standard

(Thermo Fisher Scientific).

98

3.3.2 Zebrafish embryos

The ZEM were isolated according to the same protocol as described for the

adults. However, a few changes, which will be outlined in the current section,

were made. First, during the homogenization of the embryos, no washing steps

were performed since contamination with hemoglobin was considered

negligible. Second, an additional centrifugation step at 12,000× g for 10 min at 4

°C was carried out to remove the high load of melanophores that had been

accumulated in the supernatant.


microsomes

The fluorogenic substrate benzyloxy-methyl-resorufin (Vivid® BOMR

Substrate, P2865, Thermo Fisher Scientific) (Figure 1) was used in order to assess

CYP activity in ZLM. According to the Vivid® CYP450 Screening Kits User Guide

(2012), BOMR is predominantly metabolized by human CYP3A4. Prior to activity

assessment, assay conditions were optimized for substrate and microsomal

protein concentration by testing a range of six protein concentrations of ZLM

(12.5–400 µg/mL) and seven concentrations of BOMR (0.15–9.6 µM). The optimal

microsomal protein concentration and optimal substrate concentration was 200

µg/mL and 1.2 µM, respectively, both values being situated within the linear part

of the reaction curve. All CYP activity assays were performed in non-binding

black polystyrene 96-well microplates with flat bottom and chimney wells

(655900, Greiner Bio-One International GmbH, Kremsmünster, Austria). Positive

and negative controls were included in each assay and were subjected to the same

protein and substrate concentrations as for the ZLM. Pooled human liver

microsomes (Gibco™, HMMCPL–PL050B, Thermo Fisher Scientific) and CYP3A4

Baculosomes® Plus Reagent rHuman (P2377, Thermo Fisher Scientific) were

utilized as positive control. Insect Cell Control Supersomes™ (456201, Corning

Incorporated, Corning, NY, USA), lacking CYP enzymes, were chosen as negative

control. A total incubation volume of 100 µL/well was used. The microsomal

reaction was initiated in each well by the addition of substrate solution containing

1.2 µM BOMR, 0.1 mM NADP+ (Vivid® NADP+, P2879, Thermo Fisher Scientific),

99

3.33 mM glucose-6-phosphate, 0.3 U/mL glucose-6-phosphate dehydrogenase

(Vivid® Regeneration System, P2878, Thermo Fisher Scientific) and 100 mM KPO4

buffer (pH 7.4) to the microsomal solution containing 20 µg/100 µL microsomal

protein and 100 mM KPO4 buffer (pH 7.4). Subsequently, fluorescence was

measured for 60 min with 2-min intervals using a Tecan Infinite® 200 PRO

microplate reader (Tecan Group Ltd., Männedorf, Switzerland) at λex 550 nm and

λem 590 nm. During measurement, the temperature was kept at 28 °C which is

within the zebrafish’s optimal water temperature range of 26–28.5 °C [5]. The

same temperature was utilized for the controls as, in a previous literature report

[30], similar CYP activities could be detected for HLM at 28.5 °C and 37 °C,

respectively. The concentration of resorufin (nM)—a metabolite of BOMR (Figure

1)—produced at each time point was determined from a standard curve that had

been established by using the pure fluorescent metabolite (Vivid® Red

Fluorescent Standard, P2874, Thermo Fisher Scientific). The average values of the

negative control were subtracted from the individual result values obtained for

ZLM, HLM and CYP3A4 BAC. Reaction velocities were calculated in units of

picomoles of resorufin formed per minute per milligram of microsomal protein

(pmol/min/mg MP). The lower limit of detection (LLOD) and the lower limit of

quantification (LLOQ) were 3.41 nM (0.17 pmol/min/mg MP) and 7.58 nM (0.39

pmol/min/mg MP), respectively. For each batch of ZLM, three technical replicates

of the activity assay were performed.

Figure 1: Schematic representation of the oxidative biotransformation of benzyloxy–

methyl–resorufin (BOMR) by cytochrome P450 (CYP) enzymes. The BOMR substrate

contains two potential sites of oxidation (1 and 2 in the figure) that can lead to the release

of a fluorescent metabolite, i.e. resorufin. The benzylic position (marked in green) is

suggested to be the more likely site of metabolism as hydrogen abstraction creates a more

stable radical intermediate [44].

H HH H

1 2

Benzyloxy-methyl-resorufin (BOMR) Resorufin

CYP450

+ H2CO + benzaldehyde

+ benzyl ester

100

3.5 Benzyloxy-methyl-resorufin assay in microsomes from whole

zebrafish embryo homogenates

Similar to the adult zebrafish, the fluorogenic substrate BOMR was used in

order to assess the capacity of biotransformation in zebrafish embryos from 5 to

120 hpf. Pooled human liver microsomes (Gibco™, HMMCPL–PL050B, Thermo

Fisher Scientific) and CYP3A4 Baculosomes® Plus Reagent rHuman (P2377,

Thermo Fisher Scientific) were utilized as positive control and Insect Cell Control

Supersomes™ (456201, Corning Incorporated) were chosen as negative control.

Additionally, ZLM of Batch 1 were included to be used as a reference for the

different developmental stages. Therefore, the activity assays with ZEM were

performed according to the same protocol and at the same microsomal protein

(200 µg/mL) and substrate (1.2 µM) concentration as described for the

experiments using ZLM. At the end of the assay, fluorescence was measured for

60 min with 2-min intervals at 28 °C using a Tecan Infinite® 200 PRO microplate

reader (Tecan Group Ltd.) at λex 550 nm and λem 590 nm. Resorufin concentration

and reaction velocities (pmol/min/mg MP) were calculated in a similar way as for

the ZLM. The average values of the negative control were subtracted from the

individual result values obtained for the ZEM, ZLM, HLM and CYP3A4 BAC.

For each batch of ZEM, three technical replicates of the activity assay were

performed.

3.6 Inhibition studies with adult zebrafish liver microsomes

3.6.1 Ketoconazole

Inhibition studies in ZLM were performed by co-incubation of the BOMR

substrate with ketoconazole (K1003, Sigma–Aldrich), which is known to be a non-

specific but potent inhibitor of human CYP3A. As inhibition studies require an

adequate amount of microsomal protein, Batch 1 and Batch 2 of ZLM were used

due to their large sample size. Positive and negative controls were similar to those

used in the activity assays with BOMR. Since inhibition studies show higher

sensitivity when performed at low substrate and microsomal protein

concentrations [38,45], the latter were set to 0.8 µM BOMR and 100 µg/mL

microsomal protein, both within the linear part of the reaction curve. Inhibition

101

assays were executed in non-binding black polystyrene 96-well microplates with

flat bottom and chimney wells (655900, Greiner Bio-One International GmbH)

with a total incubation volume of 100 µL/well. A range of seven ketoconazole

concentrations (0.005–20 µM) was pre-incubated with 10 µg/100 µL microsomal

protein diluted in 100 mM KPO4 buffer (pH 7.4) for 10 min. Subsequently, the

microsomal reaction was initiated in each well by the addition of 0.8 µM BOMR,

0.1 mM NADP+ (Vivid® NADP+, P2879, Thermo Fisher Scientific), 3.33 mM

glucose-6-phosphate and 0.3 U/mL glucose-6-phosphate dehydrogenase (Vivid®

Regeneration System, P2878, Thermo Fisher Scientific) diluted in 100 mM KPO4

buffer (pH 7.4) to the pre-incubated mixture. Finally, measurements of

fluorescence and calculations of resorufin concentrations and reaction velocities

were executed the same way as for the activity assays with BOMR. Additionally,

IC50 values—concentrations of ketoconazole to cause 50% inhibition of original

CYP activity—were determined for ZLM as well as for the positive controls. For

each batch of ZLM, two technical replicates of the inhibition assays were

performed.

3.6.2 CYP3cide

Inhibition studies with CYP3cide (PZ0195, Sigma–Aldrich)—a mechanism-

based and CYP3A4-specific inhibitor [39]—were carried out for ZLM of Batch 1

and Batch 2. The assays with CYP3cide were performed according to the same

protocol as described for ketoconazole. However, a few adjustments were made.

First, two separate ranges of CYP3cide concentrations (0.03–2 µM and 0.004–4

µM) were used. Second, as biotransformation of CYP3cide is required to exert its

inhibitory potential, an NADPH-regenerating system containing 1.3 mM NADP+,

3.3 mM glucose-6-phosphate, 0.4 U/mL glucose-6-phosphate dehydrogenase and

3.3 mM magnesium chloride (451220 and 451200, Corning Incorporated) was

added to the pre-incubated mixture. The microsomal reaction was then initiated

by the addition of BOMR diluted in 100 mM KPO4 buffer (pH 7.4). IC50 values of

CYP3cide were calculated for ZLM and positive controls. For Batch 1 and Batch

2 of ZLM, two technical replicates of the CYP3cide assays were performed.

102

3.6.3 Preliminary study with 1–aminobenzotriazole

In view of the preliminary in vivo studies in which zebrafish larvae are co–

incubated with 1–aminobenzotriazole and BOMR (Chapter 4), inhibition studies

with 1–aminobenzotriazole (A3940, Sigma–Aldrich) were first performed in vitro

in ZLM of Batch 2. 1–Aminobenzotriazole is widely used as a non–specific and

mechanism–based inhibitor of mammalian CYP enzymes, i.e. also called “pan–

CYP inhibitor” [39]. Similar to the CYP3cide assay, ZLM were pre–incubated with

a range of nine 1–aminobenzotriazole concentrations (3.9–1000 µM) and an

NADPH-regenerating system containing 1.3 mM NADP+, 3.3 mM glucose-6-

phosphate, 0.4 U/mL glucose-6-phosphate dehydrogenase and 3.3 mM

magnesium chloride (451220 and 451200, Corning Incorporated). However, the

pre–incubation time was extended to 30 minutes. The same microsomal protein

and BOMR concentrations were used as described for the inhibition studies with

ketoconazole and CYP3cide. As in the CYP3cide assays, the microsomal reaction

was initiated by the addition of BOMR diluted in 100 mM KPO4 buffer (pH 7.4).

The same assays were performed in HLM as a positive control. IC50 values of 1–

aminobenzotriazole were calculated for ZLM and HLM. For ZLM as well as for

HLM, two technical replicates of the 1–aminobenzotriazole assay were

performed.

3.7 Benzyloxy-methyl-resorufin assay in CYP Baculosomes®

In these activity assays, CYP1A2, CYP2B6, CYP2C9, CYP2C19, CYP2D6 and

CYP3A4 Baculosomes® Plus Reagent, rHuman (P2792, P3028, P2378, P2570, P2283

and P2377, respectively, Thermo Fisher Scientific) were used. CYP Baculosomes®

are microsomes prepared from insect cells transfected with cDNA encoding for

the above-mentioned CYP isoforms. A BOMR concentration of 3 µM was applied

as recommended by the Vivid® CYP450 Screening Kits User Guide (Life

Technologies™, Thermo Fisher Scientific). For all CYP Baculosomes®, a

microsomal protein concentration of 35 µg/mL was utilized, which represents the

mean of the protein concentrations advised for the six different CYP

Baculosomes®. Insect Cell Control Supersomes™ (456201, Corning Incorporated)

103

were included as negative control and were subjected to the same protein and

substrate concentrations as for the CYP Baculosomes®. A total incubation volume

of 100 µL/well was used. The microsomal reaction was initiated in each well

according to the same protocol as described for the BOMR assay with ZLM.

Fluorescence was measured in a similar way as for the ZLM, except for the

temperature that was kept at 37 °C (human body temperature) since this was

recommended by the manufacturer. The concentration of resorufin produced at

each time point was determined from a standard curve that had been established

by using the pure fluorescent metabolite (Vivid® Red Fluorescent Standard,

P2874, Thermo Fisher Scientific). The average values of the negative control were

subtracted from the individual result values obtained for the different CYP

Baculosomes®. The velocities of resorufin formation were calculated in

pmol/min/µg total protein. The LLOD and LLOQ for the CYP Baculosomes®

were 0.0005 pmol/min/µg total protein and 0.0016 pmol/min/µg total protein,

respectively. For each of the six CYP Baculosomes®, three technical replicates of

the BOMR assays were performed.

3.8 Benzyloxy-methyl-resorufin assay in recombinant zebrafish CYPs

Zebrafish CYP1A, CYP1B, CYP1C1, CYP1C2 and CYP1D were cloned and

co-expressed with human cytochrome P450 reductase in Escherichia coli (JM109)

and purified, all according to Scornaienchi et al. (2010) [32]. Briefly, each CYP gene

was cloned with the ompA2+ leader sequence, which targets the expressed CYPs

to the bacterial outer membrane. The ompA2+ sequence is excised after the protein

is inserted into the membrane, thereby allowing the expression of the full-length

CYP protein [32]. Each CYP gene/ompA2+ sequence was ligated into a pCW

vector, and co-transfected with the human NADPH-CYP reductase ligated into a

pACYC vector [32]. Overnight cultures were treated with ampicillin (50 µg/mL)

and chloramphenicol (25 µg/mL) (Thermo Fisher Scientific), while isopropyl β-D-

1-thiogalactopyranoside (1 mM, Thermo Fisher Scientific) was added when

cultures had reached an OD600 between 0.7 and 1.0. Expression of each CYP was

optimized with the addition of 0.1 to 1 mM δ-aminolevulinic acid (MP

Biomedicals, Santa Ana, CA, USA). Cells were allowed to express the CYP

104

proteins for 20–24 h, after which time they were harvested and the bacterial

membranes were purified. Total protein was determined for each CYP stock

using a bicinchoninic acid assay kit (Thermo Fisher Scientific).

Resorufin generation from BOMR was determined for each of the zebrafish

CYPs. The volume of all the reactions was 50 µL, which were performed in black

384-well plates (Thermo Fisher Scientific). Enzyme buffer consisted of 50 mM

TrisHCl and 100 mM NaCl (Thermo Fisher Scientific) adjusted to pH 7.8. NADPH

(Sigma–Aldrich) was prepared daily, and the final NADPH concentration in each

reaction was 150 µM. Total protein concentrations ranged between 2 and 2.5

µg/50 µL reaction for the different CYPs. Fluorescence was measured in the CYP

reactions for 8 min at 1-minute intervals using a BioTek Synergy2 microplate

reader (BioTek U.S., Winooski, VT, USA) at λex 540 nm and λem 590 nm. Reactions

were held at 29 °C. For the first experiment, eight BOMR concentrations (0.055–

40 µM, plus a 0 µM control-dimethyl sulfoxide only) were tested in duplicate

wells. The second experiment was conducted at 1.5 µM BOMR in triplicate wells,

given that resorufin generation rates decreased at BOMR concentrations above

this level for CYP1A, CYP1B and CYP1C2. The LLOD and LLOQ for the zebrafish

CYPs were 0.0008 pmol/min/µg total protein, and 0.0023 pmol/min/µg total

protein, respectively.

3.9 Luciferin-IPA assay with adult zebrafish liver microsomes

This activity assay was performed with Luciferin-IPA (P450–Glo™ CYP3A4

Assay, V9001, Promega Corporation, Madison, WI, USA), which is a highly

specific luminogenic substrate for human CYP3A4 [36,37]. Pooled human liver

microsomes (Gibco™, HMMCPL–PL050B, Thermo Fisher Scientific) and CYP3A4

Baculosomes® Plus Reagent rHuman (P2377, Thermo Fisher Scientific) were

utilized as positive control and Insect Cell Control Supersomes™ (456201,

Corning Incorporated) were used as negative control. For ZLM (Batch 1 and 2)

and HLM, the optimal microsomal protein concentration and optimal substrate

concentration was determined by testing a range of five protein concentrations

(25–400 µg/mL) and a range of six Luciferin-IPA concentrations (1–32 µM),

respectively. Since metabolite concentrations for ZLM were below the LLOQ, the

105

optimal microsomal protein concentration (200 µg/mL) and the optimal substrate

concentration (4 µM) obtained for HLM were applied in the Luciferin-IPA assays.

All assays were performed in non-treated Nunc™ F96 Microwell™ white

polystyrene plates (236205, Thermo Fisher Scientific) with a total incubation

volume of 50 µL/well. Prior to the initiation of the microsomal reaction, 10 µg/50

µL microsomal protein diluted in 100 mM KPO4 buffer (pH 7.4) was pre-

incubated with 4 µM Luciferin-IPA substrate for 10 min. Subsequently, the

microsomal reaction was initiated in each well by the addition of NADPH-

regenerating system containing 1.3 mM NADP+, 3.3 mM glucose-6-phosphate, 0.4

U/mL glucose-6-phosphate dehydrogenase and 3.3 mM magnesium chloride

(451220 and 451200, Corning Incorporated) in 100 mM KPO4 buffer (pH 7.4) to

the pre-incubation mixture. The final reaction mixture was then incubated for 10

min at 37 °C (optimal temperature for HLM) followed by the addition of 50 µL of

Luciferin Detection Reagent diluted in Reconstitution Buffer (V859A and V144A,

Promega Corporation) to each microplate well to stop the microsomal reaction.

Consequently, a luminescent signal was initiated. Subsequently, the luminescent

signal was stabilized by incubation of the mixture for 20 min at room

temperature. Finally, luminescence was measured using a Tecan Infinite® 200

PRO microplate reader (Tecan Group Ltd.). The concentration of D-Luciferin

(metabolite of Luciferin-IPA) was determined by comparing luminescence from

the microsomal reactions to that from a D-Luciferin standard curve (Beetle

Luciferin, Potassium Salt, E1601, Promega Corporation). For the positive controls

as well as for the ZLM, the average values of the negative control were subtracted

from the individual result values. Reaction velocities were calculated in

pmol/min/mg MP and the LLOD and LLOQ were 0.77 nM (0.38 pmol/min/mg

MP) and 1.74 nM (0.87 pmol/min/mg MP), respectively. For each batch of ZLM,

three technical replicates of the activity assays were performed.

3.10 Mathematical and statistical analyses

For all activity and inhibition assays, reaction velocities were calculated

within the linear part of the reaction curve. LLOD and LLOQ were determined

as described by Şengül [46]. Optimal substrate concentrations were determined

106

by nonlinear regression analysis using the substrate inhibition model in

GraphPad Prism (version 6.05; GraphPad Software, Inc., La Jolla, CA, USA).

Calculation of reaction velocities was performed in Microsoft Excel® 2010

(Microsoft Corporation, Redmond, WA, USA). The results from the CYP activity

assays with BOMR were statistically analyzed using IBM SPSS Statistics (version

23; IBM, Armonk, NY, USA). A nonparametric Levene’s test was used to test

homogeneity of variances for EM of 72 hpf and 96 hpf and for ZLM.

Subsequently, the results for these age groups were subjected to a Kruskal–Wallis

test, followed by pairwise comparisons (Mann–Whitney test) to detect differences

between the groups. Differences were considered statistically significant when p

≤ 0.05. Estimation of IC50 values was performed by a nonlinear regression analysis

with a four-parameter logistic curve in GraphPad Prism (version 6.05; GraphPad

Software, Inc.).

4 Results

4.1 Benzyloxy-methyl-resorufin assay in adult zebrafish liver micro–

somes and in microsomes from whole zebrafish embryo homogenates

CYP activity was assessed in adult zebrafish liver microsomes (ZLM) and in

microsomes from whole zebrafish embryo homogenates (ZEM) of 5–120 hpf by

means of the benzyloxy-methyl-resorufin (BOMR) assay. In these experiments,

the reaction velocities obtained for ZLM served as a reference for the values of

the ZEM. The ZLM were able to convert BOMR into the fluorescent metabolite

resorufin, i.e. mean reaction velocity of three technical replicates ± standard

deviation (S.D.): 16.28 ± 3.70, 24.95 ± 5.91, 16.63 ± 1.29, 10.52 ± 3.15, 10.12 ± 0.45

and 17.44 ± 1.35 pmol/min/mg microsomal protein (MP) for Batch 1, 2, 3, 4, 5 and

6, respectively (Figure 2). In ZEM, resorufin formation was only observed at 72

and 96 hpf, i.e. 0.42 ± 0.38 pmol/min/mg MP and 0.39 ± 0.09 pmol/min/mg MP,

for the respective developmental stages (Figure 2). These latter values were close

to the lower limit of quantification (LLOQ) and significantly lower than those of

the ZLM (p = 0.020 for both comparisons). The reaction velocity of human liver

107

microsomes (HLM) and CYP3A4 Baculosomes® (CYP3A4 BAC) (positive

controls) was 12.46 ± 1.41 pmol/min/mg MP and 6.96 ± 1.49 pmol/min/mg MP,

respectively.

Figure 2. Resorufin formation (pmol/min/mg microsomal protein) by

microsomes of zebrafish embryos (ZEM) at 72 and 96 h post-fertilization (hpf)

and by liver microsomes from adult female zebrafish (ZLM) after incubation

with benzyloxy-methyl-resorufin (BOMR). The dots are the reaction velocities

for each batch. Each dot represents the mean value of three technical replicates.

The horizontal solid line represents the mean reaction velocity of the biological

replicates for ZEM and ZLM. The mean reaction velocities for human liver

microsomes (HLM) and CYP3A4 Baculosomes® (CYP3A4 BAC) were added to

the graph as positive controls. The horizontal dotted line represents the lower

limit of quantification (LLOQ). Significant differences (p < 0.05) between age

groups are indicated by different letters (A and B).

ZE

M 7

2 h

pf

ZE

M 9

6 h

pf

ZL

M

HL

M

CY

P3A

4 B

AC

0

1 0

2 0

3 0

Re

so

ru

fin

fo

rm

ati

on

(p

mo

l/m

in/m

g M

P)

A A

B

LLO Q

108

4.2 Inhibition studies with adult zebrafish liver microsomes

4.2.1 Ketoconazole and CYP3cide

Inhibition studies with ketoconazole and CYP3cide were performed in ZLM

of Batch 1 and Batch 2 to detect whether these compounds were able to inhibit

the biotransformation of BOMR. Figure 3a,d show the mean of the results for

Batch 1 and Batch 2 of ZLM. In our study, ketoconazole strongly inhibited the

formation of resorufin in ZLM (Figure 3a) and in CYP3A4 BAC (Figure 3c),

whereas inhibition of BOMR metabolism was less pronounced in HLM (Figure

3b). In contrast to ketoconazole, CYP3cide did not inhibit the metabolism of

BOMR in ZLM (Figure 3d) and inhibition in HLM was limited (Figure 3e).

However, CYP3cide strongly inhibited CYP activity in CYP3A4 BAC (Figure 3f).

Two ranges of CYP3cide concentrations (0.03–2 µM and 0.004–4 µM) were used

in our study, showing similar results.

0 .0 0 1 0 .0 1 0 .1 1 1 0 1 0 0

0

5 0

1 0 0

1 5 0

L o g [K e to c o n a z o le ] M

% o

f c

on

tro

l v

elo

cit

y

(a)

ZLMIC50 = 0.5428 (0.3169 – 0.9299)

0 .0 0 1 0 .0 1 0 .1 1 1 0

0

5 0

1 0 0

1 5 0

L o g [C Y P 3 c id e ] M

% o

f c

on

tro

l v

elo

cit

y

ZLM

(d)

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Figure 3. The effect of various concentrations of ketoconazole—0.005; 0.02; 0.08; 0.31;

1.25; 5 and 20 µM—and CYP3cide—0.004; 0.02; 0.06; 0.25; 1 and 4—on the

biotransformation of BOMR. The dots in the graphs represent the percentage ratios

of reaction velocity in case of pre-incubation of the microsomes with the respective

inhibitor, divided by the control velocity without inhibitor. The values on the X–axis,

i.e. concentration of inhibitor, are logarithmically transformed and indicated with an

antilog numbering format. Graphs (a–c) show the results for pre-incubation with

ketoconazole with (a) demonstrating the mean of the results for Batch 1 and Batch 2

of ZLM ± standard deviation (S.D.); whereas (b,c) show the mean values of the

technical replicates ± S.D. for human liver microsomes (HLM) and CYP3A4

Baculosomes® (CYP3A4 BAC), respectively; Graphs (d–f) show the outcome for pre-

incubation with 0.004–4 µM of CYP3cide (data for 0.03–2 µM of CYP3cide not

shown) with (d) representing the mean of the results for Batch 1 and Batch 2 of ZLM

± S.D.; while (e,f) demonstrate the mean values of the technical replicates ± S.D. for

HLM and CYP3A4 BAC, respectively. In case of inhibition, the IC50 values and their

0 .0 0 1 0 .0 1 0 .1 1 1 0 1 0 0

0

5 0

1 0 0

1 5 0


% o

f c

on

tro

l v

elo

cit

yHLM

IC50 = 7.987 (0.6011 – 106.1)

(b)

0 .0 0 1 0 .0 1 0 .1 1 1 0

0

5 0

1 0 0

1 5 0


% o

f c

on

tro

l v

elo

cit

y

HLM

(e)

0 .0 0 1 0 .0 1 0 .1 1 1 0 1 0 0

0

5 0

1 0 0

1 5 0


% o

f c

on

tro

l v

elo

cit

y

BACIC50 = 0.2432 (0.1829 – 0.3234)

0 .0 0 1 0 .0 1 0 .1 1 1 0

0

5 0

1 0 0

1 5 0


% o

f c

on

tro

l v

elo

cit

y

(c)

BACIC50 = 0.0259 (0.01422 – 0.04717)

(f)

110

95% confidence intervals are added. The S.D. is not shown if the error bar is shorter

than the height of the dot.

4.2.2 Preliminary study with 1–aminobenzotriazole

A preliminary inhibition study with 1–aminobenzotriazole was

performed in ZLM of Batch 2 to detect whether this compound was able to

inhibit the biotransformation of BOMR prior to the use of this inhibitor in an

in vivo study which is described in Chapter 4. In the current in vitro study,

1–aminobenzotriazole inhibited the formation of resorufin in ZLM (IC50

value of 116.3 µM; mean of two technical replicates) and HLM (IC50 value of

74.1 µM; mean of two technical replicates), although 1–aminobenzotriazole

is a less potent inhibitor of BOMR metabolism compared with ketoconazole.

4.3 Benzyloxy-methyl-resorufin assay in cytochrome P450 (CYP)

Baculosomes® and in recombinant zebrafish CYPs

With the aim of determining whether BOMR is a CYP3A4-specific substrate,

CYP Baculosomes® expressing human CYP1A2, CYP2B6, CYP2C9, CYP2C19,

CYP2D6 and CYP3A4 were used. These CYP Baculosomes® were selected as they

represent the most important CYP enzymes involved in drug metabolism in man

(Table 1). The activity assays showed that human CYP2C9 and in particular

CYP1A2, CYP2B6 and CYP3A4 enzymes were able to convert the BOMR

substrate into the highly fluorescent resorufin (Table 2), whereas no

biotransformation of the substrate could be observed for CYP2D6 Baculosomes®.

Regarding CYP2C19, only two replicates showed values above the LLOQ, while

no resorufin was detected in the third replicate. In addition to the human CYP

Baculosomes®, activity assays with BOMR were performed in recombinant

zebrafish CYPs, which showed that the substrate was clearly biotransformed by

recombinant CYP1A and to a lesser extent by CYP1B, CYP1C1 and CYP1C2 (Table

2). Resorufin formation was below the LLOQ for CYP1D.

111

Table 2. Overview of resorufin formation by CYP Baculosomes® (BAC) with 3 µM

BOMR and by recombinant zebrafish CYPs with 1.5 µM BOMR.

Recombinant CYPs Resorufin Formation

CYP Baculosomes® 1 pmol/min/µg Total Protein

CYP1A2 BAC® 0.047 ± 0.009

CYP2B6 BAC® 0.084 ± 0.023

CYP2C9 BAC® 0.012 ± 0.001

CYP2C19 BAC® <LLOQ

CYP2D6 BAC® <LLOQ

CYP3A4 BAC® 0.042 ± 0.022

Recombinant Zebrafish CYPs 1 pmol/min/µg Total Protein

CYP1A 1.152 ± 0.068

CYP1B 0.105 ± 0.008

CYP1C1 0.004 ± 0.001

CYP1C2 0.078 ± 0.011

CYP1D <LLOQ 1 Mean value of three technical replicates ± standard deviation. LLOQ, lower limit of

quantification; BOMR, benzyloxy-methyl-resorufin; CYP, cytochrome P450.

4.4 Luciferin-IPA assay with adult zebrafish liver microsomes

The luminogenic substrate Luciferin-IPA was used to investigate whether

ZLM are able to convert this human CYP3A4-specific substrate into D-Luciferin.

However, for all batches, metabolite concentrations were below the LLOQ (mean

reaction velocity of six batches ± S.D.: 0.28 ± 0.16 pmol/min/mg MP) (Figure 4). In

contrast to ZLM, reaction velocities of the positive controls were considerably

higher: 463.90 ± 117.28 pmol/min/mg MP and 82.60 ± 43.99 pmol/min/mg MP for

HLM and CYP3A4 BAC, respectively. Since no biotransformation of Luciferin-

IPA could be observed for ZLM, luminogenic activity assays were not performed

in zebrafish embryos.

112

Figure 4. D-Luciferin formation (pmol/min/mg microsomal protein) by liver

microsomes from adult female zebrafish. Each dot represents the mean reaction

velocity (mean value of three technical replicates) ± standard deviation (S.D.) for

the corresponding batch of adult zebrafish liver microsomes. The lower

horizontal dotted line demonstrates the lower limit of detection (LLOD) and the

upper horizontal dash-dotted line represents the lower limit of quantification

(LLOQ). S.D. not shown if the error bar is shorter than the height of the dot.

5 Discussion

In view of zebrafish embryos being extensively used in developmental

toxicity studies, the present study contributes to a better understanding of the

drug-metabolizing capacity of zebrafish embryos. Since CYP enzymes are

predominantly involved in the metabolism of xenobiotics, CYP activity assays

were performed in zebrafish embryos at different developmental time points and

in liver microsomes from adult zebrafish, which served as a reference for the

embryos.

The CYP activity assays with BOMR in ZLM showed reaction velocities that

were in the same range of those of HLM. This similarity is not surprising as

Batc

h 1

Batc

h 2

Batc

h 3

Batc

h 4

Batc

h 5

Batc

h 6

0 .0

0 .5

1 .0

1 .5

Z e b ra f is h liv e r m ic ro s o m e s

D-L

uc

ife

rin

fo

rm

ati

on

(p

mo

l/m

in/m

g M

P)

L L O Q

L L O D

113

Goldstone et al. (2010) [17] identified CYP1, CYP2 and CYP3 families in adult

zebrafish and suggested that these enzymes are involved in the

biotransformation of xenobiotics as described in humans (reviewed by [15]).

Nevertheless, the human CYP3 family only consists of the CYP3A subfamily,

from which CYP3A4 plays a predominant role in drug metabolism [41], whereas

in zebrafish more CYP3 subfamilies have been characterized, i.e. the CYP3A65

isoform [17,35] and the CYP3C1–3C4 isoforms [22,34]. The CYP3A65 gene and

CYP3C1 gene were first described by Tseng et al. (2005) [35] and Corley-Smith et

al. (2006) [22], respectively, with both genes showing high expression levels in the

liver and intestine of adult zebrafish. The remaining CYP3C genes demonstrated

rather variable levels of expression in the zebrafish gastrointestinal system [34].

Since our study indicated that BOMR was metabolized by Baculosomes®

expressing human CYP3A4 and, as most of the CYP3 genes are expressed in the

zebrafish liver, one could assume that the CYP3 family also contributes to the

metabolism of BOMR in ZLM. However, caution is required since our study

showed that BOMR was clearly metabolized by recombinant zebrafish CYP1A,

which was in line with the results for the human recombinant CYPs as BOMR

was also metabolized by recombinant human CYP1A2, CYP2B6 and to a lesser

extent by CYP2C9. Our results are also in accordance with an earlier study in

which another substrate of human CYP3A4, i.e. 17β-estradiol, was clearly

metabolized by recombinant zebrafish CYP1s [32]. Moreover, we found that ZLM

were not able to metabolize the Luciferin-IPA, which is a highly specific substrate

for human CYP3A4 [36,37]. A possible explanation for the latter finding is a

difference in structure between the active site of human and zebrafish CYP3A

enzymes resulting in different drug-metabolizing capacities [47]. This hypothesis

is supported by the research of Goldstone et al. (2010) [17] who found that the

CYP3A65 gene was identical to human CYP3A4 for only 54%. Furthermore,

concerning the CYP3C family, a study of Corley-Smith et al (2006) [22] revealed

that the amino acid sequence of CYP3C1 was only 44%–49% similar to the

mammalian CYP3A. In humans, CYP3A and CYP2C subfamilies have evolved as

essentially xenobiotic metabolizing enzymes. Moreover, identification of the

surface binding–pockets of CYP3A and CYP2C showed that both enzyme

114

proteins are characterized by the haem–containing active site being exposed at

the protein surface, whereas the active site of essentially physiological CYP

families is located far from the protein surface [48,49]. Hence, the analysis of the

active site’s shape as well as its ‘buriedness’, i.e. its location towards the protein

surface, would gain insight into the role of individual zebrafish CYP enzymes in

xenobiotic metabolism [48,49]. Another explanation for Luciferin–IPA not being

metabolized by ZLM could be differences in the evolutionary tree as zebrafish

CYP3A genes do not phylogenetically cluster with mammalian CYP3A genes,

resulting in different functions of the corresponding enzymes [18,19]. Besides the

incapability of ZLM to biotransform a CYP3A4-specific substrate, ZLM and HLM

also showed different inhibition profiles when co-incubating BOMR and

ketoconazole. This dissimilarity may be due to species-differences in CYP

isoforms that interact with BOMR and/or with ketoconazole. Indeed, inhibition

by ketoconazole has already been described as being species-specific and

concentration-dependent [45,50]. In humans, ketoconazole acts as a non-specific,

but potent mixed competitive-noncompetitive inactivator of CYP3A [38]. This

observation was confirmed in the present study by the inhibition of the

biotransformation of BOMR in CYP3A4 BAC. In contrast to ketoconazole, co-

incubation of BOMR with CYP3A4-specific CYP3cide did not inhibit the velocity

of resorufin formation by ZLM. Moreover, inhibition of BOMR metabolism by

CYP3cide in HLM was less pronounced compared with inhibition by

ketoconazole. These findings imply that more than one CYP isoform is involved

in the biotransformation of BOMR in zebrafish and humans as evidenced by the

reaction phenotyping experiments with recombinant zebrafish CYPs and CYP

Baculosomes®, respectively. Based upon the results above, we can conclude that

BOMR, which was supposed to be metabolized predominantly by human

CYP3A4 [44,51], is a non-specific CYP substrate in humans and in zebrafish. In

other studies, similar conclusions were drawn for 7-benzyloxy-4-

(trifluoromethyl)-coumarin (BFC), which was assumed to be specific for

mammalian CYP3A, but appeared to be metabolized by bacterially expressed

CYP3A and CYP1 from zebrafish [32,33]. Moreover, 7–benzyloxyresorufin, of

which the chemical structure is very similar to BOMR, was shown to be

115

metabolized by human CYP1A, CYP2B and CYP3A subfamilies (reviewed by

[33]); [52]. The latter is in accordance with our study on the biotransformation of

BOMR by CYP Baculosomes® since the activity assays showed that human

CYP1A2, CYP2B6 and CYP3A4 enzymes were able to convert the BOMR

substrate. Furthermore, Scornaienchi et al. (2010) [33] showed that 7–

benzyloxyresorufin was clearly metabolized by recombinant zebrafish CYP1A,

CYP1B1, CYP1C1 and CYP1C2, which is also in accordance with the results from

the present study regarding the biotransformation of BOMR by recombinant

zebrafish CYPs. In addition, based on the structures of BOMR and 7–

benzyloxyresorufin, we might assume that the latter is converted into resorufin

and benzaldehyde, which is similar to oxidation site 1 (shown in green) in the

schematic representation of BOMR (Figure 1). Hence, we hypothesize that similar

CYP (sub)families are involved in the oxidative biotransformation of 7–

benzyloxyresorufin and BOMR into the fluorescent metabolite, i.e. resorufin.

In brief, our research showed that the adult zebrafish liver possesses CYP activity

required to metabolize the non-specific CYP substrate BOMR. Regarding the

potential contribution of CYP3A65 and/or CYP3C isoforms in BOMR

biotransformation, our study remains inconclusive mainly due to a lack of

recombinant enzymes of these isoforms. Since

Zebrafish embryos, however, were not capable of metabolizing BOMR before

72 hpf. Taking the LLOD and LLOQ parameters into account, a small amount of

resorufin could be detected at the end of organogenesis, i.e. at 72 and 96 hpf, but

the values were still negligible compared to resorufin formation by ZLM. This is

not surprising as these time points coincide with major development of the liver

and intestine, two pivotal drug-metabolizing organs. The liver gets vascularized

around 72 hpf, and reaches its adult configuration around 96 hpf [53,54]. For the

intestine, the lumen and epithelium develop craniocaudally between 54 and 102

hpf [55]. These morphological data were further substantiated by a whole-mount

in situ hybridization study in which CYP3A65 mRNA was detected in the liver at

72 hpf, followed by expression of the gene in the intestine at 84 and 96 hpf [35].

Our data are also in accordance with earlier in vitro and in vivo studies on CYP1A

activity. Indeed, in vitro assessment of basal CYP1A activity in zebrafish embryos

116

and larvae using 7-ethoxyresorufin-O-deethylase (EROD) also demonstrated low

levels of metabolite formation around 72 and 96 hpf, with even lower levels

around 120 hpf [29,56]. Even after CYP induction, activity levels remained low

and a similar temporal trend was observed for basal versus induced CYP1

activity [20,29,31]. In vivo CYP1A activity first appeared in the liver primordium

around 56 hpf, followed by the intestine at approximately 80 hpf and reaching a

peak at 104 hpf [29]. Furthermore, Alderton et al. (2010) [57] demonstrated that

zebrafish larvae of three and seven days post-fertilization (dpf) were able to

metabolize human CYP probe substrates and drugs, albeit to a small extent.

Therefore, these authors postulated that the low metabolite concentrations are

unlikely to contribute to the malformations in developmental toxicity studies [57].

In addition, Chng and colleagues (2012) [21] showed that zebrafish larvae of 120

hpf produced only two CYP metabolites of testosterone compared with the

formation of multiple CYP metabolites in liver microsomes from adult zebrafish.

Similar to our findings, these authors thus suggested a difference in function or

expression of drug-metabolizing enzymes between larval and adult zebrafish

[21]. Finally, in contrast to the in vitro CYP1A activity that has been reported for

the whole zebrafish embryo as early as at five hpf and at eight hpf [29,30], BOMR

was not metabolized by these earliest embryonic stages in our study. The

presence of CYP activity in the early zebrafish embryo before its basic body plan

has been established, is maternally derived [17] and disappears quickly. Since

BOMR is a non-specific CYP substrate and clear spatio-temporal differences in

CYP isoform expression have been reported [17], this could explain the lack of

maternally derived CYP activity in our study.

6 Conclusions

In conclusion, this in vitro study demonstrated that the non-specific CYP

substrate BOMR was metabolized by adult zebrafish liver microsomes as well as

by human liver microsomes. In contrast to the adults, zebrafish embryos were not

capable of metabolizing BOMR during a major part of organogenesis. In most

117

teratogenicity assays with zebrafish embryos, this would not be an issue since

toxicity is mainly caused by the parent compound itself. However, in case of

proteratogenic compounds, which require biotransformation to exert their

teratogenic potential, false negative results can occur if the drug-metabolizing

capacity in the zebrafish embryo is lacking. Hence, our study indicates that

zebrafish embryos have a poor CYP-related metabolizing capacity during

organogenesis, which needs to be considered in regards to their use as an

alternative animal model in developmental toxicity studies. This information

needs to be further strengthened by using other CYP substrates and investigating

other phase I reactions and the role of phase II enzymes as well.

118

7 References

1. Barros, T.P.; Alderton, W.K.; Reynolds, H.M.; Roach, A.G.; Berghmans, S.

Zebrafish: an emerging technology for in vivo pharmacological assessment to

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123

Chapter 4: From mRNA Expression of

Drug Disposition Genes to In Vivo

Assessment of CYP-Mediated

Biotransformation during Zebrafish

Embryonic and Larval Development

Adapted from:

From mRNA Expression of Drug Disposition Genes to In Vivo Assessment of

CYP-Mediated Biotransformation during Zebrafish Embryonic and Larval

Development.

International Journal of Molecular Sciences. 2018; 19 (12): 3976.

DOI 10.3390/ijms19123976

Evy Verbueken, Chloé Bars, Jonathan S. Ball, Jelena Periz-Stanacev, Waleed F. A.

Marei, Anna Tochwin, Isabelle J. Gabriëls, Ellen D. G. Michiels, Evelyn Stinckens,

Lucia Vergauwen , Dries Knapen, Chris J. Van Ginneken and Steven J. Van

Cruchten

125

1 Abstract

The zebrafish (Danio rerio) embryo is currently explored as an alternative for

developmental toxicity testing. As maternal metabolism is lacking in this model,

knowledge of the disposition of xenobiotics during zebrafish organogenesis is

pivotal in order to correctly interpret the outcome of teratogenicity assays.

Therefore, the aim of this study was to assess cytochrome P450 (CYP) activity in

zebrafish embryos and larvae until 14 d post-fertilization (dpf) by using a non-

specific CYP substrate, i.e. benzyloxy-methyl-resorufin (BOMR) and a CYP1-

specific substrate, i.e. 7-ethoxyresorufin (ER). Moreover, the constitutive mRNA

expression of CYP1A, CYP1B1, CYP1C1, CYP1C2, CYP2K6, CYP3A65, CYP3C1,

phase II enzymes uridine diphosphate glucuronosyltransferase 1A1 (UGT1A1)

and sulfotransferase 1st1 (SULT1ST1), and an ATP-binding cassette (ABC)

transporter, i.e. abcb4, was assessed during zebrafish development until 32 dpf

by means of quantitative PCR (qPCR). The present study showed that trancripts

and/or the activity of these proteins involved in disposition of xenobiotics are

generally low to undetectable before 72 h post-fertilization (hpf), which has to be

taken into account in teratogenicity testing. Full capacity appears to be reached

by the end of organogenesis (i.e. 120 hpf), although CYP1—except CYP1A—and

SULT1ST1 were shown to be already mature in early embryonic development.

2 Introduction

The thalidomide tragedy in the late fifties and early sixties resulted in the

obligatory use of a second, non-rodent, animal species in developmental toxicity

studies. This second species, in most cases the rabbit, has proven to be very

effective, as no cases of human birth defects that had not been flagged in animal

studies have been reported ever since [1,2]. However, in view of cost and time

effectiveness, and within the framework of the 3Rs—Replacement, Reduction,

and Refinement—described by Russell and Burch (1959) [3], the zebrafish (Danio

126

rerio) embryo has been proposed as an alternative non-rodent animal model for

developmental toxicity studies. Indeed, the zebrafish embryo is not considered to

be a test animal until it reaches the stage of independent feeding, i.e. at 120 h post-

fertilization (hpf) (Figure 1) [4,5]. Moreover, zebrafish are characterized by a rapid

and ex utero embryonic development and embryos may be used in medium- or

high-throughput screening because of their small size [6]. Hence, the zebrafish

embryo developmental toxicity assay (ZEDTA) considers the physiological

parameters of a whole organism together with the advantages of an in vitro

model. Due to these benefits, several pharmaceutical companies and contract

research organizations (CROs) have already adopted the ZEDTA as an early

screening method to reduce the number of compounds that need to be tested in

a mammalian model (reviewed by [7]). Further efforts are ongoing to explore

regulatory acceptance of the ZEDTA in the drug development process [8,9]. At

the same time, potential regulatory acceptance of the fish embryo acute toxicity

test (FET) for chemical toxicity testing—according to the test guideline TG 236 of

the Organization for Economic Co-operation and Development (OECD) [10]—is

under consideration as an alternative for the fish acute toxicity test, i.e. TG 203

[11,12]. The FET (chemicals) uses exposure windows between 1.5 and 96 hpf [10],

whereas the ZEDTA (pharmaceuticals) commonly uses exposure windows

between 4 and 120 hpf to ensure that the entire zebrafish organogenesis period is

covered [6,8,9,13,14].

The zebrafish liver and intestine, which are pivotal in the biotransformation

of xenobiotics, become functional towards the end of the organogenesis period,

i.e. around 96 hpf (Figure 1) [5,15-17]. Since zebrafish embryos develop ex utero,

they are directly exposed to the parent compound in developmental toxicity

assays. Only the chorion, which surrounds the embryo until 48–72 hpf [6], may

serve as a barrier for certain compounds depending on their physicochemical

properties [18]. Hence, zebrafish embryos depend on their intrinsic

biotransformation capacity for the detoxification of xenobiotics and/or

bioactivation of so-called proteratogens. In mammals, cytochrome P450 (CYP)

families CYP1, CYP2 and CYP3 are involved in the oxidative (phase I)

127

metabolism of xenobiotics as well as endogenous compounds such as steroids

(reviewed by [19-21]). Goldstone and colleagues (2010) [22] suggested that also in

adult zebrafish the CYP families 1–3 are involved in the biotransformation of

xenobiotics. In humans and several laboratory mammals, CYP-mediated

biotransformation capacity was shown to be immature during embryo-fetal

development [23,24]. However, these embryos/fetuses can rely on maternal

metabolism of the compound.

Based on the knowledge regarding the poor CYP-mediated drug metabolism

in mammalian embryos/fetuses and the relatively late functional development of

the zebrafish digestive system, i.e. liver and intestine, we hypothesize that the

intrinsic CYP-mediated biotransformation capacity in zebrafish embryos is

immature during early development. This implies that proteratogenic

compounds may lead to false negative results in developmental toxicity studies

if zebrafish embryos do not have the capacity to bioactivate those compounds.

The hypothesis has been tested by an earlier in vitro study in which intrinsic CYP

activity was assessed in microsomes—artificial subcellular fractions of

endoplasmic reticulum containing CYPs—from whole zebrafish embryo

homogenates at different time points between 5 and 120 hpf by means of a

fluorogenic non-specific CYP substrate, i.e. benzyloxy-methyl-resorufin (BOMR)

[25]. Biotransformation of BOMR into the fluorescent metabolite, i.e. resorufin, is

a measure for the CYP activity in the microsomes. Since the liver and the intestine

are the predominant sites for CYP-mediated drug metabolism (reviewed by [20]),

intrinsic CYP activity was also assessed in liver microsomes prepared from adult

female zebrafish as a reference for the embryos. In contrast to adults, zebrafish

embryos showed no CYP-mediated metabolizing capacity in vitro during a major

part of organogenesis, i.e. between 5 and 72 hpf, only poor CYP activity at 72 and

96 hpf and no CYP activity at 120 hpf [25]. Besides this in-house in vitro study,

other research groups assessed (often after exposure to CYP inducers or

inhibitors) CYP1 activity and, to a lesser extent, CYP3 activity during zebrafish

organogenesis by using substrates that are specific for the respective CYP

enzymes [26-40]. However, the overall results of these studies regarding the

128

xenobiotic-metabolizing capacity of zebrafish embryos and larvae are

contradictory, as some authors claim that zebrafish embryos show CYP-mediated

biotransformation of xenobiotics [29,40], whereas others report that the extent of

CYP-mediated biotransformation, e.g. metabolite concentrations, in zebrafish

embryos and larvae is very low and unlikely to be relevant [27,39].

Therefore, the first aim of the present study was to further investigate the

development of CYP activity in microsomes (in vitro) and in intact (in vivo)

zebrafish embryos and larvae. However, regarding zebrafish embryos “in vivo”

does not mean in vivo sensu stricto since the embryos are not considered to be a

test animal until 120 hpf. As we noted a decrease in CYP activity at the end of

zebrafish embryonic development, i.e. at 120 hpf, in a previous in vitro study [25],

we wondered whether CYP-mediated biotransformation further matures, and if

so, when. Therefore, we extended the developmental stages beyond the period of

organogenesis (120 hpf) in the present study, i.e. including 9 and 14 d post-

fertilization (dpf). At 96–120 hpf, exogenous feeding starts (Figure 1) and

exposure to environmental compounds is expected to increase in a natural

situation, which may activate the pregnane X receptor (PXR) or the aryl

hydrocarbon receptor (AhR) that regulate CYP expression [41,42]. Hence, the

onset of exogenous feeding may affect CYP activity in zebrafish larvae.

Furthermore, larvae between 8 and 15 dpf often show increased mortality due to

starvation in the period between complete yolk absorption and successful

exogenous feeding (Figure 1), and this may affect CYP activity [5,43]. Besides an

in vitro assessment, we also localized CYP-mediated biotransformation in intact

zebrafish embryos and larvae, as organ-specific concentrations of the metabolite

may be diluted when using microsomes prepared from whole organisms. Besides

a non-specific CYP substrate, i.e. BOMR, we also included the CYP1-specific 7-

ethoxyresorufin (ER) as a positive control substrate in the in vivo assay since the

ethoxyresorufin-o-deethylase (EROD) assay is a well-established method in

ecotoxicology to investigate the AhR-mediated induction of CYP1 enzymes by

ubiquitous environmental contaminants such as 2,3,7,8-tetrachlorodibenzo-p-

dioxin (TCDD) [26,44-46]. Furthermore, disposition of xenobiotics and

129

endogenous compounds not only relies on phase I CYP-mediated

biotransformation but also involves phase II reactions in which the parent

compound or phase I metabolites are conjugated with a hydrophilic moiety, and

cellular efflux by transporters in excretion organs, such as the liver, and barrier

organs, such as the intestine (reviewed by [47]). In order to get a more complete

view of the disposition in the zebrafish embryo and larvae, we decided to

investigate the developmental mRNA expression of two major phase II enzymes

in the zebrafish, i.e. sulfotransferase 1st1 (SULT1ST1) and uridine diphosphate

glucuronosyltransferase 1A1 (UGT1A1) [48-51], and the ATP-binding cassette

(ABC) transporter (abcb4). The latter was assessed since this transporter possesses

similar multixenobiotic resistance (MXR) properties as the well-characterized

mammalian ABCB1 transporter [52,53]. Since the same whole zebrafish body

samples were used as in a study of Vergauwen et al. (2018) [54], the time window

for the mRNA expression analysis was extended to 32 dpf to make the results

comparable between both studies. As in literature, data on the ontogeny of CYP1,

CYP3 and, to a lesser extent, CYP2 mRNA expression in zebrafish are limited to

approximately 6 dpf [22,26,31,32,55-63], we decided to also include the constitutive

mRNA expression of zebrafish CYP1, CYP2, and CYP3 families at different time

points between 1.5 hpf and 32 dpf in our assessment. Similar to the in vivo CYP

activity study with BOMR, the mRNA expression of most CYP enzymes, phase II

enzymes and P-glycoprotein reached maximum expression levels by the end of

zebrafish organogenesis and remained stable throughout larval development.

Hence, the present study showed general CYP-mediated biotransformation in

zebrafish embryos towards the end of organogenesis, which needs to be

considered with regards to the use of zebrafish embryos in ZEDTA and FET.

130

Figure 1. Timeline showing key events during zebrafish development, from

fertilization to juvenile stages (j.) Color bars indicate the developmental phases

with gradients representing embryo-larval and larval-juvenile transitions. The

period of embryonic development includes pre-hatching stage and

eleutheroembryo stage, i.e. the stage between hatching and onset of exogenous

feeding [5,10]. The period of organogenesis, i.e. development of brain, heart,

liver, intestine and pronephros, coincides with embryonic development.

Embryo-larval transition implies the period between the onset of exogenous

feeding and complete yolk absorption. Larval-juvenile transition reflects the

period of metamorphosis in which the larval morphology is transformed into

that of a juvenile (e.g. metamorphosis of the pigment pattern and fin

morphology) [43,64]. Developmental stages of the organogenesis period are

represented as h post-fertilization (hpf). Older developmental stages are shown

as d post-fertilization (dpf). Th: thyroid hormone. The timeline is adapted from

Vergauwen et al. (2018) [54] and based on Chang et al. (2012) [65], Drummond

et al. (1998) [66], Field et al. (2003) [15], Kimmel et al. (1995) [6], Li et al. (2000)

1 2 3 4 5 6 7 8 90 10 10 15 20 25 30 dpf

embryo larva j

organogenesis

0 24 48 72 96 120 hpf

131

[67], Ng et al. (2005) [16], Ober et al. (2003) [17], Parichy et al. (2009) [64], Strähle

et al. (2012) [5] and Wilson et al. (2012) [43].

3 Materials and Methods


larvae and adults

3.1.1 Fish maintenance and breeding

Fish maintenance and breeding: zebrafish embryos

For a description of fish maintenance and breeding with regards to zebrafish

embryos of between 5 hpf and 120 hpf, we refer to Verbueken et al. (2017) [25].

Fish maintenance and breeding: zebrafish larvae

Adult zebrafish (Danio rerio, wild-type AB zebrafish line obtained from

European Zebrafish Resource Center at Karlsruhe Institute of Technology,

Germany), which were used for spawning, were housed in enriched aquaria of

40 L at a density of ≤5 fish/L and at an automated 14/10 h light/dark cycle. Fish

were kept in fish medium, i.e. reverse osmosis water (Environmental Water

Systems Ltd., Cheddar, UK) to which commercial sea salts (Tropic Marin® Sea

Salt, Wartenberg, Germany) and NaHCO3 (Analar Normapur®, VWR

International, Leicestershire, UK) were added in order to obtain pH and

conductivity values of 8 and 350 µS/cm, respectively. The water temperature was

set to 28 °C and levels of ammonia, nitrite and nitrate were kept below the

permissible limits, i.e. NH3 < 0.25 mg/L, NO2− < 0.3 mg/L and NO3− < 20 mg/L.

Adult fish were fed freshly harvested Artemia nauplii (ZM Premium Grade

Artemia, Zebrafish Management Ltd., Winchester, UK) and tropical flake food

(TetraMin, Tetra®, Melle, Germany) twice daily.

Embryos were generated by a group spawning method as detailed in

Gustafson et al. (2012) [9]. Briefly, eggs were collected within 45 min after

132

spawning and incubated in fish medium at 28 ± 1 °C for 1–2 h. Subsequently,

embryos were treated against fungal infection using a dilute Chloramine T

bleaching solution for 60 s with gentle periodic agitation. Following bleaching,

the embryos were washed twice in fish medium with constant agitation, then

transferred into a Petri dish (50 embryos per dish) containing aerated fish

medium at 28 ± 1 °C. Embryos were staged for development according to

methods that have been previously described by Kimmel et al. (1995) [6]. At 120

hpf embryos were transferred to a crystallizing dish container (0.5 embryo/mL)

and raised until 9 or 14 dpf. Fish medium was partially (25%) renewed daily and

larvae were fed with dry feed three times a day according to the following

scheme: ZM-000 (Zebrafish Management Ltd.) for 5–8 dpf, a mix of ZM-000 and

ZM-100 (Zebrafish Management Ltd.) for 9–10 dpf and ZM-100 for 11–14 dpf.

Larvae were euthanized by an overdose of tricaine methane sulfonate (MS222; 2

mg/mL) (Sigma-Aldrich, St. Louis, MO, USA) when they reached the desired

developmental stage, i.e. 9 or 14 dpf. The terminated larvae were snap-frozen

with as less fish medium as possible in liquid nitrogen and stored at −80 °C until

processing. The animal protocols in this study were evaluated and approved by

the UK Home Office regulations and the local ethic committee for the use of

animals in scientific procedures (project number 17-002. 70/98992; August 2016;

Exeter University Animal Welfare and Ethics Review Body). In this research

paper, developmental stages of the organogenesis period are represented as h

post-fertilization (hpf), similar to the time unit used in developmental toxicity

studies. Older developmental stages are shown as d post-fertilization (dpf).

Fish maintenance: adult zebrafish

Zebrafish (Danio rerio, in-house wild-type AB zebrafish line) that were used

for the isolation of microsomes from whole adult homogenates were housed in

an aquarium of 400 L at a density of ≤5 fish/L and at an automated 14/10 h

light/dark cycle. Fish were kept in tap water at 28 °C and levels of ammonia,

nitrite and nitrate were kept below the permissible limits, i.e. NH3 < 0.25 mg/L,

133

NO2- < 0.3 mg/L and NO3- < 20 mg/L. The fish were fed three times a day with

granulated food (Biogran medium, Prodac International, Cittadella, Italy).

Adult zebrafish were euthanized by rapid cooling in ice water at 2–4 °C to

(no physical contact with ice) followed by decapitation and destruction of the

brain [68]. Subsequently, the gall bladder was removed from the body since bile

acids are detrimental for the CYP activity in the sample. The terminated fish were

snap-frozen in liquid nitrogen and stored at −80 °C until processing. Since the

adult zebrafish were used for breeding, no ethical approvement was needed for

the preparation of microsomes from whole adult homogenates.

3.1.2 Tissue collection and isolation of microsomes

Isolation of microsomes from whole zebrafish embryos

For a description of tissue collection and isolation of microsomes with

regards to zebrafish embryos of between 5 hpf and 120 hpf, we refer to Verbueken

et al. (2017) [25].

Isolation of microsomes from whole zebrafish larvae

Two biological replicates of approximately 500 larvae and three biological

replicates of approximately 700 larvae were used for microsomal protein

preparation of 9 and 14 dpf, respectively. The microsomes prepared from whole

zebrafish larvae were isolated according to the same protocol as described by

Verbueken et al. (2017) [25]. Briefly, homogenized zebrafish larvae were

centrifuged at 12,000× g which resulted in a supernatant that contains the S9-

fraction. The resulting supernatant was then subjected to two ultracentrifugation

steps at 100,000× g to render a microsomal pellet. Finally, the microsomal protein

concentration was determined by means of the microplate procedure of the

PierceTM BCA Protein Assay Kit with bovine serum albumin as a standard

(Thermo Fisher Scientific, Waltham, MA, USA).

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Isolation of microsomes from whole adult zebrafish

Three biological replicates, each consisting of seven adult zebrafish of mixed

genders, were used for the preparation of microsomes from whole adult zebrafish

homogenates. Frozen adult fish were homogenized by crushing them into a fine

powder. After the addition of homogenization buffer (10 mM KPO4 buffer

containing 1.15% KCl, 1 mM ethylenediaminetetraacetic acid (EDTA) and 1 unit

of HaltTM Protease Inhibitor Single-Use Cocktail per 10 mL buffer (the latter two

were purchased from Thermo Fisher Scientific, Waltham, MA, USA) at pH 7.4) to

the crushed tissue, an additional homogenization step was performed by means

of a Polytron® System PT 1200 E (Kinematica Inc., Bohemia, NY, USA). As a final

homogenization step, samples were subjected to ultrasonication for (5 × 5) s with

intervals of 10 s and an amplitude of 75% using an Ultrasonic Processor VCX 130

(Sonics & Materials Inc., Newton, CT, USA). The microsomes were isolated from

the whole adult zebrafish homogenates according to the same protocol as

described for the zebrafish embryos and larvae in the current study [25]. Finally,

the microsomal protein concentration was determined by means of the

microplate procedure of the PierceTM BCA Protein Assay Kit with bovine serum

albumin as a standard (Thermo Fisher Scientific, Waltham, MA, USA).

3.1.3 Benzyloxy-methyl-resorufin assay in microsomes prepared from whole

zebrafish embryos, larvae and adults

In a previous study [25], benzyloxy-methyl-resorufin (Vivid® BOMR

Substrate, P2865, Thermo Fisher Scientific) was shown to be a fluorogenic non-

specific CYP substrate in the zebrafish. Biotransformation of BOMR into resorufin

by zebrafish microsomes is a measure for the CYP activity in the respective

microsomes. In this study, BOMR was used to assess CYP activity in microsomes

prepared from (1) whole zebrafish embryo homogenates (ZEM) of 5, 24, 48, 72,

96, and 120 hpf, (2) whole zebrafish larva homogenates (ZLaM) of 9 and 14 dpf

and (3) whole adult zebrafish homogenates (ZM). The ZEM, which had been used

in a former study [25], were included in the assay in order to have a complete

overview of the development of CYP activity in zebrafish embryos and larvae as

135

well as to compare the in vitro data with the in vivo data. A microsomal protein

concentration of 200 µg/mL and a BOMR substrate concentration of 1.2 µM were

used in the activity assay as both values were situated within the linear part of

the reaction curve in an optimization study with adult female zebrafish liver

microsomes (ZLM) [25]. Insect Cell Control Supersomes™ (456201, Corning Inc.,

Corning, NY, USA), lacking CYP enzymes, were chosen as negative control. ZLM

(Batch 3, prepared from the livers of 10 adult zebrafish) that had shown positive

results in a previous study [25] were used as a positive control. Hence, only one

biological replicate of ZLM was used in the current assay. The ZEM (Batch 1–3),

which had already been used in a previous study [25] were included in the assay

to assess CYP activity throughout zebrafish development. Positive and negative

controls and ZEM were subjected to the same protein and substrate concentration

as for ZLaM and ZM. The CYP activity assays were performed in non-binding

black polystyrene 96-well microplates with flat bottom and chimney wells

(655900, Greiner Bio-One International GmbH, Kremsmünster, Austria). A total

incubation volume of 100 µL/well was used. The microsomal reaction was

initiated in each well by the addition of substrate solution containing 1.2 µM

BOMR, 0.1 mM NADP+ (Vivid® NADP+, P2879, Thermo Fisher Scientific), 3.33

mM glucose-6-phosphate, 0.3 U/mL glucose-6-phosphate dehydrogenase (Vivid®

Regeneration System, P2878, Thermo Fisher Scientific) and 100 mM KPO4 buffer

(pH 7.4) (Corning, Discovery Labware Inc., Woburn, MA, USA) to the

microsomal solution containing 20 µg/100 µL microsomal protein and 100 mM

KPO4 buffer (pH 7.4) under light-protected conditions. Subsequently,

fluorescence was measured for 72 min with 150-s intervals using a Tecan Infinite®

200 PRO microplate reader (Tecan Group Ltd., Männedorf, Switzerland) at λex

550 nm and λem 590 nm. During measurement, the temperature was kept at 28 °C

which is within the zebrafish’s optimal water temperature range of 26–28.5 °C

[69]. The concentration of resorufin (nM)—a metabolite of BOMR—produced at

each time point was determined from a standard curve that had been established

by using the pure fluorescent metabolite (Vivid® Red Fluorescent Standard,

P2874, Thermo Fisher Scientific). The average values of the negative control were

136

subtracted from the individual result values obtained for ZLaM, ZEM, ZM, and

ZLM. Reaction velocities were calculated in units of picomoles of resorufin

formed per minute per milligram of microsomal protein (pmol/min/mg MP). For

each batch of ZLaM, six technical replicates of the activity assay were performed

and for each batch of ZM, three technical replicates of the activity assay were

performed (Table 1). Only two technical replicates were included for ZEM of 5–

120 hpf as for the latter, CYP activity had already been assessed [25].

3.1.4 Mathematical and statistical analyses

The reaction velocities were calculated within the linear part of the reaction

curve. The lower limit of detection (LLOD) was 0.07 pmol/min/mg MP, and lower

limit of quantification (LLOQ) was 0.11 pmol/min/mg MP. These limits were

determined based on the mean and standard deviation of the negative control

values as described by Şengül [70]. Calculation of reaction velocities and detection

and quantification limits was performed in Microsoft Excel® 2016 (Microsoft

Corporation, Redmond, WA, USA). The results from the CYP activity assays with

BOMR that showed reaction velocities above the LLOQ were statistically

analyzed using IBM SPSS Statistics (version 25; IBM, Armonk, NY, USA). A

nonparametric Levene’s test was used to test homogeneity of variances for ZEM

of 72 hpf and 96 hpf, ZLaM of 14 dpf and ZM. Subsequently, the results for these

age groups were subjected to a Kruskal-Wallis test, followed by pairwise

comparisons (Mann-Whitney test) to detect differences between the groups.

Differences were considered statistically significant when p ≤ 0.05.

137

Table 1. Comparison of the experimental setup between in vitro and in vivo study.

Experimental Setup In Vitro In Vivo

Developmental stage

5, 24, 48, 72, 96, 120 hpf 1

9 and 14 dpf

Adults

7, 26, 50, 74, 98, 122 hpf

9 and 14 dpf

Samples Microsomes from whole

embryos/larvae/adults Intact embryos and larvae

Substrate CYP activity BOMR BOMR

Negative control/Blank Supersomes Embryos/larvae in fish

medium

Positive control Adult zebrafish liver

microsomes 1

Embryos/larvae incubated

with ER 2

Biological replicates Three (5–120 hpf; 14 dpf; adults)

Two (9 dpf) Three/developmental stage

Technical replicates

Two for 5–120 hpf

Six for 9 and 14 dpf

Three for adults

Two

Detection of resorufin

formation Microplate reader Fluorescence microscope

The microsomes prepared from whole embryos of between 5–120 hpf, which had been used

in a former study [25], were included in the in vitro assay in order to have a complete overview

of the development of CYP activity in zebrafish embryos and larvae as well as to compare the

in vitro data with the in vivo data. 1 Verbueken et al. (2017) [25]; 2 Otte et al. (2010) [34]. Hpf, h

post-fertilization; dpf, d post-fertilization; CYP, cytochrome P450; BOMR, benzyloxy-methyl-

resorufin; ER, 7-ethoxyresorufin.


larvae

3.2.1 Fish maintenance and breeding

Adult zebrafish (Danio rerio, in-house wild-type zebrafish line) were housed

in a ZebTEC zebrafish housing system (Tecniplast, Buguggiate, Italy) at an

automated 14/10 h light/dark cycle, at 28 ± 0.2 °C. Fish were kept in fish medium,

i.e. reverse osmosis water (Werner, Leverkusen, Germany) to which commercial

sea salts (Instant Ocean® Sea Salt, Blacksburg, VA, USA) and NaHCO3 (Analar

Normapur®) were added in order to obtain pH and conductivity values of 7.5 and

500 µS/cm, respectively. Around 35% of the circulating water was renewed daily

to keep levels of ammonia, nitrite and nitrate below the permissible limits, i.e.

138

NH3 < 0.25 mg/L, NO2- < 0.3 mg/L and NO3- < 20 mg/L. Adult fish were fed three

times a day: twice with 0.5% of their mean wet weight of granulated food

(Biogran medium, Prodac International, Cittadella, Italy) and once with thawed

food: alternating Artemia nauplii, Daphnia, Chaoboridae larvae and

Chironomidae larvae (Aquaria Antwerp bvba, Aartselaar, Belgium).

For the collection of zebrafish embryos, two female fish and one male fish

were transferred to a breeding tank and separated from each other the evening

before mating. The next morning, the divider was removed when the lights

turned on. After 30–40 min, eggs were collected from multiple spawning groups

and distributed into plastic beakers with an initial density of 0.4 embryo/mL.

Zebrafish embryos were raised in fish medium that had the same composition as

the water in the ZebTEC housing system and under the same environmental

conditions of light and temperature as for the adults. Dead embryos were

removed daily and the fish medium was renewed every two days until 120 hpf.

From 120 hpf until 9 dpf, fish were kept at a maximum density of 0.4 embryo/mL

and fish medium was renewed once a day and from 9 dpf until 14 dpf twice a

day. Larvae were fed twice daily with paramaecia from 4–6 dpf. From 7–9 dpf,

they were fed paramecia and SDS-100 (Special Diets Services, Essex, UK) twice

daily. From 10–13 dpf, they were additionally fed freshly harvested Artemia

nauplii twice daily. The larvae of 9 and 14 dpf were not fed on the day of the

experiment to limit the amount of food in the gastrointestinal system. For each

developmental stage—i.e. 7, 26, 50, 74, 98, and 122 hpf and 9 and 14 dpf—three

biological replicates were used in the CYP activity assays. At the end of each

assay, embryos and larvae were euthanized by transferring them to a tricaine

methane sulfonate (MS222, Sigma-Aldrich) solution with a final concentration of

1 mg/mL. Fish husbandry and all experiments were carried out in strict

accordance with the EU Directive on the protection of animals used for scientific

purposes (2010/63/EU) [71]. The animal protocols applied in this study were

evaluated and approved by the Ethical Committee of Animal Experimentation

from the University of Antwerp (Antwerp, Belgium) (ECD 2018–08; 05 March

2018).

139

3.2.2 Benzyloxy-methyl-resorufin assay in zebrafish embryos and larvae

Zebrafish embryos and larvae of 7, 26, 50, 74, 98, and 122 hpf and 9 and 14

dpf were incubated with 4 µM BOMR (Vivid® BOMR Substrate, P2865, Thermo

Fisher Scientific) dissolved in fish medium. Zebrafish embryos of 26 hpf were

manually dechorionated prior to incubation with the BOMR substrate. Since

approximately 50% of the embryos of 50 hpf had already spontaneously hatched

at this stage, manually dechorionated and spontaneously hatched embryos were

used in the assay according to a 1/1 ratio. Embryos of 7 hpf were not

dechorionated due to lower survival rates caused by the procedure at this stage

[72]. For each developmental stage, a blank group—embryos/larvae incubated in

fish medium without substrate—was included. Zebrafish embryos/larvae

incubated with 1.7 µM 7-ethoxyresorufin (ER) (Resorufin ethyl ether, Sigma-

Aldrich)—a CYP1-specific substrate—in fish medium were used as positive

control since positive results have been described in literature [30,34]. Each

embryo/larva was transferred in 150 µL of fish medium to a well of a black 96-

Well Cell Imaging Plate with clear 25 µm film bottom (Eppendorf Cell Imaging

Plates, 0030741013, Eppendorf, Hamburg, Germany). Subsequently, 50 µL of the

substrate solution (final concentration/embryo or larva: 4 µM BOMR or 1.7 µM

ER) or 50 µL of fish medium (blank) was added to the embryo/larva followed by

incubation for 60 min at 28.5 °C under light-protected conditions. Following

incubation, each embryo/larva was immobilized by the addition of tricaine

methane sulfonate (MS222, Sigma-Aldrich) with a final concentration of 0.2

mg/mL per embryo or larva. Finally, the formation of resorufin was analyzed by

means of an inverted fluorescence microscope (Olympus IX 71, Olympus

Corporation, Shinjuku, Tokyo, Japan) with a 10x objective at λex 510–550 nm and

λem ≥ 570–590 nm. Grayscale images (8 bit) were acquired by means of the

CellSens Software (Olympus Corporation) using a fixed gain and exposure

setting for all images. A qualitative and quantitative analysis of resorufin

formation was performed in the anterior and posterior trunk region of the

zebrafish embryo/larva (Figure 2). The trunk region was selected for analysis as

it contains the major CYP-containing organ, i.e. the liver, together with some

140

extrahepatic organs that are involved in drug metabolism, i.e. intestine, kidney,

cardiovascular system. Since the trunk has not been developed yet at 7 hpf

(gastrulation period [6]), analysis of metabolite formation was accomplished in

the whole embryo. For each biological replicate, at least four embryos/larvae of

each condition were evaluated from which the two best positioned

embryos/larvae were used for further analysis (Table 1).

Figure 2. Description of region of interest used for the quantitative and qualitative

analysis of resorufin formation in zebrafish embryos and larvae at 7 h post-

fertilization (hpf) (a), 26 hpf (b,c), 50 hpf (d,e), 74 hpf (f,g), 98 hpf (h,i), 122 hpf (j,k),

9 d post-fertilization (dpf) (l,m) and 14 dpf (n,o) after exposure to benzyloxy-methyl-

resorufin (BOMR) or 7-ethoxyresorufin (ER). The yellow frame indicates the region

of interest in the embryo or larva. Since for most embryos/larvae the complete trunk

region did not fit within one image, pictures of anterior and posterior trunk were

taken separately. For the quantitative analysis of resorufin formation in each

embryo/larva, average pixel intensities of anterior and posterior trunk images were

combined. Figure 2 (a) shows a vegetal pole view of the embryo. In figure 2 (b–o)

lateral views of the anterior and posterior part of the trunk region are shown. Scale

bar: 200 µm; (b,c): anterior top and dorsal right; (d–o): anterior left and dorsal top.

142

3.2.3 Preliminary inhibition study in zebrafish embryos of 98 and 122 hpf

A preliminary inhibition study with a CYP inhibitor was performed in intact

zebrafish embryos of 98 and 122 hpf to distinguish between CYP–mediated

metabolism and non–CYP–mediated metabolism. The embryonic stages of 98 and

122 hpf were chosen for this assay since both stages were able to metabolize the

BOMR substrate in the in vivo assay (4.2 of the current Chapter). In Chapter 3 of

the current thesis (Section 4.2.1), in vitro inhibition studies showed that

ketoconazole, i.e. an antifungal drug, is a potent inhibitor of BOMR

biotransformation in adult zebrafish liver microsomes. However, since

ketoconazole has embryotoxic potential as shown in some in vivo mammalian

studies as well as in the embryonic stem cell test [73], 1–aminobenzotriazole was

chosen as inhibitor in the current in vivo study. Indeed, 1–aminobenzotriazole is

widely used as a non–specific and mechanism–based inhibitor of mammalian

CYP enzymes, i.e. also called “pan–CYP inhibitor” [74], and embryotoxic effects

of this inhibitor are unknown. In this preliminary assay, we aimed to assess CYP

inhibition in zebrafish embryos of 98 and 122 hpf in case of exposure to 1–

aminobenzotriazole followed by incubation with BOMR. Zebrafish embryos of 5

hpf were exposed to 1000 µM 1–aminobenzotriazole for 1 hour or 96 hours. At 98

or 122 hpf, the exposed zebrafish embryos were washed prior to the in vivo assay

which was performed according to the same protocol as described in section 3.2.2.

Since the 1000 µM 1–aminobenzotriazole solution contains 0.46%

dimethylsulfoxide (DMSO), a solvent control containing embryos of 98 and 122

hpf exposed to 0.46% DMSO was included in the assay. Zebrafish embryos of 98

and 122 exposed to 0.1 M Tris solution (containing Tris-HCl; CaCl2.2H2O;

MgSO4.7H2O; NaHCO3 and KCl dissolved in reverse osmosis water) were

included as a blank. Only one replicate of the preliminary in vivo inhibition study

was performed.

3.2.4 Mathematical and statistical analyses

Qualitative analysis of resorufin formation was performed by visual

inspection of an overlay (grayscale/bright-field-overlay) image of the trunk

143

region of the embryo/larva. Quantitative analysis of resorufin formation was

performed in a grayscale image by measuring the average pixel intensity within

the trunk region (Figure 2) using the ImageJ software (version 1.50i; National

Institutes of Health, Bethesda, MD, USA). The corrected integrated density of

resorufin was calculated by means of a formula: (average pixel intensity of region

of interest—background average pixel intensity) × area of interest. The LLOD

(integrated density value: 522,443) and LLOQ (integrated density value:

1,421,607) were determined based on the mean and standard deviation of the

corrected integrated density for the blank embryos/larvae as described by Şengül

[70]. Calculation of corrected integrated density and detection and quantification

limits was performed in Microsoft Excel® 2016 (Microsoft Corporation). The

quantitative results from the in vivo assay with BOMR and ER that showed

values above the LLOQ were statistically analyzed using IBM SPSS Statistics

(version 25; IBM). Statistical analysis was performed on the following

developmental stages: 74, 98, 122 hpf, 9 and 14 dpf for the BOMR assay and 7, 26,

50, 74, 98, and 122 hpf for the EROD assay. A nonparametric Levene’s test was

used to test homogeneity of variances. Subsequently, the results for these age

groups were subjected to a Kruskal-Wallis test, followed by pairwise

comparisons (Mann-Whitney test) to detect differences between the groups.

Differences were considered statistically significant when p ≤ 0.05.


glycoprotein

3.3.1 Fish Maintenance and Breeding

The same samples used in the ontogeny study of Vergauwen et al. (2018) [54]

were used here. Adult zebrafish (Danio rerio, in-house wild-type zebrafish line)

were housed under the same environmental conditions as described for the in

vivo study. For the collection of zebrafish embryos, one female and one male fish

were transferred to a breeding tank and separated from each other the evening

before mating. The next morning, the divider was removed when the lights

144

turned on. Within 45 min, eggs were collected and pooled from multiple

spawning groups and randomly distributed into plastic beakers with an initial

density of 45 embryos per 100 mL. The density was gradually decreased to 7

larvae per 100 mL at 10 dpf with gentle aeration initiated at 9 dpf. Zebrafish

embryos were raised in fish medium that had the same composition as the water

in the ZebTEC housing system at 28.5 °C with 14/10 h light/dark cycle. Fish

medium was renewed daily. At 15 dpf, larvae were transferred to a ZebTEC

zebrafish housing system. Fish were fed twice daily with paramaecia from 4–6

dpf. From 7–9 dpf, they were fed paramecia and SDS-100 (Special Diets Services)

twice daily. From 10 dpf, they were additionally fed freshly harvested Artemia

nauplii twice daily. Starting at 15 dpf paramecia feeding was reduced to once

daily. From 20 to 32 dpf, they were fed Artemia nauplii once daily and SDS-100

twice daily.

Embryos/larvae/juveniles were sampled at 25 time points, each time point

containing four independent biological replicates (Table 2). Whole body samples

were snap-frozen in liquid nitrogen and stored at −80 °C until processing. Fish

husbandry and all experiments were carried out in strict accordance with the EU

Directive on the protection of animals used for scientific purposes (2010/63/EU)

[71]. The animal protocols applied in this study were evaluated and approved by

the Ethical Committee of Animal Experimentation from the University of

Antwerp (Antwerp, Belgium) (ECD 2015-51; 18 September 2015).

145

Table 2. Overview of sampling time points for analysis of mRNA expression.

Time Point Hpf Dpf Number of Organisms/Biological Replicate

1 1.5 0.06 30

2 6 0.25 30

3 14 0.58 30

4 24 1 20

5 36 1.5 20

6 48 2 20

7 60 2.5 20

8 72 3 20

9 84 3.5 20

10 96 4 10

11 120 5 10

12 144 6 10

13 192 8 10

14 240 10 10

15 288 12 10

16 336 14 10

17 384 16 10

18 432 18 10

19 480 20 10

20 528 22 10

21 576 24 10

22 624 26 10

23 672 28 10

24 720 30 10

25 768 32 10

Hpf, h post-fertilization; dpf, d post-fertilization.

3.3.2 Quantification of mRNA levels by means of qPCR

For each time point, the mRNA expression of seven phase I enzymes, i.e.

CYP1A, CYP1B1, CYP1C1, CYP1C2, CYP2K6, CYP3A65, and CYP3C1, two phase

II enzymes, i.e. sulfotransferase 1st1 (SULT1ST1) and uridine diphosphate

glucuronosyltransferase 1A1 (UGT1A1), and one P–glycoprotein, i.e. ATP-

binding cassette b4 (abcb4) transporter was analyzed by means of quantitative

polymerase chain reaction (qPCR). Except for CYP2K6, for which primers were

designed in-house, all primer sequences were obtained from literature (Table 3).

146

Most amplicons spanned two exons and the sequence of the amplicons was

confirmed using the National Center for Biotechnology Information’s Basic Local

Alignment Search Tool (NCBI, BLAST) [75] to verify specific sequence alignment

with the targeted gene in the zebrafish genome. All primers were ordered from

Eurogentec (Liège, Belgium).

RNA was extracted from homogenized whole zebrafish body samples using

the NucleoSpin® RNA isolation kit (Macherey-Nagel, Düren, Germany)

according to the manufacturer’s protocol, including a DNAse treatment. RNA

purity and integrity were confirmed using a NanoDrop spectrophotometer

(NanoDrop Technologies, Rockland, DE, USA) and a BioAnalyzer (Agilent

Technologies, Diegem, Belgium). All samples had minimal A260 nm/A280 nm ratios of

2.1 and minimal RNA integrity number (RIN) of 7.9. Complement DNA (cDNA)

was synthesized from the extracted RNA using a RevertAid H Minus First Strand

cDNA Synthesis Kit (Thermo Fischer Scientific) according to the manufacturer’s

instructions, with random hexamer primers. Subsequently, cDNA was diluted to

70 ng/µL in 0.1% diethylpyrocarbonate (DEPC)-treated water prior to its use as a

template for the qPCR reaction. Quantitative PCR reactions were performed in

an MX3005P instrument (Agilent Technologies) using the Brilliant II SYBR®

Green qPCR Master Mix (Agilent Technologies). Each qPCR reaction contained

350 ng cDNA, 10 pmol forward primer and 10 pmol reverse primer in a final

volume of 19.3 µL. Thermal cycling profiles were: an initialization step of 10 min

at 95 °C, followed by 40 cycles of a denaturation step of 20 s at 95 °C, an annealing

step of 40 s at 55 °C (51 °C for CYP2K6) and an elongation step of 30 s at 72 °C.

Melting curves were assessed to confirm specific amplification. Primer

efficiencies were determined using duplicate standard curves with four

concentrations in a 1.5-fold dilution series of a mixed cDNA sample based on

different time points. The same standard curves were included in each qPCR run

to correct for inter-run differences. 18S ribosomal RNA (18S) and beta actin 1

(actb1) (Table 3) were selected from five potential reference genes using geNorm

[76]. Both reference genes were used in further analysis of the qPCR data.

147

Table 3. Primer sequences of zebrafish target genes and reference genes used for

quantitative polymerase chain reaction analyses.

Gene Sequence (5′ to 3′) Accession

Number References

Target

CYP1A

FW:

GCATTACGATACGTTCGATAAGGAC

RV: GCTCCGAATAGGTCATTGACGAT

NM_131879.1 Goldstone et al. (2010) [22]

CYP1B1 FW: GAGCACCGAAAGACCATTTCA

RV: ATGGTCGGTGGCACAAACTC

NM_001045256.1

NM_001145708.1 Olsvik et al. (2014) [77]

CYP1C1 FW: AGTGGCACAGTCTACTTTGAGAG

RV: TCGTCCATCAGCACTCAG NM_001020610.2 Goldstone et al. (2010) [22]

CYP1C2 FW: GTGGTGGAGCACAGACTAAG

RV: TTCAGTATGAGCCTCAGTCAAAC NM_001114849.1 Jönsson et al. (2007) [78]

CYP2K6 FW: CCAGCTTTGTCCCTGTTTCTT

RV: GCAGAGAGTTCAGCCTGTGAT NM_200509.2 Designed in-house

CYP3A65 FW: CTTCGGCACCATGCTGAGAT

RV: AGATACCCCAGATCCGTCCATA NM_001037438.1 Chang et al. (2013) [79]

CYP3C1 FW: TCCAGACCTCTGGGAGTCTCCTAAT

RV: GCATGAAGGCACACTGGTTGATCT NM_212673.1 Shaya et al. (2014) [61]

SULT1ST1 FW: GTTCCTTCTTGGGTTTGTCT

RV: CTGGCAGAGTGGAATAGTTG NM_182941.1 Liu et al. (2011) [80]

UGT1A1 FW: TCCTTTGCCGCAGCATGTAT

RV: ACTCTCTGGCTTTGGCTTCG NM_001037428.2 Wang et al. (2014) [81]

abcb4 FW: TACTGATGATGCTTGGCTTAATC

RV: TCTCTGGAAAGGTGAAGTTAGG NM_001316714.1

NM_001114583.2 Fischer et al. (2013) [53]

Reference

18S

FW: CGGAGAGGGAGCCTGAGAA

RV: AGTCGGGAGTGGGTAATTTGC Biga et al. (2005) [82]

actb1 FW: AAGTGCGACGTGGACA

RV: GTTTAGGTTGGTCGTTCGTTTGA NM_131031 Gonzalez et al. (2006) [83]

hprt1 FW: GTGGCTCTATGTGTGCT

RV: CCTCCACAATCAAGACG NM_212986.1 Bio- Engineering

Com.(Shanghai, China)

rpn2 FW: TTGAGTTCAGCCAGCGT

RV: TGGCAACAAATCGGCG NM_212748.1 De Wit et al. (2008) [84]

ef1a FW: TGTCCTCAAGCCTGGTAT

RV: CATTACCACGACGGATGT NM_131263 Houbrechts et al. (2016) [85]

148

Legend table 3: FW, forward primer; RV, reverse primer; CYP, cytochrome P450; sult,

sulfotransferase; UGT, uridine diphosphate glucuronosyltransferase; abc, ATP-binding

cassette; 18S, 18S ribosomal RNA; actb1, beta actin 1; hprt1, hypoxanthine

phosphoribosyltransferase1; rpn2, ribophorin 2; ef1a, eukaryotic translation elongation factor

1 alpha 1.

3.3.3 Mathematical and statistical analyses.

The transcript abundance of each sample was divided by the geometric mean

of 18S and actb1 transcript abundances in that sample to normalize the

experimental data for reference gene expression [76]. An inter-run calibration was

performed using qbase+ software (version 3.1; Biogazelle, Zwijnaarde, Belgium).

For each gene, the resulting data were divided by the average abundance at the

time point with the lowest expression for that gene and subsequently log2

transformed to increase the resolution. The log2 relative quantities were analyzed

using R Statistical Software (version 3.4.3; RStudio Inc., Boston, MA, USA). The R

code as previously published by Vergauwen et al. (2018) (Supplementary Data in

[54]) was used in the analyses. The aim of the statistical approach was to

determine when mRNA expression data at particular time points significantly

deviate from trends in the data, thereby defining critical points (e.g. local maxima

and minima) of mRNA expression. In brief, local weighted regression (lowess)

along with residual plots were used to identify possible outliers in each dataset.

Next, local regression (loess) with the simplest fit span was utilized to estimate

the non-linear trends in responses for each gene. Selection of the loess model was

verified by confirming that the residuals had no pattern over time. Critical points

(i.e. minima, maxima, and inflection points) in the data were determined when

the derivative of the best-fit function through the data equals 0. Finally, to obtain

confidence intervals around each critical point, bootstrapping techniques were

used to find estimates of the slope of the responses.

149

4 Results


larvae and adults

CYP activity was assessed in microsomes prepared from whole zebrafish

embryo homogenates (ZEM) of between 5 hpf and 120 hpf, in microsomes

prepared from whole zebrafish larva homogenates (ZLaM) of 9 and 14 dpf and

in microsomes prepared from whole adult zebrafish (ZM) by means of the

benzyloxy-methyl-resorufin (BOMR) assay. The ZM were included in the assay

as a reference for the embryos and larvae. The ZEM of 5, 24, 48 and 120 hpf and

ZLaM of 9 dpf were not able to convert BOMR into the fluorescent metabolite, i.e.

resorufin, as metabolite concentrations were negligible. Reaction velocities for

ZEM of 72 and 96 hpf, for ZLaM of 14 dpf and for ZM were above the lower limit

of quantification (LLOQ), i.e. mean reaction velocity of three biological replicates

± standard deviation (S.D.): 0.36 ± 0.35, 0.29 ± 0.13, 0.64 ± 0.09 and 1.34 ± 0.51

pmol/min/mg microsomal protein (MP) for the respective developmental stages

(Figure 3). For the adult zebrafish liver microsomes (ZLM), which were included

as a positive control, a reaction velocity of 9.65 ± 4.23 pmol/min/mg MP (mean

value of six technical replicates for one biological replicate) was observed, which

is in line with our previous study [25]. Furthermore, the BOMR assay with ZEM

of between 5 and 120 hpf showed similar results as in a former study [25]. No

statistically significant differences were detected between 72 hpf and 96 hpf (p =

0.827) and between 72 hpf and 14 dpf (p = 0.275). Statistically significant

differences were detected between 96 hpf and 14 dpf and between ZM and the

earlier stages, i.e. 72 hpf, 96 hpf and 14 dpf (p = 0.050 for all comparisons). ZLM

and developmental stages with values below the LLOQ were not included in the

statistical analysis.

150

Figure 3. Resorufin formation (pmol/min/mg microsomal protein) by

microsomes prepared from whole zebrafish embryos (ZEM) of between 5 and

120 h post-fertilization (hpf), microsomes prepared from whole zebrafish larvae

(ZLaM) at 9 and 14 d post-fertilization and microsomes prepared from whole

adult zebrafish (ZM) after incubation with benzyloxy-methyl-resorufin

(BOMR). The dots are the reaction velocities for each biological replicate. Each

dot represents the mean value of two, three and six technical replicates for ZEM,

ZM and ZLaM, respectively. The horizontal solid line represents the mean

reaction velocity of three biological replicates for each developmental stage. The

horizontal dotted line represents the lower limit of quantification (LLOQ). The

reaction velcoties for 5–48 hpf, 120 hpf and 9 dpf could not be calculated because

of the negligible metabolite concentrations (indicated by *). No statistically

significant differences were detected between 72 hpf and 96 hpf and between 72

hpf and 14 dpf (p > 0.05). Statistically significant differences (p ≤ 0.05) between

96 hpf and 14 dpf and between ZM and the earlier stages, i.e. 72 hpf, 96 hpf and

14 dpf are indicated by different letters (A, B and C) (p = 0.050 for all

comparisons).

ZE

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larvae

Since organ-specific concentrations of resorufin may be diluted when using

microsomes prepared from whole zebrafish embryos and larvae, the aim of the

in vivo study was to localize the biotransformation of BOMR in intact zebrafish

embryos and larvae at 7, 26, 50, 74, 98, 122 hpf, 9 and 14 dpf. A quantitative and

a qualitative analysis of resorufin formation in the trunk region of each

embryo/larva was performed.

4.2.1 Quantitative analysis of resorufin formation

The BOMR substrate was not metabolized by zebrafish embryos of 7, 26, and

50 hpf, as the corrected integrated density of resorufin in the trunk region was

below the LLOQ. However, embryos of 74, 98, and 122 hpf and larvae of 9 and 14

dpf were able to biotransform BOMR (integrated density of resorufin > LLOQ)

(Figure 4a). No statistically significant differences were detected among the

different age groups (p = 0.231). However, since the integrated density of

resorufin formation also depends on the area of interest (section 3.2.4 of the

current Chapter) and since the area of interest varies between the different

developmental stages, the significance of the results is difficult to interpret.

Nevertheless, considering the research hypothesis of the current doctoral project

(Chapter 2), the main focus is on whether or not BOMR is metabolized in intact

zebrafish embryos or larvae, whereas the absolute values of resorufin formation

are of less importance.

Regarding the positive control, zebrafish larvae of 14 dpf were not able to

biotransform 7-ethoxyresorufin (ER) as the corrected integrated density of

resorufin in the trunk region was below the LLOQ. However, the

ethoxyresorufin-o-deethylase (EROD) assay showed resorufin formation in

embryos of 7, 26, 50, 74, 98, and 122 hpf (Figure 4b). Integrated density of

resorufin was significantly higher at 7 and 26 hpf compared to the other

developmental stages (p = 0.050 for all comparisons). Moreover, resorufin

formation at 7 hpf was significantly higher than at 26 hpf (p = 0.050) (Figure 4b).

152

The stage of 9 dpf was excluded from quantitative analysis since resorufin

formation could not be localized due to a technical limitation, i.e. ventral position

of the larvae.

Figure 4. Resorufin formation in the trunk region of intact zebrafish embryos and

larvae at different time points during zebrafish development between 7 h post-

fertilization (hpf) and 14 d post-fertilization (dpf) after incubation with benzyloxy-

methyl-resorufin (BOMR) (a) and 7-ethoxyresorufin (ER) (b). In both graphs,

resorufin integrated density values for 14 dpf were set to 1 (horizontal dotted line)

and the relative values are shown for the other developmental stages. At 7 hpf (b),

integrated density of resorufin was determined in the whole embryo. Each bar

represents the mean of three biological replicates ± standard deviation (S.D.). In

graph (a,b), developmental stages with values below the LLOQ were excluded from

statistical analysis (indicated by *). In graph (a), no statistically significant differences

(p > 0.05) were detected between the developmental stages that showed values above

the LLOQ. The mean corrected integrated density value for 50 hpf was below zero.

In graph (b), significant differences (p ≤ 0.05) between age groups are indicated by

different letters (A, B and C): integrated density of resorufin was significantly higher

at 7 and 26 hpf compared to the other developmental stages (p = 0.050 for all

comparisons). Moreover, resorufin formation at 7 hpf was significantly higher than

at 26 hpf (p = 0.050).

4.2.2 Qualtitative analysis of resorufin formation

Biotransformation of BOMR was localized in the liver and intestine at 74, 98,

122 hpf and at 9 dpf (Figure 5f–m). At 14 dpf, resorufin formation was only

7 h

pf

26 h

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**

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7-Ethoxyresorufin

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C

153

detected in the intestine (Figure 5n,o). Furthermore, at this stage, there is food

present and visible in the digestive tract. At 98, 122 hpf and at 9 dpf, the

metabolite of BOMR was also observed in the pronephric region and,

additionally, in the otic vesicle, which belongs to the head region (Figure 5h,j,l).

A weak fluorescent signal was localized in the otic vesicle at 74 hpf (Figure 5f).

Resorufin formation was not detected at 7, 26 and 50 hpf (Figure 5a–e). Similar to

the in vivo BOMR assay, the positive control substrate, i.e. ER, was metabolized

in the liver and intestine of zebrafish embryos of 74, 98, and 122 hpf (Figure 6f–

k). In contrast to BOMR, biotransformation of ER was also observed at 7, 26 ,and

50 hpf (Figure 6a–e) with the strongest fluorescent signal in the germ ring at 7 hpf

(Figure 6a). Since Figure 6a shows a vegetal pole view, the yolk covers the

blastoderm resulting in a fluorescent signal of the blastoderm that is less intense.

The embryo is entirely stained at 26 and 50 hpf (Figure 6b–e). Non-trunk-related

structures such as the hatching gland and the otic vesicle showed resorufin

formation at 26 hpf (Figure 6b) and at 50, 74, and 98 hpf, respectively (Figure

6d,f,h). The metabolite was not detected at 14 dpf (Figure 6l,m) and 9 dpf was

excluded from the figure since resorufin formation could not be localized due to

ventral position of the larvae.

154

Figure 5. Localization of biotransformation of benzyloxy-methyl-resorufin (BOMR)

in the trunk region of intact zebrafish embryos and larvae at 26 h post-fertilization

(hpf) (b,c), 50 hpf (d,e), 74 hpf (f,g), 98 hpf (h,i), 122 hpf (j,k), 9 d post-fertilization

(dpf) (l,m) and 14 dpf (n,o). At 7 hpf (a), qualitative analysis of resorufin formation

was performed in the whole embryo. Pictures show one embryo/larva out of six used

in the study, i.e. three biological replicates with two embryos/larvae per replicate, for

each developmental stage. Figure 5a shows a vegetal pole view of the embryo. In

figure 5b–o lateral views of the anterior and posterior part of the trunk region are

shown. The organs in which resorufin had been formed are indicated with a two-

letter combination. Since the otic vesicle is part of the head region, resorufin

formation in the respective organ is mentioned separately. S.B.: swim bladder. Scale

bar: 200 µm; anterior left and dorsal top.

156

Figure 6. Localization of biotransformation of 7-ethoxyresorufin (ER) in the trunk

region of intact zebrafish embryos and larvae at 26 h post-fertilization (hpf) (b,c), 50

hpf (d,e), 74 hpf (f,g), 98 hpf (h,i), 122 hpf (j,k) and 14 d post-fertilization (dpf) (l,m).

At 7 hpf (a), qualitative analysis of resorufin formation was performed in the whole

embryo. The stage of 9 dpf was excluded from the figure since resorufin formation

could not be localized due to ventral position of the larvae. Pictures show one

embryo/larva out of six used in the study, i.e. three biological replicates with two

embryos/larvae per replicate, for each developmental stage. Figure (a) shows a

vegetal pole view of the embryo. In figure 6 (b–m) lateral views of the anterior and

posterior part of the trunk region are shown. The organs in which resorufin had been

formed are indicated with a two-letter combination. Since the hatching gland and

otic vesicle do not belong to the trunk region, resorufin formation in the respective

organs is mentioned separately. S.B.: swim bladder. Scale bar: 200 µm; (b,c): anterior

top and dorsal right; (d–m): anterior left and dorsal top.

158

4.2.3 Preliminary inhibition study in zebrafish embryos of 98 and 122 hpf

Since the preliminary inhibition study consists of only one replicate, no

quantitative analysis of resorufin formation was performed. Regarding the

qualitative evaluation, no differences in fluorescence intensity between the 1–

aminobenzotriazole exposed embryos (1 h and 96 h of exposure) and the non–

exposed embryos were observed for both 98 hpf and 122 hpf. This finding

contradicts the in vitro inhibition study since 1–aminobenzotriazole was able to

inhibit the biotransformation of BOMR in adult zebrafish liver microsomes (IC50

= 116.3 µM) (section 4.2.2 of Chapter 3). Moreover, for both 98 and 122 hpf, the

following malformations were observed after exposure to 1–aminobenzotriazole

for 96 h (Figure 7): abnormal intestine, non–inflated swim bladder and abnormal

yolk extension. Malformations were negligible in embryos of 98 and 122 hpf

exposed to 1–aminobenzotriazole for 1 h and in embryos of 98 hpf exposed to

0.46% DMSO. However, zebrafish embryos of 122 hpf that were exposed to 0.46%

DMSO for 96 h showed the same malformations as described for the embryos

exposed to 1–aminobenzotriazole. Moreover, embryos of 122 hpf in Tris solution

(blank) also showed malformations such as abnormal yolk extension and

abnormal intestine. The malformations in the blank and the fact that no inhibition

of resorufin formation has been observed in vivo after exposure to 1–

aminobenzotriazole make it difficult to interpret the results of the current assay.

Furthermore, since only one replicate has been performed, no definite

conclusions can be drawn from this in vivo inhibition study.

159

Figure 7: Malformations in a zebrafish embryo of 122 hpf after exposure to 1–

aminobenzotriazole (ABT) for 96 h. The pictures on the left show the anterior and

posterior part of the trunk of a non–exposed zebrafish embryo of 122 hpf. The pictures on

the right show the anterior and posterior part of the trunk of a zebrafish embryo of 122

hpf after exposure to ABT for 96 h. Malformations are indicated by the following

numbers: 1: abnormal intestine; 2: non–inflated swim bladder and 3: abnormal yolk

extension. Scale bar: 200 µm; anterior left and dorsal top.


glycoprotein

The mRNA expression analysis was performed by means of a loess

regression method in order to identify key inflection points, i.e. local maxima and

minima, of transcriptional expression during zebrafish development. This

method allowed us to identify statistically significant highs and lows in the

expression profiles of phase I (Figure 8) and phase II enzymes and P–glycoprotein

(Figure 9). Data in Figure 7 and 8 are reported as log2 relative quantities—relative

Anterior part of trunk

Posterior part of trunk

2

1

3

Control 1000 µM ABT

160

to the time point with the lowest expression—, which means that a log2 relative

quantity of 2 for a particular time point corresponds to four times the expression

of the time point with the lowest expression. Consequently, data should not be

used for direct comparison of absolute expression levels among transcripts. Most

transcripts only had low expression levels at the earliest time point, i.e. 1.5 hpf

(Figures 8c–f; 9a,b). The CYP1B1 transcript was not detected at 1.5 hpf (Figure

8b), whereas relatively high expression levels could be observed for CYP1A,

CYP3C1 and abcb4 at this stage (Figures 8a,g; 9c). The high initial expression

levels of CYP1A and CYP3C1 were followed by a decline of mRNA expression

between 1.5 and 6 hpf (Note that the regression does not capture this early

decrease for CYP1A). Subsequently, transcript levels of CYP1A and CYP3C1

increased from 14 hpf until 5–6 dpf after which both transcripts started to level

out for the remaining developmental time points (Figure 8a,g). Within this period

of increasing mRNA levels, CYP1A transcript levels remained stable between 14

hpf and 84 hpf (Figure 8a). The Abcb4 transcript showed a similar expression

pattern as for CYP1A but without the short period of stable mRNA expression

during early embryonic development (Figure 9c). Regarding CYP1C1, CYP1C2,

CYP3A65, SULT1ST1 and UGT1A1, transcript levels showed a steep increase after

the first time point, reached a maximum between 120 hpf and 10 dpf and

remained stable for the remaining developmental time points (Figures 8c,d,f;

9a,b). A distinct pattern was observed for CYP2K6 and CYP1B1 since transcript

levels reached a peak at 14 hpf and 36 hpf, respectively, followed by a decrease

in expression levels until 48 hpf (Figure 8b,e). From 48 hpf onwards, CYP1B1

transcripts started to level out with a slight fluctuation (Figure 8b), whereas

CYP2K6 mRNA levels started to increase until approximately 12 dpf followed by

a decline until the end of the larval period. CYP2K6 transcript levels tended to

increase again by the beginning of the juvenile period (Figure 8e). These mRNA

expression measurements have been performed in the same samples as those

used in the study of Vergauwen et al. (2018) [54], where the ontogeny of thyroid

related genes was studied. Hence, the current results can be directly related to the

results of the previous study. The results of these studies can be directly

161

compared via interactive graphs available online

(http://zebrafishlab.be/ontogeny-explorer).

Log2

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0 5

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http://zebrafishlab.be/ontogeny-explorer

162

CYP1C1

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163

(e) CYP2K6

Log2

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164

Figure 8. Relative quantities of cytochrome P450 (CYP) 1, 2 and 3 families from whole

zebrafish bodies: (a) CYP1A, (b) CYP1B1, (c) CYP1C1, (d) CYP1C2, (e) CYP2K6, (f)

CYP3A65 and (g) CYP3C1. The graphs show log2 relative quantities which were

normalized for reference gene expression and expressed relative to the time point

with the lowest expression. Data points represent mean ± S.D. of four replicate pools

at each time point (days post-fertilization (dpf)). The red line indicates the loess fit of

the gene target and the surrounding dashed blue line indicates the 95% confidence

interval around the loess fit. The green and purple highlighted regions represent the

95% and 99% confidence intervals, respectively, of each critical point (minimum or

maximum) of mRNA expression. The color bar between 0 and 5 dpf, i.e. between 0

and 120 h post-fertilization, indicates the period of zebrafish organogenesis. The

graphs on the left are a more detailed representation of the organogenesis period for

each gene.

CYP3C1

Log2

rel

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(g)

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166

Figure 9. Relative quantities of two phase II enzymes from whole zebrafish bodies,

i.e. (a) sulfotransferase 1st1 (SULT1ST1) and (b) uridine diphosphate

glucuronosyltransferase 1A1 (UGT1A1), and one P–glycoprotein, i.e. (c) ATP-

binding cassette b4 (abcb4) transporter. The graphs show log2 relative quantities

which were normalized for reference gene expression and expressed relative to the

time point with the lowest expression. Data points represent mean ± S.D. of four

replicate pools at each time point (days post-fertilization (dpf)). The red line indicates

the loess fit of the gene target and the surrounding dashed blue line indicates the

95% confidence interval around the loess fit. The green and purple highlighted

regions represent the 95% and 99% confidence intervals, respectively, of each critical

point (minimum or maximum) of mRNA expression. The color bar between 0 and 5

dpf, i.e. between 0 and 120 h post-fertilization, indicates the period of zebrafish

organogenesis. The graphs on the left are a more detailed representation of the

organogenesis period for each gene.

Log2

rel

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Time (dpf)

Abcb4(c)

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5 Discussion

5.1 Ontogeny of in vitro and in vivo cytochrome P450 activity in

zebrafish embryos, larvae and adults

5.1.1 In vitro versus in vivo

The results of the CYP activity assays support the hypothesis that the

intrinsic CYP-mediated biotransformation capacity in zebrafish embryos is

immature during early development although differences in CYP isoforms do

occur. More specifically, biotransformation of the non-specific CYP substrate

BOMR was above the LLOQ in intact embryos from 74 hpf onwards, i.e. towards

the end of zebrafish organogenesis (Figure 1). Furthermore, these findings are in

agreement with the present in vitro data that showed no BOMR

biotransformation before 72 hpf in microsomes prepared from whole zebrafish

embryo homogenates. This onset of CYP activity at 72 hpf coincides with

vascularization of the liver, development of the intestinal epithelium and opening

of the mouth (Figure 1). By 96 hpf, the liver has reached its adult configuration

and the intestine has developed into an open-ended tube, which is reflected in

CYP activity in the respective organs of intact embryos at 98 hpf [6,15-17,86]. This

concurrence is not surprising as the liver and, to a lesser extent, the intestine are

two major organs involved in mammalian CYP-mediated metabolism of

xenobiotics [20,87]. However, there was a discordance between the in vitro and in

vivo experiments with BOMR since in vitro CYP activity was low at 72 hpf, 96

hpf and 14 dpf, whereas in vivo CYP activity was clearly observed at 74 hpf, 98

hpf and 14 dpf and even at 122 hpf and 9 dpf. For the latter two stages, no CYP

activity could be detected in vitro. The underestimation of CYP activity in the in

vitro study might be due to a dilution of the CYP enzymes, and consequently

their activity, by the presence of other microsomal proteins which are derived

from the different tissues of the whole embryos/larvae. Moreover, the reaction

velocities for BOMR biotransformation in microsomes prepared from whole

adult zebrafish (ZM) and microsomes prepared from adult zebrafish livers (ZLM)

168

were obviously different from each other, i.e. 1.34 pmol/min/mg MP versus 9.65

pmol/min/mg MP for the respective microsomes, which might confirm our

hypothesis regarding the dilution of CYP enzymes. A second hypothesis which

might explain the underestimation of in vitro CYP activity is non–specific binding

of the BOMR substrate to (liver) microsomes. Although not much is known about

the lipophilicity of BOMR, we assume that it is a lipophilic substrate since 7–

benzyloxyresorufin, which structure is very similar to BOMR, has a logPoct of ±

2.206 [88]. Due to its lipophilicity, BOMR may bind non-specifically to the lipid-

protein (non–metabolizing) milieu of the microsomal membrane [89], which

might result in lower in vitro CYP activity values compared with the in vivo CYP

activity values. Since microsomes represent a preparation of intracellular

membranes derived primarily from the endoplasmic reticulum, microsomes

prepared from whole adults (ZM) contain a broader range of microsomal proteins

in comparison with ZLM which mainly contain proteins involved in

lipid/lipoprotein biosynthesis and drug metabolism, e.g. CYPs, flavin

monooxygenases (FMOs) (phase I) and UDP glycosyltransferases (UGTs) (phase

II) [90-92]. Hence, the microsomes prepared from whole adults might be more

prone to non-specific protein binding of the BOMR substrate which might explain

their lower reaction velocities compared with ZLM. The same may be true for

microsomes prepared from whole zebrafish embryos (ZEM) and larvae (ZLaM).

However, the reaction velocities for ZEM and ZLaM cannot be compared with

the corresponding liver microsomes since we were not able to extract the livers

from the embryos and larvae. Regarding the second hypothesis, one might correct

for the non–specific binding of the substrate to the microsomes by determining

the fraction of unbound substrate (fu(mic)) in the different microsomal preparations,

e.g. by equilibrium dialysis [89].

For a comprehensive discussion of the results of the in vitro study with

BOMR in microsomes prepared from whole zebrafish embryos between 5 hpf and

120 hpf, we refer to Verbueken et al. (2017) [25].

169

5.1.2 Benzyloxy-methyl-resorufin versus 7–ethoxyresorufin

BOMR and ER biotransformation showed similar activity in the digestive

system, i.e. between 74 and 122 hpf. The detection of resorufin formation in the

digestive system at 74 hpf coincides with opening of the mouth around 72 hpf

(Figure 1) [6]. At this stage, oral ingestion of xenobiotics complements uptake of

compounds by the skin. Although Kais et al. (2017) [30] suggested that the

detection of EROD activity in the intestine of zebrafish embryos is due to

secretion of the metabolite from the liver via the bile, it should be noted that CYP

families 1, 2 and 3 were shown to be expressed in the adult mammalian and

zebrafish intestine [20,56,61,62,78,87]. Hence, the detection of resorufin formation

in the intestine of zebrafish embryos from 74 hpf onwards may be attributed to

biotransformation of the orally ingested BOMR or ER by intestinal CYP enzymes.

Because of their role in mammalian drug metabolism, intestinal CYP enzymes are

supposed to be involved in detoxification. In addition to the liver and intestine,

resorufin formation was observed in the cranial pole of the early kidney, i.e.

pronephros, of intact embryos and larvae of 98 hpf, 122 hpf and 9 dpf for BOMR

and in embryos of 98 hpf for ER. The detection of CYP activity in the pronephros,

which is involved in drug metabolism and elimination, follows the completion of

pronephric nephron and filtration barrier development by 84 hpf [66]. Hence, the

observation of CYP activity in the pronephros may be related to detoxification.

In contrast to BOMR, the present study showed that biotransformation of ER

already occurred in the germ ring (blastoderm) at 7 hpf and in the whole embryo

at 26 hpf and 50 hpf. This difference between both substrates may be explained

by the fact that BOMR and ER have a different affinity for CYP1, 2 and 3

isoenzymes. Indeed, ER is known to be specifically metabolized by CYP1

isoenzymes, whereas BOMR was shown to be a non-specific CYP substrate

according to a previous study with recombinant human CYP enzymes [25].

Although the current study does not provide a clear explanation for the presence

of CYP1 activity in the early stages of zebrafish embryonic development,

vertebrate CYP1 enzymes are known to play a role in embryonic development

since CYP1B1 is involved in the synthesis of retinoic acid (RA), an endogenous

170

signalling molecule which is essential in embryogenesis [93-95]. Furthermore,

exogenous compounds are mainly taken up by the skin until opening of the

zebrafish mouth and the onset of gill filament development, i.e. both around 72

hpf, [6]. Although not much is known about cutaneous CYPs in fish, the enzymes

can be found in adult mammalian skin (reviewed by [20]) and RA was shown to

be involved in mammalian embryonic skin development [96]. As such, CYP1

activity that was observed in the whole embryo at 26 and 50 hpf coincides with

the period in which the substrate is taken up by the embryonic skin. In contrast

to BOMR, EROD activity was not detected in larvae of 14 dpf (larvae of 9 dpf

were not included in the assay because of difficulties with positioning).

Regarding larvae of 9 and 14 dpf exposed to BOMR, the onset of exogenous

feeding around 96–120 hpf and the increased mortality between 8 and 15 dpf did

not affect the biotransformation of the substrate in the trunk region. However,

the fluorescent signal in the digestive system of BOMR-exposed larvae of 14 dpf

appeared to be less intense due to the presence of food in the digestive tract.

The trunk region was our main focus for the assessment of CYP activity since

the major CYP-containing organs are located in this area. However, BOMR and

ER were also metabolized in the otic vesicle—the zebrafish counterpart of the

mammalian inner ear—at 74, 98, 122 hpf and 9 dpf for BOMR and at 74 and 98

hpf for ER. These stages do not coincide with the development of the respective

organ as the otic vesicle and its corresponding otoliths have already been

developed around 19 hpf and 22 hpf, respectively (Figure 1) [6]. However, in

mammals, RA, and thus indirectly CYP enzymes, are suggested to be essential in

embryonic development as well as in postnatal maintenance of the mammalian

inner ear [94].

5.1.3 Literature versus current study

According to literature, studies regarding the localization of CYP activity in

intact zebrafish embryos mainly involve EROD assays [30,34], whereas to our

knowledge, in vivo studies using a non-specific CYP substrate have not yet been

described. Kais et al. (2017) [30] assessed EROD activities in intact zebrafish

171

embryos of between 24 and 120 hpf, which are in line with the results of the

present study. At 24 and 48 hpf, the authors reported biotransformation of ER in

the whole embryo, the strongest fluorescent signal being located in the head

region, i.e. brain, eyes and otic vesicle. However at 48 hpf, the fluorescent signal

decreased compared to the previous stage, which is similar to our results for

embryos of 50 hpf. From 72 hpf onwards, the authors reported EROD activity in

the digestive system, which slightly increased until 96 hpf and remained stable at

120 hpf [30]. A study from Otte et al. (2010) [34] included zebrafish embryos of 8

hpf that showed biotransformation of ER in the blastoderm, similar to the

youngest embryos in the current study. The authors also localized EROD activity

in zebrafish embryos of 32, 56, 80, 104, and 128 hpf, in similar organs as in the

current study for the corresponding stages, i.e. 26, 50, 74, 98, and 122 hpf

respectively. However, Otte and colleagues (2010) [34] were able to show a more

detailed localization of CYP1 activity e.g. in myotomes, pronephric duct, vessels

and organ primordia.. These anatomical structures were visualized by a confocal

laser scanning microscope (CLSM) which makes high resolution images possible

due to the process of optical sectioning [34]. Since in the present study and in the

one from Kais et al. (2017) [30], an epifluorescence microscope had been used,

organs like the pronephric duct and vessels could not be distinguished from the

surrounding structures.

Because of the similarities with the results described in literature, we may

conclude that ER is suited as a positive control in CYP activity assays with intact

zebrafish embryos/larvae.

5.2 Ontogeny of cytochrome P450 mRNA expression in zebrafish

embryos and larvae

As CYP transcript levels have been investigated in zebrafish embryos by

other groups before, we will focus the discussion mainly on the later stages.

172

5.2.1 Cytochrome P450 mRNA expression during zebrafish organogenesis

For all CYP enzymes that were investigated, mRNA expression levels

increased during the organogenesis period. Moreover, the increase in CYP1

transcript levels before 72 hpf was concomitant with the results of the EROD

activity assay.

In the current study, the relatively high expression levels for CYP1A and

CYP3C1 at 1.5 hpf suggest maternal transfer of the respective mRNA transcripts

since the zebrafish zygotic genome becomes gradually activated in the blastula

period throughout a window of approximately two hours, starting at cell cycle 10

(around 2.75 hpf according to Kimmel et al. (1995) [6]) (reviewed by [97]). The

maternal mRNA transcripts are produced during oogenesis and are present in

the egg at fertilization. They are considered to be essential for the development

of the earliest embryonic stages (reviewed by [97]). Moreover, a recent study

compared fertilized eggs of 1.5 hpf with unfertilized eggs for zebrafish thyroid-

related transcript levels and was not able to detect differences between both

conditions, which confirms that the detection of transcript levels at 1.5 hpf is due

to maternal transfer [54] .

No maternal transfer was detected for CYP1B1, CYP1C1, CYP1C2, CYP2K6,

and CYP3A65. However, mRNA levels increased immediately after activation of

the embryonic genome (around 6 hpf). Transcript levels of CYP1C1, CYP1C2, and

CYP3A65 showed a steep increase throughout the organogenesis, whereas a

distinct pattern was observed for CYP1B1 and CYP2K6 mRNA levels. Indeed,

CYP1B1 transcripts peaked at 36 hpf, followed by a decline until 48 hpf after

which mRNA levels started to level out for the remaining developmental time

points. This peak, which was also detected by Goldstone et al. (2010) [22] at the

same time point, coincides with the development of the eye cup and retina

(Figure 1) [6,67]. Moreover, Yin and colleagues (2008) [98] already reported basal

CYP1B1 mRNA expression in ocular cells of zebrafish at 24 hpf after which

transcription levels peaked between 30 and 48 hpf. In addition to the eye, CYP1B1

mRNA levels were detected in the zebrafish brain at 36 and 48 hpf by whole-

mount in situ hybridization [98]. Also in human fetuses, CYP1B1 mRNA was

173

abundantly expressed in the brain [99]. Regarding CYP2K6, transcript levels in

the current study peaked at 14 hpf followed by a decrease until 48 hpf. In contrast

to CYP1B1, CYP2K6 mRNA levels started to increase again after hatching until 10

dpf. In a study of Wang-Buhler et al. (2005) [63], CYP2K6 transcripts were

expressed in liver and ovary of adult zebrafish. However, the presence of CYP2K6

transcripts in adult zebrafish liver and ovary cannot explain the high transcript

level at 14 hpf in the current study since these organs develop later. Yet, the early

CYP2K6 mRNA peak coincides with the development of the brain neuromeres

(Figure 1) around 16 hpf and with the onset of heart development around 16–19

hpf [6,100]. With regards to CYP1C1 and CYP1C2, Jönsson et al. (2007) [78]

described an increase in basal mRNA levels from 8 to 96 hpf and from 8 to 72 hpf

for the respective enzymes, which is similar to the present study. However, the

same authors showed fluctuating CYP1C1 and CYP1C2 mRNA levels between 96

hpf and 7 dpf, whereas transcript levels remained stable in the current study [78].

Regarding CYP3A65, the present study and the one from Tseng et al. (2005) [62]

both reported increasing mRNA levels throughout the organogenesis period.

Moreover, CYP3A65 transcripts were detected in the liver at 72 hpf by whole-

mount in situ hybridization and subsequently in liver and intestine at 84, 96, and

120 hpf [62]. In contrast to the current study, maternal CYP3A65 transcripts were

observed at 3 hpf by Goldstone et al. (2010) [22] and a study of Glisic et al. (2016)

[101] showed low CYP3A65 mRNA expression levels until 96 hpf followed by a

peak at 120 hpf.

We can conclude that, with regards to the zebrafish organogenesis period,

the results of CYP mRNA expression analysis are in accordance with the majority

of studies described in literature.

5.2.2 Cytochrome P450 mRNA expression during zebrafish larval

development

Except for CYP1B1, all CYP transcripts that were investigated reached

maximum expression levels during embryo-larval transition, i.e. between 4 and

7 dpf, which comprises the period between the onset of exogenous feeding and

174

complete yolk absorption (Figure 1). Exogenous feeding implies increased

exposure to environmental compounds, which may cause a slight induction of

CYP mRNA expression due to PXR or AhR activation [41,42]. However, CYP1C1

and CYP1C2 transcripts reached high expression levels already around 72 hpf,

which coincides with the opening of the mouth. This implies an increased

exposure of the zebrafish embryo to exogenous compounds that are present in

the fish medium, which might result in an induction of the respective CYP

enzymes.

After reaching maximum mRNA levels during the embryo-larval transition,

CYP1A, CYP1C1, CYP1C2, and CYP3A65 transcript levels remained stable

throughout the larval period, whereas mRNA levels of CYP1B1, CYP2K6, and

CYP3C1 fluctuated to some extent. The decline of transcript levels around 10 dpf

that was observed for CYP1B1, CYP2K6, and CYP3C1 coincides with the period

of increased mortality due to starvation (Figure 1). However, the correlation

between both observations remains unclear. A more plausible explanation for the

fluctuating CYP transcript levels during the larval period might be changes in the

environment such as feeding regimen and stocking density. In a study of Wang-

Buhler et al. (2005) [63], CYP2K6 transcript levels were detected in liver and ovary

of adult zebrafish. Hence, the decline in CYP2K6 mRNA levels throughout the

larval period might be explained by the decrease in relative liver size in

proportion to the increasing body mass. The subsequent increase in CYP2K6

transcript levels at the larval-juvenile transition period may be attributed to the

onset of gonad development around 30 dpf [102]. Regarding CYP1A, CYP1C1,

CYP1C2, and CYP3A65, mRNA expression remained constant during the larval

period despite the growth burst between 9 and 51 dpf [103] and the corresponding

decline of relative organ size and organ-specific CYP mRNA expression. This

might be due to a shift or increase of organ-specific CYP mRNA expression,

which results in constant transcript levels throughout the whole larval body. In

the study of Jönsson et al. (2007) [78], transcript levels of the CYP1 family were

assessed until 57 dpf, which is still within the juvenile period of between 30 and

90 dpf. In contrast to the present study, Jönsson et al. (2007) [78] showed

175

fluctuating CYP1A and CYP1C1 mRNA levels throughout the larval period.

Regarding larval CYP1B1 and CYP1C2 mRNA expression, the results are in

accordance with the present study. To our knowledge, no other CYP mRNA

expression studies covering the whole zebrafish larval period have been

performed.

5.3. Ontogeny of mRNA expression of two Phase II enzymes and a P–

glycoprotein in zebrafish embryos and larvae

The biotransformation of xenobiotics and endogenous compounds implies

phase II reactions in which the parent compound or phase I metabolites are

conjugated with a hydrophilic moiety to enhance their water solubility and

elimination from the body. In the present study, the embryonic and larval

development of the constitutive mRNA expression of two major phase II

enzymes, i.e. sulfotransferase 1st1 (SULT1ST1) and uridine diphosphate

glucuronosyltransferase 1A1 (UGT1A1) was assessed in zebrafish. In mammals,

UGT enzymes are located predominantly in the endoplasmic reticulum of liver,

intestine, kidney, lungs, skin, brain and spleen, whereas SULT enzymes are

primarily located in the cytosol of liver, intestine, kidney, lung, platelets and

brain. Conjugation reactions comprise glucuronidation and sulfonation by UGT

and SULT enzymes, respectively (reviewed by [104]). Zebrafish UGT1A was first

identified by Huang and Wu (2010) [50] and is expressed in liver and intestine

and, to a lesser extent, in brain and testis of adult zebrafish [48]. In the current

study, UGT1A1 transcripts reached maximum expression levels during embryo-

larval transition (Figure 1) after which mRNA levels levelled off throughout the

larval period. Since embryo-larval transition coincides with the onset of

exogenous feeding and since UGT1A is supposed to be regulated through the

AhR pathway [48], we assume that the increased exposure to environmental

compounds induced UGT1A1 mRNA expression due to AhR activation. Christen

and colleagues (2014) [48] assessed UGT1A mRNA expression between 24 and 120

hpf, and showed an increase in transcript levels between 48 and 120 hpf, which

is in line with the present study. However, in contrast to the current study,

176

UGT1A mRNA levels at 24 hpf were higher than at 48 hpf [48]. Zebrafish

SULT1ST1, which was first identified by Liu et al. [51], showed maximum

transcript levels already around 72 hpf, which coincides with the first observation

of thyroid hormone synthesis (Figure 1) [65]. Moreover, zebrafish SULT1ST1

enzymes are involved in the sulfonation of endogenous thyroid hormones [85],

which might explain the maximum SULT1ST1 mRNA levels around 72 hpf. After

reaching a maximum in the current study, SULT1ST1 transcript levels remained

stable throughout the larval period. In contrast to the present study, Liu et al.

(2005) [51] showed low levels of SULT1ST1 mRNA expression in unfertilized eggs

and in embryos immediately after fertilization, suggesting maternal transfer of

the transcript.

Besides phase I and phase II enzymes, the bioavailability of xenobiotics also

depends on the presence of ATP-binding cassette (ABC) transporters that protect

cells against a wide range of xenobiotics. Zebrafish abcb4, which was first

described by Fischer et al. (2013) [53], possesses similar functional properties as

the mammalian ABCB1 transporter. The present study showed maternal transfer

of abcb4 transcripts, which suggests that Abcb4 is essential for the protection of

the early embryo against environmental compounds. Subsequently, abcb4

transcript levels declined at 6 and 14 hpf followed by an increase around 24 hpf.

Abcb4 transcripts rose until 120 hpf followed by stable mRNA levels throughout

the larval period. The temporal expression profile of abcb4 is in line with the study

of Fischer et al. (2013) [53] in which transcript expression was assessed until 48

hpf.

In the literature, not much is known about the activity of phase II enzymes

and P-glycoproteins during zebrafish development, nor about their possible role

in embryogenesis.

177

6 Conclusions

The extensive use of zebrafish embryos as an alternative animal model in

developmental toxicity studies increases the demand for a detailed investigation

of their intrinsic biotransformation capacity since the embryos cannot rely on

maternal metabolism of the xenobiotics. The present study contributes to a better

understanding of the ontogeny of metabolism and transport of xenobiotics in the

zebrafish, and suggests that, in general, the disposition of xenobiotics in zebrafish

embryos is immature during a major part of the organogenesis period, i.e. before

72 hpf. This may lead to false negative results in the case of proteratogens,

whereas the teratogenic potential might increase in the case of teratogens since

immature biotransformation might result in a higher internal concentration of the

teratogenic parent compound. Full capacity appears to be reached by the end of

organogenesis (i.e. 120 hpf), although CYP1—except CYP1A—and SULT1ST1

showed to be already mature in early embryonic development. Furthermore, the

present study showed that in vitro CYP activity assays with microsomes

prepared from whole zebrafish organisms do not always reflect the in vivo

activity and can underestimate the biotransformation capacity of the organisms.

In literature, CYP activity and expression studies mainly focus on zebrafish

embryonic development, whereas in the present study the experimental time

window has been extended to the beginning of the juvenile period. The study

showed that CYP activity and expression mainly remained stable during the

larval period. However, regarding the phase II enzymes and P-glycoprotein,

activity studies need to be performed to draw conclusions on their role in drug

metabolism during zebrafish development.

178

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Chapter 5: General discussion

189

In view of cost and time effectiveness, and within the framework of the 3Rs,

the zebrafish embryo has been proposed as an alternative animal model for

developmental toxicity screening of new drugs and environmental pollutants.

However, the externally developing zebrafish embryo cannot rely on maternal

metabolism and thus depends on its intrinsic biotransformation capacity for the

detoxification and/or bioactivation of a compound. Since knowledge of drug

disposition during zebrafish organogenesis is pivotal in order to correctly

interpret the outcome of teratogenicity assays, this doctoral thesis investigated

the developmental expression and activity of some major enzymes that are

involved in the disposition of xenobiotics. The current chapter will discuss the

main findings of the doctoral project and will put them into a broader perspective

by addressing the following questions:

1. What are the implications of the current findings for developmental

toxicity studies using zebrafish?

The first section of the ‘General discussion’ will summarize the main

findings of the doctoral project with special emphasis on the cytochrome

P450 (CYP) mRNA expression and activity data during zebrafish

embryogenesis since these findings might have implications for the

zebrafish embryo developmental toxicity assay (ZEDTA) when using

proteratogenic compounds.

2. What is known about biotransformation in other alternative test

systems used for developmental toxicity studies and what can be done

to prevent false negative results?

The findings regarding the biotransformation capacity during zebrafish

embryonic development will be compared with the xenobiotic–

metabolizing capacity of other alternative models such as the Whole

Embryo Culture (WEC) and Frog Embryo Teratogenesis Assay–Xenopus

(FETAX). Moreover, this section will provide an overview of possible

methods (i.e. exogenous metabolic activating systems) that can be applied

with the alternative test systems to improve their sensitivity and

predictivity.

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3. How does the biotransformation in zebrafish embryos/larvae relate to

adult zebrafish? And how does xenobiotic metabolism in zebrafish

relate to mammals?

In this section, we will make an intra– and interspecies comparison with

regards to the metabolites that result from the biotransformation of some

well–known xenobiotics. Moreover, a general comparison will be made

between zebrafish and humans regarding the ontogeny of

biotransformation enzymes and transport proteins.

4. Does the zebrafish embryo have the potential to be used in regulatory

developmental toxicity testing?

Based upon the research findings of this doctoral project, we will make

recommendations with regards to the use of the zebrafish embryo in

regulatory developmental toxicity testing.

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1 Implications of the current findings for developmental

toxicity studies using zebrafish embryos

The zebrafish embryo has emerged as an alternative animal model for

developmental toxicity screening during the drug development process as well

as in ecotoxicology. The exposure window of the ZEDTA and fish embryo acute

toxicity test (FET) coincides with the teratogen–sensitive period in zebrafish

development, i.e. the organogenesis [1-4]. Indeed, vital organs such as the heart,

central nervous system, intestine, pancreas, liver and pronephros have already

been developed by 96–120 hpf (Section 3.4 of Chapter 1: Introduction), whereas

in rats and rabbits, the organogenesis takes around 11–13 days (TG 414 OECD,

2001) [5]. However, the rapid developing zebrafish embryo cannot rely on

maternal metabolism unlike mammalian embryos in vivo (Figure 1). Hence, the

zebrafish embryo gets directly exposed to the parent compound and depends on

its own drug–metabolizing capacity for the detoxification or bioactivation of

xenobiotics. In view of its use as an alternative animal model in developmental

toxicity studies, the drug–metabolizing capacity of zebrafish embryos formed the

main subject of the current doctoral thesis.

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Figure 1: A schematic representation of drug disposition in pregnant mammals after

maternal drug administration. The black arrows represent the parent compound and the

white arrows represent the metabolites. The size of the arrows approximates relative

importance, although this is drug–dependent and varies with the stage of gestation. The

fetus can be exposed to either the parent compound or the metabolite after

biotransformation of the compound primarily by the maternal liver. The mammalian

placenta is mainly responsible for the passive and active (i.e. solute carrier and ATP–

binding cassette) transport of drugs and their metabolites between the maternal and

fetal circulation. Biotransformation enzymes have also been identified in the mammalian

placenta, albeit at low abundance. Hence, the contribution of placental biotransformation

enzymes to overall gestational pharmacokinetics is supposed to be minor. The figure is

reproduced from Syme et al. (2004) [6].

Based on the results of the CYP activity studies in this project, CYP–mediated

biotransformation of xenobiotics appears to be immature during a major part of

the organogenesis period, i.e. before 72 hpf. This finding is in accordance with

other literature reports on CYP activity [7-9]. As already mentioned in Chapter 4,

the detection of CYP activity from 72 hpf onwards is not surprising since it

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coincides with the development of zebrafish liver and intestine, i.e. two major

drug–metabolizing organs [10,11]. With regards to mRNA expression, most drug–

metabolizing enzymes, reached maximum expression levels by the end of

zebrafish organogenesis (Chapter 4). However, there is a discordant relationship

between the increasing mRNA expression levels of the drug disposition genes

before 72 hpf and the lack of CYP activity before this stage. The asynchronous

detection of CYP expression and CYP activity might be due to posttranscriptional

silencing, which has already been described for CYP1A in zebrafish by Mattingly

and Toscano (2001) [12]. The authors reported 2,3,7,8–tetrachlorodibenzo–p–

dioxin (TCDD)–induced mRNA expression of CYP1A at 15 hpf, whereas TCDD–

induced CYP1A activity was not detectable until 72 hpf. Other possible

explanations are posttranslational modification e.g. by phosphorylation of the

CYP enzyme, or alternative splicing, which is a complex deviation of constitutive

splicing of pre–mRNA into mature mRNA [13,14]. Both mechanisms might cause

changes in protein function and accordingly in CYP activity. Discordant

relationships between CYP expression and CYP activity have also been described

throughout human development [15].

Hence, the biotransformation capacity of zebrafish embryos is immature

during a major part of the exposure window of the FET and ZEDTA. As about

10% of the drugs approved worldwide can be classified as prodrugs, i.e. drugs

that require biotransformation to exert their effect [16], this can have a profound

impact on the predictivity of the ZEDTA for human safety assessment of drugs

in development. As such, when using proteratogens in the mammalian in vivo

developmental toxicity studies (Table 1), the embryo can be exposed to the

teratogenic metabolite due to maternal bioactivation of the compound via CYP–

mediated oxidation [17]. However, the externally developing zebrafish embryos

do not seem to have the capacity to bioactivate proteratogenic compounds during

a major part of the exposure window, which may lead to false negative results in

developmental toxicity studies [18]. On the other hand, but less problematic, the

lack of drug–metabolizing capacity might lead to false positive results in case of

teratogenic parent compounds that have non–teratogenic metabolites, as has

been shown in zebrafish embryos exposed to albendazole [19].

194

False positive and false negative results are a general problem in the

extrapolation of in vitro effects to in vivo prediction, which may arise from

differences in e.g. pharmacokinetics [20]. In this respect, the findings of this

project on the biotransformation capacity of zebrafish embryos also contribute to

the concept of adverse outcome pathways (AOPs) in which the zebrafish embryo

model may be used to predict a toxicological outcome after exposure to a

chemical [20,21]. Indeed, AOPs link adverse (toxicological) effects in individuals

or populations to a molecular initiating event (MIE) via a defined series of key

events (KEs) that are measurable through in vitro or in vivo assays (section 2.3 of

Chapter 1). The practical application of the AOPs in chemical risk assessment

requires incorporation of knowledge on ADME properties of the respective

chemical as well as on the ADME properties of the in vitro/in vivo models such

as the zebrafish embryo model [21].

Since we aimed to obtain an overall view of CYP–mediated metabolism

during zebrafish organogenesis, we used a fluorogenic non–specific CYP

substrate, i.e. benzyloxy–methyl–resorufin (BOMR). Although the classic method

of quantifying CYP enzyme activities is based on high–performance liquid

chromatography (HPLC) using conventional CYP probe substrates,

fluorescence–based assays are less time– and reagent–consuming and are suitable

for the localization of CYP activity in vivo [22]. Moreover, as BOMR showed to be

a non–specific CYP substrate (Chapter 3), it reflects the CYP–mediated

metabolism of a large number of xenobiotics since the latter often have affinity

for more than one CYP isoform [23]. In that way, the doctoral project acts as a

bridge between a) a former in vitro CYP activity study performed in our lab

where zebrafish embryonic microsomes were exposed to conventional human

CYP probe substrates, i.e. dextromethorphan, testosterone, diclofenac and

midazolam (Table 3a) [24,25] and b) an ongoing project in our lab that investigates

which mammalian proteratogens (Table 1) are metabolized by the zebrafish and

identifies which recombinant zebrafish CYP isoenzymes are involved in the

biotransformation of those proteratogens. This information will be used to

develop knockout zebrafish embryos for one or multiple CYP isoforms involved

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in the biotransformation of a particular proteratogen and assess the

morphological outcome in these knockout embryos after exposure to the

proteratogen.

Table 1: An overview of some well–known proteratogens in mammals. Adapted from

Weigt et al. (2011) [26].

Proteratogen Metabolizing

enzyme

Reactive metabolite Application

2-Acetylaminofluorene CYPs, SULTs N–hydroxy–2–

acetylaminofluorene

Aromatic

amine,

environmental

contaminant

Benzo[a]pyrene CYPs,

epoxide

hydrolases

Benzo[a]pyrene 7,8

diol–9,10 epoxide

PAH,

environmental

contaminant

Aflatoxin B1 CYPs,

epoxide

hydrolases

Aflatoxin B1–8,9–

epoxide

Mycotoxin,

food

contaminant

Carbamazepine CYPs Carbamazepine–l0,ll–

epoxide

Antiepileptic

drug

Phenytoin CYPs,

epoxide

hydrolases

5–(p–hydroxyphenyl)–

5–diphenyl-hydantoin

Antiepileptic

drug

Trimethadione CYPs Dimethadione Antiepileptic

drug

Cyclophosphamide CYPs Phosphoramide

mustard, acrolein

Cytostatic drug

Ifosfamide CYPs Ifosfamide mustard,

acrolein, dechloro–

ethyl–ifosfamide,

chloroacetaldehyde

Cytostatic drug

Tegafur CYPs 5-Fluorouracil Cytostatic drug

Thio–TEPA CYPs TEPA Cytostatic drug

CYP: Cytochrome P450; PAH: Polycyclic aromatic hydrocarbon; SULT: Sulfotransferase; TEPA:

N,N′,N′′–triethylenephosphoramide.

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Although CYP activity appears to be immature during a major part of the

(pro)teratogen–sensitive period, we were able to detect relatively high mRNA

expression levels for CYP1A and CYP3C1 already at 1.5 hpf (Chapter 4) which

suggests maternal transfer of the respective mRNA transcripts since the zebrafish

zygotic genome itself becomes gradually activated only around 2.75–4.75 hpf

(reviewed by [27]). Moreover, Goldstone and colleagues (2010) [28] were able to

detect CYP1A transcripts in unfertilized eggs, which confirms that the maternal

mRNA transcripts are produced during oogenesis and are present in the egg at

fertilization. Although maternal CYP mRNA transcripts have been detected in

several studies, they are, together with other maternal factors, considered to be

essential in early zebrafish embryonic development, rather than to play a role in

xenobiotic metabolism (reviewed by [27]).

Besides the abovementioned CYP enzymes, maternal transfer was also

observed for the abcb4 transcripts (Chapter 4). Hence, this transporter, which is

functionally similar to human P–glycoprotein [29], is considered to be essential

for the protection of the early zebrafish embryo against environmental

compounds. After a decline in transcript levels during the gastrula and

segmentation period, abcb4 mRNA levels reached a maximum by the end of

zebrafish organogenesis. The latter coincides with the maximum mRNA

expression levels of most phase I and phase II enzymes in this project as well as

with the onset of independent feeding [30]. Moreover, abcb4 transcript levels

reached a maximum by the time that liver and intestine have been fully

developed. Since these two organs appeared to contain abcb4 transcripts in adult

zebrafish (Table 5 in Chapter 1), the ABCB4 transporter might have a role in the

efflux of xenobiotics, e.g. in the bile or in the intestinal lumen, by the end of

zebrafish organogenesis. However, as the transport activity of the ABCB4

transporter has not been measured in the doctoral project, the abovementioned

assumption cannot be confirmed.

Section 2 of this chapter will elaborate on the biotransformation capacity of

other alternative test systems that have been/are being used for developmental

toxicity testing as well as on possible methods that can be applied to overcome a

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lack of metabolizing capacity and accordingly improve the sensitivity and

predictivity of these models.

2 Possible solutions to prevent false negative results in

alternative developmental toxicity testing

Besides the ZEDTA, the problem regarding the lack of biotransformation

capacity has also been described for other alternative test systems that have

been/are being used for developmental toxicity testing such as the limb bud

micromass culture, embryonic stem cell test (EST) and WEC (Table 2). Similar to

zebrafish, the drug–metabolizing capacity of embryos and tadpoles from the

African clawed frog (Xenopus laevis)—another alternative non–mammalian

model in teratogenicity studies (i.e. frog embryo teratogenesis assay–Xenopus

(FETAX))—was shown to be immature [31,32]. For a description of the

abovementioned alternative test systems, we refer to section 2.3 of Chapter 1.

Since the lack of (sufficient) biotransformation capacity seems to be a

generalized problem in the alternative developmental toxicity assays, several

solutions have been proposed and applied to address this problem. In this

respect, the addition of the metabolites of the corresponding test compound to

the culture medium might seem a logical solution. However, in case of a new

chemical entity, the metabolites might be unknown or poorly characterized [33].

Moreover, the synthetic production of known metabolites is an expensive and

labor–intensive process. Alternatively, the addition of an exogenous metabolic

activating system (MAS) to the culture medium has been proposed as a solution

to avoid false negative results in the alternative teratogenicity assays. The

following paragraph summarizes some commonly used options for MAS that

have been used in alternative test systems such as the Ames test and the WEC.

Liver preparations play a crucial role in the external MAS due to the important

drug–metabolizing capacities of this organ. For an overview of the different

metabolic activating systems that have been used in alternative assays for

developmental toxicity, we refer to Table 2.

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Co–incubation with hepatocytes

Co–incubations of mammalian embryos with hepatocytes isolated

from mouse, rat and rabbit have been described in literature (Table

2). Moreover, in a study by Hettwer et al. (2010) [34], human

hepatocytes were used as a MAS in the EST. Hepatocytes can be

obtained freshly isolated or cryopreserved. When using hepatocytes

as a MAS, the full complement of metabolic enzymes, i.e. phase I and

phase II enzymes including cofactors, and all relevant metabolic

pathways are covered [35]. Furthermore, Oglesby and colleagues

(1986) [36] showed that rat hepatocytes had the least embryotoxic

effects on the co–cultured embryo. However, there are significant

differences in culture requirements for embryos and hepatocytes [37].

Moreover, the use of hepatocytes as a MAS is a relatively expensive

and labor–intensive procedure as well as not highly amenable to

automation [35]. Due to these disadvantages, hepatocytes are not a

good choice as a MAS in the ZEDTA.

Co–incubation with liver S9 fraction

Liver S9 fraction can be obtained by differential centrifugation of liver

tissue at 9,000× g, which results in a supernatant that contains phase I

and II metabolic enzymes such as CYPs, UGTs, SULTs, aldehyde

oxidases, glutathione S–transferases, etc. [35]. S9 fractions from rat

and human livers have been used as a MAS in limb bud micromass

cultures and in WECs [38]. Moreover, S9 fractions have also been used

in the Ames test in order to examine the mutagenic potential of

procarcinogens, i.e. compounds that are enzymatically transformed

to electrophilic metabolites that may covalently bind to DNA leading

to mutation (Table 2) [39]. Although liver S9 fraction does not

represent the true metabolic profile of a hepatocyte, it is

physiologically relevant since it contains phase I as well as phase II

enzymes. However, the addition of co–factors such as NADPH (phase

I oxidation) and glutathione (phase II) to the culture medium is

199

required. In contrast to hepatocytes, the use of S9 fractions as a MAS

is much more amenable to high–throughput screening and represents

a relatively inexpensive technique [35].

Co–incubation with liver microsomes

Liver microsomes are small artificial vesicles that are obtained by

differential centrifugation of the S9 fraction–containing supernatant

at 100,000× g. Since microsomes consist of fragmented endoplasmic

reticulum, these artificial structures mainly contain CYPs, UGTs and

FMOs, but lack cytosolic enzymes and thus the majority of phase II

enzymes, including co–factors. Hence, liver microsomes do not

represent the true metabolic profile of a hepatocyte [35]. However, if

one intends to examine the role of phase I enzymes in the

biotransformation of a test compound, then the use of microsomes is

appropriate. Moreover, microsomes are cost–efficient and amenable

to high–throughput automation [35]. Due to these advantages, several

literature reports described the use of rat and human liver

microsomes as a MAS in co–culture with whole organisms, including

zebrafish embryos (Table 2). The co–incubation of zebrafish embryos

with a MAS is referred to as mDarT, i.e. Zebrafish Danio rerio

Teratogenic assay combined with an Exogenous Mammalian MAS

[18].

In developmental toxicity studies using S9 fraction or liver microsomes as a

MAS, the metabolizing enzymes are sometimes induced by pretreatment of the

animals with e.g. Aroclor 1254 or phenobarbital which both induce a broad range

of CYP enzymes [18,31,39,40].

In addition to the abovementioned metabolic activating systems, some

studies described the use of sera from dosed rats (and humans) as culture

medium for the WEC reasoning that the sera contain the respective metabolites

(reviewed by [33]). However, the use of sera as a MAS has not been well

established.

200

Although metabolic activating systems are prepared from different

mammalian species, a MAS derived from human tissue is best suited to mimic

the in vivo situation. Indeed, a non–human MAS might render different

metabolites compared to humans, which lowers the predictivity of the metabolic

teratogenicity test (Table 3a) [41]. However, the availability of human tissue is

much more limited compared with animal tissue. Moreover, exogenously added

MAS such as hepatocytes, S9 fraction and microsomes represent only a small part

of the whole in vivo system. Indeed, metabolism is always linked with passive or

active transport of compounds and metabolites across membranes and is affected

by intra– and extracellular milieu, which cannot be completely controlled [42,43].

Furthermore, individual–related factors such as underlying disease, age, gender,

co–medications, nutritional status, physical activity and genetic predisposition

affect the biotransformation capacity of an individual [42]. To this end, the

exogenous MAS often contains a pool from a large number of individual liver

tissues as a representation of the metabolizing capacity of an average individual

or population. However, the impact of interindividual variability on metabolism

might be missed as such [44].

Despite the abovementioned limitation, the use of a MAS in zebrafish

developmental toxicity studies seems to be promising to overcome the problem

regarding the immature biotransformation capacity in zebrafish embryos.

However, co–culturing embryos with a MAS has another major disadvantage: S9

fraction and liver microsomes are embryo- and cytotoxic. In the 1980s and 1990s,

cytotoxic effects have already been assigned to MAS when co–cultured with limb

bud micromass culture [45], WEC [46] and EST [34]. With regards to the zebrafish

embryo teratogenicity assay, some research groups have put effort into

optimizing the co–incubation method with rat and human liver microsomes

[40,47,48]:

In a study from Busquet et al. (2008) [18], the incubation period of the

zebrafish embryos with rat liver microsomes was restricted to 1 h

(between 2 and 3 hpf) in order to avoid toxicity of the MAS itself.

However, due to such a limited exposure window, susceptible

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developmental processes that occur during other windows of

development might not be considered. To deal with this problem,

Mattsson and colleagues (2012) [40] performed incubation with a MAS

at several developmental stages and extended the duration of the

incubation period, i.e. at 2–3 hpf (cleavage of cytoplasm), 12–14 hpf

(segmentation period) and 24–28 hpf (pharyngula period). Since the

embryos showed to be less vulnerable to MAS at 12 and 24 hpf, the

exposure duration could be extended to up to 2 and 4 h, respectively,

without impact on development [40].

In the study of Mattsson et al. (2012) [40], zebrafish embryos were put

in a microplate format in which they were individually exposed to the

MAS and the test compound so that they could not influence each

other.

A colleague from our lab found that 0.1M Tris–Hcl buffered embryo

solution was the ideal medium for co–incubation of zebrafish

embryos with liver microsomes since it showed the best balance

between no embryotoxicity and CYP activity (Unpublished data).

Pype et al. (2017) [48] and Mattsson et al. (2012) [40] found that the

microsomes themselves were embryotoxic. Moreover, Pype et al.

(2017) [48] showed that the endoplasmic reticulum seems to have

inherent embryotoxic properties that are not linked to CYP activity.

In addition, NADPH, which is an essential co–factor for CYP activity,

showed to be embryotoxic as well [40,46]. As a solution, Mattsson and

colleagues (2012) [40] included a pre–incubation step in which the

microsomes were incubated with the test compound, followed by

ultracentrifugation of the pre–incubate in order to remove the

microsomes. Subsequently, the resulting supernatant, which contains

the test compound and its metabolites, was added to the embryos.

However, this procedure did not sufficiently reduce the

embryotoxicity. With this problem in mind, a colleague from our lab

performed the same pre–incubation procedure followed by a dilution

of the supernatant prior to co–incubation with the zebrafish embryos.

202

Due to this additional dilution step, the MAS did not show any toxic

effects in the zebrafish embryos (Doctoral thesis of C. Pype, 2018). In

a recent study of Giusti et al. (2019) [49], the test compound was pre–

incubated with rat liver microsomes, followed by methanol

precipitation, centrifugation and evaporation of the supernatant. The

resulting pellet, which contains a mix of the parent compound and its

metabolites, was resuspended and added as a MAS to zebrafish

embryos at 72 hpf. Due to this method, Giusti and colleagues (2019)

[49] were able to prolong the incubation time to 48 h after which

mortality and malformations of the embryos was assessed at 120 hpf.

The co–incubation of zebrafish embryos with an external mammalian MAS

has the potential to be used in zebrafish developmental toxicity studies, although

the technique needs to be further optimized and validated before it can be used

in regulatory testing.

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Table 2: An overview of the external metabolic activating systems (MAS) that have been/are being used in alternative toxicity

testing

205

Table 2: Continued

206

3 Xenobiotic metabolism in zebrafish: an intra– and

interspecies comparison

3.1 Xenobiotic metabolite formation during zebrafish development

In this doctoral project, zebrafish embryos were able to biotransform the

fluorogenic, non–specific CYP substrate benzyloxy–methyl–resorufin (BOMR)

into the fluorescent metabolite resorufin from 72 hpf onwards. Since the detection

of resorufin is a measure for CYP activity, zebrafish embryos appear to have

CYP–mediated biotransformation capacity by the end of the organogenesis

period. Our findings are in accordance with a study from Alderton and

colleagues (2010) [58] where zebrafish embryos of 72 hpf were able to catalyze the

oxidation of chlorpromazine (an antipsychotic drug) and verapamil (a calcium

channel blocker). In addition, our lab previously showed that zebrafish embryos

were able to metabolize dextromethorphan, i.e. a human CYP2D6 probe substrate

[59], into dextrorphan from 72 hpf onwards [24]. In both studies, the respective

metabolites were detected and identified in homogenates [58] or microsomes [24]

prepared from whole zebrafish embryos by means of liquid chromatography–

mass spectrometry (LC–MS/MS), i.e. a technique which combines the physical

separation with chemical analysis of the different compounds present in the

embryo.

Some studies showed an increase in metabolite formation with increasing

zebrafish age. According to Alderton et al. (2010) [58], 10 metabolites of verapamil

were detected in zebrafish embryos at 72 hpf, whereas 12 metabolites of the drug

were detected in larvae of 7 dpf. Moreover, one of the mono–oxidized metabolites

of chlorpromazine was more abundant in zebrafish larvae of 7 dpf compared to

the embryos. In addition, Saad et al., (2017a) [24] showed higher dextrorphan

levels in zebrafish embryos at 96 hpf than at 72 hpf. However, at 120 hpf, the

levels of this dextromethorphan metabolite appeared to be significantly lower

compared with 96 hpf. A similar pattern could be observed in this doctoral project

after exposure of zebrafish embryos and larvae to BOMR since resorufin

formation appeared to be higher at 98 hpf compared with 74 and 122 hpf (Chapter

4). Although the reason for the increase in CYP activity at 96 hpf is unknown, this

207

finding seems to be irrelevant since metabolite analysis studies in zebrafish

embryos and larvae have shown that the amounts of any specific metabolite were

low and accounted for only a small percentage of the parent compound [24,58].

Furthermore, the concentrations of metabolites produced by zebrafish

embryos/larvae are generally lower compared to the same metabolites formed by

adult zebrafish. Indeed, dextrorphan levels appeared to be much lower after

incubation of dextromethorphan with microsomes prepared from whole

zebrafish embryos of 72, 96 and 120 hpf compared to dextrorphan levels

produced by adult zebrafish liver microsomes (ZLM) [24]. However, CYP activity

and accordingly metabolite formation in microsomes prepared from whole

organisms might be underestimated due to dilution of the CYP enzymes. The

finding regarding the low metabolite concentrations in zebrafish embryos/larvae

is in accordance with the in vitro data of this project since BOMR–metabolite

formation by microsomes of zebrafish embryos/larvae was significantly lower

compared to ZLM (Chapter 3) and even when compared with microsomes

prepared from whole adult zebrafish (Chapter 4). Besides lower metabolite

concentrations, zebrafish embryos/larvae appear to produce a smaller range of

metabolites in comparison with adult fish. Saad et al. (2017a) [24] showed that

ZLM were able to produce a second metabolite besides dextrorphan, i.e. 3–

methoxymorphinan (Table 3a), which was not detected after exposure of

embryonic/larval microsomes to dextromethorphan. Moreover, Chng and

colleagues (2012) [60] detected 6β–hydroxytestosterone and one putative

hydroxylated metabolite in zebrafish larvae homogenates of 120 hpf exposed to

testosterone, whereas more metabolites were identified in ZLM, i.e. 2α–, 6β–, and

16β–hydroxytestosterone and three putative hydroxylated metabolites.

However, there is a discrepancy in literature with regards to the

biotransformation of testosterone in both zebrafish embryos/larvae and adults

(Table 3a). Alderton and colleagues (2010) [58] identified one hydroxylated

testosterone metabolite in zebrafish larvae of 7 dpf, whereas Saad et al. (2017b)

[25] only detected negligible metabolite concentrations in zebrafish embryos of

96 hpf. With regards to the adults, six minor non–hydroxylated testosterone

metabolites were detected for ZLM in the study of Saad et al. (2017b) [25], whereas

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Reschly et al. (2007) [61] identified four hydroxylated metabolites, i.e. 6β–, 15α–,

16α–, and 16β–hydroxytestosterone in adult zebrafish hepatocytes.

Hence, we may conclude that CYP–mediated metabolism of xenobiotics in

zebrafish embryos and larvae is different from adults. Moreover, based on the in

vitro data of the doctoral project as well as on the findings of the abovementioned

studies, we and other authors [24,58] assume that CYP–mediated metabolism

during zebrafish organogenesis (and larval development) is low and needs to be

considered when interpreting developmental toxicity data in this model for

human safety assessment.

Phase II metabolism has also been investigated during zebrafish

development by several research groups, albeit to a lesser extent compared to

phase I metabolism. The UGT metabolite of testosterone, i.e. testosterone

glucuronide, was detected in larvae of 120 hpf [60] and 7 dpf [58], which suggests

the presence of functional UGT enzymes by the end of zebrafish organogenesis.

Moreover, zebrafish embryos of 72 hpf were able to produce paracetamol (or

acetaminophen) sulfate as a SULT metabolite at concentrations that were about

five to six times higher than the concentrations of paracetamol glucuronide [62].

The findings regarding the UGT and SULT enzyme activities are in accordance

with the mRNA expression analysis of SULT1ST1 and UGT1A1 in the doctoral

project. Indeed, SULT1st1 transcripts showed to be already mature in early

embryonic development, whereas UGT1A1 transcripts reached maximum

expression levels by the end of zebrafish organogenesis (Chapter 4).

Nevertheless, this similarity is merely an assumption since enzyme expression

does not necessarily coincide with corresponding enzyme activity. Brox and

colleagues (2016) [63] were able to detect sulfate conjugates of clofibric acid—an

environmental contaminant—already in zebrafish embryos of 7 hpf, whereas the

glucuronide–conjugated metabolites were identified from 28 or 52 hpf onwards

(incubation at 26°C). Furthermore, the internal concentrations of UGT and SULT

metabolites in zebrafish embryos increased with exposure time (up to 96 h).

However, adult zebrafish samples were not included in the study with clofibric

acid, which makes it difficult to draw a conclusion about the relevance of the

209

internal metabolite concentrations for phase II metabolism during early

embryonic development.

210

Table 3a: Interspecies comparison of the metabolites of some well–known human CYP substrates.

Substrate Zebrafish

metabolite(s)

Human

metabolite(s)

Rat metabolite(s) Rabbit

metabolites(s)

Ref.

HUMAN CYP–SPECIFIC SUBSTRATES

Dextromethorphan

(CYP2D6)

3–

Methoxymorphinan

Dextrorphan

Dextrorphan

3–Methoxymorphinan

3–hydroxymorphinan

Dextrorphan



Dextrorphan



[24,58,64

-66]

Diclofenac

(CYP2C9)

4’–hydroxydiclofenac

5–hydroxydiclofenac






[24,58,67

-69]

Midazolam

(CYP3A4/5)

None 1–hydroxymidazolam

4–hydroxymidazolam




[24,58,70,

71]

Testosterone (TST)

(CYP3A4/5)

Discrepancy in

literature*

6β–hydroxyTST

2β– hydroxyTST

Androstenedione

15β– hydroxyTST

16β–hydroxyTST

6β–hydroxyTST

16α– and 16β–

hydroxyTST

Androstenedione

7α– hydroxyTST

2α– and 2β–

hydroxyTST

6β–hydroxyTST

[24,25,60,

67,72,73]

Bold text represents the major metabolite of the respective substrate. Since human metabolites have been more extensively studied, the

list of metabolites might be longer compared to the other species. However, literature reports on metabolite formation in rabbits are scarce.

Hence, the column with rabbit metabolites might be incomplete. * see 3.1 of the current chapter for discrepancy in literature regarding

testosterone.

211

Table 3a: Continued

Substrate Zebrafish

metabolite(s)

Human metabolite(s) Rat metabolite(s) Rabbit

metabolite(s)

Ref.

OTHER HUMAN CYP SUBSTRATES

17β–estradiol

(E2)

2–hydroxyE2

4–hydroxyE2

2–hydroxyE2

4–hydroxyE2

6α– and 6β–hydroxyE2

7α–hydroxyE2

12β–hydroxyE2

15α–hydroxyE2

16α– and 16β–hydroxyE2

2–hydroxyE2

4–hydroxyE2

2–hydroxyE2

4–hydroxyE2

[74-76]

Albendazole

(ABZ)

Albendazole sulfoxide

Albendazole sulfone

ABZ–2–aminosulfone


Albendazole sulfone


ABZ–β–hydroxysulfone

ABZ–γ–hydroxysulfone


Albendazole sulfone



Albendazole sulfone

[40,77-

80]

Bold text represents the major metabolite of the respective substrate. Since human metabolites have been more extensively studied, the

list of metabolites might be longer compared to the other species. However, literature reports on metabolite formation in rabbits are scarce.

Hence, the column with rabbit metabolites might be incomplete.

212

Table 3b: Interspecies comparison of the in vitro intrinsic clearance values (expressed as µL/min/mg microsomal protein) for

some well–known human CYP substrates.

Pooled liver microsomes were used to determine the intrinsic clearance (CLint) values. The in vitro CLint values were calculated

based on the enzyme kinetic parameters, i.e. Km (Michaelis–Menten constant) and Vmax (maximal velocity) according to formula

CLint = Vmax/Km. Pooled liver microsomes contained mixed genders, except for * pooled male liver microsomes; ** and *** the

CLint values shown in the table represent the mean of the CLint values for males and females. Moreover, the CLint for testosterone

in rat (***) showed distinct gender–related differences with male rats exhibiting higher enzyme activity than females [67].

Substrate Zebrafish

metabolite

Human metabolite Rat metabolite Rabbit metabolite Ref.

HUMAN CYP–SPECIFIC SUBSTRATES

Dextromethorphan

(CYP2D6)

Dextrorphan

CLint: 13

Dextrorphan

CLint: 3.4

Dextrorphan

CLint: 271*

Unknown [24,81,

82]

Diclofenac

(CYP2C9)


CLint: 56


CLint: 215


CLint: 100*


CLint: 5**

[24,67,

82,83]

Midazolam

(CYP3A4/5)

None 1–hydroxymidazolam

CLint: 158


CLint: 767*


CLint: 271

[24,71,

82]

Testosterone (TST)

(CYP3A4/5)

Discrepancy in

literature regarding

metabolites

CLint: 31

6β–hydroxyTST

CLint: 89**

6β–hydroxyTST

CLint: 20***

6β–hydroxyTST

CLint: 42**

[24,67

]

213

3.2 The ontogeny of drug disposition enzymes and transporters in

zebrafish versus humans

Although maternal metabolism is absent during zebrafish development, a

comparison between zebrafish and humans regarding the ontogeny of drug

disposition enzymes and transporters is essential to assess the predictivity of the

zebrafish embryo as a model for human risk assessment. However, the genes of

CYP families 1–3 in zebrafish are much more diverse compared to the

“endogenous” CYP families, resulting in much less conservation of sequence

between zebrafish and human (Section 4.2 of Chapter 1: Introduction) [28]

Although there are orthologous relationships for some CYP1 and some CYP3

genes between zebrafish and human, zebrafish have 47 CYP2 genes compared to

16 in human (Table 3a of Chapter 1: Introduction) [28]. The differences between

zebrafish and human CYP genes may be explained by a.o. the remnants or so–

called “ohnologs” of whole genome duplication in the teleost line [28,84]. With

regards to phase II enzymes, zebrafish UGT1 and UGT2 genes as well as SULT1,

SULT2, SULT4 and SULT6 genes are closely related to the corresponding human

gene families. However, zebrafish contain UGT and SULT genes that do not exist

in humans and vice versa (Table 4 of Chapter 1: Introduction) [85,86]. Due to this

complexity in orthologous relationships, it is difficult to make a direct

comparison between the ontogeny of zebrafish and human drug disposition

enzymes. Hence, for this comparison, we will focus on the drug disposition and

transporter genes that have been assessed in the mRNA expression analysis of

this project (Chapter 4) and for which a human homologue has been described in

literature (Table 3a, 4 and 5 of Chapter 1).

Figure 2 shows a schematic representation of human prenatal development.

For the timeline of zebrafish development, we refer to Figure 1 in Chapter 4.

Zebrafish organogenesis, which occurs between 4 and 120 hpf, can be compared

with human embryonic development, which coincides with organogenesis and

which occurs during the first trimester of prenatal development, i.e. between

week 3 and week 9 of gestation (Figure 2) [87]. Later developmental stages are

214

difficult to compare between zebrafish and humans due to differences in life

cycle.

The whole body CYP mRNA expression analysis of this project showed that

zebrafish CYP1A transcript levels increased until the end of organogenesis after

which they started to level out for the remaining developmental time points

(Chapter 4). Zebrafish CYP1A has exon structures similar to human CYP1A1 and

CYP1A2 (Goldstone et al., 2010). Human CYP1A1 mRNA appeared to be

expressed in liver and extrahepatic tissues during the first trimester (starting from

around 6 weeks of gestation) and second trimester of gestation. In contrast to

zebrafish CYP1A, human CYP1A1 expression declined with increasing age and

was not generally detectable in adult tissues (reviewed by [88]). Regarding human

CYP1A2, the expression of this enzyme appeared to be absent during prenatal

development, but started to increase in infants around 1–3 months of age

(reviewed by [88]).

Zebrafish CYP1B1 showed a distinct pattern compared to the other

investigated zebrafish CYPs: CYP1B1 transcript levels reached a peak early in

organogenesis, followed by a decrease until 48 hpf after which CYP1B1

transcripts started to level out (Chapter 4). Zebrafish CYP1B1 has a gene structure

which is very similar to human CYP1B1 [28]. However, there is a discrepancy in

literature regarding the developmental expression of human CYP1B1 since some

authors reported CYP1B1 mRNA expression in fetal hepatic and extrahepatic

tissues during the second trimester of gestation [89], whereas other authors were

unable to detect CYP1B1 mRNA in either fetal or adult liver [90]. Hence, Hines

(2008) [91] assumed that human CYP1B1 contributes little to hepatic drug

metabolism at any age.

The mRNA expression analysis of CYP3C1 showed a similar expression

pattern as for zebrafish CYP1A (Chapter 4). It has been shown that the zebrafish

CYP3C subfamily shares synteny with the functional human CYP3A4 and fetal

CYP3A7 [28]. Human CYP3A7 starts to show significant levels of expression in

the fetal liver by the end of the first trimester of gestation after which expression

begins to decline around the first postnatal week. Simultaneously, hepatic

CYP3A4 expression begins to increase postnatally at about one week of age.

215

Although total CYP3A expression remains constant over the entire

developmental period, CYP3A7 and CYP3A4 exhibit differences in metabolic

capacity [88,92].

Zebrafish SULT1ST1 reached maximum mRNA levels already around 72 hpf

after which transcript levels remained stable throughout the larval period

(Chapter 4). This phase II enzyme shows orthologous relationships with several

members of the human SULT1A family (https://zfin.org). Several authors

reported human SULT1A protein expression in liver and extrahepatic tissues

starting from the second trimester of gestation. The expression of SULT1A protein

in human liver showed little or no significant change with age throughout pre–

and postnatal life. However, the developmental expression pattern of SULT1A

proteins appeared to vary between the different tissues (reviewed by [91,93]).

The other phase II enzyme we investigated in this project, i.e. UGT1A1,

reached maximum expression levels by the end of zebrafish organogenesis after

which mRNA levels levelled off throughout the larval period (Chapter 4). As

mentioned above, zebrafish UGT1 genes are closely related to human UGT1 genes

[85]. The expression of human UGT1A1 appears to be triggered by processes

associated with birth and mRNA seems to reach adult levels by 3–6 months of

age (reviewed by [93]).

Finally, we also included a transport protein in our mRNA expression

analysis, i.e. abcb4, which showed increasing expression levels until the end of

zebrafish organogenesis after which transcript levels started to level out for the

remaining developmental time points (Chapter 4). Zebrafish abcb4 appeared to

be functionally similar to human ABCB1 (P–glycoprotein or P–gp) [29]. According

to literature, transcript levels of human P–gp were undetectable in the first

trimester of gestation. However, P–gp mRNA expression had been observed in

fetal liver, intestine and kidney from the second trimester of gestation onwards.

After the initial expression of P–gp mRNA, intestinal transcript levels appeared

to remain stable throughout pre– and postnatal development, whereas hepatic

mRNA levels seemed to increase throughout childhood development (reviewed

by [94]).

https://zfin.org/

216

In conclusion, most of the human drug disposition enzymes—CYP1A2,

CYP3A4, UGT1A1 and enzymes that do not contain a zebrafish homologue, i.e.

CYP2C9, CYP2D6 and CYP2E1—are not expressed or are expressed at low levels

during prenatal development. For these enzymes, substantial increases in

expression are observed within the first one to two years after birth. Although

enzyme mRNA or protein expression and enzyme activity are often not

correlative, the activities of most human drug disposition enzymes also appear to

increase throughout prenatal and postnatal development (reviewed by [91,93]). In

this project we showed that a) most zebrafish phase I and phase II enzymes

reached maximum mRNA expression levels by the end of the organogenesis

period and b) CYP activity was detectable by the end of embryonic development.

Although drug disposition enzymes in zebrafish seem to be expressed and active

earlier in development compared to humans, we may assume that xenobiotic

metabolism is immature during a major part of zebrafish and human

organogenesis.

217

Figure 2: A schematic representation of human prenatal development. The period of

embryonic development coincides with organogenesis and accordingly with the

teratogen–sensitive period. The liver, which is not shown on the figure, starts developing

in the 4th week of gestation. The basic liver elements and structure are formed by the end

of the first trimester of gestation (reviewed by [95]). The figure is reproduced from

Santrock (2009) [96].

3.3 Xenobiotic metabolite formation in zebrafish versus mammals

As already mentioned in section 3.1 of the current chapter, metabolic enzyme

activity can also be assessed by means of metabolite analysis after incubation of

whole zebrafish embryos or zebrafish microsomes with the parent compound. In

Table 3a the metabolites of some well–known CYP substrates have been

compared between adult zebrafish, humans and the two traditional

developmental toxicity models, i.e. rat and rabbit. In addition, Table 3b shows

the in vitro intrinsic clearance (Clint) values for some human CYP–specific

substrates to allow a more direct comparison of the metabolic capacity across

species. As shown in Table 3a, adult zebrafish and mammals may produce

218

similar metabolites despite interspecies differences in drug–metabolizing

enzymes. Indeed, after incubation with diclofenac, i.e. a human CYP2C9 probe

substrate [59], ZLM produced 4’–hydroxydiclofenac and 5–hydroxydiclofenac at

the same ratio as for human liver microsomes (HLM), although zebrafish do not

have a CYP2C9 homologue [24,28,68]. However, the Clint values for diclofenac are

4–fold lower for adult zebrafish liver microsomes compared with human liver

microsomes (Table 3b). With regards to dextromethorphan, ZLM were able to

produce the same main metabolites as in humans, rats and rabbits, i.e. 3-

methoxymorphinan and dextrorphan, albeit at a different ratio (Table 3a).

However, the Clint values for the biotransformation of dextromethorphan into

dextrorphan are 20– to 80–fold lower for zebrafish and human liver microsomes,

respectively, compared with male rat liver microsomes (Table 3b). Although

there is a discrepancy in literature with regards to the biotransformation of

testosterone in zebrafish (section 3.1 of the current chapter), the CLint values for

testosterone are of the same order of magnitude for all the species mentioned in

table 3b. In mammals as well as in zebrafish, 2–hydroxyE2 appeared to be the

major metabolite of 17β–estradiol (E2), i.e. an endogenous estrogen steroid

hormone (Table 3a). The 2–hydroxylation of E2 in humans is mainly catalyzed by

CYP1A1, CYP1A2 and CYP3A4 [75], whereas studies with recombinant zebrafish

CYPs showed that CYP1A, CYP1C1 and to a lesser extent CYP1C2 are mainly

responsible for this reaction [76]. The similarity between humans and zebrafish

regarding the involvement of CYP1A in the 2–hydroxylation of E2 might be

explained by the fact that zebrafish CYP1A has exon structures similar to human

CYP1A1 and CYP1A2 [28]. However, zebrafish CYP1C1 and CYP1C2 do not have

a human homologue [28]. Recombinant zebrafish and recombinant human CYPs

have also been used in this doctoral project where they were incubated with the

fluorogenic substrate BOMR (Chapter 3). The BOMR assay showed that, from the

recombinant enzymes that were included in the assay, recombinant CYP1A was

one of the major enzymes involved in the biotransformation of BOMR for both

human and zebrafish. The other major human enzymes that were involved in

BOMR metabolism were CYP2B6 and CYP3A4 whereas recombinant human

CYP2C9 contributed to a lesser extent. However, in zebrafish, other recombinant

219

CYPs appeared to be involved in the biotransformation of BOMR besides CYP1A,

i.e. recombinant CYP1B, CYP1C1 and CYP1C2, albeit to a lesser extent. The

zebrafish CYP3C subfamily, which shares synteny with the functional human

CYP3A4, had not been included in the recombinant assay (Chapter 3). Hence, the

same recombinant zebrafish CYP enzymes seem to be involved in the

biotransformation of BOMR and 17β–estradiol (E2).

In contrast to the abovementioned similarities between zebrafish and

mammalian metabolites, a clear difference had been observed for midazolam, a

CYP3A4 probe substrate [59], since no metabolites could be detected after

incubation with ZLM [24] (Table 3a). The same had been observed in zebrafish

larvae of 7 dpf in a study from Alderton et al. (2010) [58]. These results are in

accordance with the Luciferin–IPA assay in this project since ZLM were not able

to metabolize this highly CYP3A4–specific substrate (Chapter 3).

Although human and zebrafish xenobiotic metabolite formation showed

some similarities, qualitative and quantitative differences between both species

have been observed. Moreover, dissimilarities in metabolite formation have also

been observed between humans and laboratory animals, albeit to a lesser extent.

Indeed, the metabolite ratio of midazolam appeared to be different between

humans and rats (Table 3a). Interspecies differences in metabolite formation are

mainly due to differences in drug disposition enzymes. Many zebrafish CYP

enzymes do not have a human orthologue due to, a.o., the retention of duplicated

genes or “ohnologs” of the third round of whole genome duplication in the teleost

line [28,84]. Even in cases where human orthologues of CYPs are present in

zebrafish, divergences may occur in relation to the major metabolites generated

or the metabolite ratio in comparison with humans. On the other hand, not

having an orthologue of human CYP in the zebrafish does not necessarily mean

that there will be no biotransformation of xenobiotics since other CYP(s) can take

over its function (reviewed by [97]). Due to interspecies differences in drug

disposition genes, the use of different species in developmental toxicity assays is

recommended in order to increase the predictivity for human risk assessment.

Moreover, it is not standard practice for drug metabolites to be evaluated

220

separately in cross–species safety assessment during preclinical drug

development. Hence, their specific contribution to the overall toxicity of the

parent compound has often remained unknown. In order to avoid delays in drug

development and marketing, the FDA Guidance on Safety Testing of Drug

Metabolites (2016) [98] encourages the industry to identify any differences in drug

metabolism between animals used in preclinical safety assessments and humans

as early as possible during the drug development process. In this respect, the

findings of the doctoral project on the ontogeny of zebrafish drug disposition

enzymes provide relevant background information with regards to its use as an

alternative model for developmental toxicity screening early in the drug

development process.

4 General conclusion and recommendations

Based on the findings of the doctoral project, we assume that the drug–

metabolizing capacity of zebrafish embryos is immature during the teratogen–

sensitive period of development. Hence, the malformations or toxic effects that

one may see in the zebrafish embryo developmental toxicity assay (ZEDTA) are

mainly due to the parent compound rather than the metabolite(s). However, the

lack of early biotransformation capacity presents a problem in case of

proteratogens since this may lead to false negative results in the ZEDTA. The

ideal solution would be to mimic maternal metabolism by adding an external

metabolic activating system (MAS) to the culture medium. Preference is given to

human liver microsomes as a MAS since it facilitates extrapolation from the

outcome of the ZEDTA to human risk assessment. Nevertheless, the availability

of human tissues is limited compared to laboratory animals, especially when

aiming at high–throughput assays. In order to prevent embryotoxicity due to the

MAS, the latter needs to be pre–incubated with the test compound after which

the pre–incubation mixture is purified to remove the embryotoxic compounds

before adding it to the culture medium. In this respect, the method of Giusti et al.

(2019) [49] seems to be promising since they were able to prolong the incubation

time with the MAS to 48 h. However, the authors exposed zebrafish embryos at

221

72 hpf, which is rather late in the organogenesis period. Moreover, Mattsson et

al.(2012) [40] showed that the vulnerability of zebrafish embryos decreases with

increasing age. Ideally, co–incubation with a MAS needs to occur during the

entire exposure window of the ZEDTA, i.e. between 4 and 120 hpf. Hence, the co–

incubation method with an external MAS needs to be further optimized,

especially with regards to the co–incubation time and period, followed by a

validation procedure in which the reproducibility of the method is evaluated by

testing (pro)teratogenic compounds as well as non–teratogenic compounds at a

range of concentrations in a number of independent runs as well as in different

laboratories [99]. The validation of the co–incubation of the ZEDTA with a MAS

is a prerequisite for acceptance in regulatory developmental toxicity testing.

This doctoral project has contributed to a better understanding of the

biotransformation capacity during zebrafish embryonic and larval development.

However, with the aim of using the ZEDTA in regulatory toxicity testing, the

knowledge gap regarding phase II enzyme and transporter activity during

zebrafish development needs to be further addressed. Similar to phase I enzymes,

phase II enzyme activity can be assessed by the analysis of metabolites after

administration of a test compound, whereas drug uptake studies may be

performed in e.g. hepatocytes by using fluorescent probes such as sodium

fluorescein (OATP family) [100], 4-[4-(dimethyl-amino)styryl]-N-

methylpyridinium iodide (ASP+) (OCT/N family) [101] and rhodamine–123

(human ABCB1 or P–gp) [102]. However, assessing transporter activity will be

more challenging as specific probes are still lacking for some individual

transporters [94].

222

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231

Summary

233

The zebrafish embryo is increasingly used as an alternative model to screen

drug candidates and environmental pollutants for developmental toxicity (i.e.

teratogenicity). Since the zebrafish is not considered to be a test animal until it

reaches the stage of independent feeding, the zebrafish embryo developmental

toxicity assay (ZEDTA) fits within the 3Rs (i.e. Replacement, Reduction and

Refinement) concept as described within laboratory animal sciences. The

externally developing zebrafish embryo cannot rely on maternal metabolism

unlike the mammalian embryo. Hence, the zebrafish embryo gets directly

exposed to the parent compound and depends on its own drug–metabolizing

capacity for the detoxification or bioactivation of xenobiotics. In this respect,

knowledge of the intrinsic biotransformation capacity during zebrafish

organogenesis, which coincides with the exposure window of the ZEDTA, is key

in order to correctly interpret the outcome of the ZEDTA. However, the overall

results of studies described in literature regarding the xenobiotic–metabolizing

capacity of zebrafish embryos are contradictory.

Hence, the main goal of this doctoral project was to characterize drug

disposition in zebrafish during organogenesis with a main focus on cytochrome

P450 (CYP)–mediated metabolism since the latter enzymes are responsible for the

oxidation of the majority of marketed drugs. To this end, the thesis investigates

the ontogeny of CYP enzymes on mRNA as well as on activity level, and to a

lesser extent also of the expression levels of two major phase II enzymes and a

drug transporter, i.e. abcb4, at different time–points during zebrafish

organogenesis and beyond.

This project mainly showed that CYP–mediated biotransformation of

xenobiotics appears to be immature during a major part of the ZEDTA exposure

window (i.e. 4–120 h post–fertilization (hpf)). Moreover, the mRNA expression

levels of the phase II enzymes and abcb4 reached maximum expression levels by

the end of zebrafish organogenesis. These findings can have a profound impact

on the predictivity of the ZEDTA for human safety assessment in the drug

development process, especially in case of proteratogenic compounds that

require bioactivation to exert their teratogenic potential.

234

A solution to overcome the immature biotransformation capacity of zebrafish

embryos is to co–incubate the ZEDTA with a human–derived external metabolic

activating system (MAS), such as human liver microsomes, during the entire

exposure window of the ZEDTA. However, the co–incubation method with the

external MAS needs to be further optimized and validated before it can be used

in regulatory developmental toxicity testing.

235

Samenvatting

237

Het zebravisembryo wordt steeds meer gebruikt als alternatief model om

nieuwe geneesmiddelen en milieuverontreinigende stoffen te screenen op

ontwikkelingstoxiciteit (i.e. teratogeniteit). Zebravissen worden volgens de

Europese wetgeving niet beschouwd als proefdieren tot het moment dat ze zich

onafhankelijk van hun dooier beginnen voeden. Bijgevolg past de

ZebravisEmbryo Ontwikkelingstoxiciteit Test (ZEDTA) binnen het concept van

de 3 V’s, i.e. Vermindering, Verfijning en Vervanging, zoals beschreven binnen

de proefdierkunde. Echter, het zebravisembryo ontwikkelt uitwendig waardoor

het geen beroep kan doen op het maternale geneesmiddelenmetabolisme zoals

bij zoogdieren. Bijgevolg wordt het zebravisembryo rechtstreeks blootgesteld aan

de teststof waardoor het embryo zelf verantwoordelijk is voor de detoxificatie of

voor de bioactivatie van geneesmiddelen. Om de resultaten van de ZEDTA op

een juiste manier te interpreteren, is het van essentieel belang om voldoende

kennis te hebben over de intrinsieke biotransformatie capaciteit van zebravissen

en dit vooral tijdens de organogenese–periode aangezien deze laatste samenvalt

met de blootstellingsperiode van de ZEDTA. Desalniettemin toont de

wetenschappelijke literatuur tegenstrijdige resultaten betreffende de capaciteit

van zebravisembryo’s om geneesmiddelen te metaboliseren.

Aldus was het hoofddoel van het huidige doctoraatsproject de karakterisatie

van de geneesmiddelendispositie in zebravissen tijdens de organogenese–

periode met de focus op cytochroom P450 (CYP)–gemedieerde omzetting. CYP

enzymen zijn immers verantwoordelijk voor de oxidatieve omzetting van het

merendeel aan geneesmiddelen op de markt. Daartoe onderzoekt de huidige

thesis de ontogenie van CYP enzymen zowel op mRNA- als op activiteitsniveau

en in mindere mate ook de expressie niveaus van twee belangrijke fase II

enzymen en een geneesmiddelen–transporter, nl. abcb4, op verschillende

tijdspunten tijdens en na de organogenese.

Het huidige project toonde voornamelijk aan dat CYP–gemedieerde

biotransformatie van geneesmiddelen immatuur is tijdens het grootste deel van

de blootstellingsperiode van de ZEDTA (i.e. 4–120 u na bevruchting). Bovendien

bereikte de mRNA expressie van de fase II enzymen en de abcb4 transporter

maximale expressieniveaus tegen het einde van de organogenese. Deze

238

bevindingen kunnen een belangrijke impact hebben op de betrouwbaarheid van

de ZEDTA voor humane risico–analyse tijdens het

geneesmiddelenontwikkelingsproces, vooral in het geval van proteratogene

stoffen die bioactivatie vereisen vooraleer ze teratogeen worden.

Een mogelijke oplossing om de immature biotransformatie capaciteit van

zebravisembryo’s op te vangen is de co–incubatie van de ZEDTA met een

humaan exogeen metabool activerend systeem (MAS), zoals humane

levermicrosomen, en dit tijdens de volledige blootstellingsperiode van de

ZEDTA. Desalniettemin dient de co–incubatie methode met een exogeen MAS

verder gevalideerd en geoptimaliseerd te worden voor het kan worden

overwogen als een regulatoire test.

239

Dankwoord

241

“I don't know where my road is going, but I know that I walk better when I

hold your hand.” (Alfred de Musset, °1810 – †1857)

Eindelijk is het zover; het proefschrift op het einde van een lange weg. Een weg

met de nodige obstakels, kronkels, hellingen en dalingen. Het doctoraat was een

waar leerproces, niet alleen op wetenschappelijk vlak, maar ook wat betreft mijn

persoonlijke ontwikkeling: grenzen werden verlegd, het zelfvertrouwen werd

aangesterkt. Deze weg heb ik echter niet alleen afgelegd. Heel wat mensen

hebben, direct of indirect, bijgedragen tot dit doctoraat en tot de persoon die ik

nu ben. Deze mensen wil ik dan ook heel graag bedanken.

Het begon allemaal in oktober 2011. De dag na mijn sollicitatie belde Prof. André

Weyns me om te zeggen dat ik kon starten als mandaat-assistent bij de

Toegepaste Diergeneeskundige Morfologie. Professor Weyns, je stond gekend als

de ‘gevreesde’ professor van de Diergeneeskunde aan de UA, maar je was een

man met een gouden hart. Professor, bedankt voor je gedrevenheid en voor je

vertrouwen. Het gaat je goed… .

Een speciaal woordje van dank breng ik graag uit naar Prof. Steven Van Cruchten,

mijn promotor. Steven, je hebt me de afgelopen jaren de nodige vrijheid en

inspraak gegeven, wat ik ten zeerste apprecieer. Het heeft bijgedragen in mijn

zelfontplooiing en me gemaakt tot wie ik nu ben. Je hebt me een halt toegeroepen

of een andere richting uitgeduwd wanneer ik even ‘vast’ zat. Je hebt mijn

motivatie terug naar boven gehaald wanneer ik het allemaal even niet meer zag

zitten. Dankjewel, Steven, om de afgelopen jaren mijn promotor te zijn en voor je

vertrouwen. Ik wens je heel veel succes in alles wat je onderneemt, op het werk

en daarbuiten.

Een tweede woordje van dank gaat naar Prof. Chris Van Ginneken. Bedankt,

Chris, om me deze kans te geven bij de Toegepaste Diergeneeskundige

Morfologie, voor je feedback op de artikels en het proefschrift, voor je luisterend

oor wanneer het op persoonlijk vlak wat minder ging en voor je steun. Ik wens je

veel succes en geluk toe, zowel op professioneel als op persoonlijk vlak.

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De afgelopen jaren stond ik er gelukkig niet alleen voor. Heel wat mensen hebben

een belangrijke bijdrage geleverd aan dit doctoraat.

In eerste instantie wil ik graag de mensen van het Zebrafishlab bedanken. Lucia

en An, jullie hebben me alle knepen van het ‘zebravisvak’ geleerd. Bedankt om

jullie expertise met mij te delen. Bedankt ook aan Prof. Dries Knapen, hoofd van

het zebravislabo én voorzitter van mijn individuele doctoraatscommissie.

Dankjewel om deze voorzittersrol op jou te nemen, voor al jouw tips en

opbouwende feedback tijdens het doctoraat en bij het finaliseren van dit

proefschrift en voor het gebruik van jullie vissen wanneer de onze weigerden mee

te werken.

Thanks to dr. Derek Alsop and Prof. dr. Joanna Wilson from the Department of

Biology of the McMaster University in Canada to perform the experiments with

the recombinant zebrafish CYP enzymes.

Also thanks to dr. Jonathan Ball and his lab from the University of Exeter in the

UK to provide us with the zebrafish larvae of 9 and 14 days post–fertilization

which we used in the in vitro study.

Waleed, thanks a lot for helping me out with the fluorescence microscope and for

patiently answering all my questions regarding the fluorescence measurements

and calculations. I really appreciate it!

Dikke merci ook aan Gunther om al die jaren onze visjes zo goed te verzorgen.

Het was steeds een heel karwei, dag in dag uit. Ik zou niet weten wat we zonder

jou hadden moeten beginnen. Daarnaast stond je ook steeds klaar om te helpen

of om een gezellig babbeltje te slaan. Ik wens je veel geluk toe met alles wat op je

pad komt!

Bedankt Prof. Pieter Annaert, Prof. Juliette Legler en dr. Luc Van Nassauw om

deel uit te maken van mijn doctoraatsjury, voor het grondig en kritisch nalezen

van het proefschrift en voor jullie waardevolle feedback.

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Gelukkig kon ik de afgelopen jaren ook terugvallen op de lieve collega’s van de

Toegepaste Diergeneeskundige Morfologie.

Hans, Bart en Christophe, wij zijn allemaal op dezelfde dag gestart bij de

Morfologie. Hans en ikzelf als de twee naïeve doctoraatsstudentjes, Bart en

Christophe als de ervaren post–doc en docent. Bedankt voor de gezellige

lunchmomenten in de Resto en voor jullie unieke humor. Ik wens jullie veel geluk

toe, op professioneel gebied, maar eens zoveel daarbuiten… .

Maartje, Véronique, Nathalie, Marleen, Gilbert en Christel, jullie vormden reeds

een vaste waarde bij de Morfologie op het moment dat ik begon. Bedankt voor

jullie aangename ontvangst en om me wegwijs te maken in het labo. Gilbert,

bedankt voor de gezellige babbeltjes en om ervoor te zorgen dat alles zo netjes

klaar lag voor de practica. We zijn elkaar ondertussen wat uit het oog verloren,

maar ik hoop dat je het goed stelt en wens je het allerbeste toe. Dankjewel Christel

voor het regelen van alle administratieve zaken. Marleen, dikke merci voor je

eindeloze behulpzaamheid. Ik heb enorm veel van je geleerd. Heel fijn dat je

momenteel mijn bureaugenootje bent!

Een dikke merci aan Els en Sofie, mijn twee vroegere bureaugenootjes. Het klikte

meteen tussen ons gedrieën. Wat hebben we gelachen en (soms iets teveel)

getetterd. Maar ook op momenten dat het minder goed ging konden we bij elkaar

terecht. Sofie en Tom, jullie trouwfeest zal ik niet snel vergeten. Dansen tot in de

vroege uurtjes, zalig. Jullie zijn een topkoppel! Els, na Sofies doctoraat bleven we

met z’n tweeën over. Je bent op vele vlakken een enorme steun voor mij geweest.

Je hebt me de ‘wereld’ van de microsomen en de CYP’s leren kennen, stond altijd

klaar om te helpen en was steeds een luisterend oor wanneer ik dit nodig had.

Bedankt ook voor de talrijke boeiende (soms diepgaande) gesprekken. Els en

Sofie, twee topmadammen, ik wens jullie het allerbeste toe. We stay in touch!

From the second office in the corridor, I moved to the sixth office, the ‘zebafish-

office’, with ‘zebrafish-people’ Casper and Chloé. Besides being zebrafish-people

talking about zebrafish-stuff, we could get along well with each other. Casper,

thanks for your help, your support, your valuable insights. The ETS conferences

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were never boring with you around! Dear, dear Chloé, I’ve never met such a nice,

warm, helpful and friendly person as you. Thanks for your everlasting smile,

your support, your great help with the qPCRs and so much more…. I’ll never

forget our trip to Brussels to see (and off course hear) our musical hero, Ludovico.

We should definitely do this again!

Denise, Marjan en An, de bureau ‘next door’, jullie zijn stuk voor stuk

topmadammen! Bedankt voor jullie steun, peptalk en gezellige babbeltjes.

Dankzij jullie is het nooit saai in het ietwat verborgen pathologiegebouwtje.

Denise, dikke merci om altijd voor iedereen in de bres te springen en voor je

eeuwige optimisme. Niets is jou teveel. Ik kijk uit naar onze verdere

samenwerking!

Bedankt ook aan Laura, Kevin, Charlotte, Sara, Miriam, Falk, Allan, Jente, Katty

en Steve, de huidige Comparative Perinatal Development–clan. Bedankt voor al

jullie hulp en voor de bemoedigende woorden (die de afgelopen maanden zeker

van doen waren). Het was/is fijn samenwerken met jullie! Ik wens jullie het

allerbeste toe, op en naast het werk. Een speciale dankjewel ook aan Katty. Merci

voor de toffe babbeltjes, jouw goede raad, je hulp, troost en steun. Ons labo mag

haar twee handjes kussen met een gouden collega als jij!

Dikke merci ook aan de doctoraatsstudenten van het zebravislabo voor jullie

hulp, tips en gezelschap! Evelyn en Nathalie, jullie waren er als toenmalige

thesisstudenten bij tijdens de opstart van het zebravislabo in het UC-gebouw op

CDE. Het klikte meteen tussen ons gedrieën. Dat maakte dat we momenteel nog

steeds contact hebben. Evelyn, jij bent blijven ‘plakken’ in het zebravislabo en je

springt af en toe eens binnen in gebouw U voor een gezellig babbeltje. In het

nieuwe jaar moeten we zeker opnieuw werk maken van onze regelmatige

‘lunchdates’! Ik wens jou heel veel succes bij het afwerken van je doctoraat, je

verdere carrière en uiteraard met alles daarbuiten. Je bent een heel sterke

persoonlijkheid en hebt een doorzettingsvermogen om ‘U’ tegen te zeggen. I’m

sure: You can do it! Nathalie, jij hebt ondertussen ook al een hele weg afgelegd:

een doctoraat behaald, getrouwd met ‘onze’ Casper,… . De gezellige etentjes in

de Sole zorgen ervoor dat we op tijd en stond kunnen bijbabbelen.

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Bedankt ook Ellen, de recentste doctor, voor de talrijke fijne babbeltjes.

Bekommernissen om het onderzoek en renovatie-troubles zijn de revue

gepasseerd. Maar gelukkig hebben we ook heel wat afgelachen. Ik wens je veel

geluk toe op alle vlakken. We keep in touch! Isabelle, ik vond het heel fijn

samenwerken met jou voor de practica Proefdierkunde. We hebben het er toen

toch heel goed vanaf gebracht. Heel veel succes met alles wat je onderneemt. Also

thanks to Jelena for your help with the qPCR. I wish you all the best!

Dankjewel aan mijn huidige lieve collega’s van het Decanaat FBD die me met

oppeppende, bemoedigende woorden hebben bijgestaan de afgelopen maanden.

Jullie zijn een topteam en ben blij dat ik er deel van mag uitmaken. Bedankt Kim

om me de kans te geven voor deze boeiende job!

En last but not least wil ik graag mijn familie en vrienden bedanken voor hun

eeuwige steun en vertrouwen, hun bemoedigende woorden en hun begrip voor

mijn ietwat beperkt sociaal leven de afgelopen maanden.

Melissa, Joke, Amelia, Stijn, An, Ivo en Petra, bedankt voor de gezellige en lekkere

etentjes, dagtripjes, shopdagen, enz. Merci Kenny, Kelly, Ivan, Peter, Joris en

Liesbet, alias ‘de Kempervenners’, voor de superleuke weekends in de Ardennen,

Centerparcs, camping Witters,…. Het is altijd ambiance met jullie!

Dikke merci aan de ‘Dierenartsvriendinnetjes’, Dominique, Nicole, Annelies,

Annemie, Goedele, Lonne, Barbara en Marijse. We zijn samen aan het

dierenartsenavontuur begonnen, hier aan de UA. En jawel, ondertussen hebben

we al exact 10 jaar (!) ons diploma op zak. Dankjewel voor jullie

onvoorwaardelijke steun tijdens al die jaren. We waren (en zijn nog steeds) een

geweldig team! Barbara, je bent er altijd geweest voor mij, zelfs op momenten dat

je het zelf moeilijk had. Dankjewel voor alle afgelopen jaren en voor de jaren die

nog gaan komen. Je bent, oprecht, een fantastische madam! Dominique, lieve

vriendin, al van in 1ste Kan kunnen we het heel goed met elkaar vinden. Steeds

kan ik op je rekenen, waar je ook bent, al is het 6000 km van hier. Bedankt voor

alles, om te zijn wie je bent.

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Eeuwige dank aan mijn lieve familie. Mama, papa, bomma en bompa, bedankt

voor jullie onvoorwaardelijke steun en liefde, voor de waarden en normen die

jullie me hebben meegegeven. Deze hebben me gebracht tot waar ik nu sta.

Mama, ik ben ervan overtuigd dat je dit speciale moment van hierboven

aanschouwt en dat je me van daaruit jouw eeuwig positieve energie toestuurt.

Bedankt dat je er altijd was voor mij. Dankjewel papa om altijd zo nauw

betrokken te zijn bij mijn studies. Je was, en bent nog steeds, mijn grootste

supporter. Ik zie jullie graag!

Dikke merci ook aan mijn schoonfamilie. Jullie hebben de voorbereidingen van

deze thesis van dichtbij meegemaakt. Bedankt Véronique, Luc en Gilberte voor

alle lekkere maaltijden die jullie ons hebben voorgeschoteld, voor jullie steun en

bemoedigende woorden en voor zoveel meer. Dankjewel ook Jolien en Arno voor

jullie enthousiasme en aanmoedigingen tijdens de laatste loodjes. Jullie kaartje

met bemoedigende spreukjes heeft al goed dienst gedaan!

Lieve Stijn, al meer dan 12 jaar ben je mijn steun en toeverlaat. We hebben al heel

wat watertjes doorzwommen, maar dat heeft onze band net sterker gemaakt.

Bedankt voor je enorme geduld, je zorgzaamheid, je liefde en trouw, je

oprechtheid. Kortom, bedankt om mijn lief te zijn. Ik zie je graag!

Curriculum Vitae Evy Verbueken

Personalia

Name: Evy Verbueken

Date of birth: 11/12/1985

Address Waterkrekel 155, 2235 Houtvenne

Nationality: Belgian

Working experience

Teaching assistant Sept. 2018 - present

Laboratory of Applied Veterinary Morphology

University of Antwerp – Campus Drie Eiken, Wilrijk

Student counselor and mentor Sept. 2018 - present



Academic assistant in Veterinary Sciences Oct. 2011 – Sept. 2018


This position comprises two main tasks, creating a variable job content:

Research: preparing a doctoral thesis

Teaching practical courses to veterinary bachelor students

(mainly anatomy and embryology)

Doctoral thesis entitled: Drug disposition in the zebrafish embryo and larva:

focus on cytochrome P450 activity

Practitioner for companion animals Jul. 2009 – Sept. 2011

At veterinary practice ‘Aan de Vesten’, Lier

Job description: first-line veterinary medicine in companion animals

As a veterinary practitioner, I was able to develop and refine my social

and communication skills in interaction with clients and colleagues.

Moreover, during my work as a practitioner, I have learned to work

independently and to take responsibility.

Education

Master in the Veterinary Sciences 2006 - 2009

Faculty of Veterinary Sciences

University of Ghent, Belgium

Bachelor in the Veterinary Sciences 2003 - 2006


University of Antwerp

Secondary school: Modern languages – Sciences 1997 - 2003

Sint-Ursula Lyceum

Lier, Belgium

Courses

FRAME Training School in the Experimental 30 Mar. – 1 Apr. 2015

Design and Statistical Analysis of Biomedical Experiments

University of Coimbra, Portugal

Laboratory Animal Science: certificate of Feb. 2014

experimenter Cat. C


English level 5 Jul. 2012

Linguapolis, Language Institute


Bibliography

Publications in international peer-reviewed journals

Verbueken E, Bars C, Ball JS, Periz-Stanacev J, Marei WFA, Tochwin A,

Gabriëls IJ, Michiels EDG, Stinckens E, Vergauwen L, Knapen D, Van Ginneken

CJ, Van Cruchten SJ. From mRNA expression of drug disposition genes to in

vivo assessment of CYP-mediated biotransformation during zebrafish

embryonic and larval development.

International Journal of Molecular Sciences. 2018; 19(12).

Saad M, Bijttebier S, Matheeussen A, Verbueken E, Pype C, Casteleyn C, Van

Ginneken C, Maes L, Cos P, Van Cruchten S. UPLC/MS MS data of

testosterone metabolites in human and zebrafish liver microsomes and whole

zebrafish larval microsomes.

Data in brief. 2017; 16.

Pype C, Verbueken E, Saad MA, Bars C, Van Ginneken CJ, Knapen D, Van

Cruchten SJ. Antioxidants reduce reactive oxygen species but not

embryotoxicity in the metabolic Danio rerio test (mDarT).

Reproductive Toxicology. 2017; 72.

Saad M, Matheeussen A, Bijttebier S, Verbueken E, Pype C, Casteleyn C, Van

Ginneken C, Apers S, Maes L, Cos P, Van Cruchten S. In vitro CYP-mediated

drug metabolism in the zebrafish (embryo) using human reference

compounds.

Toxicology In Vitro. 2017; 42.

Verbueken E, Alsop D, Saad MA, Pype C, Van Peer EM, Casteleyn CR, Van

Ginneken CJ, Wilson J, Van Cruchten SJ. In vitro biotransformation of two

human CYP3A probe substrates and their inhibition during early zebrafish

development.

International Journal of Molecular Sciences. 2017; 18(1).

Saad MA, Verbueken E, Pype C, Casteleyn CR, Van Ginneken CJ, Maes L, Cos

P, Van Cruchten SJ. In vitro CYP1A activity in the zebrafish: temporal but low


adult stage.


Saad M, Cavanaugh K, Verbueken E, Pype C, Casteleyn C, Van Ginneken C,

Van Cruchten S. Xenobiotic metabolism in the zebrafish: a review of the


families 1 to 3.

Journal of Toxicological Sciences. 2016; 41(1).

Pype C, Verbueken E, Saad MA, Casteleyn CR, Van Ginneken CJ, Knapen D,

Van Cruchten SJ. Incubation at 32.5°C and above causes malformations in the

zebrafish embryo.


Van Peer E, Verbueken E, Saad M, Casteleyn C, Van Ginneken C, Van Cruchten

S. Ontogeny of CYP3A and P-Glycoprotein in the Liver and the Small

Intestine of the Göttingen Minipig: An Immunohistochemical Evaluation;.

Basic and Clinical Pharmacology and Toxicology. 2014; 114 (5).

Oral presentations at international conferences

Localization of cytochrome P450 activity in the zebrafish embryo and larva.

At the 46th Annual Meeting of the European Teratology Society, 10-13th

September 2018, Berlin, Germany


At the BelTox annual meeting 2017, 1st December 2017, Leuven, Belgium.


At the Research Day of the faculty of Pharmaceutical, Biomedical and

Veterinary Sciences of the University of Antwerp, 27th November 2017, Wilrijk,

Belgium.

Is CYP3A activity in the liver of adult zebrafish influenced by tricaine

methanesulfonate (MS-222)?

At the 43rd Annual Meeting of the European Teratology Society, 31st August –

2nd September 2015, Amsterdam, The Netherlands.

Poster presentations at international conferences


At the 45th Annual Meeting of the European Teratology Society, 4-7th September

2017, Budapest, Hungary.

In vitro characterization and ontogeny of CYP3A activity in the zebrafish.

At the 44th Annual Meeting of the European Teratology Society, 11-14th

September 2016, Dublin, Ireland.

The effect of tricaine methanesulfonate (MS-222) on CYP3A activity in the

liver of adult zebrafish.

At the Fish and Amphibian Embryos as Alternative Models in Toxicology and

Teratology, 1-2nd December 2014, Aulnay-sous-Bois (Paris), France.

Lack of in vitro CYP3A activity in early zebrafish embryos.

At the 42nd Annual Meeting of the European Teratology Society, 1-4th September

2014, Hamburg, Germany.

Cytochrome P450 activity in the gastrointestinal system of the zebrafish

embryo.

At the 8th European Zebrafish Meeting, 9-13th July 2013, Barcelona, Spain.

The zebrafish as a model for developmental toxicity.

At the 18th National Symposium on Applied Biological Sciences, 8th February

2013, University of Ghent, Belgium.

Award

The BelTox Young Scientists Award for best platform presentation regarding:


At the BelTox annual meeting 2017, 1st December 2017, Leuven, Belgium.

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