drug disposition in the zebrafish embryo and larva: focus on … · 2020. 1. 7. · universiteit...
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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
Diergeneeskundige Wetenschappen
Universiteit Antwerpen
Prof. Dr. C.J. Van Ginneken
Faculteit Farmaceutische, Biomedische en
Diergeneeskundige Wetenschappen
Universiteit Antwerpen
Members of the PhD Steering Committee
Prof. Dr. D. Knapen Faculteit Farmaceutische, Biomedische en
Diergeneeskundige Wetenschappen
Universiteit Antwerpen
Dr. Luc Van Nassauw
Faculteit Geneeskunde en Gezondheids-
wetenschappen
Universiteit Antwerpen
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
4.1 Benzyloxy-methyl-resorufin assay in adult zebrafish liver
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
4.1 In vitro study on cytochrome P450 activity in zebrafish embryos,
larvae and adults…………………………………………………….….148
4.2 In vivo study on cytochrome P450 activity in zebrafish embryos and
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
4.3 mRNA Expression of Phase I and Phase II enzymes and P–
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
6
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
10
11
Chapter 1: General introduction
12
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.
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).
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
± 30 hpf: somite dev.
± 54 hpf: heart–beat visible
± 160 hpf: hatching
± 22 hpf: somite dev.
± 27 hpf: heart–beat visible
± 120 hpf: hatching
±18 hpf: somite dev.
± 26 hpf: heart–beat visible
± 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.
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.
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.
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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Chapter 2: Aims of the doctoral
project
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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).
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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.
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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
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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
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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
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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.
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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.
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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).
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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.
3.4 Benzyloxy-methyl-resorufin assay in adult zebrafish liver
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),
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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
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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.
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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)
109
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
L o g [K e to c o n a z o le ] M
% 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
L o g [C Y P 3 c id e ] M
% 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
L o g [K e to c o n a z o le ] M
% 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
L o g [C Y P 3 c id e ] M
% 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
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Zebrafish: an emerging technology for in vivo pharmacological assessment to
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pharmacology 2008, 154, 1400-1413.
2. Kari, G.; Rodeck, U.; Dicker, A.P. Zebrafish: an emerging model system for
human disease and drug discovery. Clinical pharmacology and therapeutics 2007, 82,
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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
124
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
3.1 In vitro study on cytochrome P450 activity in zebrafish embryos,
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
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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,
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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
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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
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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.
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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.
3.2 In vivo study on cytochrome P450 activity in zebrafish embryos and
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.
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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).
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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
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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.
141
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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.
3.3 mRNA Expression of Phase I and Phase II enzymes and P–
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).
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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.
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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
4.1 In vitro study on cytochrome P450 activity in zebrafish embryos,
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
M 5
- 4
8 h
pf
ZE
M 7
2 h
pf
ZE
M 9
6 h
pf
ZE
M 1
20
hp
f
ZL
aM
9 d
pf
ZL
aM
14
dp
f
ZM
0
2
4
Re
so
ru
fin
fo
rm
atio
n (
pm
ol/
min
/mg
MP
)
L L O Q
A B
A
B
C
* **
151
4.2 In vivo study on cytochrome P450 activity in zebrafish embryos and
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
pf
50 h
pf
74 h
pf
98 h
pf
122 h
pf
14 d
pf
0
5
1 0
1 5
3 0
6 0
9 0
E R
7 h
pf
26 h
pf
50 h
pf
74 h
pf
98 h
pf
122 h
pf
9 d
pf
14 d
pf
0
1
2
3
4
B O M RBenzyloxy-methyl-resorufin
Re
soru
fin
form
atio
n in
tru
nk
regi
on
re
lati
ve to
14
dp
f
Zebrafish developmental stage
**
Re
soru
fin
form
atio
n in
tru
nk
regi
on
re
lati
ve to
14
dp
f
Zebrafish developmental stage
7-Ethoxyresorufin
*
A
*
A
A
A
A
A
B
C CC
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.
155
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.
157
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.
4.3 mRNA Expression of Phase I and Phase II enzymes and P–
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
rel
ativ
eq
uan
tity
CYP1A(a)
Time (dpf)
0 5
02
46
8
Log2
rel
ativ
eq
uan
tity
Time (dpf)
Log2
rel
ativ
eq
uan
tity
Time (dpf)
CYP1B1(b)
0 5Time (dpf)
Log2
rel
ativ
eq
uan
tity
-20
24
162
CYP1C1
Time (dpf)
Log2
rel
ativ
eq
uan
tity
(c)
Time (dpf)
Log2
rel
ativ
eq
uan
tity
02
46
81
0
0 5
CYP1C2
Log2
rel
ativ
eq
uan
tity
Time (dpf)
(d)
Time (dpf)
Log2
rel
ativ
eq
uan
tity
02
46
163
(e) CYP2K6
Log2
rel
ativ
eq
uan
tity
Time (dpf)
Time (dpf)0 5
02
46
8
Log2
rel
ativ
eq
uan
tity
CYP3A65
Time (dpf)
Log2
rel
ativ
eq
uan
tity
(f)
50Time (dpf)
Log2
rel
ativ
eq
uan
tity
05
10
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
ativ
eq
uan
tity
Time (dpf)
(g)
Log2
rel
ativ
eq
uan
tity
Time (dpf)
0 5
01
23
165
SULT1ST1
Time (dpf)Lo
g2 r
elat
ive
qu
anti
ty
(a)
50Time (dpf)
Log2
rel
ativ
eq
uan
tity
02
46
Log2
rel
ativ
eq
uan
tity
Time (dpf)
UGT1A1(b)
Time (dpf)
Log2
rel
ativ
eq
uan
tity
0 5
05
10
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
ativ
eq
uan
tity
Time (dpf)
Abcb4(c)
Log2
rel
ativ
eq
uan
tity
Time (dpf)0 5
05
10
167
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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187
Chapter 5: General discussion
188
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.
190
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.
191
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.
192
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
193
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
195
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.
196
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
197
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.
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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
201
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.
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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.
203
204
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
208
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
3–Methoxymorphinan
3–hydroxymorphinan
Dextrorphan
3–Methoxymorphinan
3–hydroxymorphinan
[24,58,64
-66]
Diclofenac
(CYP2C9)
4’–hydroxydiclofenac
5–hydroxydiclofenac
4’–hydroxydiclofenac
5–hydroxydiclofenac
4’–hydroxydiclofenac
5–hydroxydiclofenac
4’–hydroxydiclofenac
[24,58,67
-69]
Midazolam
(CYP3A4/5)
None 1–hydroxymidazolam
4–hydroxymidazolam
4–hydroxymidazolam
1–hydroxymidazolam
1–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 sulfoxide
Albendazole sulfone
ABZ–2–aminosulfone
ABZ–β–hydroxysulfone
ABZ–γ–hydroxysulfone
Albendazole sulfoxide
Albendazole sulfone
ABZ–2–aminosulfone
Albendazole sulfoxide
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)
4’–hydroxydiclofenac
CLint: 56
4’–hydroxydiclofenac
CLint: 215
4’–hydroxydiclofenac
CLint: 100*
4’–hydroxydiclofenac
CLint: 5**
[24,67,
82,83]
Midazolam
(CYP3A4/5)
None 1–hydroxymidazolam
CLint: 158
1–hydroxymidazolam
CLint: 767*
1–hydroxymidazolam
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]).
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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Summary
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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.
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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.
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Samenvatting
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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
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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
240
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
244
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.
246
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
Faculty of Pharmaceutical, Biomedical and Veterinary Sciences
University of Antwerp – Campus Drie Eiken, Wilrijk
Academic assistant in Veterinary Sciences Oct. 2011 – Sept. 2018
University of Antwerp – Campus Drie Eiken, Wilrijk
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
Faculty of Pharmaceutical, Biomedical and Veterinary Sciences
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
University of Antwerp
English level 5 Jul. 2012
Linguapolis, Language Institute
University of Antwerp
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
metabolite levels during organogenesis and lack of gender differences in the
adult stage.
Reproductive Toxicology. 2016; 64.
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
spatiotemporal distribution, modulation and activity of Cytochrome P450
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.
Reproductive Toxicology. 2015; 56.
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
Localization of cytochrome P450 activity in the zebrafish embryo and larva.
At the BelTox annual meeting 2017, 1st December 2017, Leuven, Belgium.
Localization of cytochrome P450 activity in the zebrafish embryo and larva.
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
Localization of cytochrome P450 activity in the zebrafish embryo and larva.
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:
Localization of cytochrome P450 activity in the zebrafish embryo and larva.
At the BelTox annual meeting 2017, 1st December 2017, Leuven, Belgium.