Zebrafish Breeding in the Laboratory Environment
Andrzej Nasiadka and Matthew D. Clark
Abstract
The zebrafish, Danio rerio, has become a major model organism
used in biomedical studies. The widespread use of Danio
rerio in research laboratories requires a comprehensive understanding of the husbandry of this species to ensure efficient propagation and maintenance of healthy and genetically
diverse colonies. Breeding is a key element in zebrafish husbandry. It is a complex process influenced by a number of
factors. Mate choice and mating behavior depend, for example, on olfactory cues, visual stimuli, and social interactions. Spawning is affected by the age and size of fish, interval
at which fish are used for egg production, light cycle, diet, and
fish health status. A number of breeding strategies, based on
either single-pair matings or group crosses, are commonly
employed in the laboratory to propagate lines and to identify
carriers of specific mutations and/or transgenes. Propagation
of zebrafish lines, in particular wild-type-derived strains, is
closely monitored to ensure that genetic diversity and vigor
are maintained. A robust zebrafish line typically carries a
large number of polymorphic variations, which may interfere with reproducibility of experiments. To get a better
insight into these variations, a wild-type hybrid Sanger AB
Tübingen line has been generated from sequenced homozygous founders.
Key Words: breeding; Danio rerio; hybrid line; inbreeding;
SAT; spawning; zebrafish
Introduction
Z
ebrafish, Danio rerio, is a tropical freshwater teleost
belonging to the Cyprinidae family. The species has
become a major research model used in biomedical
studies to investigate, for example, vertebrate development
genetics, physiology, and behavior (Grunwald and Eisen
2002). The popularity of zebrafish as a model organism can
Andrzej Nasiadka, PhD, is a research associate at the Zebrafish International
Resource Center, University of Oregon, Eugene. Matthew D. Clark, PhD, is
a staff scientist in the Vertebrate Development and Genetics Department,
Wellcome Trust Sanger Institute, Cambridge, and is the Sequencing
Technology Development Team Leader, The Genome Analysis Centre,
Norwich, United Kingdom.
Address correspondence and reprint requests to Dr. Andrzej Nasiadka,
Zebrafish International Resource Center, 5274 University of Oregon, 1307
Franklin Boulevard, Eugene, OR 97403 or email [email protected].
Volume 53, Number 2
2012
be accounted for by a number of attributes, including the
relative ease of rearing and breeding in captivity, high fecundity, rapid development, short generation time, and availability of genomic resources, including the complete zebrafish
genome sequence. An additional attribute is the transparency
of the embryo, which allows for live imaging of transgenic
lines expressing products of fluorescent reporter genes in living cells and developing tissues.
The widespread use of D. rerio in research laboratories
requires a solid knowledge of the husbandry of this species
to ensure efficient propagation and maintenance of healthy
and genetically diverse colonies. Investigation of D. rerio
reproduction in the wild as well as in the laboratory setting
is of high importance for husbandry. In recent years, understanding of zebrafish reproductive biology and behavior has
greatly increased. Advances have been made with regard
to gonad development (Rodriguez-Mari et al. 2010;
Siegfried and Nüsslein-Volhard 2008) and the role of olfactory cues, visual stimuli, and social interactions in reproduction, including mate choice and courtship (Gerlach 2006;
Gerlach and Lysiak 2006; Skinner and Watt 2007; Spence
and Smith 2005, 2006).
Propagation of zebrafish wild-type-derived lines in the
laboratory is based on specific breeding schemes whose goal
is to assure not only that next fish generations are produced
but also that the vigor of these strains is preserved. Because
decreased fertility and viability are often associated with
inbreeding depression (Mrakovcic and Haley 1978, 1979),
sustaining line vigor may require maintaining genetic heterogeneity of these strains or generating genetically diverse
hybrid lines. Genetically diverse lines are often highly polymorphic, which may interfere with reproducibility of scientific experiments. To get a better insight into variability
associated with polymorphisms, complete genomic sequence information was obtained for homozygous wild-type
AB and Tübingen (TU1) individuals and a hybrid Sanger AB
Tübingen (SAT1) line was generated by crossing them.
In this article, we review some aspects of D. rerio breeding in the laboratory environment. In particular, we describe
zebrafish sexual dimorphism and the effects of olfactory
cues, visual stimuli, and social interactions on D. rerio reproduction. We also focus on factors influencing zebrafish
spawning. Then we describe breeding strategies commonly
used in the laboratory environment and discuss the need and
that appear ≥3x throughout this article: SAT, Sanger AB
Tübingen; SNP, single nucleotide polymorphism; TU, Tübingen
1Abbreviations
161
consequences of maintaining genetically diverse colonies.
This information can be used to maximize breeding efficiency in laboratory settings and, ultimately, make scientific
research more productive and successful. It is important to
emphasize that it may not always be possible to incorporate all
of the factors affecting zebrafish breeding into breeding protocols performed in the laboratory. Deciding which factors
should be taken into account will depend, for example, on
whether the line needs to be outcrossed or incrossed, whether
single-pair matings or group crosses have to be set up, and
whether a large or limited number of fish are available. Thus,
each breeding event should be individually evaluated with regard to how embryo production can be optimized. Detailed
step-by-step protocols that describe zebrafish breeding are not
provided in this article; such protocols can be found elsewhere (Harper and Lawrence 2011; Nüsslein-Volhard and
Dahm 2002; Westerfield 1993).
Sexual Dimorphism in Zebrafish
The mechanism of sex determination in D. rerio is unknown.
Zebrafish have no apparent heteromorphic chromosomes,
which indicates that no clear sex-determining chromosome
exists (Amores and Postlethwait 1999; Pijnacker and Ferwerda
1995; Schreeb et al. 1993; Traut and Winking 2001; Wallace
and Wallace 2003). It has been observed that sex ratios in
propagated colonies are often variable, depending on rearing
density, hypoxia, or food availability (Lawrence et al. 2008;
Shang et al. 2006). This demonstrates that, in addition to genetic factors, sex determination in zebrafish is also influenced by environmental inputs. Indeed, environmental changes
that involve hormones (Hill and Janz 2003; Westerfield
1993) and temperature (Uchida et al. 2004) have been shown
to affect sex differentiation.
Zebrafish adults lack strong sexual dimorphism. Individuals of this species can be sexed by body shape and color.
Females have a larger, light silver belly that protrudes from
the body in the anterior region. Males typically lack a protruding belly and are therefore more streamlined in shape (a
torpedo-like shape). Males also display a gold-reddish hue
between blue stripes, particularly in the anal and caudal fins.
Female’s blue stripes alternate with silver stripes, and their
dorsal fin shows a stronger yellow hue.
Zebrafish secondary sex characteristics are driven by the
sex of the gonad. Interestingly, gonad development for both
females and males is preceeded by a juvenile ovary stage
(Maack and Segner 2003; Takahashi 1977). This juvenile
ovary develops into a mature ovary in females but degenerates in males by means of apoptosis (Rodriguez-Mari et al.
2010; Uchida et al. 2004). The gonad in males is restructured
to form a testis. This transformation process is usually completed by the 3rd month of D. rerio development (Devlin and
Nagahama 2002; Maack and Segner 2003). The precise timing of stages of gonad development varies, depending on
husbandry conditions or type of analyzed strain (Maack and
Segner 2003; Takahashi 1977).
162
The Role of Olfactory Cues in D. rerio
Reproduction
Zebrafish are oviparous organisms with external fertilization
and no parent care. They are asynchronous spawners (Breder
et al. 1966), scattering mature, nonadhesive eggs in batches.
Fecundity is typically high, and a single female may release
clutches of several hundred eggs in a single spawning session. The presence of males stimulates ovulation in females
and oviposition (Hisaoka and Firlit 1962; van den Hurk and
Resink 1992).
Olfactory cues have been shown to play a role in mediating zebrafish reproduction (van den Hurk and Lambert 1983;
van den Hurk and Resink 1992). One of their functions is to
synchronize mating behavior. Females require several hours
to come into spawning condition, and they typically ovulate
overnight. It has been demonstrated that male gonad pheromones stimulate ovulation in females (van den Hurk and
Resink 1992). After ovulation, females release gonad pheromones that consist of a mixture of steroid glucuronides produced in the ovaries to attract males, and they stimulate them
to perform courtship (van den Hurk and Lambert 1983).
Females exposed to male odors have been shown to
produce more eggs than nonexposed females (Gerlach
2006). The eggs of exposed females were also of higher
quality. Some pheromones released by females, in turn,
have been reported to inhibit spawning of other females,
therefore playing a crucial role in competitive interactions
between females (Gerlach 2006).
Olfactory cues have also been shown to mediate kin recognition, with adult females preferring odor cues of unrelated males to unfamiliar relatives (Gerlach and Lysiak
2006). The preference of adult females for nonkin males has
most likely developed to prevent mating with close relatives
and thus avoid inbreeding, which can result in reduced
fertility and decreased survival (Mrakovcic and Haley 1978,
1979). Interestingly, adult males appear to remain neutral
when allowed to choose between kin females and nonkin
females (Gerlach and Lysiak 2006). Also, juvenile zebrafish differ significantly from adult females in that they
show a strong preference for the olfactory cues of their relatives (Gerlach and Lysiak 2006).
Mate Choice
Although olfactory cues seem to play a role in induction of
breeding condition and kin recognition, additional factors
such as visual stimuli and behavior cues could participate in
selection of an individual mating partner.
Evidence has been presented that zebrafish females prefer (Pyron 2003; Skinner 2004) and strategically allocate
their eggs differentially toward larger males (Skinner and
Watt 2007). Pritchard (2001) has suggested that phenotypic
traits such as male carotenoid coloration, longitudinal melanophore stripes, and symmetry of caudal fin pattern may influence mate choice (Pritchard 2001). The importance of visual
ILAR Journal
determinants has also been investigated by Engeszer and
colleagues (2004), who analyzed aggregation preferences of
wild-type and color-mutant zebrafish. This study demonstrated that zebrafish are able to distinguish visually between
alternative pigment pattern phenotypes and that the preferences for specific phenotypes are based on experience acquired during early development.
The effect of visual stimuli and morphology on mate
choice can be potentially confounded by social interactions.
These interactions in D. rerio result in the establishment of
dominance hierarchies in which dominant individuals
generally behave aggressively toward subordinate individuals (Pritchard 2001). Mating behavior within zebrafish social
hierarchies seems to be affected by both intrasexual competition and female mate preferences. Paull and colleagues
(2010) have observed that, although social rank within dominance hierarchies is established in a sex-independent manner, dominant females appear less aggressive in the morning
whereas aggression of dominant males decreases in the
afternoon. Given that spawning in D. rerio typically takes
place at dawn, this temporal difference in behavior may suggest that the function of dominance interactions in males is
to control access to spawning with females.
Dominant and subordinate individuals differ in gonadspecific expression of some genes (Filby et al. 2010). Moreover, ovaries in dominant females are larger relative to the
body mass in comparison with ovaries in subordinate females (Filby et al. 2010). There are also gonad-specific differences between dominant and subordinate males. In
particular, evidence has been presented that dominant males
have a higher number of spermatids in their testes compared
with subordinate males (Filby et al. 2010).
Dominant males have been reported to have greater reproductive success than subordinate males (Paull et al. 2010;
Spence et al. 2006). It is important to emphasize that reproduction is not exclusive to dominant males and subordinate
individuals also contribute to fertilization, although the degree of this contribution is significantly lower (Paull et al.
2010; Spence et al. 2006). Interestingly, dominant females
do not seem to have greater overall reproduction success
compared with subordinate females (Paull et al. 2010). However, dominant females have been found to monopolize
spawnings with dominant males (Paull et al. 2010). This
preference has been observed in the presence of male-male
competition. When female mate preferences have been
tested in single-pair matings in which females are allowed
access to only one male during the spawning session, no
correlation between female breeding success and male social rank has been found (Spence and Smith 2006). Thus,
zebrafish mate choice may be strictly determined by malemale competition, with dominant males excluding other
males rather than females actively choosing mates. It is also
possible that females’ preference for dominant males only
manifests itself when these males are presented simultaneously (Paull et al. 2010) or in sequence (Skinner and Watt
2007) with subordinate individuals during the same daily
mating period.
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2012
Visual stimuli and social interactions will most likely
have a more pronounced effect on group crosses than singlepair matings. These interactions may limit the number of fish
contributing to the progeny. Thus, if the goal is to maintain
genetic diversity of the line, it may be more beneficial to set
up more small-group crosses rather than fewer big crosses.
Also, most of the experiments that have investigated effects
of social interactions on zebrafish reproductive success were
carried out on fish obtained from commercial breeders.
These fish might have displayed more robust social behavior
than stocks propagated for many generations in the laboratory where the selection for specific behavior traits was not
applied. It therefore would be interesting to investigate
whether laboratory stocks have retained most of their social
behavior or whether the majority of these traits have been
permanently lost.
Zebrafish Courtship and Oviposition
Courtship behavior in the zebrafish male is triggered by female pheromones (van den Hurk and Lambert 1983; van den
Hurk et al. 1987). Initially, zebrafish courtship behavior was
described as consisting of abrupt turns and an elliptical pattern made by the male around the female (Guthrie and Muntz
1993). This description has been further broken down into
three general phases: initiatory, receptive/appetitive, and
spawning (Darrow and Harris 2004).
At the initiatory stage, males follow or swim alongside
females, which, at that time, typically perform abrupt
swimming movements. The receptive/appetitive stage is
characterized by males touching the female’s side or tail with
their noses or heads, circling around or in front of the female,
or tail sweeping and circling along the female’s body. Females at this stage swim alongside males or remain still while
being courted. Finally, at the spawning stage, a male and a
female swim side by side in the same direction close to each
other so that their genital pores can be aligned. The male then
performs rapid tail oscillations against the female’s side, triggering oviposition in the female and simultaneously releasing
sperm.
Females generally do not scatter all their eggs in a single
spawning (Skinner and Watt 2007). Typically, 5 to 20 eggs
are released at a time. Occasionally, a female couples together in one spawning session with different males.
Factors Affecting Spawning
A number of factors affect egg production and spawning. They
include the age and size of fish, interval at which fish are used
for egg production, light cycle, diet, and fish health status.
Laboratory zebrafish typically attain sexual maturity at
the 3rd month of their development. Initial spawns can be
observed in fish that are 2.5 months old. However, these
early spawns may produce eggs that are of lower quality and
quantity. The use of fish younger than 2.5 months for reproduction may be possible through in vitro fertilization.
163
Once sexual maturity is reached, prime reproductive performance is maintained for several months, and then it decreases with age. The gametes in older fish start deteriorating,
and animals that are, for example, older than 1.5 years spawn
fewer eggs that are also often of inferior quality (C. Carmichael,
Zebrafish International Resource Center (ZIRC), University
of Oregon, personal communication, 2012). It has also been
observed that the average sperm cell count peaks when males
are 10 months old (C. Carmichael and J. Matthews, ZIRC,
personal communication, 2012). These observations suggest
that the optimal zebrafish reproduction through natural mating
occurs when the fish are aged 6 months to 1 year.
Reproductive maturity and spawning efficiency also depend on the size of the fish (Eaton and Farley 1974a). Fish
reared under circumstances that result in lower growth rates
may require more than 3 months to reach sexual maturity. It has
been shown that large females spawn more frequently and produce larger clutch sizes than small females (Paull et al. 2008;
Spence and Smith 2006). Interestingly, small females have been
reported to generate larger eggs in terms of egg diameter (UusiHeikkila et al. 2010). However, eggs produced by small fish
show higher mortality rates and are of lower quality than the
eggs derived from large individuals (Uusi-Heikkila et al. 2010).
This suggests that size-dependent maternal effects contribute to
viability through egg matter composition rather than egg size.
Thus, egg size may not be a good indicator of quality.
Sexually mature zebrafish can spawn in the laboratory
continuously all year at a frequency of two or three times a
week (Eaton and Farley 1974b). When optimal conditions
are provided, females can even spawn daily for a limited period of time (Spence and Smith 2005). Overspawning, however, decreases the quantity and quality of eggs, and, if this
is the case, fish should be given at least 1 week rest to restore
resources before they are used in the next spawning event. It
has been reported that an optimal breeding frequency for
zebrafish is every 10 days (Niimi and LaHam 1974). If fish
are not spawned often, eggs are reabsorbed in the female
fish. Females not exposed to males for a prolonged period of
time often develop a plug, consisting of necrotic clumped
eggs, which clogs the oviduct and prevents any further oviposition (Spence et al. 2008). Such females are usually referred to as “eggbound.” To maintain a good quality of eggs,
females should be housed with males and spawned at least
once a month (Niimi and LaHam 1974).
Zebrafish reproduction depends strongly on photoperiod.
Mating is initiated at the onset of light, and spawning typically takes place over a short period thereafter (Breder et al.
1966; Spence et al. 2007; Westerfield 1993). Zebrafish in the
laboratory can occasionally breed throughout the day. If
a reliable egg production is required later during the day,
several approaches can be used to circumvent this limitation. For example, isolation cabinets with shifted light:dark
cycles could be used. Also, males and females could be
separated in crossing cages by dividers that are removed
shortly before the eggs are required. Finally, eggs can be obtained by squeezing females, and the eggs can then be fertilized in vitro.
164
Given that zebrafish are continuous spawners, maintenance of a good breeding colony requires constant replacement of nutrients used in reproduction. For this purpose, the
fish should be provided with a balanced diet consisting of a
variety of foods (Westerfield 1993). Markovich and colleagues (2007) have evaluated the effect of different diets on
spawning and embryo survival. The results of this study have
demonstrated that adult fish fed three times a day with
higher-fat diets, such as Artemia, laid significantly more
eggs than those fed a flake diet. Interestingly, however, the fish
fed a flake diet produced a higher percentage of viable
progeny than the fish fed higher-fat diets (Markovich et al.
2007). It has also been reported that fertilization rates are
affected by not only the percentage but also the types of fatty
acids in the diet (Meinelt et al. 1999).
Spawning and reproductive success are greatly influenced
by the stress (Ramsay et al. 2009a) and health status of the fish
(Matthews 2004; Ramsay et al. 2009b). Unhealthy fish are less
likely to produce viable offspring. Thus, any fish with signs of
disease should not be bred but removed from the system.
Setting Up Mating Crosses
In the laboratory environment, crosses are set up in static
water in plastic containers often referred to as crossing
cages. Originally, breeding in these cages was performed
over marbles whose main function was to prevent the fish in
the tank from consuming freshly released eggs because the
eggs would fall into spaces between marbles that are inaccessible to fish. Zebrafish females have been described as
choosy with respect to sites for oviposition (Spence et al.
2008), preferring a gravel substrate to silt (Spence et al.
2007). Because marbles mimic gravel, they could, in addition to protecting the eggs, contribute to a higher number of
ovipositions.
Most laboratories have stopped using marbles. Instead,
crossing cages have been designed with an additional plastic
container that holds the fish and is inserted inside the cage.
The bottom of this container is perforated, which allows
freshly released eggs to fall down into the outer cage and be
protected from predation. Imitation plastic plants or green
mesh are often placed in the inner container to provide artificial spawning sites as well as places of refuge to decrease
the effects of aggression related to antagonistic social behavior. Compared with marbles, this breeding setup is more
convenient for egg collection.
Crossing cages are available in different sizes and designs
from a number of manufacturers (e.g., Aquaneering, Aquatic
Habitats, Thoren Aquatics, ZebTEC). Goolish and colleagues
(1998) have evaluated chamber volume requirements for
zebrafish spawning. In their study, six adult zebrafish were
tested in volumes ranging from 100 ml to 500 ml. The results
demonstrated that egg production is decreased in breeding
volumes of 200 ml or less (Goolish et al. 1998). Thus,
chamber volume needs to be considered an important factor
affecting breeding.
ILAR Journal
Mating crosses are optimally set up in the afternoon or
early evening and left undisturbed until the following morning. If collected embryos need to be precisely synchronized
at the same development stage, a male and a female in the
crossing cage can be separated with a plastic divider that is
removed in the morning before spawning. Sometimes, if no
eggs are present on the first morning, the fish can be left in
the crossing cage for one more day until the eggs are laid.
Spawning can be frequently initiated by placing mating fish
in fresh and shallow water.
Single-Pair Matings and Group Crosses
Although zebrafish are group spawners (Spence et al. 2008),
both single-pair matings and group crosses are used in the
laboratory (Westerfield 1993). One application for singlepair breeding is to identify carriers of recessive mutations.
An incross between two heterozygous fish gives rise to progeny in which 25% of individuals show mutant phenotype.
Pair-wise outcrosses are also common. They are used to identify carriers of dominant traits that are visible only during early
development, and thereby screening of the progeny is required for carrier identification. Examples of fish analyzed
this way are those carrying a transgene whose product is expressed and visually detected during early embryogenesis. A
single-pair mating could also be used for embryo production
for experiments in which genetically homogenous individuals are needed.
Populations derived from group crosses are usually more
genetically diverse than those obtained from a single-pair
mating because, in such crosses, more than two individuals
contribute to the offspring. Maintaining genetically diverse
colonies and therefore avoiding inbreeding is crucial for fish
line propagation (Mrakovcic and Haley 1978, 1979). For this
reason, group crosses are preferably used to establish next
generations. To make sure that multiple individuals indeed
contribute to the offspring, a new generation of fish is established from embryos derived from a number of small group
crosses (containing, for example, two females and three
males) rather than from one large group cross.
Population density and sex ratio in group crosses have a
significant impact on mating behavior and reproductive success (Paull et al. 2008; Pritchard 2001; Spence and Smith
2005). Some male zebrafish are territorial during mating, restricting their activity to a few body lengths of a spawning
site (Spence and Smith 2005). Although both territorial and
nonterritorial males show the same courtship behavior, territorial males pursue females only within a specific territory,
chasing other males away (Spence et al. 2008).
A study analyzing the effects of density and sex ratio on
the behavior of territorial males showed that aggression rates
increased with density (Spence and Smith 2005). Courtship
behavior also increased in more dense populations, but only
in those with a female-biased sex ratio. In populations with a
male-biased sex ratio, courtship behavior decreased with
density.
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2012
The reproductive success of territorial males has also been
shown to be density dependent (Spence et al. 2006). When
adult fish were combined at low densities, territorial males
sired significantly more offspring than nonterritorial males.
In contrast, when fish were combined at high densities, territorial males were no more successful than nonterritorial
males. Based on these observations, it has been proposed
that zebrafish males show two distinct mating strategies—
territorial defense and active pursuit of females (Spence
et al. 2008). Population density seems to have an effect on
which of these two strategies is selected.
The density of the population also affects female reproductive success. It has been reported that, at higher densities,
average egg production per capita decreases (Spence and
Smith 2005). This decrease is apparently because, at higher
densities, rather than some females being excluded from
spawning, females spawn smaller clutches (Spence and
Smith 2006). Spence and Smith (2006) have proposed that
this effect most likely results from increased aggression at
high density by territorial males toward rivals, which interferes negatively with ovipositions by females.
Propagation of Wild-Type-Derived Lines
A variety of wild-type zebrafish lines is available (Johnson and
Zon 1999; Trevarrow and Robison 2004; Westerfield 1993), including the most commonly used laboratory strains: TU, AB,
WIK, and Tupfel long fin. These and other wild-type lines can
be obtained from the Zebrafish International Resource Center
(ZIRC). Because line propagation by incrossing can lead to
inbreeding depression that results in decreased fertility
and viability (Mrakovcic and Haley 1978, 1979), most lines
are preserved as heterogeneous, either by maintaining large
populations propagated through small group matings or by
round robin matings2 and removal of early lethal carriers from
the pool (Trevarrow and Robison 2004). To increase line heterogeneity and vigor, hybrid strains have been generated between
polymorphic wild-type lines (e.g., the Hopkins laboratory made
a hybrid line between TU and AB designated TAB [available
from ZIRC]).
Because maintenance of wild-type line heterogeneity
could require propagation of large populations through
elaborate breeding techniques, such protocols may not be
feasible for some zebrafish laboratories. These laboratories
may still use genetically diverse lines by ordering embryos
and/or adult fish from ZIRC, where all of the wild-type
lines are propagated with a goal to maintain line
heterogeneity.
The use of outbred lines with a high level of heterogeneity
can be advantageous because these lines may be viewed
as more representative of the species and could, therefore,
give broader validity for experiments than a single inbred
2A
detailed description of the round robin mating technique can be found
on the Zebrafish Information Network website dedicated to the AB
wild-type line: http://zfin.org/action/genotype/genotype-detail?zdbID=
ZDB-GENO-960809-7 (accessed November 5, 2012).
165
line (Festing 1995). Still, outbred lines characterized by high
genetic variability may affect reproducibility of experiments
and pose a significant challenge for research in which sequencespecific reagents such as primers, morpholinos, or even zinc
finger nucleases are used. Before these reagents are designed, it
is strongly recommended that researchers examine how polymorphic the region of interest is. This may even involve
resequencing the region. Because of a high number of
polymorphisms, the zebrafish reference genomic sequence,
prepared based on the TU line, may not always serve as a reliable source of sequence for other wild-type lines or even other
TU sublines. Indeed, comparison of different TU sequence
reads generated as part of the Zebrafish Sequencing Project at
the Wellcome Trust Sanger Institute revealed 645,088 singlenucleotide polymorphisms (SNPs1) (Bradley et al. 2007).
Inbreeding
High sequence variability of a line can be decreased by inbreeding. The Committee on Standardized Genetic Nomenclature for Mice specified in 1952 that 20 generations of
full-sibling (brother × sister) mating were necessary for a
line to be referred to as inbred (Nomenclature Committee
1952). It was amended later that other breeding schemes
were also acceptable provided that the inbreeding was equivalent to 20 successive generations of sibling mating (i.e., a
coefficient of inbreeding [F] of 98.6%) (Doolittle 1983).
Because zebrafish do not appear to have sex chromosomes
(Wallace and Wallace 2003) and sex ratios in colonies vary
(Lawrence et al. 2008) (possibly under genetic influence;
Bradley et al. 2011) and because inbreeding significantly
reduces fertility and survival (Mrakovcic and Haley 1978,
1979), generation of zebrafish isogenic lines seems to be a
long and challenging project. Currently, there are no zebrafish
inbred lines in which all individuals are identical and homozygous, as has been described for mouse isogenic lines. Some
near-homozygous zebrafish lines, such as C32 and SJD, have
been made using a combination of inbreeding and heat shock
(to generate homozygous gynogenetic diploids) and early
pressure (to generate heterozygous gynogenetic diploids), but
these lines were difficult to maintain and eventually had to be
outcrossed to increase viability (Johnson and Zon 1999).
It may yet be possible to generate true isogenic lines using conventional sibling matings with a large starting population, especially if this population has already been partially
inbred and cleared of some recessive lethal mutations. However, if gynogenetic techniques are not employed, it will take
a long time to complete this process. A better understanding
of the genetic and environmental factors affecting sex determination would aid with inbreeding strategies.
A Sequenced Hybrid Line
The SAT derives from two wild-type lines, AB and TU. A
male doubled haploid AB and a female doubled haploid TU
fish were generated in John Postlethwait’s laboratory using
166
heat shock. Both fish were sequenced using Illumina GA sequencing technology to approximately ×40 coverage (M.D.
Clark, unpublished results).
The sequenced fish have been outcrossed to each other
and the resultant F1 individuals incrossed to generate
F2 progeny for mapping using a custom SNP array of
more than 200,000 SNPs. Adult F2 pairs were also
shipped to ZIRC, where they are available as F3 generation
onwards.
The AB and TU G0 sequences have been assembled into
de novo contigs and can be searched for polymorphisms at
the Sanger Institute’s Zebrafish BLAST server.3 Short
sequences, such as 25–base pair morpholino sequences, will
not produce BLAST hits with default parameters. Longer
sequences (e.g., 50 base pairs around the start codon) will,
however, generate BLAST hits.
The SAT line should be largely free of embryonic lethal
mutations because these should have been selected against
during the doubled haploid process. Note that some of these
mutations might have been complemented by wild-type maternal messenger RNA or protein. Similarly, trans-heterozygous
lethal alleles should not prevail in the F1 individuals.
The genome of the F2 progeny and subsequent generations
can be reconstructed by dense genotyping and imputation of the
intervening genotypes (Ellinghaus et al. 2009; Li et al. 2011)
using, for example, a 200k SNP array, calculating the crossovers, and identifying the pieces of sequenced AB and TU
genomes present in a given fish. Once the genomes are reconstructed with high-density genotyping, low-density genotyping
(e.g., 96 or 384 well-spaced SNPs) can be used for several
subsequent generations. Thereafter, when a large proportion
of the sequence becomes ambiguous because of novel recombination sites, high-density genotyping will need to be reemployed to construct the genome sequence anew.
The SAT line is thus genetically defined, and individual
genomes can be reconstructed. It can be maintained and
propagated as other wild-type strains (i.e., as a large population with group or round robin matings). Importantly, the
line is largely free of lethal and deleterious alleles and should
possess hybrid vigor, enabling it to be used as a wild-type
line and maintained as a strain. It is highly recommended
that this line be used for experiments for which knowledge
of precise genome sequence is necessary. Examples include
morpholino and polymerase chain reaction primer design as
well as reverse genetics techniques such as targeting induced
local lesions in genomes and zinc finger nuclease targeting.
Conclusion
Breeding is an important part of zebrafish husbandry. Its understanding is crucial for optimizing scientific research in
which D. rerio is used as a model organism. Mate choice,
courtship, and spawning are affected by a number of factors.
3Available
from www.sanger.ac.uk/cgi-bin/blast/submitblast/d_rerio (accessed
on September 26, 2012).
ILAR Journal
It would be important to investigate their interdependence
and describe conditions and circumstances in which individual factors become primary determinants affecting breeding. Also, a better insight into zebrafish reproduction and
reproductive behavior will come from more comprehensive
understanding of molecular mechanisms underlying these
processes.
One of the major concerns related to zebrafish breeding
and line propagation in the laboratory is preservation of the
vigor of a line. To achieve this, current breeding strategies
avoid inbreeding and focus on maintaining genetic diversity
of the line. This, however, often results in a high degree of
polymorphic variations, which affect effectiveness of the research and its reproducibility. Thus, attempts should be
made to develop breeding schemes that would ultimately
give rise to relatively robust inbred lines characterized by a
high degree of genetic homogeneity.
Successful inbreeding strategies could be applied to lines
such as SAT to produce recombinant inbred lines. Such
lines are powerful tools for mapping traits and are created by
conventional intercross matings to generate the recombination
breakpoints, followed by incrossing to homozygosity (Peirce
et al. 2004). In zebrafish, heat shock and/or early pressure
gynogenetic diploids could be used to shorten time to achieve
homozygosity. This is a significant project and, if recombinant inbred lines were indeed established in zebrafish, they
could be used for association mapping, as has been done in
maize (McMullen et al. 2009).
Acknowledgments
A. Nasiadka thanks Katrina Murray, Jen Matthews, April
Freeman, Carrie Carmichael, and Dagmara Marston for reading this manuscript and offering valuable insights and advice.
M. Clark thanks Elisabeth Busch-Nentwich and Derek Stemple for discussions and advice. The Zebrafish International
Resource Center is supported by the National Institutes of
Health, National Center for Research Resources (grant P40
RR012546). The D. rerio Sequencing Project at the Wellcome
Trust Sanger Institute is funded by a Wellcome Trust grant.
References
Amores A, Postlethwait JH. 1999. Banded chromosomes and the zebrafish
karyotype. Methods Cell Biol 60:323-338.
Bradley KM, Breyer JP, Melville DB, Broman KW, Knapik EW, Smith JR.
2011. An SNP-based linkage map for zebrafish reveals sex determination loci. G3 (Bethesda) 1:3-9.
Bradley KM, Elmore JB, Breyer JP, Yaspan BL, Jessen JR, Knapik EW,
Smith JR. 2007. A major zebrafish polymorphism resource for genetic
mapping. Genome Biol 8(4):R55.
Breder CM, Rosen DE, American Museum of Natural History. 1966. Modes
of reproduction in fishes. Garden City (NY): Natural History Press.
Darrow KO, Harris WA. 2004. Characterization and development of courtship in zebrafish, Danio rerio. Zebrafish 1(1):40-45.
Devlin RH, Nagahama Y. 2002. Sex determination and sex differentiation
in fish: An overview of genetic, physiological, and environmental influences. Aquaculture 208:191-364.
Volume 53, Number 2
2012
Doolittle DP. 1983. Genetics and probability in animal breeding experiments Green El. Bioscience 33:285.
Eaton RC, Farley RD. 1974a. Growth and reduction of depensation of the
zebrafish, Brachydanio rerio, reared in the laboratory. Copeia 1:204209.
Eaton RC, Farley RD. 1974b. Spawning cycle and egg production in zebrafish,
Brachydanio rerio, reared in the laboratory. Copeia 1:195-204.
Ellinghaus D, Schreiber S, Franke A, Nothnagel M. 2009. Current software
for genotype imputation. Hum Genomics 3:371-380.
Engeszer RE, Ryan MJ, Parichy DM. 2004. Learned social preference in
zebrafish. Curr Biol 14:881-884.
Festing MF. 1995. Use of a multistrain assay could improve the NTP carcinogenesis bioassay. Environ Health Perspect 103:44-52.
Filby AL, Paull GC, Bartlett EJ, Van Look KJ, Tyler CR. 2010. Physiological and health consequences of social status in zebrafish (Danio rerio).
Physiol Behav 101:576-587.
Gerlach G. 2006. Pheromonal regulation of reproductive success in female
zebrafish: Female suppression and male enhancement. Anim Behav
72:1119-1124.
Gerlach G, Lysiak N. 2006. Kin recognition and inbreeding avoidance in
zebrafish, Danio rerio, is based on phenotype matching. Anim Behav
71:1371-1377.
Goolish EM, Evans R, Okutake K, Max R. 1998. Chamber volume requirements for reproduction of the zebrafish Danio rerio. Prog Fish Cult
60:127-132.
Grunwald DJ, Eisen JS. 2002. Headwaters of the zebrafish—Emergence of
a new model vertebrate. Nat Rev Genet 3(9):717-724.
Guthrie DM, Muntz WRA. 1993. Role of vision in fish behavior. In: Pitcher TJ,
ed. Behavior of Teleost Fishes. Baltimore: Chapman and Hall. p 87-128.
Harper C, Lawrence C. 2011. The Laboratory Zebrafish. Boca Raton FL:
CRC Press.
Hill RL, Janz DM. 2003. Developmental estrogenic exposure in zebrafish
(Danio rerio): I. Effects on sex ratio and breeding success. Aquat Toxicol 63(4):417-429.
Hisaoka KK, Firlit CF. 1962. Ovarian cycle and egg production in the zebrafish,
Brachydanio rerio. Copeia 1962:788-792.
Johnson SL, Zon LI. 1999. Genetic backgrounds and some standard stocks
and strains used in zebrafish developmental biology and genetics. Methods Cell Biol 60:357-359.
Lawrence C, Ebersole JP, Kesseli RV. 2008. Rapid growth and out-crossing
promote female development in zebrafish (Danio rerio). Environ Biol
Fish 81:239-246.
Li Y, Sidore C, Kang HM, Boehnke M, Abecasis G. 2011. Low coverage
sequencing: Implications for the design of complex trait association
studies. Genome Res 21:940-951.
Maack G, Segner H. 2003. Morphological development of the gonads in
zebrafish. J Fish Biol 62:895-906.
Markovich ML, Rizzuto NV, Brown PB. 2007. Diet affects spawning in
zebrafish. Zebrafish 4:69-74.
Matthews JL. 2004. Common diseases of laboratory zebrafish. Methods
Cell Biol 77:617-643.
McMullen MD, Kresovich S, Villeda HS, Bradbury P, Li H, Sun Q, FlintGarcia S, Thornsberry J, Acharya C, Bottoms C, Brown P, Browne C,
Eller M, Guill K, Harjes C, Kroon D, Lepak N, Mitchell SE, Peterson B,
Pressoir G, Romero S, Oropeza Rosas M, Salvo S, Yates H, Hanson M,
Jones E, Smith S, Glaubitz JC, Goodman M, Ware D, Holland JB,
Buckler ES. 2009. Genetic properties of the maize nested association
mapping population. Science 325(5941):737-740.
Meinelt T, Schulz C, Wirth M, Kurzinger H, Steinberg C. 1999. Dietary
fatty acid composition influences the fertilization rate of zebrafish
(Danio rerio Hamilton-Buchanan). J Appl Ichthyol 15:19-23.
Mrakovcic M, Haley LE. 1978. Genetic effect of inbreeding in warm water
fish Brachidanio rerio. Can J Genet Cytol 20:450-450.
Mrakovcic M, Haley LE. 1979. Inbreeding depression in the zebra fish
Brachydanio rerio (Hamilton Buchanan). J Fish Biol 15:323-327.
Niimi AJ, LaHam QN. 1974. Influence of breeding time interval on egg
number, mortality, and hatching of zebra fish Brachydanio rerio. Can J
Zool 52:515-517.
167
Nomenclature Committee. 1952. Committee on Standardized Nomenclature for Inbred Strains of Mice. Cancer Res 12:602-613.
Nüsslein-Volhard C, Dahm R, eds. 2002. Zebrafish: A Practical Approach.
New York: Oxford University Press.
Paull GC, Van Look KJ, Santos EM, Filby AL, Gray DM, Nash JP, Tyler
CR. 2008. Variability in measures of reproductive success in laboratorykept colonies of zebrafish and implications for studies addressing
population-level effects of environmental chemicals. Aquat Toxicol 87:115126.
Paull GC, Filby AL, Giddins HG, Coe TS, Hamilton PB, Tyler CR. 2010.
Dominance hierarchies in zebrafish (Danio rerio) and their relationship
with reproductive success. Zebrafish 7:109-117.
Peirce JL, Lu L, Gu J, Silver LM, Williams RW. 2004. A new set of BXD
recombinant inbred lines from advanced intercross populations in mice.
BMC Genet 5:7.
Pijnacker LP, Ferwerda MA. 1995. Zebrafish chromosome banding. Genome 38:1052-1055.
Pritchard L. 2001. Behaviour and morphology of the zebrafish, Danio rerio.
PhD dissertation, University of Leeds.
Pyron M. 2003. Female preferences and male-male interactions in zebrafish
(Danio rerio). Can J Zool 81:122-125.
Ramsay JM, Watral V, Schreck CB, Kent ML. 2009a. Husbandry stress
exacerbates mycobacterial infections in adult zebrafish, Danio rerio
(Hamilton). J Fish Dis 32:931-941.
Ramsay JM, Watral V, Schreck CB, Kent ML. 2009b. Pseudoloma neurophilia infections in zebrafish Danio rerio: Effects of stress on survival,
growth, and reproduction. Dis Aquat Organ 88:69-84.
Rodriguez-Mari A, Canestro C, BreMiller RA, Nguyen-Johnson A,
Asakawa K, Kawakami K, Postlethwait JH. 2010. Sex reversal in
zebrafish fancl mutants is caused by Tp53-mediated germ cell apoptosis. PLoS Genet 6:1-14.
Schreeb KH, Groth G, Sachsse W, Freundt KJ. 1993. The karyotype of the
zebrafish (Brachydanio rerio). J Exp Anim Sci 36:27-31.
Shang EHH, Yu RMK, Wu RSS. 2006. Hypoxia affects sex differentiation
and development, leading to a male-dominated population in zebrafish
(Danio rerio). Environ Sci Technol 40:3118-3122.
Siegfried KR, Nüsslein-Volhard C. 2008. Germ line control of female sex
determination in zebrafish. Dev Biol 324:277-287.
Skinner AMJ. 2004. Sexual selection in the zebra fish (Danio rerio) and the
guppy (Poecilia reticulata) PhD dissertation, University of Sheffield.
Skinner AMJ, Watt PJ. 2007. Strategic egg allocation in the zebra fish, Danio rerio. Behav Ecol 18:905-909.
168
Spence R, Smith C. 2005. Male territoriality mediates density and sex ratio
effects on oviposition in the zebrafish, Danio rerio. Anim Behav 69:
1317-1323.
Spence R, Smith C. 2006. Mating preference of female zebrafish, Danio
rerio, in relation to male dominance. Behav Ecol 17:779-783.
Spence R, Jordan WC, Smith C. 2006. Genetic analysis of male reproductive
success in relation to density in the zebrafish, Danio rerio. Front Zool 3:5.
Spence R, Ashton R, Smith C. 2007. Oviposition decisions are mediated by
spawning site quality in wild and domesticated zebrafish, Danio rerio.
Behaviour 144:953-966.
Spence R, Gerlach G, Lawrence C, Smith C. 2008. The behaviour and ecology of the zebrafish, Danio rerio. Biol Rev Camb Philos Soc 83:13-34.
Takahashi H. 1977. Juvenile hermaphroditism in the zebrafish, Brachydanio rerio. Bulletin of Faculty of Fisheries of Hokkaido University 28:
57-65.
Traut W, Winking H. 2001. Meiotic chromosomes and stages of sex chromosome evolution in fish: Zebrafish, platyfish and guppy. Chromosome
Res 9:659-672.
Trevarrow B, Robison B. 2004. Genetic backgrounds, standard lines, and
husbandry of zebrafish. Methods Cell Biol 77:599-616.
Uchida D, Yamashita M, Kitano T, Iguchi T. 2004. An aromatase inhibitor
or high water temperature induce oocyte apoptosis and depletion of
P450 aromatase activity in the gonads of genetic female zebrafish
during sex-reversal. Comp Biochem Physiol A Mol Integr Physiol
137:11-20.
Uusi-Heikkila S, Wolter C, Meinelt T, Arlinghaus R. 2010. Size-dependent
reproductive success of wild zebrafish Danio rerio in the laboratory.
J Fish Biol 77(3):552-569.
van den Hurk R, Lambert JGD. 1983. Ovarian-steroid glucuronides function as sex-pheromones for male zebrafish, Brachydanio rerio. Can J
Zool 61:2381-2387.
van den Hurk R, Resink JW. 1992. Male reproductive-system as sexpheromone producer in teleost fish. J Exp Zool 261:204-213.
van den Hurk R, Schoonen WG, van Zoelen GA, Lambert JG. 1987. The
biosynthesis of steroid glucuronides in the testis of the zebrafish,
Brachydanio rerio, and their pheromonal function as ovulation inducers. Gen Comp Endocrinol 68:179-188.
Wallace BM, Wallace H. 2003. Synaptonemal complex karyotype of zebrafish.
Heredity 90:136-140.
Westerfield M. 1993. The Zebrafish Book: A Guide for the Laboratory
Use of Zebrafish (Brachydanio rerio). Eugene: University of
Oregon Press.
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