Aspects of Larval Rearing
Carole Wilson
Abstract
Fish—and zebrafish (Danio rerio) in particular—are now
the second-most used biomedical model in the United
Kingdom. The use of fish in research rose by 23% in 2011,
primarily reflecting a rise in the use of zebrafish. Despite the
increasing importance of zebrafish as a biomedical model
system, there are currently no legislative guidelines or requirements for larval husbandry in the United Kingdom,
the European Union, or the United States. This has led to
a variety of procedures and methods being developed
for larval rearing, many of which are not derived from
peer-reviewed protocols. This article reviews published
work relating to larval rearing and some unpublished
protocols to establish optimized and standardized husbandry procedures.
Key Words: Danio rerio; fry; husbandry; larval rearing;
zebrafish
Introduction
L
ong established as an aquarium fish, the zebrafish (Danio
rerio) has been used as a biomedical model since the
1930s (Laale 1975). The zebrafish was introduced as a
model system for developmental genetics in the late 1970s
by George Streisinger and his colleagues (Nüsslein-Volhard
and Dahm 2002; Westerfield 2007) and was subsequently
used for the first large-scale mutagenic screens in a vertebrate model (Granato and Nüsslein-Volhard 1996).
The popularity of zebrafish is a result of the many advantages in maintenance and manipulation of this organism. Adults
are small, tolerant of a wide range of environmental parameters, fecund, and inexpensive to keep compared with other
biomedical models, such as the mouse. Zebrafish breed to
give rise to large clutches of externally fertilized embryos,
often in excess of 200, which develop fast and are transparent.
With the technical advances of the last decade, there has
been an explosion in the use of the zebrafish as a vertebrate
model in a multitude of diverse disciplines, including live
Carole Wilson, Molecular Genetics (Hons), is Head of Fish Facilities,
Division of Biosciences, University College London, United Kingdom.
Address correspondence and reprint requests to Carole Wilson, Division
of Biosciences, University College London, Gower Street, London WC1E
6BT, UK or email [email protected].
Volume 53, Number 2
2012
imaging of developmental processes (Cyejic et al. 2008; van
Ham et al. 2010; Yaniv et al. 2006), drug and small molecule
screens (MacRae and Peterson 2003; Tamplin et al. 2012;
Taylor et al. 2010), cancer research (Amatruda et al. 2002;
Feitsma and Cuppen 2008; Liu and Leach 2011), behavioral
studies (Barth et al. 2005; Gerlai et al. 2000; Miklósi and
Andrew 2006), and toxicology (Hill et al. 2005; Spitsbergen
and Kent 2003).
Despite the widespread use of zebrafish, there are many
areas of their husbandry that need improvement (Lawrence
2007; Spence et al. 2007), one of which is larval rearing.
Here, protocols have been developed that might owe little to
knowledge of the true requirements of larval zebrafish and
more to the constraints and pressures of fish facilities and
laboratories. Maximal survival rates are vital for animal welfare and research and critical, for example, for the success of
widely used mutagenesis screens.
Many aspects of zebrafish early larval physiology, anatomy,
and genetics are well studied (Haffter et al. 1996), but this
knowledge has seldom been used systematically to improve
husbandry. The lack of published information and adaptation of
protocols that are available from both fish hobbyist and, more
recently, aquaculture literature, aimed at larger-scale production
of fish, is reflected in the large degree of variation in approaches
used. This article presents a formal review of some published
literature and an overview of some unpublished protocols and
methods of larval husbandry in Europe and the United States to
arrive at a more standardized approach.
Consideration of Adult Husbandry for
Successful Larval Production
Many factors influencing larval productivity and quality affect processes prior to spawning, and thus the condition of
the breeding stock and their breeding environment need to
be taken into consideration for successful larval rearing. A
facility setting will be different from a wild habitat, and
many strains of zebrafish will have undergone elements of
domestication, including larger body size and, in some
strains, lessened aggression (Moretz et al. 2007). It may be
beneficial to try to allow fish to display as much natural behavior as possible. Studies of zebrafish in their natural habitat
(Engeszer et al. 2007; McClure et al. 2006; Spence et al.
2006, 2008) can be used to understand their general environment. In a wild and/or natural habitat, zebrafish are generally
considered seasonal breeders; however, the finding of females
with mature ova outside of the season (Spence et al. 2006)
169
suggests that mating may be more dependent on available nutrition than initially assumed. Studies reporting zebrafish larval
growth in their natural habitats are scarce but suggest that adult
zebrafish live in streams and come to spawn in flooded, vegetative areas, where the larval forms grow protected from predators
before moving into the margins of slow-moving rivers and
streams as adults in the rainy season (Engeszer et al. 2007).
Zebrafish have specific courtship behaviors, including
the male chasing the female, swimming around her, and
swimming between her and the spawning site (Darrow and
Harris 2004; Pyron 2003; Spence and Smith 2005); social
preferences; and both male and female dominance hierarchies (Delaney et al. 2002; Engeszer et al. 2004; Paull et al.
2010). Unfortunately, the facility setting may not always be
conducive to allowing such natural behaviors. For example,
small tank size can affect natural courtship, and lack of substrates may have an effect on oviposition and therefore reproductive success because females may prefer to spawn
over gravel (Spence et al. 2007). Another factor that has an
effect on reproductive success and is more specific to facilities is inbreeding depression, which affects both fecundity
and fertility (Monson and Sadler 2010). Broodstock nutrition also affects spawning, as well as the quality of the eggs
and embryos produced. Lipid fatty acids and vitamins have
all been identified as critical factors in broodstock diet for
fecundity and fertility of the broodstock and the quality of
the embryonic yolk sac (Alsop et al. 2008; Izquierdo et al.
2001; Markovich et al. 2007; Watts et al. 2012, in this issue).
The genetic status of the broodstock should also be taken
into account because certain genetic lines will always show
a propensity to produce offspring that have either lower survival rates or less-robust offspring (C. Wilson, unpublished
observations). Fertility and fecundity drop with age, and, as
the laboratory zebrafish reach ages greater than 18 months,
there is a marked reduction in fertility and fecundity (C. Wilson,
personal observations) and increase in health problems. Thus,
even though fish in a research setting may survive in excess of
3 years, it is unlikely they will be of breeding quality for so
long (Nasiadka and Clark 2012, in this issue).
Factors Affecting Developmental Stages
Developmental stages of fish can be defined in several ways.
As an approximation, early stages of zebrafish are usually expressed in hours or days postfertilization at a standard temperature of 28.5ºC; lower or higher rearing temperatures will affect
this staging process (Harper and Lawrence 2010; Kimmel 1995;
Nüsslein-Volhard and Dahm 2002) by slowing down or accelerating, respectively, the speed of development. Morphological
landmarks, such as the number of somites or the migration of
the lateral line organ, can be used in preference to hours or days
postfertilization, similar to the staging used in aquaculture
(Koubbi et al. 1990; Nikolioudakis et al. 2010).
Temperature is one factor affecting development, but
there are many others, including rearing density, water quality
(Lawrence and Mason 2012, in this issue), broodstock, and,
170
at later stages of development, nutritional factors. Several studies have shown differences in larval-rearing success between
different diet regimens. Lawrence (2007) showed a varied mean
growth rate and varied survival rate across several artificial
diets he reviewed. A possible explanation for this variance is
that manufactured diets cater for several species of fish larvae
(Sales and Janssens 2003) and the nutrimental needs of different larval stages of zebrafish are not met by these foods. In trials, larval stages able to take Artemia appeared to have higher
survival rates than those fed on artificial diets (Carvalho et al.
2006; Hensley and Leung 2010), suggesting that Artemia, although not part of a natural zebrafish diet, has a close approximation of the nutritional needs of larval zebrafish.
The Relationship between Developmental
Stages and Larval-Rearing Protocols
Protocols
Successful larval rearing depends on matching the physiological needs of the larval fish to the rearing protocol
(Table 1; Figure 1). However, there are many factors governing the overall running of zebrafish facilities, not all of
which relate directly to meeting the biological needs of the
fish. Availability of space, financial resources, and technical
help have led to a wide variety of ways in which husbandry
is administered in facilities that range from small facilities comprising only a few tanks, a correspondingly low
number of fish, and little in the way of technical support, to
very large multirack facilities with dedicated technical staff.
The data collated below represent an overview of the
ways in which different establishments conduct larval rearing and were derived from both anecdotal evidence and
larval-rearing protocols from eight different facilities in the
United Kingdom, Europe, and the United States. Additional
data were taken from a larval-rearing survey conducted by the
Zebrafish Husbandry Association (ZHA1) in 2009 (I. Addatto,
ZHA, personal communication, 2011) to which there were 48
respondents. By building a picture of current husbandry
practice, striking differences emerged between facilities
in the way in which larval rearing is conducted (Table 2;
Appendix). This diversity may reflect the diverse needs
within the zebrafish community and adaptation of protocols
from both hobbyist and aquaculture sources. For example,
Paramecium is more commonly used by the hobbyist for
small-scale fish breeding. Rotifer and phytoplankton-rich
water techniques are often used in large-scale aquaculture
production. Various methods of zebrafish larval rearing and
protocols have been described and can be found in reference
books (Nüsslein-Volhard and Dahm 2002; Westerfield
2007), articles (Best et al. 2010), and web resources (such as
the protocols from the Parichy laboratory2).
that appear ≥3x throughout this article: DPF, days
postfertilization; ZHA, Zebrafish Husbandry Association
2 http://protist.biology.washington.edu/dparichy/ProtocolsPage.htm
(accessed October 4, 2012).
1Abbreviations
ILAR Journal
Table 1 Comparison of stages of larval development and generalized larval-rearing protocols
Time
Stage
0-72 HPF
Embryos
Morphology
Gut
Other
42 HPF: gut develops
into hollow tube
Development reliant on
yolk sac
Petri dish, hold at 28.5°C
4-5 DPF: swim bladder
inflates; first feeding;
gape size ≈100 µm
4-9 DPF: static/slow drip
water flow
48 HPF: esophagus,
liver, pancreas,
and pharynx join to gut
72 HPF-13 DPF
Early larvae
72 HPF: mouth
and anus open
5-6 DPF: digestive tract
opens, digestive
enzymes secreted
7 DPF: complete yolk
absorption
Body length:
5 DPF: 4 mm
10 DPF: 8 mm
(TL/AB F1) UCL
14-29 DPF
Midstage
larvae
Body length:
15 DPF: ≈13 mm
28 DPF: ≈24 mm
(TL/AB F1) UCL
15-20 DPF
Metamorphosis
30 days to 3 or
4 months
Juveniles
End of metamorphosis;
complete fin
complementation and
full adult pigmentation
2-4 months: sexual
maturity
Generalized rearing
protocol
>200 embryos per dish (90 ×
200 mm) is likely to cause
asynchronous growth
5-13 DPF: slow drip
Exogenous feeding begins 4-5
days before full yolk
absorption
Particle size must correspond
with gape size
Live foods: Paramecium, rotifer
with gradual transition to
Artemia at ≈9-10 DPF
Increase water flow rate
as larval body measurement and
mass increase
Feed Artemia or powder food
of equivalent nutritional
status to encourage growth
(continued use of
Paramecium at this stage
likely to slow growth)
Feed as adult, perhaps more
frequently; usual to continue
Artemia
Increase water flow rate
See Table 2 for more detailed larval-rearing protocols. DPF, days postfertilization; HPF, hours postfertilization; TL/AB F1, hybrid cross of zebrafish
and wild-type lines TL and AB; UCL, University College London
Embryonic Stage (Zero to 48 Hr)
Zebrafish are egg-layers, providing no parental aftercare.
The developing embryos are held within a chorion, a membrane that provides some protection, for the first 48 hours of
life. Hatching time from the chorion may vary slightly according to the strength of the chorion and the muscular
movement of the embryo inside (Laale 1975).
The embryonic intestine develops from an endodermal
rod into a hollow tube at about 42 hours postfertilization, and
at around 48 hours postfertilization, the esophagus, liver,
pancreas, and pharynx are joined to the gut. At around 72 hours
postfertilization, the mouth opens, followed by the anus
(Holmberg et al. 2004; Wallace et al. 2005). Posthatching,
the embryo develops an attachment gland, composed of
small secretory cells around the mouth and under the eyes,
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2012
that allows the early larval form to become attached to either
hard surfaces or plants and to reach the surface for air intake
by inflating the swim bladder (Laale 1975). Through these
stages, embryonic development is still reliant on the yolk for
nutrients (Jardine and Litvak 2003; Wallace et al. 2005).
Because the embryo is still reliant on the yolk sac at this
stage, most organizations begin larval rearing between 4 and
7 days postfertilization (DPF1), with 5 days being favored in the
United Kingdom. Prior to this stage, most larval zebrafish will
be in incubators held at approximately 28.5ºC, either in fish water taken directly or filtered from a main system or in embryo
media (Nüsslein-Volhard and Dahm 2002; Westerfield 2007).
Embryos are kept in standard petri dishes (90 × 20 mm) at
stocking densities between 16 and 50 but up to 100 fry per petri
dish. High stocking densities during embryogenesis (more than
200 in a standard Petri dish, 90 × 20 mm) can promote the
171
Figure 1 Different stages of larval development
growth of fungi and protozoa in the dish and can result in asynchronous growth within the clutches (Harper and Lawrence
2010; C. Wilson, personal observations).
Early- to Midlarval Stage (72 HPF to 13 DPF)
At 4 to 5 DPF, the swim bladder inflates, and the larval fish
begins swimming in the water column. At 5 to 6 DPF, the
digestive tract opens, and digestive enzymes are secreted,
suggesting the larval fish can begin exogenous feeding even
though the yolk sac is not yet completely depleted (Holmberg
et al. 2004). At this point, the intestinal tract is still very
underdeveloped, and the larva will not be able to absorb
many nutrients; by 7 DPF, the yolk is completely absorbed,
and it seems determinant for the survival of the larva that the
period from endogenous to complete exogenous feeding
overlap. Indeed, delayed exogenous feeding at this point results in low survival rates and starvation at around 10 DPF.
As the larvae start feeding, it is usual to move them from
the petri dishes into a larger volume of water. At this stage,
the larvae should only be in static or very slow-moving water
so they can maintain position in the water column. Substantial
water changes on static tanks should be avoided because
they are likely to cause stress and may lower both growth
and survival rates (C. Wilson, personal observation). At this
point, some aspects of water quality do not seem as important because larval zebrafish seem more tolerant of poorer
water quality with respect to free ammonia than adult fish
(Best et al. 2010), although temperature will still affect the
rate of larval development and thus staging.
Thirty-five of 46 respondents in the ZHA survey begin
exogenous feeding between 4 and 7 DPF, usually at 5 DPF,
suggesting that it is widely accepted that an exogenous food
source must be provided by 7 DPF, when the yolk reserve is
completely depleted (Jardine and Litvak 2003). At this stage,
most facilities provide a diet rich in live foods, popularly
Paramecium or rotifer, complemented with powdered food
suitable for an early, first feeding gape size of approximately
100 µm. Paramecium, sized 100 to 150 µm, and the smaller
rotifer species, upwards of 150 to 220 µm (Lawrence 2007;
C. Wilson, personal observation), thus make ideal first live
foods. In addition, Best and colleagues (2010) showed both
high survival rates and growth rates at these early larval
stages with a rotifer diet. The use of live foods also seems to
stimulate hunter/prey behavior that utilizes both visual and
olfactory cues (Borla et al. 2002; Ostrander 2000), which
may be useful in stimulating feeding behaviors in a way that
172
an exclusively dry diet might not. A small number of facilities
provide no live food at this stage, beginning live food with
Artemia at a later stage. This might reflect time constrictions.
Smaller facilities, especially those with no dedicated technical
support, may not have the time to culture live foods.
Midlarval Stage, Including Metamorphosis
(14 to 29 DPF)
Metamorphosis occurs between the larval and juvenile stages,
and in zebrafish it reflects the transition between larval form
and adult form; the end of metamorphosis is marked, among
other things, by a complete fin complement and full adult
pigmentation (Parichy 2003).
By 14 DPF, most facilities have larvae on a drip-through
or running water system. Stocking densities are, however,
very diverse: 29 of 42 respondents in the ZHA survey have
densities above 30 larvae per liter at first introduction, with
just under 30% introducing larvae to the system at 40 to 50
larvae per liter. None of the UK facilities consulted put larvae
onto a system at stocking densities greater than 20 per liter.
At midlarval stage, the feeding regimens become very
disparate; although Artemia is a well-known foodstuff for
both adult zebrafish and larval forms with corresponding
gape size, usually from 8 to 9 DPF (C. Wilson, personal observation), some facilities do not provide it until 14 to 15
DPF. A wide variety of powder food and flaked food is given,
which reflects both a difference in availability of foods in
different countries and also a dearth of food products specifically manufactured for the nutritional needs of zebrafish.
Unfortunately, the complete nutritional requirements of both
adult and larval zebrafish are currently unclear (Lawrence
2007).
Paramecium may not have a sufficient nutritional value
to push larval development to and through metamorphosis
(Lawrence 2007), and it is important to move the larvae onto
a food of higher nutritional status; otherwise zebrafish may
have smaller larval size and higher mortality rates (Maley
et al. 2008; C. Wilson, personal observation). For this reason, toward the end of the early larval stage (8-9 DPF) and
into the midlarval stage (10-15 DPF), larger particle size
food of a greater nutritional quality should be offered. Commonly Artemia is offered at this point, and although not
naturally occurring in the diet of zebrafish, it appears to have
most of the nutritional complement that is required for larval
stages, when gape size is large enough to take either whole
Artemia or smaller pieces of Artemia.
ILAR Journal
Juvenile and Adult Stage (29 DPF and Beyond)
When the fish enter the juvenile stage, they are usually
treated in a similar way to adult fish, with the exception of
additional feeding to promote growth and slower water flow
rates, reflecting their smaller size. Husbandry and welfare
issues of adult fish are discussed elsewhere in this issue.
Considerations of the Effects of LarvalRearing Protocols on Larvae Quality
Different larval-rearing protocols can have an important impact on larval size and mass, survival rates, and sex determination and ratio of larvae.
Feeding zebrafish Paramecium alone during the early larval stages can result in smaller larvae and lower survival rates
(Maley et al. 2008), suggesting that Paramecium alone may be
an insufficient nutritional source. Paramecium may not promote growth as well as Artemia, thus the average length was
7.3 to 7.6 mm at 21 days after Paramecium-only feeding compared with 14.3 mm at the same point after Artemia-only feeding (Carvalho et al. 2006). Even at 28 DPF, fry had grown to
only 9.9 mm on a Paramecium-only diet (Maley et al. 2008).
Survival rates are also affected by diet, but the relationship with other factors such as water quality and genetic factors must be considered, because different publications
describe different findings, ranging from 22.6% survival on
a live food diet—Paramecium and Artemia (Maley et al.
2008)—to 86% survival on Artemia (Carvalho et al. 2006).
Survival Rates
In the ZHA survey, 39 of 44 respondents reported survival
rates of 50% or higher, with 17 reporting survival rates of
75% or higher. In correspondence with facilities in the
United Kingdom, most also reported survival rates in excess
of 70% on most strains. Of the three reported protocols
(Table 2), protocol 1 had average survival rates of 79%, with
a range of 0-100% survival (n = 40). Protocol 1 also had average wild-type strain survival rate of 82% (n = 7) and a range
of 42-100%, counted from 4 DPF. Protocol 2 reported average survival rates of 87% (n = 10) across all lines, counted
from 5 DPF. Protocol 3 reported probable survival rates of
80-85%. Specific lines were reported as having lower survival rates; these include some transgenic lines, including
injected embryos, and some mutant lines, such as albino b3,
which because of their genetic background can have low
survival rates (C. Wilson, personal observation). These
problems are unlikely to be caused by factors related to larvalrearing protocols but rather by factors such as broodstock
problems, genetics, or manual embryonic manipulation.
The majority of reported mortalities (21 of 35 respondents, ZHA survey) occur around 11 to 16 DPF, when larvae
are at their most vulnerable: the yolk sac has been completely depleted and the larvae are dependent on exogenous
feeding. High mortality rates at this point may be explained
Volume 53, Number 2
2012
by lack of correct nutrition or starvation. Many facilities feed
ad libitum. When feeding live foods such as Paramecium, it
is vital, however, that both quality and quantity are checked
prior to feeding; if either is poor it can result in starvation (C.
Wilson, personal observation). Feeding stressors include
under- and overfeeding, continuation of Paramecium-only
diet for too long, feeding of diet inconsistent with gape size,
sudden changes of diet, and no overlapping dietary regimen
at times of developmental changes.
Extensive and frequent water changes affecting water
chemistry will create another stressor. Large changes can affect temperature and pH. Siphoning debris from larval tanks
seems to be a common practice in the United States, where 28
of 48 respondents to the ZHA survey indicated siphoning at
least once a week. Siphoning is uncommon in the United
Kingdom (personal observation). It can cause water disturbance, but remaining food debris might lead to depressed
oxygen levels and elevated ammonia leading to larval death
(C. Wilson, personal observation). Siphoning and manual
water changes are not practical for larger facilities because
they are very labor intensive.
A small percentage of mortality occurs at later stages—16
DPF and beyond. At this point, metamorphosis should be
complete and the larvae less vulnerable. One possibility for
this later-stage mortality may be delayed metamorphosis or
genetic factors inherent to the strain. Some facilities may not
remove homozygous lethal mutant embryos before putting
them into the nursery, and this would lower perceived survival rates. Embryos that have been exposed to physical manipulation may also be candidates for nonsurvival and, again,
may give lower survival rates than expected on average.
Survival rates are likely an indicator of good or bad larvalrearing technique but may also reflect, to some extent, the
research goals of a laboratory.
Sex Determination
In addition to affecting survival rate, larval-rearing protocols
and research goals can have an effect on sex determination.
Both genetic and environmental components likely influence sex determination in zebrafish (Lawrence et al. 2008).
However, unlike in many other species, in which sex determination is determined by chromosomal background, no such
genetic mechanism has been identified in zebrafish, although
several candidate genes are implicated in the process, which
is likely to be a complex interaction of several loci interacting
with environmental factors (Bradley et al. 2011; von Hofsten
and Olsson 2005). All zebrafish initially exhibit gonad plasticity and develop immature ovaries, but the point at which
spermatogenesis is determined in fish that will develop as
males is currently unknown (Orban et al. 2009). However,
one study showed that somatic genes were expressed indifferently at 10 to 17 DPF and then became sexually dimorphic at
3 weeks (Tong et al. 2010). Thus, practices employed in larval rearing are likely to have an effect on the sex of the adult
fish. Anecdotally, high stocking densities and limited food
173
174
ILAR Journal
14-18
19-30
17-30
≥29
31-60
31-60
2
3
1
2
3
Maximum 7 fish per liter
Maximum 15 fish per liter
Maximum 60 fish per liter
Maximum 5 fish per liter
Maximum 7 fish per liter
Dripa
Rapid drip, 0.5 ml/sec
Flow through
8 ml/sec
Rapid dripa
3 ml/sec
Maximum 17 fish per liter
Maximum 7 fish per liter
Maximum 60 fish per liter
Maximum 16 fish per liter
Maximum 60 fish per liter
Maximum 17 fish per liter
Petri dish, maximum 50
Petri dish, maximum 50
Petri dish, maximum 50
Maximum 17 fish per liter
Paramecium or fry powderd
Rotifer: Nannochloropsis
Instar I and II Artemia
Instar I Artemia and
powderc food
Paramecium or fry powderd
Artemia at 72 Instar II
Instar I and II Artemia
Instar I Artemia and adult
powder food
Artemia and flake food
Instar I and II Artemia
None
None
None
Paramecium and
fry powderc
Paramecium or fry powderd
Rotifer: Nannochloropsis
Paramecium, fry powderc,
and Instar I Artemia
Stocking density per liter Feed type
Drip, 1.2 ml/ sec
Slow dripa
Slow drip, 0.1 ml/sec
Static
Static
Static
Slow drip,
0.3 ml/sec
Static
Static
Slow drip,
0.45 ml/sec
Water flow rate
NA
NA
NA
Paramecium × 2 daily
Fry powder × 1
×2b
Ad libitum
Paramecium and
Artemia × 2
Powder food × 1
×2b
Rotifer × 2
Artemia × 3
Artemia × 2
Powder food × 1
×2b
×2b
4 times daily
AM and PM
regimen as adult
AM and PM
AM and PM
Feed frequency
See Appendix for more details of individual protocols. Protocol 1: University College London. Protocol 2: Cancer Research UK. Protocol 3: Children's Hospital Boston. DPF, days postfertilization;
NA, not applicable.
aPrecise speed of drip not measured.
bReduced to one feeding per day at weekend.
cProfile of powder food: crude protein, minimum 50%; crude lipid, minimum 12%; ash, maximum 10%; crude fiber, maximum 2.5%; moisture, maximum 10%. Raw ingredients: marine proteins,
plant algae/yeast, fish oil, cholesterol, plant starches, vitamin and vitamin premixes, antioxidants, pigments, biodegradable binders.
dProfile of powder food: protein, 55%; lipid, 14%; ash, 12%; fiber, 1%; vitamin A, 30,000 IU/Kg; vitamin D3, 2500 IU/Kg; vitamin E, 700 IU/Kg; vitamin C, 2000 IU/Kg; ␻3 HUFA, 30 Mg/g. Raw
ingredients: fish products, cereal grain products and byproducts, oils and fats, vitamins, minerals, antioxidants.
Juvenile
14-28
9-3
10-16
2
3
Midlarval
1
5-8
5-9
9-13
2
3
1
Early larval
Midlarval through
metamorphosis
0-3
0-4
0-4
4-8
1
2
3
1
Embryonic
Age (DPF)
Protocol
Stage
Table 2 Comparison of three currently used larval-rearing protocols
resources produce more males than females, and lower stocking densities and high food availability produce more females.
Some reports support the idea of growth rate as the main factor in environmental determination of sex (Lawrence et al.
2008). Other environmental factors such as hypoxia and temperature may also have an effect on sex determination (Orban
et al. 2009), as do estrogen compounds (Maack and Segner
2004). Watts and colleagues (2012) discuss sex differences
and differentiation elsewhere in this issue.
Summary
From modest origins, the use of the zebrafish as a biomedical
model has increased rapidly; fish are now the secondmost used species in the United Kingdom (after mice).
More animal facilities, used to catering for the husbandry
needs of mammalian species, now find themselves providing
husbandry for aquatic species, especially zebrafish, on a
scale unimaginable 20 years ago. Other facilities are
catering exclusively for zebrafish, housing tens of thousands, if not hundreds of thousands, of fish. Although the
zebrafish has been the object of much scientific research
and many aspects of its biology are well understood,
many of the husbandry requirements still remain elusive.
Many practices are still based largely on anecdotal
evidence or on aspects of husbandry procured from other
fish species, originally from the ornamental fish trade and
more recently from aquaculture. Groups and individuals
find it difficult to obtain specific zebrafish husbandry
information because it is often published in journals that
they may not have access to or be aware of. There are
many areas of zebrafish husbandry that still require a
great deal of research, and this includes larval rearing.
The anecdotal evidence of larval-rearing protocols and
techniques described here suggests that a wide variety of
techniques are in use. This lack of standardization may
have an important impact on the reproducibility of results
from research under such variable conditions. For example,
the three protocols in Table 2 vary considerably in water
flow rates and diet, but all appear to result in similar survival
rates. It is difficult to determine what effect each factor in
different protocols has on larvae and on larvae as they
turn adult. For example, how do the different protocols
affect sex ratios, fertility, fecundity, and larval morphology?
More trials will be needed to rigorously examine each
aspect of larval rearing on survival, robustness, and sex
ratio to ascertain the importance of adhering to specific
larval-rearing practices.
Although the interest in zebrafish husbandry has increased notably in recent years—from animal technologists
through managers and directors to veterinarians, legislators,
and welfare groups—there are no standardized practices
based on sound scientific evidence. Perhaps now is the time
to really consider how to drive research in this direction
and to begin implementing protocols and standards that
everyone can follow.
Volume 53, Number 2
2012
Acknowledgments
I thank Carly Nicholls and Heather Calloway for their help
with the images and tables and Florencia Cavodeassi and
Anukampa Barth for their patience and help with the manuscript. I also owe thanks to Isaac Adatto and the board of directors at Zebrafish Husbandry Association for allowing me
to use information they have collected about larval-rearing
practices and to all the anonymous people and institutions in
the United Kingdom, Europe, and the United States who
shared their larval-rearing protocols and practices with me. I
also thank Christian Lawrence, Gary Childs, and Darren Martin
for allowing me to publish their larval-rearing protocols.
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ILAR Journal
Appendix: Larval-Rearing Procedures of
Three Different Facilities
Stage 3: Days 17 to 30 (and up to 45)
Color code: Yellow
The larval-rearing procedures of three different facilities—the
Aquatic Resources Program at Children’s Hospital Boston,
Cancer Research UK, and University College London—are
outlined below. Please note that all days are days
postfertilization.
Aquatic Resources Program, Children’s
Hospital Boston
Flow: 0.5 ml/sec. This is a rapid drip, just below a stream.
Screen: Medium-grade mesh
Feed types: Artemia nauplii/metanauplii
Feed amounts: Artemia: To satiation (all the nauplii the fish
will eat within 5 min)
Feed frequencies: Artemia: 8 AM, 10:30 AM, 12 PM, 3:30 PM
Stage 1: Days 5 to 9
Color code: Green
Stocking densities: 40 to 60 fish per liter. This would be 80
to 120 larvae, respectively.
Tank type/system: 2 L mouse cage/nursery
Flow: None. Fish go on nursery in 100 ml of water. Through
additions from feedings volume may increase over stage to
400 to 500 ml.
Stage 4: Days 30 (45) to 60
Flow: 3.0 ml/sec
Screen: None required; in practice the smallest-grade mesh
required for stage 2 is recommended,
Feed types: Rotifers, fed out in 5 ppt salinity greenwater
(Nannochloropsis).
Feed amounts: Ad libitum. There should be enough at each
feeding such that the rotifers live and reproduce in the tanks
at very high densities for the duration of this stage.
Feed frequencies: 9 AM, 3 AM
Stocking densities: Up to 40 to 60 fish per liter. This would
be 80 to 120 larvae, respectively.
Screen: None
Feed types: Artemia nauplii/metanauplii
Feed amounts: Artemia: Feed each tank to satiation (all the
nauplii the fish will eat within 5 min)
Feed frequencies: Artemia: 8:30 AM, 3:30 PM
Stocking densities: 10 to 15 fish per liter
Tank type/system: 9 L tank/juvenile
Cancer Research UK
Tank type/system: 2 L mouse cage/nursery.
Day 0
Stage 2: Days 10 to 16
Color code: White
Approximately 16 fertile embryos are sorted (dead/infertile
eggs removed) into a petri dish. A larger amount of embryos per
dish will result in small, poor-quality fish that are mainly male.
Flow: 0.1 ml/sec. This is a slow, steady drip.
Screen: Smallest-grade mesh
Feed types: Artemia nauplii/metanauplii; rotifers if
available.
Feed amounts: Artemia: To satiation (all the nauplii the fish
will eat within 5 min). For practical purposes, estimate 5 to 10
cysts per larva, per tank at each feeding. Rotifers: Ad libitum.
Feed frequencies: Artemia: 8:30 AM, 12 PM, 3:30 PM. Rotifers: 9 AM, 3 PM, if available.
Stocking densities: Up to 40 to 60 fish per liter. This would
be 80 to 120 larvae, respectively.
Tank type/system: 2 L mouse cage/nursery
Volume 53, Number 2
2012
Day 5
Fry are moved to “sandwich” boxes. They are fed on ZM000 fry powder or paramecium.
Day 9
Fry are transferred to small plastic tanks under drippers. A
blue sticker is used to indicate that they are to be fed ZM-000
fry powder or paramecium.
Day 14
A blue sticker is used to indicate that they are still to be
fed ZM-000 fry powder. An orange sticker is added to
177
indicate that the fry are to be fed live brine shrimp as
well.
Day 19
An orange sticker is used to indicate that the fry are to be fed
live brine shrimp. The blue sticker is removed.
One Month
The fry are transferred to the larger plastic tanks. This is recorded in the Aquarium Diary for later transfer to the Fish
Database. They are fed live brine shrimp only.
Day 8
Feed: 3 ml filtered paramecium per 1 L fish water, once in
the morning and once in the evening; powdered fry food at
lunch-time feed
Day 9
Feed: 2 drops of concentrated 24-hr Artemia first feed of
morning (check fry to see if it has been eaten); 3 ml filtered
paramecium per 1 L fish water, once in the morning and
once in the evening; powdered fry food at lunch-time feed
Two Months
Day 10
Old aquarium 2B 31a fry are transferred to the glass tanks.
An orange sticker placed on the tank indicates that as young
fish an additional feed of live brine shrimp can be given.
These fish are now fed Tetrafin fish flake twice a day.
New aquarium 2B 31e fry are transferred to the small
and/or large plastic tanks depending on number and size of
fish. (Dividing the fish into two plastic tanks appears to improve growth.) These fish are now fed fish flake in the morning and live brine shrimp in the afternoon. In both cases, this
is recorded in the Aquarium Diary for later transfer to the
Fish Database.
Feed: 2 drops of concentrated 24-hr Artemia first feed of
morning (check fry to see if it has been eaten); 3 ml filtered
paramecium per 1 L fish water, once in the morning and
once in the evening; powdered fry food at lunch-time feed
University College London
The fry are on a slow drip, with increased water flow as the
fry become older.
Day 4
Fry into the nursery (slow drip).
Day 11
Feed: 0.5 ml of concentrated 24-hr Artemia first feed of
morning (check fry to see if it has been eaten); 2 ml paramecium in morning feed with brine shrimp; powdered fry
food in solution of fish water at lunch-time feed; 3 ml
filtered paramecium per 1 liter fish water, once in the
afternoon
Day 12
Feed: 1 ml of concentrated 24-hr Artemia first feed of morning (check fry to see if it has been eaten); powdered fry food
and very finely crushed adult food in solution of fish water at
lunch-time feed; 3 ml filtered paramecium per 1 liter fish
water, once in the afternoon
Stocking density: No more than 15 fry per 1 L fish water
Feed: 10 ml filtered paramecium
Day 5
Feed: 3 ml filtered paramecium per 1 L fish water, once in
the morning and once in the evening; powdered fry food in
solution of fish water at lunch-time feed
Day 6
Feed: 3 ml filtered paramecium per 1 L fish water, once in
the morning and once in the evening; powdered fry food at
lunch-time feed
Day 13
Feed: 1 ml of concentrated 24-hr Artemia first feed of morning; powdered fry food and finely crushed adult food at
lunch-time feed; 0.5 ml concentrated 24-hr Artemia and 1 ml
paramecium per 1 liter fish water once in the afternoon
Day 14
Feed: 1 ml of concentrated 24-hr Artemia first feed of morning; powdered fry food and finely crushed adult food at
lunch-time feed; 1 ml concentrated 24-hr Artemia per 1 liter
fish water once in the afternoon
Day 7
Days 15 to 28
Feed: 3 ml filtered paramecium per 1 L fish water, once in the
morning and once in the evening; powdered fry food at lunchtime feed
178
Feed: As for day 14, but increase the amount of powdered
food, working toward adult powdered food
ILAR Journal