Genetic analysis of vertebrates, especially mice, has tradition-
ally focused on traits that are expressed postnatally, and do not
lead to embryonic lethality (Lyon and Searle, 1989). Zygotic-
effect mutations leading to intrauterine embryonic lethality are
difficult both to identify and to analyze in mammals.
Therefore, there have been few mutations found to interfere
with the establishment of the body plan and early steps of
organogenesis. This situation has changed since targeted gene
disruption makes it possible to follow the segregation of
mutant alleles using molecular analysis, and to determine
whether a given embryo is homozygous for the induced
mutation (Capecchi, 1989). This technology has made a
tremendous impact on our understanding of vertebrate devel-
opment (McMahon and Bradley, 1990; Thomas and Capecchi,
1990; Joyner et al., 1991; Carpenter et al., 1993; Urbanek et
al., 1994; Wurst et al., 1994). Two important approaches
remain very difficult in mice. First, forward genetics to identify
novel genes involved in embryogenesis is prohibitively
expensive. Second, experimental and embryological studies of
mutations acting early in embryogenesis are greatly inhibited
by the intrauterine mode of development.
It is exactly the combination of the last two approaches that
has been the experimental basis for the current detailed under-
standing of invertebrate and plant development. Large scale,
systematic genetic screens led to the characterization of genetic
pathways that pattern the C. elegans, Drosophila and Ara-
bidopsis embryos (Hirsh and Vanderslice, 1976; Nüsslein-
Volhard and Wieschaus, 1980; Mayer et al., 1991).
Zebrafish (Danio rerio), a small tropical freshwater teleost,
was recognized as a genetic system in which similar approaches
would be feasible (Streisinger et al., 1981). The high fecundity,
short generation time and rapid development of the externally
fertilized, translucent embryos make it an excellent vertebrate
genetic model system (reviewed by Driever et al., 1994). Devel-
Development 123, 37-46
Printed in Great Britain © The Company of Biologists Limited 1996
embryonic phenotypes have been instrumental in the
understanding of invertebrate and plant development.
Here, we report the results from the first application of
such a large-scale genetic screening to vertebrate develop-
Male zebrafish were mutagenized with N-ethyl N-
nitrosourea to induce mutations in spermatogonial cells at
an average specific locus rate of one in 651 mutagenized
genomes. Mutations were transmitted to the F1generation,
and 2205 F2families were raised. F3embryos from sibling
crosses within the F2 families were screened for develop-
mental abnormalities. A total of 2337 mutagenized
genomes were analyzed, and 2383 mutations resulting in
abnormal embryonic and early larval phenotypes were
identified. The phenotypes of 695 mutants indicated
involvement of the identified loci in specific aspects of
embryogenesis. These mutations were maintained for
further characterization and were classified into categories
according to their phenotypes. The analyses and genetic
genome-wide mutagenesis screens forcomplementation of mutations from several categories are
reported in separate manuscripts. Mutations affecting pig-
mentation, motility, muscle and body shape have not been
extensively analyzed and are listed here. A total of 331
mutations were tested for allelism within their respective
categories. This defined 220 genetic loci with on average 1.5
alleles per locus. For about two-thirds of all loci only one
allele was isolated. Therefore it is not possible to give a
reliable estimate on the degree of saturation reached in our
screen; however, the number of genes that can mutate to
visible embryonic and early larval phenotypes in zebrafish
is expected to be several-fold larger than the one for which
we have observed mutant alleles during the screen. This
screen demonstrates that mutations affecting a variety of
developmental processes can be efficiently recovered from
Key words: zebrafish Danio rerio, mutagenesis, genetic control,
A genetic screen for mutations affecting embryogenesis in zebrafish
W. Driever*, L. Solnica-Krezel, A. F. Schier, S. C. F. Neuhauss, J. Malicki, D. L. Stemple, D. Y. R. Stainier†,
F. Zwartkruis‡, S. Abdelilah, Z. Rangini§, J. Belak and C. Boggs
Cardiovascular Research Center, Massachusetts General Hospital and Harvard Medical School, 149 13th Street, Charlestown,
MA 02129, USA
*Author for correspondence (e-mail: Driever@Helix.MGH.Harvard.EDU or: Driever@ruf.uni-freiburg.de)
†Present address: Department of Biochemistry and Biophysics, School of Medicine, UCSF, San Francisco, CA 94143-0554, USA
‡Present address: Laboratory for Physiological Chemistry, Utrecht University, Universiteitsweg 100, 3584 CG Utrecht, The Netherlands
§Present address: Department of Oncology, Sharett Institute, Hadassah Hospital, Jerusalem 91120, Israel
opment of the zebrafish embryo (Kimmel et al., 1995) has been
studied in detail, from pre-gastrula and gastrula stages
(reviewed by Driever, 1995; Kuwada, 1995; Solnica-Krezel et
al., 1995; Woo et al., 1995) through organogenesis (Schmitt and
Dowling, 1994; Stainier and Fishman, 1994), lending a solid
base of knowledge for the detection and interpretation of mutant
phenotypes. Prior to our screen, nine zebrafish mutations
affecting embryogenesis have been published and their effect
on development studied in detail, providing important insights
into mechanisms of vertebrate development (Grunwald et al.,
1988; Kimmel et al., 1989; Felsenfeld et al., 1990; Westerfield
et al., 1990; Hatta et al., 1991; Halpern et al., 1993; Abdelilah
et al., 1994; Stainier et al., 1995; Talbot et al., 1995).
Chemical mutagens are preferred over gamma- or X-rays for
genome-wide screens, since they predominantly induce lesions
limited to single genes (Singer and Grunberger, 1983). N-
ethyl-N-nitrosourea (ENU), which had been demonstrated pre-
viously to be the most potent mutagen of the mouse germ line
(Russell et al., 1979), can induce mutations in the proliferating
germ line of male zebrafish with high efficiency. Specific-locus
rates ranging from one newly induced allele per 300 to 2000
mutagenized genomes were achieved for four different pig-
mentation loci. Methods have been developed to induce and
recover mutations in zebrafish on a large scale, with the goal
of saturating the genome for mutations affecting embryogen-
esis (Mullins et al., 1994; Solnica-Krezel et al., 1994; Riley
and Grunwald, 1995).
Here, we report the results of one of the two first large-scale
screens for embryonic visible mutations in a vertebrate (for
details of the other, parallel screen see Haffter et al., 1996).
Our screen is based on the following concepts. (1) Detailed
visual inspection of living zebrafish larvae during several
stages of embryonic and early larval development will identify
mutants deficient in genes that are required for patterning and
differentiation of the embryo. The translucent zebrafish
embryo is ideally suited for such a study, since from fertiliza-
tion to late organogenesis, development can be observed in
vivo. (2) Recessive embryonic lethal mutations can be effi-
ciently recovered in a two-generation breeding scheme
(Haldane, 1956). (3) The analysis of large F3egg clutches (20-
90 embryos) makes it possible to distinguish polygenic or non-
genetic defects from single-gene mutations, which will
segregate in a Mendelian fashion.
Here and in the accompanying manuscripts (Brockerhoff et
al., 1995; Abdelilah et al., 1996; Neuhauss et al., 1996; Pack
et al., 1996; Schier et al., 1996; Solnica-Krezel et al., 1996;
Stainier et al., 1996; Stemple et al., 1996; Weinstein et al.,
1996; Malicki et al., 1996a,b), we present 577 mutations that
were found to affect specific aspects of early zebrafish
MATERIALS AND METHODS
Fish stocks and fish keeping
Fish husbandry and genetic crosses were performed as previously
described (Solnica-Krezel et al., 1994), except that the fish water is
now based on reversed osmosis water (Ionpure Reversed Osmosis
system, Millipore), with the addition of 157 mg/l calcium sulfate
(Plaster of Paris), 20 mg/l sodium bicarbonate and 100 mg/l Instant
Ocean salt mix (Aquarium Systems, Mentor, Ohio). The pH in the
recirculating water system equilibrates to about 6.9, due to acid pro-
duction in tanks and filters.
The mutagenesis was performed using fish of the ‘AB’ line
(Chakrabarti et al., 1983). The AB line was maintained in our facility
by inbreeding, and fish stocks were tested every generation for the
absence of embryonic- or larval-lethal mutations.
Mutagenesis and breeding of F2families
The screen was designed as a two-generation breeding screen with
analysis of mutant phenotypes in F3 embryos (Solnica-Krezel et al.,
1994). Adult male AB zebrafish were incubated at weekly intervals for
1 hour each in solutions of the mutagen ENU. Nine sets of males were
mutagenized using ENU at concentrations between 2 mM and 3.5 mM,
and the dose was repeated two to six times (see Table 1). About 240
out of 266 males survived the mutagenic regimens. Mutations were
recovered from mutagenized proliferating germ line, such that
mutations were fixed in the G0male. The mutagenized males were bred
several times with females heterozygous for four unlinked pigmenta-
tion mutations to determine the efficiency of mutagenesis (Table 1; and
Solnica-Krezel et al., 1994). Between 4 weeks and 2 months after muta-
genesis, males were bred with AB females, except for the NB 11 set,
for which about half of the progeny was obtained from crosses with
AB/HK hybrid females. The F1generation comprised 17,000 progeny
of G0males from 346 crosses. Survival of F1larvae to adulthood was
low, and ranged between 10% and about 70% for individual families.
From August 1992 to March 1994, up to 50 F2 families were
generated every week. Only crosses with more than 60 larvae with
inflated swim bladders by 5 dpf (days postfertilization) were kept and
up to 125 larvae were raised. 2312 families from F1×F1 crosses, and
487 from F1×AB were raised, representing a total of 5111 mutage-
nized genomes. Of these 2799 F2families, 594 were discarded before
the age of 4 months, since the number of fish had dropped below 20,
which made efficient screening impossible. Of the 2205 F2 families
raised to adulthood, 397 could not be screened because either they
contained fish of one sex only, or no productive crosses could be
obtained. Thus, 1808 F2families (317 from F1×AB, 1491 from F1×F1
crosses) representing 3299 genomes, were screened. The overall
screen is depicted in Fig. 1.
At the time the screen was designed, the only possibility for mapping
of mutations appeared to be based on a RAPD map (Postlethwait et al.,
1994), so care was taken to perform the mutagenesis and generate F1
and F2families in an AB-strain inbred background. With the availabil-
ity of an SSR-based map, however, mapping of mutations recovered
from outbred backgrounds has become feasible (Knapik et al., 1996).
Since specific locus rates had been determined for each of the
separate mutagenic regimes, we were able to compare the specific
locus rate (Solnica-Krezel et al., 1994) with the number of lethal
mutations induced per mutagenized genome (Table 1), and found that
the correlation was good.
Organization of the screening procedure
For each F2family, up to 25 single pair crosses were set up in breeding
traps, in the late afternoon. Starting at noon on the next day, eggs were
harvested into Petri dishes, and parents of successful crosses were
transferred pairwise into containers with about 1.5 l of fish facility
water (disposable plastic containers, 1.9 l, Fisher). Each container with
F2 parents, as well as the Petri dish with their progeny, was given a
unique label for later reidentification. Fish were stored in these con-
tainers for up to 7 days without feeding and survived as well as fish in
tanks. If fewer than six crosses with more than 20 fertilized embryos
each were obtained, the F2family was crossed again 1 or 2 weeks later
before fish were discarded. A minimum of 20 and up to 90 fertilized
embryos from each cross were sorted, at 6-12 hours postfertilization
(hpf), into Falcon 6-well tissue culture plates, 30 embryos per well.
As soon as specific phenotypes were observed, parental fish were
transferred from containers to facility 1l tanks and fed. The same pair
was crossed again 1-2 weeks later, and phenotypes were analyzed in
more detail and documented photographically. If the phenotype was
confirmed, and segregated in about one quarter of the embryos, inter-
W. Driever and others
39 Genetic screen in zebrafish
esting and specific mutations were assigned an allele number (m). For
preservation of mutations, usually three sperm samples were frozen
from the heterozygous F2male (see below). Further, heterozygous fish
were outcrossed with an ‘AB’ strain fish, and sometimes with a ‘HK’,
‘Tübingen’ or ‘AB-gol’ strain fish. The outcrossed F3 progeny was
raised, and segregation of the mutation in the F3as well as F4gener-
ation was determined.
We tried to screen at least six egglays per F2family to optimize the
chance of finding a mutation against the effort involved and facility
space required. The probability of finding a mutation in a family is
P=(1−0.75n), where n is the number of successful crosses. For n=6
(4;8), P gives 0.82 (0.68;0.9), respectively. Thus, by screening a
minimum of six crosses for a family, we would lose at most 18% of
the mutations. The rationale behind analyzing at least 20 embryos per
cross was twofold. First, the probability of finding at least one mutant
among 20 embryos if both parents are heterozygous is 99.6%. More
importantly, on average there are five mutant embryos of a given
mutation among the 20 embryos, thus allowing the segregation of a
mutant phenotype to be determined during the initial screening.
The actual number of mutagenized genomes analyzed per F2family
was calculated according to the following equation:
G = (1−0.75n)×a,
where G=genomes analyzed; n=number of crosses with more than 20
embryos; a=number of mutagenized genomes per family: a=1 for
F1×AB- and 2 for F1×F1-derived F2families).
Visual screening of embryonic and early larval
development of F3
Each cross was analyzed under a dissecting microscope (Wild M5
with transmitted light base; 5-50× and 10-100× optics) at five stages
during embryonic and early larval development. At each time point,
phenotypes were documented on a protocol form, and the percentage
of mutant embryos noted.
(1) At 6-12 hpf, morphology of the gastrula was examined during
sorting of fertilized embryos. Progress of epiboly, formation of the
embryonic shield and quality of the yolk cell were recorded.
(2) At 24-36 hpf the major focus of screening was the central
nervous system (CNS) morphology. During later stages of develop-
ment, the morphology of the brain is more difficult to screen, since the
brain becomes more compact and obscured by pigmentation. The
following were analyzed: shape of diencephalon, telencephalon,
epiphysis, tectum, cerebellum, midbrain-hindbrain border and
hindbrain; size of brain ventricles; shape of the spinal cord, presence
of floor plate; shape of eyes and otic vesicle. In addition, the shape and
proper differentiation of notochord and somites, the overall shape of
the body and yolk, and the presence of a touch response, were noted.
(3) At 48-60 hpf the major focus was on morphology and function
of the cardiovascular system. By this time, the heart and vasculature
are fully functional. Screening for cardiovascular defects was
performed at this early stage because pericardial and general edema
and/or secondary cardiovascular defects can obscure the phenotype at
3 dpf or later. Morphology of the heart, beat rate, blood flow and the
major vessels were analyzed. Additionally, pigmentation and body
shape were evaluated.
(4) At 72-84 hpf the following features were analyzed: general body
shape and pigmentation; morphology of the brain, eyes, ears and spinal
cord; formation of jaw and branchial arches; form and function of heart,
blood and the circulatory system, presence of edema; and differentiation
of notochord, somites, muscle, pectoral and tail fins. Furthermore, touch
response and motility of the larvae were tested by tapping them with a
needle at one side of the trunk. Wild-type larvae respond with a typical
fast-start escape response (Eaton and Bombardieri, 1978). Normally, by
72 hpf all larvae are hatched. The percentage of non-hatched larvae was
recorded, and larvae that did not hatch were tested for motility by
tapping the chorion, or dechorionation.
(5) After 5 dpf morphological analysis and tests performed at 3 dpf
were repeated. Following observation of touch response and motility,
fish were anesthetized for detailed analysis of late differentiating
structures. Specifically, the morphology of the jaw and branchial
region was inspected. Number and location of lateral line organs was
analyzed. Presence of intestinal organs (gut, liver, pancreas) and anus
was verified, and larvae were inspected for formation of edematous
kidney cysts. Inflation of the swim bladder was scored for crosses with
mutant phenotypes. In a subset of mutagenized lines (corresponding
to about 500 mutagenized genomes), a detailed screen of the mor-
phology of gut, liver and pancreas was performed utilizing a Wild
M10 dissecting scope and 50-100× magnification (Pack et al., 1996).
Mutations that lead to extensive cellular degeneration, and/or block
circulation, before 72 hpf (when the hatching period begins; Kimmel
et al., 1995), are called embryonic lethal. Mutations that result in
defects manifest between 72 hpf and 5 dpf are classified as early larval
mutations. Most of the latter are also lethal, as indicated by a failure
to inflate the swim bladder and/or to feed.
Maintenance of stocks and mutant alleles
Mutant alleles are maintained as outcrossed stocks and/or frozen
sperm samples. Care was taken to have one to three samples frozen
from the original heterozygous AB male(s) isolated during the screen.
Up to six additional samples were frozen from the first and second
generation outcrosses into AB, Tübingen or HK strain backgrounds.
About 2900 sperm samples have been processed this way. More than
half of the mutations reported from this screen were initially main-
tained as frozen samples only, and subsequently recovered by in vitro
fertilization for further characterization of the mutant phenotype.
The protocol for cryopreservation of sperm was modified from that
Table 1. Mutagenesis conditions and the recovery of embryonic and larval lethal mutations
ENU treatment Mutagenesis Specific locus rateGenomes screenedLethal mutations Lethals per genome
6×1 hour, 3 mM
6×1 hour, 3 mM
2×1 hour, 3 mM
4×1 hour, 3 mM
4×1 hour, 2.5 mM
4×1 hour, 2 mM
4×1 hour, 3 mM
4×1 hour, 3.5 mM
4×1 hour, 3 mM
The specific locus rate for NB4 to NB11 was determined for the pigmentation loci golden, brass, sparse and albino, and is reported in Solnica-Krezel et al.
*Screening for NB12 was less efficient since a nematode infection affected survival of fish.
described by Westerfield (1994). The detailed protocol is available,
for brevity of printed space, at our WWW server sites
‘http://zebrafish.mgh.harvard.edu’ or ‘http://zebrafish.uni-freiburg.de’.
Genetic nomenclature and availability of mutations
Mutant alleles and genetic loci are named according to the nomen-
clature rules for zebrafish (Mullins, 1995). Allele designations include
the letter m indicating our laboratory as the source of the mutation,
and a number, consecutively assigned to mutations as they were
recovered during the screen.
If complementation within a phenotypic class of mutations has been
completed, locus names have been assigned to complementation
groups. Locus names and three-letter abbreviations are used in the
text. Therefore, each locus reported from this screen should represent
a separate gene. However, if alleles at a locus express unrelated phe-
notypes, complementation would not have been performed and
separate names assigned. Genetic mapping of mutant alleles will
resolve these rare problems. All mutations for which complementa-
tion has not been performed or not completed are referred to by their
allele designations only.
All mutations reported by allele designation in this and the
accompanying manuscripts are available on request. Requests should
be directed to ‘Driever@helix.mgh.harvard.edu’. Since mutant
stocks will soon be also maintained at other sites, in the future the
address where a specific mutation can be obtained will be made
public on the Oregon (http://zfish.uoregon.edu) or MGH
(http://zebrafish.mgh.harvard.edu) or Freiburg (http://zebrafish.uni-
freiburg.de) zebrafish www Internet servers.
Due to limited space and resources, for most of the mutations we
will only be able to send out batches of larvae from outcrosses of het-
erozygous fish. Mutant stocks that are not currently maintained in our
laboratory will require us to schedule the recovery of the mutant allele
from frozen sperm, raise the mutant stock in our facility, identify the
heterozygous fish, replenish the depleted number of frozen sperm
samples, and prepare a cross for shipment. Thus it might take several
weeks to more than 6 months before stocks carrying a
mutation can be shipped, depending on availability of
heterozygous fish in our facility. A stock center for the
zebrafish community is being planned to expedite this
Design of the genetic screen
Our screen was designed to recover zygotic-effect
mutations in genes involved in a broad range of devel-
opmental processes, namely establishment of the
embryonic axes; early pattern formation and region-
alization of the germ layers during gastrulation;
differentiation of mesodermal derivatives; brain and
sense organ formation; and finally, various aspects of
organogenesis. The rationale for designing the screen
to study such a wide range of developmental
processes was threefold. Firstly, a broad screen can
help answer important questions about the vertebrate
genome, for example: Which developmental
processes are sensitive to mutations? What type of
embryonic-lethal and early larval lethal phenotypes
can be observed? Secondly, we reasoned that the
resources and time necessary to perform a screen for
mutations affecting a single aspect of development
would be similar to one of broader design. The same
number of fish would have to be raised and the same
number of crosses performed to screen the required
number of mutagenized genomes. Moreover, we wanted to
establish a genetic resource that would attract researchers to exploit
the advantages of the zebrafish system and use a genetic approach
to study aspects of development that are difficult to investigate in
A classic two-generation breeding scheme (Haldane, 1956)
was employed in the screen. With this approach, the segregation
of recessive mutations can be easily tested, and the F2 parents
of a cross producing mutant F3 embryos are identified as het-
erozygous carriers for the mutation. We decided against an F1
screen based on haploid embryos to avoid the obfuscation of
mutant phenotypes by the abnormal development of haploid
embryos. Additionally, we chose not to utilize the ‘early
pressure’ approach to generate diploid gynogenic embryos, since
the fraction of mutant embryos of the developing half tetrad F2
would depend on the gene centromere distance, and bias against
isolation of mutations in genes close to the telomeres (Streisinger
et al., 1981; Streisinger et al., 1986; Driever et al., 1994).
We raised 2799 F2families, a total of about 224,000 larvae,
representing 5111 mutagenized genomes. Only 1808 F2
families finally produced successful crosses for F3embryos to
be screened. More than 30,000 crosses were set up, 10,811 of
which produced a sufficient number of fertilized embryos for
screening. Nearly 500,000 F3embryos were visually analyzed
at five different developmental stages for abnormalities in
development. Based on the number of crosses analyzed for
each family, we calculate that we effectively screened 2337
mutagenized genomes of the 3299 genomes represented in the
Based on the specific locus rates determined after ENU muta-
genesis, we had previously argued that a screen of 1600 genomes
would be sufficient to identify mutations in 87% of all genes of
W. Driever and others
with ethyl nitrosourea
4 times at weekly interval
( for example, 3 mM ENU
for 1 hr)
analyze specific locus
rate in about 10,000
progeny: Specific locus
rate is 1 in 300-2,000
grow 30 times 100 F1
set up 200 crosses
every month to
generate the F2
grow 200 F2 lines
(~60 fish each)
cross F2 siblings
to identify mutant
If there is one lethal mutations induced in every gamete, on
average every second F2 sibling cross will give rise to 1/4 progeny
mutant in one of these genes.
+ + + +
> 4 weeks after mutagenesis
gol, brs, alb, spa
total screen (9 sets):
266 G0 males mutagenized
240 G0 males survive
346 G0 x wild type
ca. 17,000 F1 fish
2,799 F2 families started
1,808 F2 families screened
10,811 F2 x F2 screen
ca. 500,000 F3 embryos
948 mutations with
Fig. 1. F2screen to identify embryonic lethal mutations in zebrafish.
41 Genetic screen in zebrafish
average mutability. For genes of less than average mutability, we
estimated that 5,000-10,000 genomes would have to be screened
(Solnica-Krezel et al., 1994). Based on this estimate, initially F2
families representing 5111 mutagenized genomes were
generated. The number of genomes actually screened was still
well above the number of genomes estimated to be necessary to
saturate for mutations in loci that mutate at average rates.
Given that only a small number of mutagenized genomes
(on average about 150) was screened from each mutagenized
male, the chance that alleles at any of the identified loci
represent clonal events is small (see also Discussion in Solnica-
Krezel et al., 1994). For most loci with several mutant alleles,
we can trace mutant alleles to different sets of mutagenized
males (data not shown).
Detection of mutant phenotypes
Detailed visual inspection of living zebrafish embryos at five
stages during embryogenesis and early larval development was
the main strategy to identify new mutations. Two additional
small-scale screens were carried out in parallel. In one screen,
F3 crosses representing 241 mutagenized genomes were
screened for abnormal optokinetic behavior. This screen was
based on analysis of the optokinetic nystagmus (OKN), and led
to the isolation of two mutations with absent or abnormal
OKN, both being morphologically normal. An additional
sixteen mutations with both morphological and OKN defects
were identified. Results from this screen are published
elsewhere (Brockerhoff et al., 1995).
In a second small-scale screen, we wanted to assess whether a
visual inspection of brain morphology is sufficient to detect most
abnormalities in organization of the zebrafish brain. Embryos
from crosses representing 101 mutagenized genomes were
screened for abnormal distribution and levels of acetylcholine
esterase (AchE) activity in the zebrafish brain at 24 hpf. At 24 hpf,
AchE is expressed in a characteristic pattern in early neuronal
clusters of the fore-, mid- and hindbrain (Ross et al., 1992); the
rhombomeric organization of the hindbrain is especially well
visualized. Our pilot screen did not reveal any specific mutation
that was not already detected by morphological criteria (two
mutations affecting brain development, and five mutations
affecting neuronal survival were recovered per 100 genomes;
Table 4). Frequently reduced AchE levels were encountered in
25% of embryos due to delay of embryonic development. In all
cases, live siblings maintained for visual inspection indicated a
mutation-induced delay of development that was not specific to
the CNS. Thus, the effort to perform a AchE screen was not con-
sidered an efficient way to recover subtle CNS mutations.
Classes of mutations isolated during the screen
During the screen, 2383 mutations that lead to abnormal
embryonic and early larval development were found (Table 2).
Thus, about one mutation with a visible phenotype was
induced per mutagenized haploid genome. Due to lack of good
cytogenetics in zebrafish, we were not able to determine
whether all of these mutations represent lesions in single
zygotic genes, or if some of them are caused by chromosomal
abnormalities. However, we applied the strict criterion that
only mutant phenotypes segregating at 25% were considered
for our statistics. Further, since we screened on average more
than five crosses per F2 family, we determined that a given
such mutation was recovered on average from one of four F2
crosses (data not shown). Therefore, we consider it likely that
most of the mutations presented here do not represent chro-
mosomal rearrangements producing aneuploid embryos, or
haploinsufficient maternal or zygotic effect mutations. Pheno-
types were classified into four major categories.
(1) 494 mutations resulted in a widespread, apparently non-
tissue-specific onset of degeneration between 1 and 6 dpf. In
most cases, delay of developmental progress was detectable as
early as 24 to 48 hpf. We assume that these genes are essential
for general cellular survival.
(2) 587 mutations resulted in onset of retarded development
between 2 and 5 dpf. The embryos developed a syndrome of
defects, including small eye and brain size; under-differenti-
ated and small jaw and gill region; short and underdeveloped
pectoral fins; and delayed differentiation of internal organs.
The trunk and tail region appeared to be largely normal. The
extent of delay, and the degree to which different parts of the
embryo were affected, were variable. We hypothesize that this
group of mutations affects basic functions of all cells in the
embryo. The late onset of the phenotype could argue that
maternally derived gene products sustain development of the
embryo for the first few days. We note that tissues most
affected are late differentiating/proliferating tissues, while
somitic muscle, for instance, is not initially affected.
(3) 351 mutations lead to degeneration of the entire CNS.
Table 2. Categories of embryonic and larval lethal mutations detected in the screen
Widespread, apparently not tissue-specific; onset of degeneration 1-6 dpf
Normal early development, delayed development obvious only later
than 24 hpf, late differentiating structures like jaw, pectoral fins, eyes,
intestine very small and underdeveloped
Degeneration of CNS, not localized, onset 1-5 dpf. Death of embryo
usually within two days
Abnormal late development, ‘organogenesis’: embryos develop normally
during the first day, as judged by visual inspection
Abnormal development visually detectable during the first day of
development: pattern formation and early differentiation
Total number of mutations
Mutations maintained for further characterization after screen
Mutations reported here
Onset of degeneration was between 1 and 5 dpf. In most cases,
degeneration spread within 24-48 hours to the rest of the body.
These mutations could affect loci specifically required in the
neural lineages, or, perhaps, the phenotypes reflect a more
stringent requirement for metabolic gene products in the neural
Mutations from the above three categories were not main-
tained, since they did not appear to affect specific aspects of
development and occurred too frequently to be maintained.
However, all these mutations were characterized well enough
during the screen to determine that they are very likely to be
zygotic effect, recessive lethal mutations. Only mutations that
segregate in 25% of the F3crosses were included in this count.
During the first third of the screen, F2crosses were repeated to
confirm segregation for these mutations. Frequently, the
mutations were recovered in several (about every fourth) of the
crosses within a given F2family.
(4) 261 mutations lead to region-, tissue- or organ-specific
abnormalities during the first day of development, and 687
after the first day of development. Among these 948 mutations,
695 were maintained for further characterization. The other
253 mutations were of one of the following groups: (1)
abnormal body curvature (bent toward dorsal, ventral or
lateral) with no other visible morphological defects; (2) subtle
deviations from normal pigmentation pattern or intensity of
pigmentation; (3) mutations that were lost after phenotypes had
been confirmed, and therefore neither outcrosses nor frozen
sperm samples generated. For the first two groups, a subset of
mutations was maintained (see below).
The 695 mutations maintained for further characterization
are categorized in Table 4. Of the 695 initially characterized
mutations, 114 were lost during attempts to recover the
mutations, or were found to be non-specific, i.e. displaying a
phenotype similar to one of the first three classes described
above. Most misassigned mutations were in the eye and cran-
iofacial groups. Often a developmental delay was initially mis-
interpreted as a region-specific defect. Many of the lost car-
diovascular mutants were initially characterized as valve or
functional phenotypes of variable penetrance, and difficult to
recover from outcrosses.
Complementation analysis and number of genetic loci
Complementation analysis was performed within phenotypi-
cally related groups of mutations. The number of crosses
performed to isolate a sufficient number of heterozygous fish
for allelism tests, and the actual complementation crosses, rep-
resented an effort comparable to the initial screen. Embryos
from more than 17,000 crosses were analyzed to prepare and
perform the complementation analysis reported in the above
Complementation within the phenotypic classes was
completed for 331 mutations. These mutations define 220 genetic
loci. On average 1.5 alleles per locus were recovered (Table 3).
Mutations affecting several aspects of zebrafish development
have not yet been extensively analyzed. Classification of these
mutations and their isolation numbers are listed in Table 5.
More detailed information on these mutations is available on
the World Wide Web at ‘http://zebrafish.mgh.harvard.edu’,
including a brief description of each mutation, as well as
several photographs depicting the mutant phenotype.
Sources for updates on genetic information
Complementation of mutations isolated in this screen with
those performed in other laboratories, further characterization
of phenotypes and genetic mapping of mutant loci, will con-
tinuously increase the usefulness of these mutations in the
study of vertebrate development. Under the lead of M. West-
erfield and the University of Oregon computer science group,
a searchable database will be established at the University of
Oregon that will contain frequently updated information on
zebrafish genetic resources (http://zfish.uoregon.edu).
In the meantime, updated and extended information will
be available on the
‘http://zebrafish.mgh.harvard.edu’. For mutations presented in
this and the accompanying manuscripts, phenotypic descrip-
tions and sets of microphotographs will be available. Further,
a database will be established and routinely updated to search
all published mutations by locus name and locus abbreviation,
allele designation, and phenotype keywords.
Mutational analysis of vertebrate development
Our genetic screen demonstrates that it is possible to efficiently
induce and recover mutations in various aspects of zebrafish
development. Mutations affecting early developmental events,
such as patterning of the embryonic axes and gastrulation
movements (Solnica-Krezel et al., 1996), as well as mutations
in genes controlling organ development (Stainier et al., 1996),
were recovered. Moreover, since the screen was performed on
living embryos and larvae at several stages of development,
mutations affecting not only the form, but also functional aspects
of early larvae, were identified. Examples include mutations
affecting heart beat rate (Stainier et al., 1996), touch response or
optokinetic behavior (Brockerhoff et al., 1995). Additionally,
genetic pathways controlling the formation of certain structures,
such as the notochord, may be described (Stemple et al., 1996).
A relatively small number of mutations has been isolated
and shown to be involved in formation of the embryonic axes
at the beginning of gastrulation (Solnica-Krezel et al., 1996).
The same is true for anterior-posterior patterning of the brain
(Schier et al., 1996). Whether this finding reflects a strong
maternal contribution, or redundant genetic pathways, or the
inability of this screen to isolate additional mutations involved
W. Driever and others
Table 3. Distribution of frequencies at which alleles for
individual loci have been recovered
Alleles/locusLoci Number of mutations
Only loci from phenotypic classes for which complementation has been
completed are listed.
43 Genetic screen in zebrafish
in these processes due to haplo-insufficient effects (see below),
remains to be determined.
Estimates on saturation and essential gene number
We have recovered new alleles in each of the six previously
published zebrafish zygotic-effect genes, ntl, flh, cyc, fub, clo
and spt. While it is tempting to use this fact as an argument
that alleles in a large portion of all developmental control genes
were recovered during the screen, it may indicate that most of
the previously identified genes are mutational ‘hot spots’, and
more effort is needed to saturate the genome for mutations
Indeed, a statistical analysis of the distribution of allele
number indicates that the 220 loci defined here represent only
a small portion of all loci mutable to an observable develop-
mental phenotype in the zebrafish genome. Since more than
two-thirds of all loci are represented by only one allele it is
impossible to provide a reliable estimate of the total number
of genes required specifically for the control of development.
The data on allele distribution reported here do not fit a Poisson
distribution, which would require genes to be mutable at a
similar rate. Fitting the data to a negative binomial distribution
indicates that a large class of loci exists for which no alleles
were recovered. It is possible that the number of developmen-
tal control genes is several-fold higher than the estimated 400
genetic loci (577 mutations, 1.5 alleles per locus average) for
which alleles were recovered.
Similar conclusions would apply to estimates of the total
number of genes essential for embryonic and early larval
survival. If one applies the same rate, 1.5 alleles per locus, to
the total number of 2383 embryonic and early larval lethal
mutations identified in the screen, one can estimate that they
represent about 1600 essential loci. Again, the total number of
essential genes would be expected to be several-fold higher.
Such estimates are close to data obtained from mutational
analysis of the mouse genome, for which estimates of the total
number of essential genes range from about 5,000 to 10,000
(Carter, 1957; Shedlovsky et al., 1986).
These estimates are significantly larger than earlier estimates
of the total number of loci yielding embryonic lethal mutations
in zebrafish (Mullins et al., 1994; Solnica-Krezel et al., 1994). It
appears that many genes mutate at rates significantly lower than
the pigmentation loci used to determine the specific locus rates
after ENU mutagenesis. One possibility is that the albino, golden
and sparse loci used to determine the specific locus rates are
mutational hot spots, which in turn could explain why they were
the first genetic pigmentation defects characterized in zebrafish
(Chakrabarti et al., 1983). The brass locus, which mutates at two-
to fivefold lower rates, may in fact be more representative of the
zebrafish genome. Further, the specific locus test is more
Table 4. Phenotypic classes of mutations isolated during the genetic screen
per locus Class of mutationsIsolatedLost1
Gastrulation, tail formation
Early onset and neural degeneration
Late onset neural degeneration
Heart and vasculature
Muscle and somite development
General body shape
1Lost either due to inability to freeze sperm prior to death of founder fish, or to inability to recover the mutant allele from an outcross obtained from a frozen
2Pending completion of complementation for about 5% of loci.
3Efforts are still undertaken to recover six potential maternal effect mutations from these ten.
4Data to differentiate between lost and nonspecific mutations not available.
5Three mutations were lost; however the phenotype was well enough documented to be reported.
6Not including cyc, oep and boz.
7Some mutations are listed in several groups, since the phenotypes are pleiotropic. Therefore, the sum of the numbers listed for the individual classes is in most
cases larger than the total.
8Not including sly, bal, gup.
9Number of loci does not include 18 mutations affecting otoliths, for which complementation has not been completed.
n.d. not determined
References: a, Solnica-Krezel et al. (1996); b, Stemple et al. (1996); c, Schier et al. (1996); d, Abdelilah et al. (1996); e, M. Rodriguez et al. (unpublished); f,
Malicki et al. (1996a,b); g, Neuhauss et al. (1996); h, Weinstein et al. (1996); i, Stainier et al. (1996); j, Pack et al. (1996); k, I. Drummond and W. Driever,
unpublished; l, M.-A. Akimenko and W. Driever, unpublished; m, Brockerhoff et al. (1995); n, this manuscript.
sensitive when compared to the F2screen, since even weak alleles
can be recovered when trans-heterozygous with strong alleles.
During the F2screen, a homozygote of a weak allele may result
in a phenotype too subtle or penetrance too low to be recovered.
While saturation for loci mutable to visible phenotypes has
not been reached, it is important to note that our screen
indicates that a large number of developmental pathways, and
many of their components, are amenable to genetic analysis.
This holds true both for early, gastrulation phenotypes as well
as for late organogenesis phenotypes. A large number of loci,
probably in the thousands, can be defined by their phenotype
during embryogenesis and early larval development.
Limitations of this genetic screen
Several limitations on the recovery of developmental control
genes are intrinsic to the design of the screen. Maternal-effect
genes, which in Xenopus have been shown to play an important
role in pattern formation (Kessler and Melton, 1994), would
not have been recovered from our screen. A screen for
maternal-effect genes would have required raising the F3gen-
eration, and phenotypes would only be expressed in F4
embryos. Dominant lethal maternal-effect mutations would
have been detected in the F2generation of the screen, but it is
impossible to establish genetic lines bearing these mutations.
Dominant lethal zygotic-effect mutations would be expressed
in the F1 generation, and heterozygotes eliminated. Haploin-
sufficient, or low-penetrance dominant mutations, would be
eliminated to a large extent in the F1 generation. A recent
prominent example of such a phenotype in mice is the targeted
disruption of the HNF-3β gene, for which a haploinsufficient
effect reduces the viability of heterozygous carriers (Ang and
Rossant, 1994; Weinstein et al., 1994).
Added limitations stem from the organization of the ver-
tebrate genome. Genes are often represented in large families,
with partially or completely overlapping developmental
functions. Therefore, mutations in individual genes might have
only subtle or no phenotypes, while the developmental impor-
tance of a set of genes can only be uncovered by double or
triple mutants. The functional redundancy of MyoD and Myf-
5 in muscle determination is one such example (Rudnicki et
al., 1993). In light of the growing evidence for genetic redun-
dancy in the vertebrate, our screen is a first functional survey
of a vertebrate genome for genes required uniquely for specific
developmental processes. Our findings indicate that many loci
are indeed essential and not part of completely redundant sets
of genes. In the future, second-site non-complementation F1
screens will be a powerful tool for unraveling genetic
(I) Future genetic screens
The main limitation of our screening strategy is the detection of
subtle phenotypes. We tried to assess this problem by screening
a subset of lines utilizing a simple histological stain for acetyl-
choline esterase activity. Those results indicate that we would
not have missed many mutations affecting the overall architec-
ture of the brain during the morphological screen. However,
regional identity, neuronal connectivity and differentiation could
only be assessed with more sophisticated sets of molecular
markers, employing in situ or antigen expression analysis.
Screens focused to specific aspects of development, and
employing sets of markers to identify specific cell types, will
undoubtedly identify a large number of additional mutations in
novel genetic loci. A first such screen using immunohistochem-
istry to detect formation of dorsal root ganglia in zebrafish has
recently been performed (Henion et al., 1996). The small size of
the zebrafish embryo and early larvae make whole-mount
immunohistochemistry as well as RNA in situ hybridization
feasible even on the large scale needed for genetic screens.
We chose a two-generation breeding scheme for our screen,
W. Driever and others
Table 5. Mutations isolated during the genetic screen and not studied in detail
Short bodies (viable)
Early larvae have dorsally curved body
Early larvae have ventrally curved body
m451, m737, m763
m71, m72, m85, m130, m177, m252, m408, m415, m439, m542, m570, m575, m577, m593, m609, m611,
m648, m679, m695, m705, m742, m746, m773
Lateral line organs:
Number of lateral line organs reduced or
missing in trunk and tail
No touch response
Reduced or abnormal touch response
Abnormal shape or differentiation
Pigmentation lighter or no dark pigment
m92, m167, m355, m418, m434, m493, m519, m654, m624
m127, m154, m246, m385 m386, m4421 m455, m522, m588, m596, m665, m723, m749
m143, m195, m260, m446, m505, m558, m774
m26, m65, m104, m106, m164, m192, m261, m272, m279, m324, m334, m356, m374, m396, m397, m398,
m449, m457, m460, m556, m589, golm592, m621, m682, m693, m731, m732, m741
Pigmentation darker or dilated melanophores
Iridophores, xanthophores abnormal or absent
m266, m368, m417, m761
m95, m187, m464, m540, m571, m615, m652, m666, m722, m756, m783
m121, m136, spam141, m179,m204, m280, spam442, spam443, m467, m500, m717
Descriptions of the phenotypes as well as pictures can be retrieved on the Internet/WWW at ‘http://zebrafish.mgh.harvard.edu’or ‘http://zebrafish.uni-freiburg.de’
45 Genetic screen in zebrafish
since it is the only method presently available with a very low
background in non-genetic developmental abnormalities. Fur-
thermore, it provides the least bias against certain types of phe-
notypes, like those resembling haploid embryos obtained in
haploid or early pressure diploid screens (Streisinger et al.,
1981; Kimmel et al., 1989; Driever et al., 1994). It also does
not bias against genes in certain chromosomal regions, like
early pressure half-tetrads, which segregate mutant phenotypes
in loci close to the ends of the chromosomes at low rates.
However, haploid and early pressure gynogenetic F2 progeny
are much faster, easier and cheaper to generate than F3
progeny, and therefore will in the future be used extensively
for rapid surveys of the zebrafish genome for mutations
affecting defined aspects of development.
(II) Molecular analysis – mapping and cloning
It will be important to link the mutant loci isolated in zebrafish
to the repertoire of genes isolated and characterized at a molecular
level in other vertebrate systems. This will involve either the
isolation of candidate genes, or positional cloning, both of which
require a good genetic map for zebrafish. More than one thousand
marker (RAPDs and SSLPs) have been placed on the zebrafish
genetic map (Postlethwait et al., 1994; Johnson et al., 1996;
Knapik et al., 1996), and these laboratories expect to reach a
marker density of one marker per centiMorgan (with an average
of 590 kbp per cM in zebrafish). The availability of the map has
already facilitated the identification of the gene for the floating
headmutation, the zebrafish homologue of XenopusXnot (Talbot
et al., 1995). The identification of candidate genes will be further
accelerated as soon as synteny relationships between the
zebrafish and mammalian genomes are characterized by mapping
of genes cloned from zebrafish and mammals (Nadeau, 1989;
Nadeau et al., 1995; Postlethwait et al., personal communication).
Large insert libraries necessary for positional cloning are
available (Genome Systems, St Louis, USA). Given the excellent
mapping resolution one can obtain in zebrafish (segregation
analysis of marker in one thousand progeny by PCR should result
in better than 0.1 cM, or 50-100 kbp resolution), and efficient
ways to screen genomic regions of this size for sequences
expressed in the affected structures of the embryo by in situ
hybridization, cloning of mutant genes will hopefully soon be a
routine procedure similar to those in invertebrate genetic model
Will these efforts lead to the identification of new genes, or
will most of them be homologues of genes previously identi-
fied in Drosophila and other invertebrates? The analysis of the
human genome reveals that thus far about half of all open
reading frames have no previously known homologues, and
several hundred previously unknown gene families can be
predicted (Adams et al., 1995). We hope that zebrafish genetics
will significantly contribute to our understanding of vertebrate-
specific genetic functions. Many features of vertebrate embryo-
genesis, such as the neural crest or the notochord, and organo-
genesis have no obvious counterparts in invertebrates, and a
detailed molecular understanding of vertebrate development
will have to rely on the combination of forward genetic
approaches, molecular characterization and experimental
analysis in vertebrate model systems like the zebrafish.
We would like to thank Mark Fishman for constant support, Susan
Brockerhoff for enlightening us about the potentials of optokinetic
screens, and actually performing one in our laboratory. Thanks to
Yosef Gruenbaum for help during optimization of the cryoprotection
procedure for zebrafish sperm and, along with Abraham Fainsod, for
communicating to us the composition of the I-buffer. Thanks to
Richard Feichtinger for advice on statistical analysis. We thank Lisa
Vogelsang, Jeanine Downing, Laike Stewart, Pamela Cohen, Thomas
Binder, Kristen Dieffenbach, Xiaorong Ji, Heather Goldsboro and
Snorri Gunnarson for technical help during the various stages of the
screen. Thanks also to Brant Weinstein, Erez Raz and Eliza Mount-
castle-Shah for critical reading of the manuscript. This work was
supported in part by NIH RO1-HD29761 and a sponsored research
agreement with Bristol Myers-Squibb (to W. D.). The establishment
of the zebrafish database on the Internet is funded by an NSF grant
(to Monte Westerfield, Eugene and W. D.). Further support in the
form of fellowships came from HFSP and the Fullbright Program (to
Z. R.), EMBO and Swiss National Fund (to A. S.), Helen Hay
Whitney Foundation (to D. L. S. and D. Y. S.), and the Damon
Runyon-Walter Winchell Cancer Research Fund (to J. M.).
Abdelilah, S., Mountcastle-Shah, E., Harvey, M., Solnica-Krezel, L., Schier,
A. F., Stemple, D. L., Malicki, J., Neuhauss, S. C. F., Zwartkruis, F.,
Stainier, D. Y. R., Rangini, Z. and Driever, W. (1996). Mutations affecting
neural survival in the zebrafish, Danio rerio. Development 123, 217-227.
Abdelilah, S., Solnica-Krezel, L., Stainier, D. Y. and Driever, W. (1994).
Implications for dorsoventral axis determination from the zebrafish mutation
janus. Nature 370, 468-71.
Adams, M., Kerlavage, A. R., Fleischman, R. D., Fulder, R. A., Bult, C. J.,
others and Venter, J. C. (1995). Initial assessment of human gene diversity
and expression patterns based upon 83 million nucleotides of cDNA
sequence. Nature 377 Suppl., 3-174.
Ang, S.-L. and Rossant, J.(1994). HNF-3βis essential for node and notochord
formation in mouse development. Cell 78, 561-574.
Brockerhoff, S. E., Hurley, J. B., Driever, W., Neuhauss, S. and Dowling, J.
E. (1995). A behavioral screen for isolating zebrafish mutants with visual
system defects. Proc. Natl. Acad. Sci. USA 92, 10545-49.
Capecchi, M. R. (1989). Altering the genome by homologous recombination.
Science 244, 1288-1292.
Carpenter, E. M., Goddard, J. M., Chisaka, O., Manley, N. R. and
Capecchi, M. R. (1993). Loss of Hox-A1 (Hox-1.6) function results in the
reorganization of the murine hindbrain. Development 118, 1063-1075.
Carter, T. C. (1957). Recessive lethal mutations induced in the mouse by
chronic gamma irradiation. Proc. R. Soc. B. 147, 109-123.
Chakrabarti, S., Streisinger, G., Singer, F. and Walker, C. (1983).
Frequency of γ-ray induced specific locus and recessive lethal mutations in
mature germ cells of the zebrafish, Brachydaniorerio. Genetics103, 109-123.
Driever, W. (1995). Axis formation in zebrafish. Curr. Opin. Genet. Dev. 5,
Driever, W., Stemple, D., Schier, A. and Solnica-Krezel, L.(1994). Zebrafish:
genetic tools for studying vertebrate development. Trends Genet. 10, 152-9.
Eaton, R. C. and Bombardieri, R. (1978). Behavioral functions of the
Mauthner neuron. In Neurobiology of the Mauthner Cell (ed. D. S. Faber and
H. Korn). New York, Raven Press.
Felsenfeld, A. L., Walker, C., Westerfield, M., Kimmel, C. and Streisinger,
G. (1990). Mutations affecting skeletal muscle myofibril structure in the
zebrafish. Development 108, 443-59.
Grunwald, D. J., Kimmel, C. B., Westerfield, M., Walker, C. and
Streisinger, G. (1988). A neural degeneration mutation that spares primary
neurons in the zebrafish. Dev. Biol. 126, 115-28.
Haffter, P., Granato, M., Brand, M., Mullins, M. C., Hammerschmidt, M.,
Kane, D. A., Odenthal, J., van Eeden, F. J. M., Jiang, Y.-J., Heisenberg,
C.-P., Kelsh, R. N., Furutani-Seiki, M., Vogelsang, E., Beuchle, D.,
Schach, U., Fabian, C. and Nüsslein-Volhard, C. (1996). The
identification of genes with unique and essential functions in the
development of the zebrafish, Danio rerio. Development 123, 1-36.
Halpern, M. E., Ho, R. K., Walker, C. and Kimmel, C. B. (1993). Induction
of muscle pioneers and floor plate is distinguished by the zebrafish no tail
mutation. Cell 75, 99-111.
Hatta, K., Kimmel, C. B., Ho, R. K. and Walker, C. (1991). The cyclops
mutation blocks specification of the floor plate of the zebrafish central
nervous system. Nature 350, 339-41.
46 Download full-text
Henion, P. D., Raible, D. W., Beattie, C. E., Stoesser, K. L., Weston, J. A.
and Eisen, J. S. (1996). A screen for mutations affecting development of
zebrafish neural crest. Dev. Genet. (in press).
Hirsh, D. and Vanderslice, R. (1976). Temperature sensitive developmental
mutants of Caenorhabditis elegans. Dev. Biol. 49, 220-235.
Johnson, S. L., Gates, M. A., Johnson, M., Talbot, W. S., Horne, S., Baik,
K., Rude, S., Wong, J. R. and Postlethwait, J. H. (1996). Half-Tetrad
Analysis in zebrafish II: Centromere-linkage Analysis and Consolidation of
the Zebrafish Genetic Map. Genetics (in press).
Joyner, A. L., Herrup, K., Auerbach, B. A., Davis, C. A. and Rossant, J.
(1991). Subtle cerebellar phenotype in mice homozygous for a targeted
deletion of the En-2 homeobox. Science 251, 1239-1243.
Kessler, D. S. and Melton, D. A. (1994). Vertebrate embryonic induction:
Mesodermal and neural patterning. Science 266, 596-604.
Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B. and Schilling,
T. F. (1995). Stages of embryonic development of the zebrafish. Dev.
Dynam. 203, 253-310.
Kimmel, C. B., Kane, D. A., Walker, C., Warga, R. M. and Rothman, M. B.
(1989). A mutation that changes cell movement and cell fate in the zebrafish
embryo. Nature 337, 358-62.
Knapik, E. W., Goodman, A., Atkinson, O. S., Roberts, C. T., Shiozawa,
M., Sim, C. U., Weksler-Zangen, S., Trolliet, M. R., Futrell, C., Innes, B.
A., Koike, G., McLaughlin, M. G., Pierre, L., Simon, J. S., Vilallonga, E.,
Roy, M., Chiang, P.-W., Fishman, M. C., Driever, W. and Jacob, H. J.
(1996). A reference cross DNA panel for zebrafish (Danio rerio) anchored
with simple sequence length polymorphisms. Development 123, 451-460.
Kuwada, J. Y. (1995). Development of the zebrafish nervous system: genetic
analysis and manipulation. [Review]. Curr. Opin. Neurobiol. 5, 50-4.
Lyon, M. F. and Searle, A. G. (1989). Genetic Variants and Strains of the
Laboratory Mouse. Oxford, Oxford University Press.
Malicki, J., Neuhauss, S. C. F., Schier, A. F., Solnica-Krezel, L., Stemple, D.
L., Stainier, D. Y. R., Abdelilah, S., Zwartkruis, F., Rangini, Z. and
Driever, W. (1996a). Mutations affecting development of the zebrafish
retina. Development 123, 263-273.
Malicki, J., Schier, A. F., Solnica-Krezel, L., Stemple, D. L., Neuhauss, S.
C. F., Stainier, D. Y. R., Abdelilah, S., Rangini, Z., Zwartkruis, F. and
Driever, W. (1996b). Mutations affecting development of the zebrafish ear.
Development 123, 275-283.
Mayer, U., Torres-Ruiz, R. A., Berleth, T., Misera, S. and Jürgens, G.(1991).
Mutations affecting body organization in the Arabidopsis embryo. Nature353,
McMahon, A. P. and Bradley, A. (1990). The Wnt-1 (int-1) proto-oncogen is
required for development of a large region of the mouse brain. Cell62, 1073-85.
Mullins, M. (1995). Zebrafish. In Trends in Genetics: Genetic Nomenclature
Guide (ed. A. Stewart). Cambridge: Elsevier Trends Journals.
Mullins, M. C., Hammerschmidt, M., Haffter, P. and Nusslein-Volhard, C.
(1994). Large-scale mutagenesis in the zebrafish: in search of genes
controlling development in a vertebrate. Curr. Biol. 4, 189-202.
Nadeau, J. H. (1989). Maps of linkage and synteny homologies between
mouse and man. Trends Genet. 5, 82.
Nadeau, J. H., Grant, P. L., Mankala, S., Reiner, A. H., Richardson, J. E.
and Eppig, J. T. (1995). A Rosetta Stone of mammalian genetics. Nature
Neuhauss, S. C. F., Solnica-Krezel, L., Schier, A. F., Zwartkruis, F.,
Stemple, D. L., Malicki, J., Abdelilah, S., Stainier, D. Y. R. and Driever,
W. (1996). Mutations affecting craniofacial development in zebrafish.
Development 123, 357-367.
Nüsslein-Volhard, C. and Wieschaus, E. (1980). Mutations affecting
segment number and polarity in Drosophila. Nature 287, 795-801.
Pack, M., Solnica-Krezel, L., Malicki, J., Neuhauss, S. C. F., Schier, A. F.,
Stemple, D. L., Driever, W. and Fishman, M. C. (1996). Mutations
affecting development of zebrafish digestive organs. Development 123, 321-
Postlethwait, J. H., Johnson, S. L., Midson, C. N., Talbot, W. S., Gates, M.,
Ballinger, E. W., Africa, D., Andrews, R., Carl, T., Eisen, J. S. and et al.
(1994). A genetic linkage map for the zebrafish. Science 264, 699-703.
Riley, B. B. and Grunwald, D. J. (1995). Efficient induction of point
mutations allowing recovery of specific locus mutations in zebrafish. Proc.
Natl Acad. Sci. USA 92, 5997-6001.
Ross, L. S., Parrett, T. and Easter, S. S., Jr. (1992). Axonogenesis and
morphogenesis in the embryonic zebrafish brain. J. Neurosci. 12, 467-82.
Rudnicki, M. A., Schnegelsberg, P. N. J., Stead, R., Braun, T., Arnold, H.-
H. and Jaenisch, R. (1993). MyoD or Myf-5 is required for the formation of
skeletal muscle. Cell 75, 1351-1359.
Russell, W., Kelly, E., Hunsicker, P., Bangham, J., Maddux, S. and Phipps,
E. (1979). Specific-locus test shows ethylnitrosourea to be the most potent
mutagen in the mouse. Proc. Natl. Acad. Sci. USA 76, 5818-5819.
Schier, A. F., Neuhauss, S. C. F., Harvey, M., Malicki, J., Solnica-Krezel,
L., Stainier, D. Y. R., Zwartkruis, F., Abdelilah, S., Stemple, D. L.,
Rangini, Z., Yang, H. and Driever, W. (1996). Mutations affecting the
development of the embryonic zebrafish brain. Development 123, 165-178.
Schmitt, E. A. and Dowling, J. E. (1994). Early eye morphogenesis in the
zebrafish, Brachydanio rerio. J. Comp. Neurol. 344, 532-42.
Shedlovsky, A., Guenet, J., Johnson, L. and Dove, W. (1986). Induction of
recessive lethal mutations in the T/t-H-2 region of the mouse genome by a
point mutagen. Genet. Res. Camb. 47, 135-142.
Singer, B. and Grunberger, D. (1983). Molecular Biology of Mutagens and
Carcinogens. New York, Plenum Press.
Solnica-Krezel, L., Schier, A. F. and Driever, W.(1994). Efficient recovery of
ENU-induced mutations from the zebrafish germline. Genetics 136, 1401-20.
Solnica-Krezel, L., Stemple, D. L. and Driever, W. (1995). Transparent
things – cell fates and cell movements during early embryogenesis of
zebrafish. BioEssays 17, 931-939.
Solnica-Krezel, L., Stemple, D. L., Mountcastle-Shah, E., Rangini, Z.,
Neuhauss, S. C. F., Malicki, J., Schier, A. F., Stainier, D. Y. R.,
Zwartkruis, F., Abdelilah, S. and Driever, W. (1996). Mutations affecting
cell fates and cellular rearrangements during gastrulation in zebrafish.
Development 123, 67-80.
Stainier, D. Y. R. and Fishman, M. C.(1994). The zebrafish as a model system
to study cardiovascular development. Trend Cardiovasc. Med. 4, 207-212.
Stainier, D. Y. R., Fouquet, B., Chen, J.-N., Warren, K. S., Weinstein, B. M.,
Meiler, S., Mohideen, M.-A. P. K., Neuhauss, S. C. F., Solnica-Krezel, L.,
Schier, A. F., Zwartkruis, F., Stemple, D. L., Malicki, J., Driever, W. and
Fishman, M. C. (1996). Mutations affecting the formation and function of the
cardiovascular system in the zebrafish embryo. Development 123, 285-292.
Stainier, D. Y. R., Weinstein, B. M., Detrich III, H. W., Zon, L. I. and
Fishman, M. C. (1995). Cloche, an early acting zebrafish gene, is required by
both the endothelial and hematopoietic lineages. Development121, 3141-3150.
Stemple, D. L., Solnica-Krezel, L., Zwartkruis, F., Neuhauss, S. C. F.,
Schier, A. F., Malicki, J., Stainier, D. Y. R., Abdelilah, S., Rangini, Z.,
Mountcastle-Shah, E. and Driever, W. (1996). Mutations affecting
development of the notochord in zebrafish. Development 123, 117-128.
Streisinger, G., Walker, C., Dower, N., Knauber, D. and Singer, F. (1981).
Production of clones of homozygous diploid zebra fish (Brachydanio rerio).
Nature 291, 293-296.
Streisinger, G., Singer, F., Walker, C., Knauber, D. and Dower, N. (1986).
Segregation analyses and gene-centromere distances in zebrafish. Genetics
Talbot, W. S., Trevarrow, B., Halpern, M. E., Melby, A. E., Far, G.,
Postlethwait, J. H., Jovett, T., Kimmel, C. B. and Kimelman, D.(1995). A
homeobox gene essential for zebrafish notochord development. Nature 378,
Thomas, K. R. and Capecchi, M. R.(1990). Targeted disruption of the murine
int-1 proto-oncogene resulting in severe abnormalities in midbrain and
cerebellar development. Nature 346, 847-850.
Urbanek, P., Wang, Z. Q., Fetka, I., Wagner, E. F. and Busslinger, M.
(1994). Complete block of early B-cell differentiation and altered patterning
of the posterior midbrain in mice lacking Pax5/BSAP. Cell 79, 901-912.
Weinstein, D. C., Ruiz i Altaba, A., Chen, W. S., Hoodless, P., Prezioso, V.
R., Jessel, T. M. and Darnell, J. E. (1994). The winged-helix transcription
factor HNF-3β is required for notochord development in the mouse embryo.
Cell 78, 757-588.
Weinstein, B. M., Schier, A. F., Abdelilah, S., Malicki, J., Solnica-Krezel,
L., Stemple, D. L., Stainier, D. Y. R., Zwartkruis, F., Driever, W. and
Fishman, M. C. (1996). Hematopoietic mutations in the zebrafish.
Development 123, 303-309.
Westerfield, M. (1994). The Zebrafish Book. Eugene, University of Oregon
Westerfield, M., Liu, D. W., Kimmel, C. B. and Walker, C. (1990).
Pathfinding and synapse formation in a zebrafish mutant lacking functional
acetylcholine receptors. Neuron 4, 867-74.
Woo, K., Shih, J. and Fraser, S. (1995). Fate maps of the zebrafish embryo.
Curr. Op. Genet. Dev. 5, 439-443.
Wurst, W., Auerbach, A. B. and Joyner, A. L. (1994). Multiple
developmental defects in Engrailed-1 mutant mice: an early mid-hindbrain
deletion and patterning defects in forelimbs and sternum. Development 120,
(Accepted 10 April 1996)
W. Driever and others