Mutations affecting xanthophore pigmentation in the zebrafish, Danio rerio

Abstract

In a large-scale screen for mutants with defects in embryonic development we identified 17 genes (65 mutants) specifically required for the development of xanthophores. We provide evidence that these genes are required for three different aspects of xanthophore development. (1) Pigment cell formation and migration (pfeffer and salz); (2) pigment synthesis (edison, yobo, yocca and brie) and (3) pigment translocation (esrom, tilsit and tofu). The number of xanthophore cells that appear in the body is reduced in embryos with mutations in the two genes, salz and pfeffer. In heterozygous and homozygous salz and pfeffer adults, the melanophore stripes are interrupted, indicating that xanthophore cells have an important function in adult melanophore pattern formation. Most other genes affect only larval pigmentation. In embryos mutant for edison, yobo, yocca and brie, differences in pteridine synthesis can be observed under UV light and by thin-layer chromatography. Homozygous mutant females of yobo show a recessive maternal effect. Embryonic development is slowed down and embryos display head and tail truncations. Xanthophores in larvae mutant in the three genes esrom, tilsit and tofu appear less spread out. In addition, these mutants display a defect in retinotectal axon pathfinding. These mutations may affect xanthophore pigment distribution within the cells or xanthophore cell shape. Mutations in seven genes affecting xanthophore pigmentation remain unclassified.

Full text

INTRODUCTION
Skin coloration in fish and amphibia is created by the interac-
tion of light with pigmented cells (melanophores, xanthophores
and iridophores), that are derived from the neural crest
(DuShane, 1934; Rawles, 1944). Within these cells, pigments
are located in specialized pigment organelles such as
melanosomes, which contain melanin (Charles and Ingram,
1959; Drochmans, 1960; Birbeck, 1963); pterinosomes, which
contain pteridines in xanthophores (Matsumoto, 1965a;
Kamei-Takeuchi and Hama, 1971; Bagnara, 1976); and the
reflecting platelets which contain purines in iridophores
(Bagnara and Stackhouse 1961; Bagnara et al., 1979). Pteri-
nosomes, like melanosomes, are derived from the Golgi
complex (Obika, 1993). They contain a species-specific set of
pteridines that appear simultaneously with the differentiation
of xanthophores (Matsumoto et al., 1960; Hama, 1963; Obika,
1963; Matsumoto, 1965b). Sepiapterins, drosopterins and
several colourless pteridines can be detected as yellow pig-
mentation that first becomes visible within pterinosomes
(Obika, 1963; Kamei-Takeuchi and Hama, 1971). These
pteridines are largely the same as the pteridines described for
Drosophila eye pigmentation (Matsumoto, 1965a; Bagnara,
1966; Matsumoto and Obika, 1968; Frost and Malacinski,
1980).
Pigment organelles in individual chromatophores can be
transported along uncharacterized paths radiating from the cell
center, so that the organelles are either dispersed throughout
the cytoplasm or aggregated in the center of the cell (Taylor,
1992). The distribution of pigment organelles within pigment
cells is important in determining the general colouration of the
body. Cells with dispersed organelles give rise to the general
colour pattern of the animal, whereas chromatophores with
aggregated organelles appear less coloured (reviewed by Luby-
Phelps and Schliwa, 1982; Obika, 1986).
Several mutations have been described in different
amphibian species that affect different aspects of vertebrate pig-
mentation (Frost and Malacinski, 1980; Frost et al., 1982;
Droin, 1992). To gain a more detailed insight into the different
aspects of vertebrate pigmentation it would be helpful to study
a collection of mutations in one species. The zebrafish is an
ideal system for using the powerful tools of genetics to study
different aspects of vertebrate pigmentation (Mullins et al.,
1994; Mullins and Nüsslein-Volhard, 1993; Haffter et al.,
391
Development 123, 391-398
Printed in Great Britain © The Company of Biologists Limited 1996
DEV3364
In a large-scale screen for mutants with defects in
embryonic development we identified 17 genes (65
mutants) specifically required for the development of xan-
thophores. We provide evidence that these genes are
required for three different aspects of xanthophore devel-
opment. (1) Pigment cell formation and migration (pfeffer
and salz); (2) pigment synthesis (edison, yobo, yocca and
brie) and (3) pigment translocation (esrom, tilsit and tofu).
The number of xanthophore cells that appear in the body
is reduced in embryos with mutations in the two genes, salz
and pfeffer. In heterozygous and homozygous salz and
pfeffer adults, the melanophore stripes are interrupted,
indicating that xanthophore cells have an important
function in adult melanophore pattern formation. Most
other genes affect only larval pigmentation. In embryos
mutant for edison, yobo, yocca and brie, differences in
pteridine synthesis can be observed under UV light and by
thin-layer chromatography. Homozygous mutant females
of yobo show a recessive maternal effect. Embryonic devel-
opment is slowed down and embryos display head and tail
truncations. Xanthophores in larvae mutant in the three
genes esrom, tilsit and tofu appear less spread out. In
addition, these mutants display a defect in retinotectal axon
pathfinding. These mutations may affect xanthophore
pigment distribution within the cells or xanthophore cell
shape. Mutations in seven genes affecting xanthophore pig-
mentation remain unclassified.
Key words: pigment cells, chromatophores, xanthophores, pigment
pattern, pigment translocation, pteridines, maternal effect
SUMMARY
Mutations affecting xanthophore pigmentation in the zebrafish,
Danio rerio
Jörg Odenthal*, Karin Rossnagel, Pascal Haffter, Robert N. Kelsh†, Elisabeth Vogelsang‡, Michael Brand§,
Fredericus J. M. van Eeden, Makoto Furutani-Seiki, Michael Granato, Matthias Hammerschmidt¶,
Carl-Philipp Heisenberg, Yun-Jin Jiang, Donald A. Kane, Mary C. Mullins** and Christiane Nüsslein-Volhard
Max-Planck Institut für Entwicklungsbiologie, Spemannstrasse 35/III, 72076 Tübingen, Germany
*Author for correspondence
†Present address: Institute of Neuroscience, University of Oregon, Eugene, Oregon 97403, USA
‡Present address: Institut für Genetik der Universität Köln, Weyertal 121, 50931 Köln, Germany
§Present address: Institut für Neurobiologie, Universität Heidelberg, Im Neuenheimer Feld 364, 69120 Heidelberg, Germany
¶Present address: Harvard University, Department of Molecular and Cellular Biology, 16 Divinity Avenue, Cambridge, MA 02138, USA
**Present address: Department of Biology, University of Pennsylvania, Philadelphia PA 19104, USA

392
1996). Mutations affecting the development of the adult
pigment pattern in the zebrafish have been described previously
(Kirschbaum, 1975; Johnson et al., 1995).
Zebrafish xanthophore pigmentation first becomes apparent
after 42 hours of development (high pec stage) in the dorsal head
as a very pale cast of yellow (Kimmel et al., 1995). However,
individual xanthophore cells are difficult to distinguish. After 3
days of development (protruding mouth stage), strong yellow
xanthophore pigmentation is apparent in the entire dorsal half of
the body in regions devoid of melanophores. Xanthophores
become granular after 5 days of development and are now seen
throughout the body, with greater numbers dorsally. Adult
zebrafish have five alternating blue-black (melanophores and iri-
dophores) and silvery-yellow (xanthophores and iridophores)
stripes aligned parallel to the long axis of the body. This stripe
pattern extends into the caudal and anal fins, but not into the
dorsal and the paired lateral fins (Kirschbaum, 1975). Studies of
adult pigmentation mutants revealed interactions between
melanophores and iridophores in pigment pattern formation
(Johnson et al., 1995). In contrast, little is known about the inter-
action of the xanthophores with other chromatophores
(Epperlein and Löfberg, 1984; Goodrich et al., 1954).
In a screen for mutants with defects in embryonic develop-
ment, many embryonic pigmentation mutants were isolated
that allow the study of different aspects of pigment cell devel-
opment (Kelsh et al., 1996). In this paper we describe
mutations affecting the development of xanthophore pigmen-
tation. We provide evidence that genes involved in three
different aspects of xanthophore development were isolated in
the screen. Mutations in salz and pfeffer lead to an adult
pigment pattern phenotype that is probably due to a reduced
number of xanthophore cells in the body. Mutations in edison,
yobo, yocca and brie affect the pteridine content and so
probably function in pigment synthesis or uptake. Mutations in
esrom, tilsit and tofu affect retinotectal axon pathfinding as
well as xanthophore pigmentation, and might be required for
pigment translocation or for the generation of normal xan-
thophore cell shape.
MATERIALS AND METHODS
Fish raising, screen and crosses
Fish stocks were maintained as previously described by Mullins et al.
(1994), the isolation of the mutants and the complementation analysis
was performed as described by Haffter et al. (1996).
Ascending chromatography
Thin-layer chromatography was carried out as described by Epperlein
and Claviez, (1982a). After 3 days of development 45 embryos were
extracted with 100 µl diluted ammonia, 0.1% β-mercaptoethanol, pH
10.0. 25 µl were applied to heat-activated silica (Silica gel IB2, J. T.
Baker) and cellulose (Cellulose F1440, Schleicher and Schuell) plates.
Chromatography was allowed to proceed for 3 hours using n-
propanol/1% ammonia (2:1) as a solvent. RFis the distance migrated
by the substance divided by the distance migrated by the solvent.
Fluorescence pictures
Embryos were mounted in methyl cellulose with 1 drop of 0.2% 3-
aminobenzoic acid ethyl ester (Sigma), 1 drop of dilute ammonia,
0.1% β-mercaptoethanol, pH 10.0, and were illuminated with UV
light (DAPI-filter 395-420). The dilute ammonia liberates pteridines
from their protein carriers at high pH. These are then visualized as
light blue fluorescence (Ziegler, 1965; Epperlein and Claviez, 1982a).
Photographs were taken with 160 ASA tungsten films.
RESULTS
Genes required for xanthophore development
In a large-scale screen for mutations leading to defects during
the first 6 days of development, we screened for pigmentation
phenotypes on the 3rd and 6th day of development (Haffter et
al., 1996). A total of 284 mutations affecting larval pigmenta-
tion were isolated (Kelsh et al., 1996). Of these, 124 mutants
(41 genes) show xanthophore pigmentation defects (Table 1
and Kelsh et al., 1996). 20 mutants, defining 14 complemen-
tation groups, have defects in all three types of pigment cells,
that is the melanophores, the iridophores and the xanthophores.
Xanthophores and melanophores are affected in 19 mutants,
which make up four complementation groups. Xanthophore
and iridophore pigmentation is affected by mutations in at least
six genes (20 mutants). In this paper we focus on mutations in
17 genes (64 mutations), which lead to defects in xanthophores
without an effect on larval melanophore or iridophore pig-
mentation. Xanthophore specification or migration are
probably affected in salz (sal) and pfeffer (pfe) (Class II,
missing cell type in Kelsh et al., 1996), whereas the other genes
are probably required for xanthophore differentiation.
Mutations affecting xanthophore pigmentation lead to pale
J. Odenthal and others
Table 1. Classes of mutations affecting xanthophore pigmentation
Number Number Unresolved
Affected pigment cell type of mutants of genes (alleles) Gene names (alleles) (mutants)
Xanthophores, Melanophores 21 14 (20) blanched (1), bleached (3), bleich (1), colourless (2), 1 (tc233b)
and Iridophores pech (1), puzzle (1), sahne (1), sallow (1),
sunbleached (1), stonewashed (1), u-boot (1),
weiss (1), washed out (4), white tail (1)
Xanthophores and Melanophores 19 4 (17) cold-light (2), polished (2), stars and stripes (12), 2 (tc249, tz294)
tinte (1)
Xanthophores and Iridophores 20 6 (14) choco (2), cookie (2), milky (3), matt (1), pistachio (5), 6 (tj266c, tm107,
vanille (1) tq262a, tu235b,
tz284, ty105g)
Xanthophores 64 17 (62) bressot (1), brie (4), esrom (14), clorix (1), 2 (tn14, ta53b)
edison (11), feta (1), kefir (2), non blond (1),
pfeffer (6), quark(6), ricotta (1), salz (7), tartar(1),
tilsit (1), tofu (1), yobo (3), yocca (1)

393Zebrafish xanthophore mutants
yellow, whitish or granular cells (Class VI.J, pale xan-
thophores in Kelsh et al., 1996). Of the 17 genes required for
xanthophore pigmentation, larvae mutant for any of six genes,
bressot (bst), clorix (clx), edison (edi), esrom (esr), non blond
(nob) and tilsit (til), die in larval stages, whereas mutant larvae
for any of the other 11 genes survive to adulthood.
Xanthophore specification and migration
In sal and pfe mutant embryos a partial absence of xan-
thophores in the dorsal head becomes apparent after 4 days of
development whereas melanophore and iridophore pigmenta-
tion appears normal. 1 day later, large areas devoid of granular
xanthophores can be seen, and only a few individual yellow
spots in the dorsal head are present (arrows in Fig. 1C,E). The
overall number of xanthophores is reduced in pfe larva, as visu-
alized by their fluorescence under UV light (Fig. 1D). In sal
mutant larvae the number of xanthophores is strongly reduced
in lateral and ventral regions, whereas many xanthophores
remain dorsal to the neural tube (Fig. 1F and data not shown).
The strength of the defect in mutant larvae of both genes is
dependent on the allele; weak alleles produce a slightly
reduced number of xanthophores (data not shown), whereas in
embryos with strong alleles only a very small number of xan-
thophores is detectable in the body. For the strong allele
pfetq211, we observed a slight reduction in the number of xan-
thophore cells, and also in heterozygous larvae (data not
shown).
Heterozygous adult fish carrying a mutation in sal or in pfe
show irregular and interrupted melanophore stripes (shown for
pfe in Fig. 2B), thus indicating a defect in the formation or
maintenance of the adult melanophore pattern. Xanthophores
in the adult heterozygous fish are detectable at a slightly lower
density (data not shown; Haffter, personal communication).
The general appearance is slightly less yellow than wild type
fish. We compared the dominant adult phenotypes to determine
the relative strength of different alleles, which gave the same
results as counting the xanthophore number in the larva
(Table 2).
Homozygous sal and pfe fish can be raised to adulthood.
After 3 to 4 weeks of development these fish can be distin-
guished from wild type by a pigment pattern phenotype: the
spotted melanophore stripe pigmentation (blue-black stripes)
fades out towards the tail and shows a ‘salt and pepper’-like
appearance. This phenotype is very prominent in the adult fish,
and no xanthophores are detectable in the body of embryos
with the strong alleles (Fig. 2C and data not shown). The
weakest allele, pfetg283a, does not produce any obvious
phenotype in the heterozygous or homozygous adults.
Defects in pteridine synthesis in
edison
,
tartar
,
brie
,
yocca
and
yobo
In edison (edi) larvae no yellow pigmentation is detectable
(Fig. 3C). edi mutant embryos can be detected by the strong
blue fluorescence of embryonic cells including the yolk under
Table 2. Genes required for xanthophore pigmentation but not for melanophore or iridophore development
Adult Other Other
Gene Alleles viability phenotypes descriptions
Pigment cell number:
pfeffer (pfe) tm236b>tq211>tg17>tc227b>te220 >tg283a Viable Dominant and recessive a,b
adult phenotype
salz (sal) tl241>tf34>tp71c>tt254a>tm246b>tb213c>tf238b Viable Dominant and recessive a,b
adult phenotype
Pigment synthesis:
edison (edi) tc245c, tk232, tl35a, tl245, to255b, tp62, tp67b, tr276, Lethal Fluorescence in entire embryo
tt232, tv04, tz253
tartar (tar) td09 Viable Fluorescence in entire embryo
Recessive adult phenotype b
brie (bri) tg211b, tj226a, tm42c, tu269 Viable
yobo (yob) tc251, tk13, ty44d Viable Maternal effect b
Small iris in adult fish
yocca (yoc) tm86 Viable
Pigment distribution:
esrom (esr) tb241a, te250, te275, te279, te376, tf04z, tg05, tg265, Lethal Retinotectal axon pathfinding c
th222, th36a, tj236, tn207, tp203, ts208
tilsit (til) ty130b Lethal Retinotectal axon pathfinding c
tofu (tof) tq213c Viable Retinotectal axon pathfinding c
Not further classified:
bressot (bst) tp223b Lethal
feta (fet) ty107 Viable
kefir (kef) ta65b, tf229 (lost) Viable
quark (qua) tc276, tg239d, tk236, to241b, tp72g, tv46 Viable
ricotta (ric) tb212 Viable
clorix (clx) tj244 Lethal Small otoliths d
Motility defect e
non blond (nob) tt288 Lethal Small otoliths d
General retardation
Unresolved:
tn14 tn14 Lethal Jaw defects
ta53b ta53b Lethal Tectum necrosis
Alleles of different strength are listed according to descending strength separated by >.
References: a, Kelsh et al. (1996); b, Haffter, personal communication; c, Karlstrom et al. (1996); d, Whitfield et al. (1996); e, Granato et al. (1996).

394
illumination with UV light after 24 hours of development (Fig.
4). This fluorescence subsequently decreases, and by day 6 flu-
orescence is mainly located in the gut and is excreted. A single
allele for tartar (tar) was isolated and this shows a similar but
slightly weaker phenotype to that of edi. Weak yellow pig-
mentation can be detected in tar embryos after 5 days of devel-
opment (data not shown). Similarly to edi, tar embryos show
blue fluorescence in the entire embryo. In mutant edi and tar
embryos abnormally shaped and brightly fluorescing xan-
thophores are present in the body (shown for edi in Fig. 3D).
In contrast to edi, homozygous tar fish survive to adulthood
and show pale melanophore stripes in the adult (data not
shown).
As a first attempt to investigate the synthesis of yellow
pigments in the mutants, we performed thin-layer chromat-
ography of larvae to analyze the pteridine components in one
allele each of brie, esrom, edison, feta, pfeffer, quark, salz,
tartar, yobo and yocca. In wild-type larvae, eight different flu-
orescent bands can be visualized by UV light on the chro-
matogram (Table 3). Two bands show visible yellow colour
(bands 3 and 7) and therefore are most likely to be sepiapterins
(Matsumoto and Obika, 1968; reviewed by Bagnara, 1966).
The other six bands, which are colourless under visible light,
show fluorescent colours under UV light, are probably colour-
less pteridines such as biopterin and neopterin (light blue flu-
orescence), xanthopterin (green fluorescence) and isoxan-
thopterin (dark blue fluorescence). Correlations between the
known pteridines and the eight bands detectable in the wild-
type larvae are so far not clear.
Larvae mutant for any of three genes, brie (bri), yocca (yoc)
and yobo (yob), show similar pteridine patterns that differ from
wild type. The intensity of three fluorescent bands is increased
(band 2 increased, band 4 blue instead of green, band 5 strongly
increased), and the intensity of two other bands is reduced
(bands 6 and 8). For the other seven genes no changes were
found.
Xanthophores, as judged by morphological criteria, are
different in the three mutants, which show unusual pteridine
composition. In bri, cells appear feathery and whitish with
normal granularity after 5 days of development (Fig. 3E and
data not shown); under fluorescent light these cells appear
greenish (Fig. 3F). Xanthophores in yoc mutant larvae appear
brownish in the head. Under UV light these cells show a
diffuse greenish fluorescence (data not shown). Faintly
granular xanthophores with reduced yellow pigmentation are
detectable in the head of yob mutant larvae, and fluorescent
cells are not detectable in the body (data not shown). Xan-
thophores in homozygous adult yob, yoc and bri fish appear
J. Odenthal and others
Fig. 1. The number of xanthophore cells is reduced in salz and
pfeffer homozygous larvae. Dorsal views of day-6 larvae in the head
region posterior to the eyes, viewed with nomarski optics (A,C,E)
and lateral view of ammonia-induced fluorescence in the trunk under
UV light epi-illumination (B,D,F). Wild type (A,B) and homozygous
mutant larva of pfetq211 (C,D) and saltl241 (E,F). M, melanophores; X
and arrow, xanthophores.
Fig. 2. Adult fish that are heterozygous or homozygous for salz or
pfeffer show pigment pattern defects. (A) Wild-type, (B)
heterozygous pfetg17 and (C) homozygous saltt254a fish in a lateral
view.
Fig. 3. Xanthophore pigment is reduced in edison and brie. Day 6
larva, dorsal view of the head region posterior to the eyes, viewed
with nomarski optics (A,C,E) and lateral view of ammonia-induced
fluorescence in the trunk under UV light epi-illumination (B,D,F).
Wild type (A,B) and homozygous mutant larvae of edito255b (C),
editc245c (D), and britj226a (E,F). M, melanophores; X and arrow,
xanthophores.

395Zebrafish xanthophore mutants
normal. Homozygous yob fish do show a seemingly unrelated
adult phenotype; the iris in the eye is small (Haffter, personal
communication).
Maternal effect phenotype of
yob
Females homozygous for yob produce embryos that develop
more slowly than embryos derived from heterozygous females
or from females homozygous for any of the other viable xan-
thophore mutations. By day 2, head and tail truncations of
variable strengths are observed. The strength of this maternal
effect phenotype varies amongst the different alleles. A slow
epiboly phenotype was found for embryos derived from a
female homozygous for the strongest allele yobtk13 (data not
shown). In embryos derived from a female with the inter-
mediate allele yobty44d, epiboly is normal but the antero-
posterior extension is reduced, whereas the axis is broadened
laterally (Fig. 5B,D). This leads to a similar but slightly weaker
phenotype (Fig. 5F,H) than that produced by the strongest
allele described above. In many embryos the eyes are fused in
front of the head and the tail is completely missing (Fig. 5F).
In the case of the weakest allele, yobtc251, some embryos
derived from homozygous females are indistinguishable from
wild type. The strength of the embryonic phenotype is deter-
mined by the mother and is independent of the zygotic
genotype (data not shown), indicating that the zygotic function
of yob is restricted to xanthophore development.
esr
,
til
and
tof
show defects in xanthophore
pigmentation and in the retinotectal axon
pathfinding
Overall xanthophore pigmentation is reduced in mutant larvae
of esrom (esr), tilsit (til) and tofu (tof) after 5 days of devel-
opment, although distinct yellow cells are apparent (shown for
til in Fig. 6C). Xanthophores appear more compact and less
granular compared to wild type (arrow in Fig. 6C). Under UV
light, the characteristic fluorescent spot at the center of each
xanthophore appears brighter and more compact in larvae
mutant for each of these three genes than in wild type, whereas
the dendritic extensions of the xanthophores appear more
spread out in esr and til larvae (shown for til in Fig. 6D).
In addition to the pale xanthophore pigmentation phenotype,
mutant esr, til and tof larvae show a defect in retinotectal axon
pathfinding (Karlstrom et al., 1996). The occurrence of both
phenotypes in esr, til and tof mutants suggests that these genes
Fig. 4. Mutant edison embryos show strong autofluorescence under
UV-light illumination. Embryos at the pharyngula period derived
from two heterozygous carriers for editk232. Homozygous mutant edi
embryos show strong blue fluorescence in the entire embryo,
whereas wild-type siblings show only background levels.
Fig. 5. Embryos derived from homozygous mutant yobty44d females
show a maternal effect phenotype. Lateral view of (A) a wild-type
embryo and (B) an embryo derived from a yob homozygous female
at the 5-somite stage, showing the reduced length of the body.
(C) Dorsal view of a 10-somite stage wild-type embryo and (D) a 10-
somite stage embryo derived from a yob homozygous female
showing the wider axis. Lateral view of a wild-type embryo (E) and
an embryo derived from a yob homozygous female (F) at the
pharyngula period, showing the head and tail defects. Lateral view of
a wild-type embryo (G) and an embryo derived from a yob
homozygous female (H) at the hatching period.
Fig. 6. In tilsit mutants xanthophore pigment appears condensed
whereas the cells appear more spread out. Dorsal view of day-6 larva
in the head region posterior to the eyes, viewed with Nomarski optics
(A,C), and lateral view of ammonia-induced fluorescence in the
trunk under UV-light epi-illumination (B,D). (A,B) Wild type and
(C,D) homozygous mutant larva of tilty130b. M, melanophores; X,
and arrow, xanthophores.

396
may encode components of the cytoskeleton required for both
pigment cell morphology and axonal pathfinding of retinal
axons.
Mutants with other xanthophore defects
Mutations in two genes, kefir (kef) and feta (fet), result in
changed colours of the xanthophores. In 5-day-old kef
embryos, whitish cells with faint granularity appear through-
out the xanthophore regions, whereas in fet mutant embryos
brownish cells can be detected in these positions (data not
shown). In quark (qua) mutant embryos the granularity of the
xanthophores is strongly reduced, whereas mutations in ricotta
(ric) and bressot (bst) lead to reduced or no xanthophore pig-
mentation.
In addition to the xanthophore pigmentation phenotype,
further defects are detectable in non blond (nob) and clorix
(clx) mutant larvae. Mutations in both genes lead to small
otoliths (Whitfield et al., 1996). Whereas nob larvae are
generally retarded, clx mutant embryos show an additional
motility (Granato et al., 1996) and melanophore pigmentation
defect.
DISCUSSION
Genes required for xanthophore development
In our screen for mutations with morphologically visible
embryonic and larval phenotypes, we identified 17 genes
required for larval xanthophore pigmentation, but without any
effect on melanophores and iridophores. Multiple alleles were
isolated for nine genes, ranging from two alleles (kef) to 14
alleles (esr). For eight genes only a single allele was isolated
for each, leading to an average allele frequency of 3.6. From
the average allele frequency of this phenotypic class we
conclude that we probably identified most of the genes required
specifically for xanthophore pigmentation in our screen. We
might have missed additional alleles of feta, tilsit, and yocca,
because the phenotypes of the single alleles are very weak.
Pigment pattern formation is different in the larva
and in the adult
One surprising result of this study is the finding that most
mutants with strongly reduced xanthophore pigmentation in
the larva show no obvious xanthophore phenotype as adults (as
observed under a dissecting microscope), indicating that larval
and adult xanthophore pigmentation in the zebrafish require
different genes. Mutations in only three genes (salz, pfeffer and
tartar) cause adult phenotypes. Whereas xanthophores and
melanophores are pale in tar homozygous adults, xanthophores
are not detectable in sal and pfe homozygous adults.
Larval xanthophores in fish occupy the zones not occupied
by melanophores (Epperlein and Claviez, 1982b). In sandy
mutant larvae unpigmented melanophores form unpigmented
gaps between pigmented xanthophores in the dorsal head
(sandy in class VI.A, Fig. 6 in Kelsh et al., 1996). However, if
the number of melanophores is reduced, as in sparse mutant
larvae, xanthophores fill the vacant positions of the
melanophores (sparse in class III, Fig. 3 in Kelsh et al., 1996).
The observation of unpigmented spaces between melanophores
and iridophores in sal and pfe larvae indicates that in the wild-
type situation xanthophores fill the gaps between the
melanophore and iridophore stripes.
Whereas xanthophores are apparently not required for the
proper formation of the larval melanophore stripes, they have
an important function in adult pigment pattern formation.
Partial absence of xanthophores in sal and pfe heterozygous
adult fish leads to irregular blue-black stripes, and a ‘salt and
pepper’ pattern of melanophore pigmentation fading out
towards the tail. A similar but stronger phenotype can be
observed in homozygous mutant fish. The correlation of a
reduced number of xanthophores with an irregular
melanophore pattern in the adult is in agreement with earlier
findings, which indicated that the xanthophores play an
important role in pigment pattern formation in the body
(Epperlein and Claviez, 1982a; Epperlein and Löfberg, 1984)
and in the anal fin (Goodrich and Nichols, 1931; Goodrich et
al., 1954). The correlation between xanthophore and
melanophore pattern can also be seen in adult zebrafish that are
mutant for leopard, where the number of xanthophores in the
anal fin is reduced and the melanophore pigment pattern is
disrupted as well (Kirschbaum et al., 1975; Haffter, personal
communication). A similar influence on the adult melanophore
pigment pattern by iridophores has been found (Johnson et al.,
1995; Haffter, personal communication). In mutants where the
number of iridophores is reduced (transparent, rose, shady),
the melanophore pattern in the adult body is disrupted. Again
this influence is restricted to the adult pigment pattern and has
no effect on larval pigment pattern formation. At present it is
unclear whether xanthophores have a function in the initiation
or in the maintenance of the melanophore stripes. A more
J. Odenthal and others
Table 3. Pteridine fluorescence in mutant larvae at day 6
Colour Mutant larvae
UV light Visible light RFwt yoc bri yob edi pfe sal esr tof fet qua
1) Orange No 0.75 − − − − − − − − − − −
2) Light blue No 0.52 −+ + + − − − − − − −
3) Orange Yellow 0.48 + + + + + + + + + + +
4) Light green No 0.45 −+ (b) + (b) + (b) − − − − − − −
5) Light green No 0.24 −+ + + − − − − − − −
6) Dark blue No 0.19 + + − − + + + + + + +
7) Orange Yellow 0.11 + + + + + + + + + + +
8) Dark blue No 0.06 + − − − + + + + + + +
RFis the distance migrated by the substance divided by the distance migrated by the solvent.
The intensity of the bands was divided into two classes, strong (+) and weak (−). (b) indicates a blue-coloured instead of a green-coloured band. The alleles
used were: yoc (tm86), bri (tj226a), yob (ty44d), edi (to255b), pfe (tg17), sal (tt254a), esr (th222), tof (tq213c), fet (ty107) and qua (tk236).

397Zebrafish xanthophore mutants
detailed analysis of the transition from the larval to the adult
pigment pattern in these mutants is required to address this
point.
Pteridine composition is different in
bri
,
yob
and
yoc
Mutations in three genes (edi, bri and qua) lead to the complete
loss of yellow colour, whereas in all other xanthophore mutants
the yellow colour is only reduced. It has been previously shown
that xanthophores contain different kinds of pteridine pigments
(Matsumoto and Obika, 1968; reviewed by Bagnara, 1966).
They appear almost simultaneously with the differentiation of
xanthophores (Kamei-Takeuchi and Hama, 1971). Specific sets
of pteridines are described for many species. Most frequently
xanthophores contain sepiapterine (visible yellow), drosopter-
ine (visible red) and colourless pteridines (biopterin).
As a first step towards an analysis of the pteridine bio-
chemistry, we performed thin-layer chromatography. We iden-
tified eight different bands that are visible under UV light. Two
of these bands are visibly yellow and are therefore probably
sepiapterines, whereas the others are colourless under normal
illumination; no red drosopterins were found in this analysis
(Matsumoto and Obika, 1968; reviewed by Bagnara, 1966). A
detailed analysis will be necessary to determine which of the
bands correspond to known pteridines. Mutations in three
genes, bri, yob and yoc, produce similar differences in
pteridine composition compared to wild type and may encode
enzymes required to produce specific pteridines. Further chro-
matographical analysis of wild-type and mutant larvae is
required to resolve the pathway for pteridine synthesis.
Mutations in two genes, edison and tartar, lead to blue flu-
orescence in the yolk and the entire embryo. We propose that
edi and tar encode enzymes required for early steps in pteridine
synthesis. The lack of edi and tar function in mutant embryos
leads to the accumulation of fluorescent intermediates that, in
the case of edi, seem to be toxic for the larvae. This phenotype
would predict a change in pteridine composition. Surprisingly,
we have not yet found a difference in the pteridine pattern for
edi using chromatography. A more detailed analysis needs to
be carried out to allow further characterization of the bio-
chemical defects in these mutants.
Yobo
has similarities to the
Drosophila
mutation
deep orange
The yob gene product has at least two biological functions
during development. The first function is in pteridine synthesis,
since pteridine composition is altered in yob mutant embryos.
The second function, provided maternally, is required in the
embryo for normal early development. Several genes with a
similar combination of zygotic and maternal requirement are
known in Drosophila. For example, mutations in the deep
orange (dor) gene lead to both a maternal effect phenotype and
a change in eye colour. The phenotype is accompanied by alter-
ations in pteridine composition (Counce, 1957). The strength
of the embryonic yob phenotype is independent of the zygotic
genotype, indicating that zygotic yob function is not required
during early development. This is in contrast to dor, for which
expression is also required zygotically for embryonic develop-
ment.
Xanthophores and retinotectal projection
Three mutants (esr, til, tof) were isolated with defects in xan-
thophore pigmentation and retinotectal axon pathfinding.
Yellow pigmentation is concentrated towards the cell center in
mutants of all three genes and the morphology of the xan-
thophores under UV light is abnormal. The distribution of the
coloured organelles within the pigment cell is important for the
general apparence of the body colour. Dispersed organelles
lead to a strong colour, whereas pigment cells with aggregated
organelles are weakly coloured (reviewed by Luby-Phelps and
Schliwa, 1982; Obika, M., 1986). The pigmentation phenotype
in esr, til and tof could be explained by a defect in pigment
translocation. Retinotectal axons stop prematurely on their way
to the tectum and show mapping defects if they reach the
tectum (Karlstrom et al., 1996). The phenotypes observed in
seemingly unrelated cell types suggests that esr, til and tof may
encode components of the cytoskeleton. It further suggests that
these genes are required in a common pathway for both
pigment cell morphology and axonal pathfinding of retinal
axons. This hypothesis will have to be tested by a careful
analysis of the cytoskeleton in the xanthophores and the retinal
axons.
In summary, the collection of mutants described here and by
Kelsh et al. (1996) will be helpful tools to gain insight into
different processes involved in vertebrate pigmentation.
We thank Silke Rudolph for technical assistance, Joel Wilson and
Cornelia Fricke for their help in the fish work and the thin layer chro-
matography, and Darren Gilmour for help with the manuscript.
REFERENCES
Bagnara, J. T. and Stackhouse, H. L. (1961). Purine components of
guanophores in amphibians. Anat. Rec. 139, 292.
Bagnara, J. T. (1966). Cytology and cytophysiology of non-melanophore
pigment cells. Intern. Rev. Cytol. 20, 173-205.
Bagnara, J. T. (1976). Colour change. In Physiology of the Amphibia, Vol. 3
(ed. B. Lofts), pp. 1-52. New York: Academic Press.
Bagnara, J. T., Matsumoto, J., Ferris, W., Frost, S. K., Turner, W. A., Jr.,
Tchen, T. T. and Taylor, J. D. (1979). Common origin of pigment cells.
Science 203, 410-415.
Birbeck, M. S. C. (1963). Electron microscopy of melanocytes: the fine
structure of hair-bulb premelanophores. Ann. New York Acad. Sci. 100, 540.
Charles, A. and Ingram, J. T. (1959). Electron microscope observations of the
melanocyte of the human epidermis. J. Biophys. Biochem. Cytol. 6, 41-44.
Counce, S. J. (1957). A female-sterile mutant (Deep orange) of Drosophila
melanogaster: Increasing isoxanthopterine content. Experientia 13, 354-356.
Drochmans, P. (1960). Electron microscope studies of epidermal melanocytes,
and the fine structure of melanin granules. J. Biophys. Biochem. Cytol. 8,
165-180.
Droin, A. (1992). Genetic and experimental studies on a new pigment mutant in
Xenopus laevis. J. Exp. Zool. 264, 196-205.
DuShane, G. P. (1934). The origin of pigment cells in amphibia. Science 80,
620-621.
Epperlein, H. H. and Claviez, M. (1982a). Changes in the distribution of
melanophores and xanthophores in Triturus alpestris embryos during their
transition from the uniform to banded pattern. Wilhelm Roux’s Arch. Dev.
Biol. 192, 5-18.
Epperlein, H. H. and Claviez, M. (1982b). Formation of pigment cell patterns
in Triturus alpestris embryos. Dev. Biol. 91, 497-502.
Epperlein, H. H. and Löfberg, J. (1984). Xanthophores in chromatophore
groups of the premigratory neural crest initiate the pigment pattern of the
axolotl larva. Wilhelm Roux’s Arch. Dev. Biol. 193, 357-369.
Frost, S. K. and Malacinski G. M. (1980). The developmental genetics of
pigment mutants in the mexican axolotl. Dev. Genet. 1, 271-294.
Frost, S. K., Ellinger, M. S. and Murphy, J. A. (1982). The pale mutation in
Bombina orientalis: Effects on melanophores and xanthophores. J. Exp.
Zool. 221, 125-129.
Goodrich, H. B. and Nichols, R. (1931). The development and the

398
regeneration of the colour pattern in Brachidanio rerio. J. Morph. Physiol.
52, 513-518.
Goodrich, H. B., Marzullo, C. M. and Brownson, W. H. (1954). An analysis
of the formation of colour patterns in two fresh-water fish. J. Exp. Zool. 125,
487-505.
Granato, M., van Eeden, F. J. M., Schach, U., Trowe, T., Brand, M.,
Furutani-Seiki, M., Haffter, P., Hammerschmidt, M., Heisenberg, C.-P.,
Jiang, Y.-J., Kane, D. A., Kelsh, R. N., Mullins, M. C., Odenthal, J. and
Nüsslein-Volhard, C. (1996). Genes controlling and mediating locomotion
behavior of the zebrafish embryo and larva. Development 123, 399-413.
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.
Hama, T. (1963). The relation between the chromatophores and pterin
compounds. Ann. New York Acad. Sci. 100, 977-986.
Johnson, S. L., Africa, D., Walker, C. and Weston, J. A. (1995). Genetic
control of adult pigment stripe development in zebrafish. Dev. Biol. 167, 27-
33.
Kamei-Takeuchi, I. and Hama, T. (1971). Structural changes of pterinosome
(pteridine pigment granule) during the xanthophore differentiation of
Oryzias fish J. Ultrastruct. Res. 34, 452-463.
Karlstrom, R. O., Trowe, T., Klostermann, S., Baier, H., Brand, M.,
Crawford, A. D., Grunewald, B., Haffter, P., Hoffmann, H., Meyer, S.
U., Müller, B. K., Richter, S., van Eeden, F. J. M., Nüsslein-Volhard, C.
and Bonhoeffer, F. (1996). Zebrafish mutations affecting retinotectal axon
pathfinding. Development 123, 415-426.
Kelsh, R. N., Brand, M., Jiang, Y.-J., Heisenberg, C.-P., Lin, S., Haffter, P.,
Odenthal, J., Mullins, M. C., van Eeden, F. J. M., Furutani-Seiki, M.,
Granato, M., Hammerschmidt, M., Kane, D. A., Warga, R. M., Beuchle,
D., Vogelsang, L. and Nüsslein-Volhard, C. (1996). Zebrafish
pigmentation mutations and the processes of neural crest development.
Development 123, 369-389.
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.
Kirschbaum, F. (1975). Untersuchungen über das Farbenmuster der
Zebrabarbe Brachidanio rerio (Cyprinidae, Teleostei). Wilhelm Roux’s
Arch. Dev. Biol. 177, 129-152.
Luby-Phelps, K. J. and Schliwa, M. (1982). Pigment migration in
chromatophores: A model system for intracellular particle transport. In
Axoplasmic Transport, pp. 15-26. Berlin, Heidelberg: Springer-Verlag.
Matsumoto, J., Kalishima, T. and Hama, T. (1960). Relation between the
pigmentation and pterin derivatives of chromatophores during development
in the normal black and transparent scaled types of goldfish (Carassius
auratus). Genetics 45, 1177-1189.
Matsumoto, J. (1965a). Studies on fine structure and cytochemical properties
of erythrophores in swordtail, Xiphophorus helleri, with special reference to
their pigment granules (Pterinosomes). J. Cell Biol. 27, 493-504.
Matsumoto, J. (1965b). Role of pteridines in the pigmentation of
chromatophores in cyprinid fish, Japan. J. Zool. 14, 45-94.
Matsumoto, J. and Obika, M. (1968). Morphological and biochemical
characterization of goldfish erythrophores and their pterinosomes. J. Cell
Biol. 39, 233-250.
Mullins, M. C. and Nüsslein-Volhard, C. (1993). Mutational approaches to
study embryonic pattern formation in the zebrafish. Curr. Biol. 3, 648-654.
Mullins, M. C., Hammerschmidt, M., Haffter, P. and Nüsslein-Volhard, C.
(1994). Large-scale mutagenesis in the zebrafish: In search of genes
controlling development in a vertebrate. Curr. Biol. 4, 189-202.
Obika, M. (1963). Association of pteridines with amphibian larva
pigmentation and their biosynthesis in developing chromatophores. Dev.
Biol. 6, 99-112.
Obika, M. (1986). Intracellular transport of pigment granules in fish
chromatophores. Zool. Sci. 3, 1-11.
Obika, M. (1993). Formation of pterinosomes and carotinoid granules in
xanthophores of the teleost Oryzias latipes as revealed by the rapid-freezing
and freeze-substitution method. Cell Tissue Res. 271, 81-86.
Rawles, M. E. (1944). The migration of melanoblasts after hatching into
pigment-free skin grafts of the common fowl. Pysiol. Zool. 17, 167-183.
Taylor, J. D. (1992). Does the introduction of a new player, the endoplasmatic
reticulum, create more or less confusion in understanding the mechanism(s)
of pigmentary organelle translocation. Pigment Cell Research 5, 49-57.
Whitfield, T. T., Granato, M., van Eeden, F. J. M., Schach, U., Brand, M.,
Furutani-Seiki, M., Haffter, P., Hammerschmidt, M., Heisenberg, C.-P.,
Jiang, Y.-J., Kane, D. A., Kelsh, R. N., Mullins, M. C., Odenthal, J. and
Nüsslein-Volhard, C. (1996). Mutations affecting development of the
zebrafish inner ear and lateral line. Development 123, 241-254.
Ziegler, I. (1965). Pterine als Wirkstoffe und Pigmente. Ergeb. Physiol.
Biochem. Exp. Pharmakol. 56, 1-66.
(Accepted 9 April 1996)
J. Odenthal and others