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doi: 10.1242/10.1242/dev.00307
Section of Integrative Biology, Section of Molecular, Cell and Developmental Biology, Institute for Cellular and Molecular Biology, University of Texas at Austin, 1 University Station C0930, Austin, TX 78712, USA
* Author for correspondence (e-mail: dparichy{at}mail.utexas.edu)
Accepted 25 November 2002
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SUMMARY |
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 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key words: Melanophore, Xnathophore, Pigment pattern, Zebrafish, fms, Csf1r, Neural crest
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INTRODUCTION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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;
Twitty, 1936;
Jackson, 1994). In amniotes,
coat and plumage patterns result largely from the spatial and temporal pattern
of melanocyte differentiation and the transfer of melanin to developing hair
or feathers, respectively (Nordlund et
al., 1998). By contrast, pigment patterns of ectothermic
vertebrates such as fishes and amphibians largely reflect the spatial
arrangements of several classes of pigment cells, or chromatophores, that
retain their pigments intracellularly
(Bagnara, 1998). These include
black melanophores, yellow or orange xanthophores and iridescent iridophores.
The combinations of these and other chromatophore classes generate an
extraordinary diversity of pigment patterns that serve a variety of roles
across species, from crypsis and predator avoidance to schooling and mate
recognition (e.g., Keenleyside,
1955; Endler, 1987;
Houde, 1997;
Couldridge and Alexander,
2002).
Fishes of the genus Danio exhibit a range of pigment patterns
including horizontal stripes and vertical bars, as well as mottled and uniform
patterns (Fang, 1997;
Fang, 1998;
Fang, 2000;
Quigley and Parichy, 2002).
Some insights into the mechanisms underlying these patterns can be gained by
analyzing wild-type and mutant zebrafish, D. rerio
(Parichy and Johnson, 2001).
During normal development, zebrafish develop an embryonic and early larval
pigment pattern comprising several stripes of melanophores with widely
scattered xanthophores. This pattern persists for about 2 weeks, at which time
an adult pattern begins to form. Over the following 2-3 weeks, a
juvenile/early adult pattern develops consisting initially of two dark
`primary' melanophore stripes with an intervening light stripe. Subsequently,
additional `secondary' melanophore stripes are added as the fish grows
(Fig. 1A)
(Goodrich and Nichols, 1931;
Kirschbaum, 1975;
Johnson et al., 1995;
Parichy et al., 2000a;
Parichy and Johnson, 2001).
Dark stripes comprise principally melanophores and iridophores, though
occasional xanthophores can be found within these stripes as well
(Fig. 1C,E). Light stripes lack
melanophores and include only xanthophores and iridophores.
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Of particular interest for understanding pattern-forming mechanisms and
their evolution are mutants that alter the normal adult striped pattern,
either by perturbing the development of particular classes of chromatophores
or by affecting the extracellular environment in which these cells reside.
Molecular analyses identified one such mutant (previously, panther)
as an orthologue of the fms (Csf1r) locus
(Parichy et al., 2000a).
fms encodes a type III receptor tyrosine kinase known previously in
mammals for roles in reproduction as well as the development of macrophages
and osteoclasts (e.g., Marks and Lane,
1976; Motoyoshi,
1998; Dai et al.,
2002; Tagoh et al.,
2002). In zebrafish, fms- mutants exhibit a
normal pattern of embryonic and early larval melanophores. In contrast, the
pattern of adult melanophore stripes is severely disrupted
(Fig. 1B). This defect is
associated with fewer melanophores, disorganized melanophore movements, and
both increased and disorganized melanophore death [which occurs normally in
developing interstripe regions (Parichy et
al., 2000a)].
Previous analyses suggest that fms and its homologue,
kit, promote the development of temporally distinct populations of
adult stripe melanophores in zebrafish. kit is essential for the
migration and maintenance of melanophores and their precursors, and
kit mutants lack early metamorphic melanophores that normally arise
in a dispersed pattern over the flank and ultimately migrate into stripes
(Johnson et al., 1995;
Parichy et al., 1999) (see
also Rawls and Johnson, 2001;
Quigley and Parichy, 2002). In
contrast, fms- mutants retain these cells and instead
exhibit an increasingly severe deficit of stripe melanophores beginning during
middle metamorphic stages and extending into late metamorphosis and adult life
(Parichy et al., 2000a). These
observations suggested that fms promotes the development of a
late-appearing metamorphic melanophore population, distinct from an
early-appearing metamorphic melanophore population that requires kit.
Consistent with this model, fish doubly mutant for both fms and
kit exhibit additive effects of the two mutations and lack nearly all
melanophores. A direct role for fms in establishing or maintaining
melanophore precursors was also suggested by the observation that many cells
migrating from the embryonic neural crest express both fms and either
mitfa or endothelin receptor b1 (ednrb1), two genes
associated with melanophore development [though all of these loci are also
co-expressed at early stages with xanthophore lineage markers, suggesting
these cells may not yet be specified for one or another cell lineage (for
details, see Quigley and Parichy,
2002)].
Despite genetic evidence for a fms-dependent population of adult
melanophores, fms is not detectably expressed in melanophores at the
time when the migration and survival of these cells differs between wild-type
and fms- mutants (though both melanophores and
melanoblasts express kit; Fig.
1F,G) (Parichy et al.,
1999; Parichy et al.,
2000a). Thus, it remains unclear how, or when, fms
promotes melanophore development and adult stripe formation. A possible
explanation comes from the finding that, in contrast to the situation in
melanophores or late stage melanoblasts, fms is expressed in yellow
xanthophores and their precursors, xanthoblasts. Moreover,
fms- mutants lack xanthophores and xanthoblasts in both
embryos and adults, indicating an essential role for fms in the
development of the xanthophore lineage
(Fig. 1D)
(Parichy et al., 2000a). In
turn, these and other observations (e.g.
Goodrich et al., 1954;
Goodrich and Greene, 1959)
suggested a model in which fms (i) acts directly to promote the
establishment or maintenance of a subpopulation of stripe melanophore early
precursors (above); and (ii) also acts indirectly through the xanthophore
lineage to promote melanophore morphogenesis. Thus, interactions between
melanophores and fms-dependent xanthophores were hypothesized to
contribute to adult stripe formation. Such interactions could have broad
phylogenetic significance, as alternating patterns of melanophores and
xanthophores are found in a diverse array of ectothermic vertebrates,
including fishes, frogs, salamanders and reptiles (e.g.
Brodie, 1992;
Seehausen et al., 1999;
Parichy, 2001a;
Parichy, 2001b).
In this paper, we investigate how and when fms activity is essential for the generation of an adult pigment pattern. Using cell transplantations, we show that fms acts through the xanthophore lineage to promote the arrangement of melanophores into stripes. To assess when fms is required for xanthophore development and melanophore stripe formation, we isolated temperature sensitive fms alleles and used reciprocal temperature shift experiments to enhance or curtail Fms activity at a range of stages from embryo to adult. These analyses show that fms is essential throughout development for maintaining cells of the xanthophore lineage and also for maintaining the integrity of melanophore stripes. Our results further suggest that fms is not required for establishing a population of precursor stem cells during embryogenesis (though it may serve to maintain precursors once established); rather, Fms activity is essential for recruiting precursor cells to the xanthophore lineage during later post-embryonic and adult development.
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MATERIALS AND METHODS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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;
Lister et al., 1999).
fms1 and fmsblue are recessive
homozygous viable, and are each predicted to encode proteins with
substitutions in the functionally important and phylogenetically conserved
kinase domains; these are likely to be null or severe loss of function alleles
(Parichy et al., 2000a).
nacrew2 is recessive homozygous viable and acts as a null
allele (Lister et al., 1999).
Fish transgenic for green fluorescent protein (GFP) under the control of the
ß-actin promoter were generously provided by K. Poss and have been
maintained in the ABUT genetic background.
Cell transplantation
Cell transplantations were performed on 3.3- to 3.8-hour embryos using a
Narishige IM-9B micrometer-driven microinjection apparatus mounted on a
Narishige micromanipulator. For operations, embryos were placed in wells
formed in agar-lined dishes containing 10% Hanks solution
(Westerfield, 1993) plus 1%
penicillin/streptomycin. Typically 50-100 blastomeres were transplanted per
embryo. To identify donor cells in host backgrounds, we used donors that were
homozygous transgenic for GFP under the control of the ß-actin promoter
for wild-type 
fms-, fms-
wild-type, and nacre- fms-
chimeras. To identify donor melanophores in wild-type
nacre- and fms-
nacre- chimeras, we used the endogenous melanin of donor
melanophores as an autonomous marker of cell lineage, as
nacre- hosts are unable to produce melanophores owing to a
mutation that acts autonomously to the melanophore lineage
(Lister et al., 1999) see
below); donor cell types other than melanophores were not assessed in these
chimeras. For transplants involving ß-actin-GFP transgenic donors, we
identified GFP+ donor cells under epifluorescent illumination using
an EGFP filter set on a Zeiss Axioplan 2i microscope. Although xanthophores
autofluoresce green as well (Raible and
Eisen, 1994; Parichy et al.,
2000a), GFP fluorescence is markedly brighter and color-shifted
relative to endogenous xanthophore autofluorescence. To prevent melanin from
quenching GFP fluorescence in melanophores, we treated fish with 2-3 drops of
10 mg/ml epinephrine prior to viewing, thereby causing melanin-containing
melanosomes to be contracted towards cell centers; GFP fluorescence could then
be clearly observed in cell peripheries. Individuals completely lacking
GFP+ cells, or comprising greater than 
75% GFP+
cells were discarded and are not included in the analyses below; typically,
however, chimeras exhibited relatively low percentages of donor cells
(<25%) that were often distributed widely in the adult fish with patches of
donor cells consisting of only one or a few cell lineages (see below). Cell
transplants employing both fms1 and
fmsblue yielded equivalent results; thus both genotypes
are referred to below as fms-.
Mutagenesis and non-complementation screening
To isolate temperature-sensitive alleles of fms, we crossed
homozygous fms1 or fmsblue females to
ABUT males that had been mutagenized three times over the course of
three weeks with 3 mM N-ethyl-N-nitrosourea (Sigma)
(Solnica-Krezel et al., 1994).
Embryos were incubated at 33°C until hatching at which time individuals
lacking xanthophores were transferred to 28.5°C and reared through sexual
maturity. To test whether newly isolated mutants were allelic to fms,
and to test for temperature sensitivity, mutant fish were backcrossed to
fms1 or fmsblue and sibships were
split between 24°C and 33°C. Mutants were considered alleles of
fms if at 33°C none of the offspring developed wild-type
phenotypes; mutants were identified as temperature-sensitive if at 24°C
approximately half of the offspring developed fms null phenotypes
(presumptive fms-/fms-) and half of the
offspring developed less severe or wild-type phenotypes (presumptive
fmsTS/fms-). At least 100 offspring
were examined at each temperature.
Sequencing and genotyping
Sequencing of mutant alleles was performed following RT-PCR of fms
cDNA from haploid embryos. Sequencing reactions were performed with BigDye dye
terminator sequencing chemistry and resolved on an ABI-377 automated
sequencer. Resulting sequences were compared to those of unmutagenized
ABUT harboring the ancestral unmutagenized chromosome and wild-type
fms sequence. For primer extension genotyping of
fmsut.r4e174A, forward and reverse primers (fms174a-F:
TCGAGTTCTCTTTGTTTCTCCGAG; fms174a-R: CTCCGATTCTAGCGCAGCAAATG) flanking the
mutant lesion were used to amplify genomic DNA and excess primers were
digested using shrimp alkaline phosphatase and exonuclease (Amersham). Primer
extension reactions were performed in 20 µl volumes with 10 µl PCR, 0.5
U Thermosequenase (Amersham), 50 µM ddGTP, 50 µM dATP, 50 µM dTTP,
12.5 pmol fms174a-R and supplied reaction buffer. After denaturing (94°C,
2 minutes) reactions were run for 50 cycles at: 94°C, 5 seconds; 43°C,
15 seconds; 60°C, 1 minute
(Hoogendoorn et al., 1999).
After denaturing (94°C, 1 minute), primer extension products were resolved
by HPLC on a Transgenomic WAVE DNA Fragment Analysis System at 80°C.
Wild-type, fms1, or fmsblue alleles
result in addition of 3 nucleotides (ddGAT) whereas the
fmsut.r4e174A allele results in addition of 2 nucleotides
(ddGT) to the extension primer.
Histology
mRNA in situ hybridization employed riboprobes for fms as well as
GTP-cyclohydrolase (gch) and xanthine dehydrogenase
(xdh), which have been described previously
(Parichy et al., 2000a). mRNA
in situ hybridization of embryos followed the method of Jowett and Yan
(Jowett and Yan, 1996).
To test for apoptosis, TUNEL assays were performed as described
(Zhang and Galileo, 1997;
Parichy et al., 1999). After
TUNEL staining, embryos were examined in whole mounts and numbers of
TUNEL+ cells were counted along the dorsal neural tube and in
neural crest migratory pathways from just anterior to the midbrain-hindbrain
junction to a point one-third of the distance along the tail (extensive
genotype and temperature-independent cell death in posterior tail tips
precluded accurate counts in this region). Stained cells ventral to the
ventral margins of the myotomes (and likely to include macrophages rather than
neural crest-derived cells) (Herbomel et
al., 2001) were not included in counts. Embryos were fixed and
examined in whole mount. To identify dead chromatophores in adults, fish were
fixed in 4% paraformaldehyde in PBS. After washing in PBS, fins were removed
and mounted on glass slides and trunks were embedded in OCT and
cryosectioned.
Morphometrics, image analyses and statistical methods
Fish sizes were quantified by measuring the length from the tip of the
snout to the caudal peduncle (standard length, SL), as well as the height of
the flank at the anterior and posterior margins of the anal fin. To quantify
patterns, fish were imaged using a combination of incident and transmitted
illumination with a Zeiss Axiocam digital camera mounted on either an Olympus
SZX-12 epifluorescence stereomicroscope, or a Zeiss Axioplan 2i equipped with
differential interference contrast optics. Both xanthophores and melanophores
could be readily identified under these conditions. To allow accurate cell
counts in temperature shift experiments, fish were treated with epinephrine
(above) prior to imaging, and images were acquired using the square root
transformation and color enhancement feature in Zeiss Axiovision 3.0.
Xanthophore and melanophore densities were calculated for a representative
region of the flank bordered by: anteriorly, the anterior margin of the dorsal
fin; posteriorly, the posterior margin of the anal fin; ventrally, the ventral
margin of the flank; and dorsally, a position just ventral to the dorsal
margin of the flank (this position was determined by estimating the total
height of the dorsal flank at the posterior margin of the anal fin, then
selecting a point 10% of this distance from the dorsal edge; this restriction
reduced variation among individuals owing to curvature of the fish in this
region). Pigment cells in Adobe Photoshop images were identified by eye and
marked digitally; the IPTK 4.0 software package (Reindeer Graphics) was then
used to automatically count marks and to calculate total measurement areas.
For imaging of some whole-mount embryos following in situ hybridizations, the
Extended Focus module of Zeiss Axiovision 3.0 software was employed to flatten
multiple focal planes into a single plane (which results in characteristic
fringing at the edges of some features, such as melanophores).
All statistical analyses were performed using standard methods
(Sokal and Rohlf, 1981) in the
JMP Statistical Analysis Package for Macintosh (SAS Institute, Cary, NC).
Analyses of xanthophore and melanophore numbers and distributions were
performed by treating size at temperature shift as both a continuous and
ordinal factor and after controlling for variation in individual size at the
time of imaging. Analyses of stripe breaks were done using maximum likelihood
estimation and significance of effects were estimated using likelihood ratio
tests. Nearest neighbor distances among melanophores were assessed initially
as hierarchical mixed model analyses of covariance, with individual
melanophore measurements nested within individuals (a random effect), and
individuals nested within temperature treatments (a fixed effect); this
approach avoids pseudoreplication from analyzing individual melanophores as
independent data points. Methodological details and complete statistical
analyses are available on request.
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RESULTS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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As a first step in testing how fms promotes adult pigment pattern
development, we asked whether fms acts autonomously to pigment cell
lineages during xanthophore development and melanophore stripe formation. In
principle, xanthophore development and melanophore stripe patterning could
depend on several potential sources of fms activity: early (possibly
unspecified) precursor cells that coexpress fms and genes associated
with melanophore development (e.g., mitfa, ednrb1); cells of the
xanthophore lineage that express and require fms and in turn
influence melanophore behavior; or melanophores themselves, if these cells in
fact express fms at functionally significant but undetectable levels.
To address these issues, we constructed chimeras between wild-type and
fms- mutant embryos [for a general discussion of this
approach in the context of analyzing functions of identified genes see Rossant
and Spence (Rossant and Spence,
1998)]. We predicted that if fms acts autonomously to
pigment cell lineages in promoting xanthophore development and melanophore
stripe formation, then donor wild-type fms+ melanophores
and xanthophores should develop in fms- mutant hosts, and
these cells should be capable of forming a wild-type stripe pattern (that
might or might not include host fms- mutant melanophores;
see below). If fms acts non-autonomously to pigment cell lineages,
then donor wild-type fms+ melanophores and xanthophores
might not develop in fms- mutant hosts, stripes might not
develop, or both. To test these predictions, we reared wild-type
fms- (as well as fms-
wild-type) chimeric embryos 8-12 weeks through metamorphosis and adult pigment
pattern formation. To distinguish donor from host cells, we used donors that
expressed GFP ubiquitously under the control of the ß-actin promoter.
For all genotypes examined (Table
1, and below), chimeras reared to adult stages exhibited donor
cells in a variety of derivatives, including muscle, gut, lateral line,
epidermis, dermal bone and endoskeleton, similar to chimeras typically
examined at embryonic stages (e.g. Ho and
Kimmel, 1993; Halpern et al.,
1993). A subset of chimeras exhibited donor-derived pigment cells,
and these frequently occurred in the absence of other donor-derived cell types
(see below).
Results from wild-type 
fms- chimeras suggested
that fms acts autonomously to promote xanthophore development. When
wild-type cells were transplanted to fms- hosts,
donor-derived (fms+ GFP+) melanophores and
xanthophores could be observed on the flank in both embryos and adults
(Fig. 2A,B,E,F;
Table 2). Of the chimeras
examined at adult stages, 40% exhibited donor-derived pigment cells,
including donor xanthophores. Frequently, these cells were distributed widely
over the flank and apparently independently of other donor-derived cell types
(Fig. 2F and data not shown).
The wide distribution of donor-derived melanophores and xanthophores is not
unexpected given the migratory nature, invasiveness, and proliferative
capabilities of these cells [particularly xanthophores (e.g.
Tucker and Erickson, 1986a;
Parichy, 1996a;
Parichy, 1996b;
Wilkie et al., 2002) (D. M.
P., unpublished data)]. Conversely, when fms- cells were
transplanted into wild-type hosts, donor fms- cells
contributed to a range of non-pigment cell derivatives, but only a single
individual (of 15 surviving chimeras) had a few donor
(fms- GFP+) melanophores
(Fig. 2I,J); donor
fms- xanthophores were not observed. Although the paucity
of donor melanophores in fms- wild-type chimeras
raises the possibility of an autonomous role for fms in melanophore
development (consistent with genetic analyses; see Introduction), we cannot
yet exclude the formal possibility that differences in genetic background
unrelated to fms- might have biased the differentiation of
donor cells away from melanophore fates.
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Pigment patterns of wild-type fms- chimeras
further suggested that adult melanophore stripes result from fms
acting autonomously to pigment cell lineages, but non-autonomously relative to
the melanophore lineage. In wild-type fms- chimeras
with donor (fms+ GFP+) melanophores and
xanthophores, stripes developed that frequently resembled wild-type stripes
and were considerably more organized than patterns of fms-
mutants (Fig. 2C; compare with
Fig. 1A,B). The degree of
melanophore stripe organization appeared to depend on the distribution of
donor wild-type (fms+) cells, as individuals with more
donor melanophores and xanthophores had more distinctive stripes than
individuals with few donor melanophores and xanthophores
(Fig. 2D). Similar variation in
stripe morphology could be observed in individuals with donor melanophores and
xanthophores only in discrete regions (Fig.
2G,H). (These qualitative interpretations are supported by
quantitative analyses of temperature shift experiments below.) Importantly,
melanophore stripes that developed in wild-type
fms- chimeras always included host
fms- melanophores interspersed with donor
fms+ melanophores. Moreover, in regions with donor pigment
cells and stripes, host fms- melanophores were not found
in xanthophore-rich interstripe regions
(Fig. 2E,F), suggesting that
wild-type fms+ donor cells were able to organize
fms- mutant host melanophores into stripes. Owing to the
difficulty of examining GFP expression in single cells of adult fish, it was
not possible to quantitatively assess whether wild-type
fms+ melanophores or xanthophores that affected the
arrangement of fms- mutant melanophores also affected the
numbers of fms- mutant melanophores that differentiated.
In reciprocal fms- wild-type chimeras, the only
individual that developed donor fms- melanophores
exhibited these cells only within the melanophore stripes
(Fig. 2I,J). Taken together,
these results suggest that: (i) fms acts autonomously to pigment cell
lineages to promote xanthophore development (and possibly melanophore
development); and (ii) fms acts non-autonomously on the melanophore
lineage to promote the arrangement of both fms+ and
fms- melanophores into stripes.
Xanthophore-autonomous role for fms in promoting melanophore
stripe development revealed by transplants between nacre-
and fms- mutants
Pigment cell distributions in wild-type fms-
chimeras suggested that melanophore stripe development depends in part on
fms acting non-autonomously relative to individual melanophores,
since even fms- mutant melanophores were arranged in
stripes in the presence of fms+ pigment cells. We reasoned
that at least two explanations could account for such non-autonomous effects
on melanophore distributions. First, fms could be expressed at
undetectable but functionally important levels by a subset of melanophores
that, in turn, organize other melanophores into stripes. This model would be
consistent with previous genetic analyses that identified distinct
fms-dependent and kit-dependent melanophores constituting
adult stripes (Parichy et al.,
2000a). Second, fms-dependent cells of the xanthophore
lineage could be essential for promoting the organization of melanophores into
stripes. To distinguish between these models, we constructed chimeras between
fms- mutants and nacre- mutants.
nacre- mutants lack melanophores because of a mutation in
mitfa, which encodes a microphthalmia-like transcription factor that
acts cell autonomously during melanophore specification
(Lister et al., 1999). Since
nacre- mutants retain xanthophores, however, comparisons
of cell distributions between wild-type fms- and
nacre- fms- chimeras allows
the isolation of a potential role for fms in promoting the
organization of melanophore stripes via interactions among melanophores
themselves. Thus, we predicted that if fms acts through the
melanophore lineage to promote the organization of melanophores into stripes,
then nacre- fms- chimeras
should not form melanophores stripes, since fms+
melanophores would not be present. Conversely, if fms acts through
the xanthophore lineage to promote the organization of melanophores into
stripes, then nacre- fms-
chimeras should be capable of developing melanophore stripes, since
fms+ xanthophores would be contributed by the
nacre- mutant background.
As previously, nacre- cells transplanted into
fms- hosts contributed to a range of tissues including
muscle, epidermis, nerves and others. As expected, and consistent with
previous analyses of the nacre- mutation
(Lister et al., 1999), none of
the nacre- fms- chimeras
exhibited donor (fms+ nacre-
GFP+) melanophores. However, 24% of chimeras reared to adult
stages exhibited donor (fms+ nacre-
GFP+) xanthophores and these fish were invariably striped
(Fig. 3A,B). By contrast,
nacre- fms- chimeras that
lacked xanthophores failed to develop distinctive, well-organized melanophore
stripes and instead exhibited melanophore patterns indistinguishable from
fms- mutant controls; this was true of chimeras in which
nacre- donor cells developed as non-pigment cell lineages,
as well as chimeras in which nacre- donor cells developed
as the third major class of pigment cells, iridescent iridophores
(Fig. 3C,D). In reciprocal
experiments, fms- nacre- (as
well as wild-type nacre-) chimeras also developed
regions of well-formed melanophore stripes
(Fig. 3E,F). Together, these
results demonstrate that non-autonomous roles for fms in promoting
melanophore stripe development act largely or entirely via the xanthophore
lineage.
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Isolation of temperature-sensitive fms alleles
As mRNA in situ hybridization reveals fms expression in cells of
the xanthophore lineage throughout embryonic and larval stages, as well as in
embryonic cells expressing markers of multiple pigment cell lineages (see
Introduction), we asked whether critical periods exist for Fms activity in
promoting xanthophore development and adult stripe formation. To this end, we
screened for temperature-sensitive alleles by non-complementation of
fmsblue or fms1. We isolated 75
new alleles and tested 42 for temperature sensitivity. Three alleles
(fmsut.r4e174A, fmsut.r4e536,
fmsut.r4e564) resulted in presumptive fms null
phenotypes (Parichy et al.,
2000a) at a restrictive temperature (33°C) and wild-type
phenotypes at a permissive temperature (24°C) when in trans with
fmsblue or fms1. Sequence analyses of
fmsut.r4e174A (hereafter, fms174A)
identified a TC transversion resulting in an amino acid substitution
(Y185H) within the predicted second immunoglobulin-like domain of the protein,
a region likely to be essential for ligand binding
(Jiang et al., 2000)
(Fig. 4A). Primer extension
assays (Fig. 4B) confirmed
cosegregation of this lesion and the temperature-sensitive phenotype (data not
shown).
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To further evaluate the phenotype of fms174A we reared homozygous individuals at 33°C and 24°C. fms174A homozygotes reared at 33°C completely lacked xanthophores as both larvae and adults, and failed to develop normal adult melanophore stripes (Fig. 4C,D). In contrast, fms174A homozygotes reared at 24°C had approximately wild-type numbers of xanthophores throughout development and formed wild-type adult melanophore stripes (Fig. 4E,F). Given the strong temperature sensitivity of this allele, all subsequent analyses were performed using homozygous fms174A, unless otherwise indicated.
Since xanthophore precursors fail to disperse from the vicinity of the
neural crest in fmsblue and fmse1
embryos (Parichy et al.,
2000a), we asked whether a similar defect was present in
fms174A embryos. In situ hybridizations show that
xanthophore precursors are not found on the flank of
fms174A embryos when reared at 33°C, as evidenced by
an absence of cells expressing fms as well as the xanthophore lineage
markers, gch and xdh
(Fig. 4G-I).
Essential roles for Fms in maintaining xanthophore lineage and
melanophore stripes throughout development
As an initial step in evaluating the temporal requirements for fms
in promoting pigment pattern formation, we asked whether a critical period
exists beyond which xanthophores and melanophore stripes become independent of
Fms activity. Thus, we reared homozygous fms174A mutant
larvae at 24°C and upshifted individuals to 33°C to curtail Fms
activity at a range of stages from embryo to early adult (standard length,
SL=3.5-17.0 mm, n=95). Since the presence or absence of xanthophores
is causally related to adult melanophore stripe formation (above), we
quantified the densities of xanthophores at a stage when an adult stripe
pattern would have been present in wild-type individuals. These analyses
demonstrated that xanthophores were completely eliminated by restricting Fms
activity in individuals through late metamorphic stages (<10 mm SL;
Fig. 5A,
Fig. 6A,B). This corresponds to
the period by which the initial two adult melanophore stripes have formed and
are becoming increasingly regular in their outlines, but prior to the
development of adult scales, or the formation of secondary adult melanophore
stripes (Quigley and Parichy,
2002) (D. M. P. and J. M. T., unpublished data). In contrast, in
larvae upshifted at larger sites, we observed higher densities of residual
xanthophores when formation of primary adult melanophore stripes was
essentially completed and secondary melanophore stripes had started to develop
(Fig. 6C; final xanthophore
density vs. size when shifted, partial regression coefficient=5.13; s.e.=0.46;
F1,92=125.47, P<0.0001).
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Correlated with the loss of xanthophores was a severe perturbation of adult
melanophore stripes, with resulting phenotypes resembling the pigment pattern
of fms1 and fmsblue mutants (e.g.
Fig. 6A',B').
Melanophore stripes depend on both melanophore numbers and arrangements. Thus,
to quantify effects on melanophore stripe morphology, we assessed both
melanophore densities and organization. This analysis revealed an 11%
reduction in melanophore densities in upshifted individuals as compared to
control siblings left at 24°C (respective
means±s.d.=126±30.9, 142±30.8; n=95, 56;
F1,148=4.96, P<0.05). In contrast to
xanthophores (above), we detected only a marginal difference in melanophore
densities among individuals upshifted at different sizes, with a slightly more
severe deficit in smaller individuals upshifted when smaller (P=0.06;
data not shown).
To assess melanophore organization quantitatively, we examined nearest neighbor distances among melanophores. Well-defined stripes are associated with low variability in the distances between adjacent melanophores, whereas poorly defined stripes are associated with increased variability in the distances between adjacent melanophores; thus, coefficients of variation for melanophore nearest neighbor distances are a sensitive measure of melanophore distributions (D. M. P. and J. M. T., unpublished data). We examined coefficients of variation for mean melanophore nearest neighbor distances between individuals upshifted to 33°C and control siblings left at 24°C. This analysis revealed increased variability in melanophore positions in upshifted individuals compared to controls (F1,141=99.8, P<0.0001; means±s.d.: 41.6±4.67, 31.0±7.56; n=93, 51). Given results of cell transplantation experiments that suggested a role for xanthophores in promoting the organization of melanophores in stripes, we further asked whether differences in xanthophore densities among upshifted individuals were associated with variation in melanophore spacing. Fig. 5C shows that lower xanthophore densities were directly related to increased coefficients of variation for melanophore nearest neighbor distances (partial regression coefficient=-0.11; s.e.=0.02; F1,90=22.74, P<0.0001). Thus, continuous Fms activity is essential for maintaining normal numbers of melanophores, as well as the normal spacing of melanophores within stripes, in a manner that directly correlates with xanthophore densities.
During terminal stages of metamorphosis and during juvenile development
(>10 mm SL), initial experiments resulted in a severe reduction of
xanthophores, but not a complete elimination of these cells
(Fig. 5A,
Fig. 6C). To assess whether
these weaker phenotypes reflect a period of independence from Fms activity, we
reared fish to adult stages (18 mm SL) at 24°C, shifted these
individuals to 33°C, and examined the patterns regularly for approx. 6
weeks. These individuals lost approx. 98% of xanthophores and exhibited a
severe degeneration of melanophore stripes within 5 weeks of curtailing Fms
activity (similar changes were not observed in fms174A/+
individuals shifted at the same time; data not shown). Repeated imaging of
individual fish revealed that loss of xanthophores occurred gradually over a
period of weeks (Fig. 6D-G; for
animated time-lapse images, see Supplemental Data:
http://dev.biologists.org/supplemental/),
with the onset of loss varying considerably among individuals.
These analyses show that Fms activity is essential for maintaining differentiated cells of the xanthophore lineage and for maintaining the striped arrangement of adult melanophores throughout development.
Chromatoblast and chromatophore death when Fms activity is
curtailed
Given the loss of xanthophores and some melanophores upon shifting
fms174A fish to a restrictive temperature, we asked
whether these losses might be accounted for partly by cell death. In
fms174A embryos shifted to 33°C, we observed a rapid
(within 2 hours) increase in unpigmented TUNEL+ cells in
neural crest migratory pathways (Fig.
7A, Fig. 8A). In
fms174A juveniles, we did not observe a gross increase in
xanthophore or melanophore death immediately upon shifting to 33°C, though
unpigmented apoptotic cells were observed frequently within the dermis
(Fig. 7B) in locations where
fms-expressing cells normally are found
(Parichy et al., 2000a) (D. M.
P., unpublished data). Within 2-3 days of shifting fish to 33°C, however,
we observed extensive xanthophore and melanophore debris within the fins of
fms174A mutant but not wild-type individuals
(Fig. 7C). This pigmented
debris could also be identified within exclusion bodies that are extruded
through the epidermis by unknown mechanisms
(Parichy et al., 1999;
Sugimoto et al., 2000;
Sugimoto, 2002)
(Fig. 7D-F). We confirmed that
exclusion bodies contained xanthophore-derived pteridine pigments by their
autofluorescence (Fig.
7D'). An analysis of the number of such extrusions in adult
fish shifted to 33°C for 3 days revealed a dramatic increase in fins of
fms174A mutants as compared to wild-type
(Fig. 8B). Similar pigmented
debris and exclusion bodies were observed on the trunk at lower frequencies
(e.g., Fig. 6G, and data not
shown). Since fms mutants are deficient in macrophages at embryonic
stages (Herbomel et al.,
2001), it might be argued that pigmented debris and exclusion
bodies in upshifted fms174A individuals reflects an
abnormal manifestation of normal pigment cell turnover, revealed by a loss of
macrophages upon temperature upshift and a subsequent failure to clear dying
cells. This appears not to be the case, however, since histological staining
and time-lapse imaging reveals phagocytic and motile macrophages that persist
even after temperature upshift of fms174A adults (D. M.
P., unpublished data). Together, these findings suggest that Fms is required
for maintaining pigment cell precursors, and that at least some differentiated
xanthophores and melanophores that disappear when Fms activity is curtailed
are lost by death rather than dedifferentiation.
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Late activation of Fms allows recovery of xanthophores and
melanophore stripes through adult stages
Temperature shift experiments revealed an essential role for Fms in
maintaining cells of the xanthophore lineage and maintaining the striped
pattern of adult melanophores thoughout development. These findings do not
exclude the possibility that a critical period for Fms activity exists during
early development; for example, Fms might be required to establish a
population of precursor cells during embryogenesis that is recruited to
differentiate only much later, during pigment pattern metamorphosis. To
investigate whether a critical period exists after which xanthophores and
adult melanophore stripes can no longer be rescued, we reared homozygous
fms174A individuals at 33°C and shifted them to
24°C at a range of sizes from embryo to early adult (SL=3.6-16.3 mm,
n=67). These analyses showed that shifting fish to 24°C allowed
extensive recovery of xanthophores and melanophore stripes through late stages
of metamorphosis (Fig. 5B,
Fig. 9A,B; see below and
Supplemental Data:
http://dev.biologists.org/supplemental/)
We did not observe marked recovery of xanthophores in individuals downshifted
at larger sizes (>10 mm SL, but see below; final xanthophore density vs.
size when shifted, partial regression coefficient=-15.67; SE=1.581;
F1,63=98.19, P<0.0001;
Fig. 5B,
Fig. 9C).
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In addition to restoring xanthophores, providing Fms activity typically resulted in the recovery of an adult melanophore stripe pattern (Fig. 9A,B). Quantitation of melanophore densities and organization did not reveal a significant difference in melanophore densities between individuals downshifted at any stage and control individuals left at 33°C (P=0.9), though variability in melanophore nearest neighbor distances was reduced in downshifted individuals with greater xanthophore densities (partial regression coefficient=-5.13; s.e.=0.46; F1,92=125.47; P<0.0001; Fig. 5C). Thus, melanophores were more organized in individuals that recovered higher densities of xanthophores. Since some downshifted individuals recovered only irregular and broken melanophore stripes, as an additional indicator of stripe morphology we assessed the frequencies of individuals that developed complete stripes (defined as stripes having no more than one break per side; e.g., Fig. 10). Fig. 5D shows that complete melanophore stripes were more frequent in individuals downshifted by middle metamorphic stages: whereas individuals reared either at 24°C or downshifted prior to 8.0 mm SL typically exhibited no more than one stripe break per side (mean=0.2 breaks, s.d.=0.53, n=95), individuals downshifted at larger sizes exhibited from 3-10 breaks, or severely disrupted melanophore stripes resembling control fish maintained at 33°C. Thus, melanophore stripe organization is largely restored when Fms activity is provided at stages through middle to late metamorphosis and this restoration was associated with — and presumably caused by — the concomitant restoration of xanthophore densities.
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We observed only partial recovery of xanthophore numbers and stripe
morphology during late metamorphic stages and beyond (>8-10 mm SL;
Fig. 5B,D,
Fig. 9C). This difference from
earlier stages could reflect either a critical point beyond which xanthophores
and stripes can no longer be recovered fully; or simply insufficient time
between temperature downshift and scoring of the resulting patterns for
complete recovery to occur. To address these possibilities, we repeated the
initial experiments by rearing fish to juvenile stages (12 mm SL) at
33°C, shifting them to 24°C, and examining pattern development over
several months. These individuals gradually recovered xanthophores over
several weeks (Fig. 9D-I; for
animated time-lapse images, see: Supplementary Data). New xanthophores
appeared first in the fins and subsequently were found in ventral regions of
the flank, and to a lesser extent on dorsal scales. Gradually, the
distributional limits of these cells extended until they had completely
covered the flank. As xanthophores occupied new regions, melanophores in these
regions adopted increasingly spread morphologies and became increasingly well
organized; partially formed adult melanophore stripes developed within approx.
6 weeks of temperature down-shift. Xanthophore distributions and melanophore
patterns on the body were indistinguishable from wild-type by approx. 4 months
(see: Supplementary Data:
http://dev.biologists.org/supplemental/).
Intriguingly, however, the orientations of fin stripes were haphazard
(sometimes even within a fin) and were frequently perpendicular to the
wild-type orientations. Such perturbations to fin stripes were evident in both
caudal fins (Fig. 9J-L) and
anal fins (data not shown). These results demonstrate that Fms activity is
sufficient to recruit xanthophores (though not melanophores) and to rescue the
organization of melanophore stripes through metamorphic and juvenile
stages.
Fms is not essential for establishing a population of chromatophore
precursors during embryogenesis
Xanthophore recovery and pattern regulation in fms174A
homozygotes following temperature down-shift (above) is consistent with a
model in which Fms is not essential for establishing a population of
chromatophore precursor during early development. Nevertheless, we reasoned
that any residual activity by fms174A at the restrictive
temperature might allow the escape of some otherwise Fms-dependent cells
during embryogenesis, and these cells would then be able to repopulate the
flank. In light of this possibility, we repeated the temperature down-shift
experiments using individuals transheterozygous for
fms174A and fmsblue [which is likely
to act as a null allele (Parichy et al.,
2000a)], and placed embryos at a higher initial temperature of
35°C until they had reached the feeding stage. When larvae were
downshifted to 24°C, these individuals rapidly recovered approximately
normal numbers of fms-expressing cells and xanthophores (similar
results were obtained by knocking-down Fms activity in wild-type embryos with
a morpholino oligonucleotide; data not shown). Interestingly, however,
fms174A/fmsblue transheterozygotes exhibited
frequent breaks in adult stripes, far in excess of heterozygous
fms174A/+ or fmsblue/+ individuals
reared initially at 35°C, or
fms174A/fmsblue individuals reared throughout
development at 24°C (Fig.
10). These data do not support an essential early role for Fms in
establishing precursor cells that are solely responsible for generating later
adult stripes. Nevertheless, these findings are consistent with an initial
reduction in Fms-dependent cells contributing to irregularities in the
patterning of adult melanophore stripes.
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DISCUSSION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Independence of embryonic and metamorphic xanthophore populations
revealed by modulation of Fms activity
Many organisms undergo a metamorphosis in which an embryonic or larval
morphology is transformed into that of an adult. Among vertebrates, such
changes are especially pronounced in anuran amphibians; similar albeit less
dramatic changes also occur in salamanders and teleosts. In zebrafish, a
v
8 mm, reflecting slower changes at late
metamorphic and juvenile stages; DMP and JMT, manuscript in preparation). (A)
Temperature up-shift ablates xanthophores through middle stages of pigment
pattern metamorphosis though residual xanthophores persist in individuals
shifted during late pigment pattern metamorphosis or beyond. Mean densities of
xanthophores were significantly reduced in upshifted individuals as compared
to sibling controls left at 24°C (F1,148=500.0,
P<0.0001). (B) Temperature down-shift allows substantial
xanthophore recovery through middle stages of pigment pattern metamorphosis,
but less marked recovery during later metamorphic and juvenile stages (see
text for details). Mean densities of xanthophores for downshifted individuals
were significantly greater overall than sibling controls left at 33°C
(F1,74=9.27, P<0.005). (C) Melanophore
organization is correlated with xanthophore density. Reduced variation in
nearest neighbor distances between melanophores is associated with increased
xanthophore densities in both temperature upshift (red points) and downshift
(green points) experiments. Red diamond indicates the mean for individuals
completely lacking xanthophores at 33°C (upshift and control, pooled);
green diamond, the mean for control individuals reared exclusively at
24°C. Note that variability in melanophore nearest neighbor distances is
increased among individuals with partially disrupted stripes, whereas the most
severe phenotypes at 33°C have somewhat lower coefficients of variation,
reflecting a more uniform dispersion of melanophores once xanthophores and
stripes have been lost. Regression shown includes only individuals with
partial xanthophore deficits compared to controls. Individual and pooled
values shown are based on 98,643 melanophores, 47,067 xanthophores. (D)
Complete melanophore stripes are more common when Fms activity is provided by
temperature downshift prior to late metamorphic stages. Shown are percentages
of individuals downshifted at different sizes that exhibited complete
melanophore stripes (as defined by 
1 200 µm gap per side). Thus,
individuals shifted at sizes 
2=65.2, d.f.=13, P<0.0001). The
orange line indicates the percentage of individuals with complete stripes
among controls reared at 24°C.
