|
|
|
|
|
|
|
||||
| Home Help Feedback Subscriptions Archive Search Table of Contents | ||||
First published online 19 November 2008
doi: 10.1242/dev.030981
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Research Report |
1 Department of Molecular, Cell and Developmental Biology, University of
California Santa Barbara, Santa Barbara, CA 93106, USA.
2 Department of Zoology, Graduate School of Science, Kyoto University, Sakyo-ku,
Kyoto, 606-8502, Japan.
Author for correspondence (e-mail:
w_smith{at}lifesci.ucsb.edu)
Accepted 28 October 2008
SUMMARY
We report the isolation of a recessive ENU-induced short-tailed mutant in the ascidian Ciona intestinalis that is the product of a premature stop in the brachyury gene. Notochord differentiation and morphogenesis are severely disrupted in the mutant line. At the larval stage, variable degrees of ectopic endoderm staining were observed in the homozygous mutants, indicating that loss of brachyury results in stochastic fate transformation. In post-metamorphosis mutants, a uniform defect in tail resorption was observed, together with variable defects in digestive tract development. Some cells misdirected from the notochord lineage were found to be incorporated into definitive endodermal structures, such as stomach and intestine.
Key words: Notochord, Endoderm, Metamorphosis, Ascidian, Ciona intestinalis
INTRODUCTION
The T-box transcription factor brachyury has an essential role in
chordate mesoderm development (Herrmann and
Kispert, 1994
; Showell et al.,
2004). The structure and function of brachyury has been
extensively characterized in a number of chordates, including the ascidian
(Satoh, 2003;
Showell et al., 2004). In
ascidians, brachyury is expressed in the notochord precursor and the
notochord (Corbo et al., 1997;
Imai et al., 2000;
Yasuo and Satoh, 1993), and
experimental manipulations have confirmed its role in notochord development
(Di Gregorio et al., 2002;
Satou et al., 2001;
Yamada et al., 2003;
Yasuo and Satoh, 1998). Both
the molecular pathway inducing brachyury expression in the ascidian
notochord precursor, and direct and indirect downstream targets of
brachyury have been well characterized
(Hotta et al., 2000;
Hotta et al., 2008;
Imai et al., 2006;
Nishida, 2005;
Takahashi et al., 1999;
Yagi et al., 2004).
Functional study of brachyury in ascidians has relied largely on
the misexpression of DNA constructs, the downregulation of brachyury
expression by targeting upstream factors, and morpholino knockdown
(Di Gregorio et al., 2002;
Satou et al., 2001;
Takahashi et al., 1999;
Yamada et al., 2003). Although
these approaches have been fruitful, they frequently result in mosaic and/or
incomplete knockdown. Here, we report an N-ethyl-N-nitrosourea (ENU)-induced
null mutant allele of brachyury in Ciona intestinalis.
Several new and surprising results come from the study of this mutant.
Significantly, despite the lack of brachyury function, the homozygous
mutants show a highly variable misexpression of endoderm markers in the
transfated notochord lineage. The mutant line has allowed us to follow the
consequences of loss of brachyury through metamorphosis, where we
find that re-directed cells of the notochord lineage are able to contribute to
the definitive endoderm of the juvenile.
MATERIALS AND METHODS
Animals
Adult C. intestinalis were collected at the Santa Barbara Yacht
Harbor. ENU treatment and culturing of animals was as described
(Hendrickson et al., 2004;
Moody et al., 1999).
SNP linkage mapping
Larvae from crossed heterozygous mutant 411 adults were segregated by
phenotype and then pooled in groups of 50 to 200. Genomic DNA was isolated
from pooled larvae as described
(Hendrickson et al., 2004).
Forty-six PCR primer sets that amplify loci on the various chromosome arms
were designed based on the C. intestinalis genome sequence. The full
sequences of the panel of primers are available on request.
Sequence of mutant brachyury allele
Genomic DNA for dideoxy sequencing of the brachyury gene from
mutant 411 was amplified by specific primers that covered the entire ORF:
5'-ATGACGTCATCAGATAGTAAGTTAGC-3' (bra1F) and
5'-TCACAAAGAAGGTGGCGT-3' (bra1R); or bra1F and
5'-GGTTCGTATTTATGCAGAGAGT-3' (bra4R).
Microarray analysis
Microarray analysis was performed using the C. intestinalis
Oligoarray ver.1 (Yamada et al.,
2005). Two hundred nanograms of Trizol-isolated (Invitrogen) total
RNA from each sample were used. Replicates included swapping of the
fluorescent dyes between the samples.
Immunohistochemical staining and whole-mount in situ hybridization
Embryos were stained with BODIPY-FL phallacidin (Invitrogen), DAPI (Sigma)
and rabbit anti-GFP (Invitrogen). Alexa 488 (Invitrogen) was used as a
secondary antibody for the anti-GFP antibody. Immunostaining was as described
previously (Veeman et al.,
2008). Alkaline phosphatase (AP) activity was detected as
described (Whittaker and Meedel,
1989). AP staining reactions were allowed to proceed for 40-60
minutes at room temperature.
For whole-mount in situ hybridization, antisense RNA probes were
transcribed from cDNA clones in the C. intestinalis gene collection
release 1 (Satou et al.,
2002). In situ hybridization was as described
(Satou et al., 1995).
Single tadpole PCR
Single larvae were genotyped for the brachyury locus after AP
staining or in situ hybridization. Genomic DNA was isolated for PCR from
single larvae as described previously
(Veeman et al., 2008). Four
µl samples of the digested larvae or negative controls (from 11 µl
total) were PCR-amplified for 40 cycles using the bra1F and
bra4R primers (see above).
Reverse transcription-PCR (RT-PCR)
RNA was isolated with Trizol at the following stages: 110-cell, early
tailbud, larva, and stage 3 juvenile (Chiba
et al., 2004). The SMART RACE cDNA amplification kit (Clontech)
was used for cDNA synthesis. The cDNA samples were amplified with primers
bra1F and bra1R for brachyury, or with
5'-CGTTTTCCCATCCATCGTAG-3' and
5'-CCAGCAGATTCCATACCAAG-3' for cytoplasmic actin.
|
RESULTS AND DISCUSSION
Isolation of an ENU-induced Ciona intestinalis brachyury mutant
Five hundred and twenty-nine F1 adults were screened and 14 mutants in nine
complementation groups were isolated. Several of the mutants had short tail
phenotypes, including mutant 411 shown in
Fig. 1A. At a gross level, the
head and trunk of the mutant 411 larvae appeared largely normal, while the
tail appeared to lack vacuolated notochord cells. Single nucleotide
polymorphism (SNP)-based linkage analysis using 46 sets of PCR primers
targeting the various chromosome arms was used to link mutant 411 to
chromosome arm 12p (Fig. 1B).
Chromosome arm 12p contains at least three notochord genes (data not shown),
including brachyury (Shoguchi et
al., 2008). The brachyury gene from homozygous 411
mutants was found to have six nucleotide substitutions in exons compared with
the published C. intestinalis cDNA sequence (NM_001078478). The most
significant of these substitutions was a cytosine for thymine in exon 2 that
changed amino acid 49 from an arginine to a stop codon
(Fig. 1C). The predicted
protein from this mutant gene would be truncated at the beginning of the
DNA-binding T-box (Fig. 1C).
The fact that the mutant 411 phenotype is similar to previous reports for
Ciona embryos with downregulated or knocked-down brachyury
(Di Gregorio et al., 2002;
Satou et al., 2001;
Yamada et al., 2003) allowed
us to confidently conclude that the mutant phenotype was due to a
loss-of-function brachyury allele. In addition, we observed that four
genes previously shown to be downstream of brachyury [noto4,
noto8, prickle (pk) and tropomyosin
(Hotta et al., 2000)] and
laminin alpha 3/4/5 were not expressed in homozygous mutant 411
embryos (data not shown). Because the predicted protein is so severely
truncated, the mutation is likely to be a null. In the remainder of this
manuscript we will refer to this mutant as brachyury-
(bra-).
brachyury mutant at tailbud stages
To investigate the morphology and fates of the notochord lineage in the
homozygous mutants (bra-/-), the line was crossed to a
stable line expressing GFP under the control of the C. intestinalis
brachyury 5' regulatory region
(Joly et al., 2007). At the
early tailbud stage (Fig.
1D,E), the GFP-fluorescing cells in bra-/-
embryos were organized into a mass of tissue that was nearly as long and
narrow as the wild-type notochord, and the overall anterior/posterior (A/P)
length of the embryo was similar to wild type. Despite this, the convergent
extension (C/E) of the notochord lineage was severely disrupted. In embryos
double stained with DAPI and phallacidin
(Fig. 1F,G), the disrupted
morphology of the notochord lineage was evident and, most noticeably, the
cells of the notochord lineage failed to intercalate into a single column,
unlike in wild-type embryos (Fig.
1G, arrow).
brachyury occupies a key position in the gene regulatory network
that specifies notochord fate (Showell et
al., 2004). Approximately forty candidate brachyury
targets have been identified in C. intestinalis
(Hotta et al., 2000;
Takahashi et al., 1999). The
null mutation reported here provides a new tool for investigating
brachyury function. Microarray analysis was used to identify three
new notochord genes of unknown function that are downregulated in the
homozygous bra-/- mutant larvae (see Fig. S1 in the
supplementary material).
|
;
Imai et al., 2002a;
Kumano et al., 2006;
Minokawa et al., 2001).
However, morpholino knockdown of brachyury in Ciona savignyi
resulted in a short tail phenotype, but with no reported ectopic endoderm
(Satou et al., 2001). The
mutant line described here offers a stable, non-mosaic background in which to
study the effects of loss of brachyury function. On examination of
the pan-neural gene ETR-1 and muscle actin, there was no indication
of notochord cells transfating to neural or endodermal tissues in
bra-/- larvae (data not shown). By contrast, the
examination of alkaline phosphatase (AP) activity, as a marker of endoderm,
revealed definitive, but highly variable, transfating of notochord to
endoderm-like cells in the bra-/- larvae. Approximately
one-half of the bra-/- larvae had ectopic AP staining in
the core of the tail (Fig.
2B,C). Because of the variability in AP staining, the genotypes of
individual larvae were determined by genomic PCR and dideoxy sequencing
(Fig. 2B,C, lower panels).
Identical results were obtained by in situ hybridization using an
endoderm-specific marker, kyotograil2005.572.7.1
(Fig. 2E,F). Finally, AP
activity colocalized with brachyury promoter-driven GFP fluorescence
in bra-/- larvae, but not in wild-type larvae, indicating
that the notochord cells were transfated to endoderm in the homozygous mutant
(Fig. 2G,H).
The fate of cells in the notochord lineage of bra-/-
embryos not expressing ectopic endoderm markers is unknown, although by
morphology and gene expression they do not become notochord. Previous reports
show that ascidian Zic can prevent notochord cells from becoming endoderm, but
Zic is not expressed in the notochord until after gastrulation
(Imai et al., 2002b;
Kumano et al., 2006). Thus,
one possibility is that transfating of notochord cells in
bra-/- embryos is repressed late in the process of
determination, and that only a fraction of the embryos are able to overcome
the repression and express ectopic endoderm markers.
Although genes required for tail elongation, including pk and
laminin alpha 3/4/5 are not expressed in bra-/-
mutants, tail elongation was not completely disrupted, as was observed in
aimless/chongmague double homozygous mutants in C. savignyi
(Veeman et al., 2008). In
fact, the A/P axis in bra-/- mutants was similar to that
of wild-type embryos at early tailbud stage. One possibility is that the
transfated notochord cells are following the morphogenetic pathway of endoderm
in bra-/- mutants. Cells of the endodermal strand do form
a single-file row underlying the notochord in the wild type, although the
morphogenetic mechanisms are unknown. Alternatively, there may be an intrinsic
and cell fate-independent program in the notochord lineage to undergo at least
the initial steps of C/E.
Transfated notochord lineage contributes to definitive endoderm
Following the larval stage, ascidians undergo metamorphosis. In wild-type
ascidians, the tail is largely reabsorbed and most adult organs are present by
juvenile stage 4 (Fig. 3A,D)
(Chiba et al., 2004).
bra-/- larvae also undergo metamorphosis, but the
resulting juveniles are highly abnormal
(Fig. 3B,C). As with larvae,
bra-/- juveniles were variable
(Table 1), with some showing a
more moderate phenotype with well-developed adult organs, such as endostyle
and protostigmata (e.g. Fig.
3B), while others were more severely disrupted (e.g.
Fig. 3C). Even in the less
severe examples the digestive tracts were abnormal and did not appear to make
a complete tract (e.g. Fig.
3E). Nearly all bra-/- juveniles arrested at
this stage, presumably because of these defects in the digestive system.
|
|
Despite the fact that the brachyury gene is not transcribed past
the tailbud stage, the perdurance of GFP allows us to follow the notochord
lineage well into the juvenile stage (Fig.
3F-J) (Deschet et al.,
2003). As has been described previously
(Cloney, 1978), the remnants of
the notochord are coiled at one end of the developing juvenile at stage 3
(Fig. 3F). By stage 4, only a
small clump of GFP-positive cells is found between the esophagus and the
stomach (Fig. 3H). In contrast
to wild type, most GFP-positive cells in the mutant were found more centrally
located in the trunk region at stage 3
(Fig. 3G). Most significantly,
at stage 4 there was a much greater persistence of GFP-positive cells, many of
which had been incorporated into definitive endodermal organs, such as the
stomach and the intestine (Fig.
3I,J, arrows), suggesting that the cells were not following the
apoptotic pathway of normal notochord cells but rather were transfated.
Rescue of metamorphosis by tail ablation
The above observations that transfated notochord cells in
bra-/- animals could contribute to definitive digestive
organs suggested that gut development was abnormal due to an excess of cells.
As a possible mechanism to correct this defect, the tails of larvae were
removed and the remaining trunks were allowed to settle and undergo
metamorphosis. Tail ablation resulted in an increase from 1% to 11% of
bra-/- animals that had well-formed digestive tracts,
indicating that the persistence of the tail remnants in the juvenile disrupts
normal development (Table
1).
In conclusion, the mutant line reported here will provide a useful tool for future studies on notochord specification and brachyury function. Among the unresolved issues is the variability in the expression of endoderm markers in bra-/- mutants. Investigation in this area may provide an insight into the quantitative effects of other loci and mechanisms of embryonic robustness.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/136/1/35/DC1
Footnotes
This work was supported by grants from the NIH (HD38701 and GM075049) to W.C.S., and a Grant-in-aid for Scientific Research from MGXT to N.S. (No.17018018). We thank Dr Yasunori Sasakura, Maizuru Fisheries Research Station of Kyoto University, Mrs Yasuyo Kasuga, Dr Michael T. Veeman, Erin L. Mulholland and the Santa Barbara Ascidian Stock Center for their help with this project. Deposited in PMC for release after 12 months.
* Present address: Sars International Centre for Marine Molecular Biology,
NO-5008 Bergen, Norway 
REFERENCES
Chiba, S., Sasaki, A., Nakayama, A., Takamura, K. and Satoh,
N. (2004). Development of Ciona intestinalis juveniles
(through 2nd ascidian stage). Zool. Sci.
21,285
-298.[CrossRef][Medline]
Cloney, R. A. (1978). Ascidian metamorphosis:
review and analysis. In Settlement and Metamorphosis of Marine
Larvae (ed. F. S. Chia and M. E. Rice), pp.255
-282. Amsterdam: Elsevier.
Corbo, J. C., Levine, M. and Zeller, R. W.
(1997). Characterization of a notochord-specific enhancer from
the Brachyury promoter region of the ascidian, Ciona intestinalis.
Development 124,589
-602.[Abstract]
Deschet, K., Nakatani, Y. and Smith, W. C.
(2003). Generation of Ci-Brachyury-GFP stable transgenic lines in
the ascidian Ciona savignyi. Genesis
35,248
-259.[CrossRef][Medline]
Di Gregorio, A., Harland, R. M., Levine, M. and Casey, E. S.
(2002). Tail morphogenesis in the ascidian, Ciona intestinalis,
requires cooperation between notochord and muscle. Dev.
Biol. 244,385
-395.[CrossRef][Medline]
Hendrickson, C., Christiaen, L., Deschet, K., Jiang, D., Joly,
J. S., Legendre, L., Nakatani, Y., Tresser, J. and Smith, W. C.
(2004). Culture of adult ascidians and ascidian genetics. In
Development of Sea Urchins, Ascidians, and Other Invertebrate
Deuterostomes: Experimental Approaches, vol.74
, pp. 143-169. San Diego:
Elsevier.[CrossRef]
Herrmann, B. G. and Kispert, A. (1994). The T
genes in embryogenesis. Trends Genet.
10,280
-286.[CrossRef][Medline]
Hotta, K., Takahashi, H., Asakura, T., Saitoh, B., Takatori, N.,
Satou, Y. and Satoh, N. (2000). Characterization of
Brachyury-downstream notochord genes in the Ciona intestinalis embryo.
Dev. Biol. 224,69
-80.[CrossRef][Medline]
Hotta, K., Takahashi, H., Satoh, N. and Gojobori, T.
(2008). Brachyury-downstream gene sets in a chordate, Ciona
intestinalis: integrating notochord specification, morphogenesis and chordate
evolution. Evol. Dev.
10, 37-51.[Medline]
Imai, K., Takada, N., Satoh, N. and Satou, Y.
(2000). (beta)-catenin mediates the specification of endoderm
cells in ascidian embryos. Development
127,3009
-3020.[Abstract]
Imai, K. S., Satoh, N. and Satou, Y. (2002a).
An essential role of a FoxD gene in notochord induction in Ciona embryos.
Development 129,3441
-3453.[Medline]
Imai, K. S., Satou, Y. and Satoh, N. (2002b).
Multiple functions of a Zic-like gene in the differentiation of notochord,
central nervous system and muscle in Ciona savignyi embryos.
Development 129,2723
-2732.
Imai, K. S., Levine, M., Satoh, N. and Satou, Y.
(2006). Regulatory blueprint for a chordate embryo.
Science 312,1183
-1187.
Joly, J. S., Kano, S., Matsuoka, T., Auger, H., Hirayama, K.,
Satoh, N., Awazu, S., Legendre, L. and Sasakura, Y. (2007).
Culture of Ciona intestinalis in closed systems. Dev.
Dyn. 236,1832
-1840.[CrossRef][Medline]
Kumano, G., Yamaguchi, S. and Nishida, H.
(2006). Overlapping expression of FoxA and Zic confers
responsiveness to FGF signaling to specify notochord in ascidian embryos.
Dev. Biol. 300,770
-784.[CrossRef][Medline]
Minokawa, T., Yagi, K., Makabe, K. and Nishida, H.
(2001). Binary specification of nerve cord and notochord cell
fates in ascidian embryos. Development
128,2007
-2017.
Moody, R., Davis, S. W., Cubas, F. and Smith, W. C.
(1999). Isolation of developmental mutants of the ascidian Ciona
savignyi. Mol. Gen. Genet.
262,199
-206.[CrossRef][Medline]
Nishida, H. (2005). Specification of embryonic
axis and mosaic development in ascidians. Dev. Dyn.
233,1177
-1193.[CrossRef][Medline]
Satoh, N. (2003). The ascidian tadpole larva:
comparative molecular development and genomics. Nat. Rev.
Genet. 4,285
-295.[Medline]
Satou, Y., Kusakabe, T., Araki, S. and Satoh, N.
(1995). Timing of initiation of muscle-specific gene expression
in the ascidian embryo precedes that of developmental fate restriction in
lineage cells. Dev. Growth Differ.
37,319
-327.[CrossRef]
Satou, Y., Imai, K. S. and Satoh, N. (2001).
Action of morpholinos in Ciona embryos. Genesis
30,103
-106.[CrossRef][Medline]
Satou, Y., Yamada, L., Mochizuki, Y., Takatori, N., Kawashima,
T., Sasaki, A., Hamaguchi, M., Awazu, S., Yagi, K., Sasakura, Y. et al.
(2002). A cDNA resource from the basal chordate Ciona
intestinalis. Genesis
33,153
-154.[CrossRef][Medline]
Shoguchi, E., Hamaguchi, M. and Satoh, N.
(2008). Genome-wide network of regulatory genes for construction
of a chordate embryo. Dev. Biol.
316,498
-509.[CrossRef][Medline]
Showell, C., Binder, O. and Conlon, F. L.
(2004). T-box genes in early embryogenesis. Dev.
Dyn. 229,201
-218.[CrossRef][Medline]
Takahashi, H., Hotta, K., Erives, A., Di Gregorio, A., Zeller,
R. W., Levine, M. and Satoh, N. (1999). Brachyury downstream
notochord differentiation in the ascidian embryo. Genes
Dev. 13,1519
-1523.
Veeman, M. T., Nakatani, Y., Hendrickson, C., Ericson, V., Lin,
C. and Smith, W. C. (2008). Chongmague reveals an essential
role for laminin-mediated boundary formation in chordate convergence and
extension movements. Development
135, 33-41.
Whittaker, J. and Meedel, T. (1989). Two
histospecific enzyme expressions in the same cleavage-arrested one-celled
ascidian embryos. J. Exp. Zool.
250,168
-175.[CrossRef][Medline]
Yagi, K., Satou, Y. and Satoh, N. (2004). A
zinc finger transcription factor, ZicL, is a direct activator of Brachyury in
the notochord specification of Ciona intestinalis.
Development 131,1279
-1288.
Yamada, L., Shoguchi, E., Wada, S., Kobayashi, K., Mochizuki,
Y., Satou, Y. and Satoh, N. (2003). Morpholino-based gene
knockdown screen of novel genes with developmental function in Ciona
intestinalis. Development
130,6485
-6495.
Yamada, L., Kobayashi, K., Satou, Y. and Satoh, N.
(2005). Microarray analysis of localization of maternal
transcripts in eggs and early embryos of the ascidian, Ciona intestinalis.
Dev. Biol. 284,536
-550.[Medline]
Yasuo, H. and Satoh, N. (1993). Function of
vertebrate T-gene. Nature
364,582
-583.
Yasuo, H. and Satoh, N. (1998). Conservation of
the developmental role of Brachyury in notochord formation in a urochordate,
the ascidian Halocynthia roretzi. Dev. Biol.
200,158
-170.[CrossRef][Medline]
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Reddit 
Technorati 
Twitter What's this?
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
