Department of Neurology and the Ernest Gallo Clinic and Research Center, University of California at San Francisco, 5858 Horton Street, Emeryville, CA 94608, USA
* Author for correspondence (e-mail: andpete{at}itsa.ucsf.edu)
Accepted 30 September 2003
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SUMMARY |
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Key words: Atrophin, Forebrain, Co-repressor, Mouse, Notochord, ANR
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Introduction |
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The anterior neural ridge (ANR), located at the rostral margin of the
neural plate, is a signaling center required for forebrain development,
specifically for elaboration of pattern in the most anterior subregion of the
brain, the telencephalon (Eagleson and
Dempewolf, 2002; Shimamura and
Rubenstein, 1997
). One of the signals secreted by the ANR
beginning at E8.0 is Fgf8 (Crossley and
Martin, 1995
). Mutants with reduced Fgf8 expression in
the ANR and its derivatives, such as Hex/,
Hesx1/ and
oto/ embryos, exhibit incomplete
telencephalic development
(Martinez-Barbera and Beddington,
2001
; Martinez-Barbera et al.,
2000
; Zoltewicz et al.,
1999
). In addition, mouse embryos expressing Fgf8 from a
hypomorphic allele develop reduced telencephalic structures
(Meyers et al., 1998
). These
studies together define a prominent role for Fgf8 in ANR function.
The study of mutant alleles is a powerful approach for understanding gene
function. We have carried out a random chemical mutagenesis screen in mice,
aimed at identifying recessive mutations affecting early embryonic patterning
(Hentges et al., 1999). In
this screen, we uncovered an embryonic lethal mutation in an atrophin family
member. We have named the mutant allele openmind (om), and
the gene atrophin-2 (Atr2). Atr2, known in human as Arginine
(R) Glutamic Acid (E) Repeat Encoding or RERE, was first described as an
atrophin 1 (Atr1)-related protein that could heterodimerize with Atr1
(Waerner et al., 2001
;
Yanagisawa et al., 2000
). Atr1
(Drpla Mouse Genome Informatics) has been well studied because it
causes a human neurodegenerative disease known as dentatorubral-pallidoluysian
atrophy (DRPLA) when its polyglutatmine tract is abnormally expanded. Atr2 is
distinguished from Atr1 in that it does not have a polyglutamine tract, but it
does bind tightly to glutamine-expanded Atr1
(Yanagisawa et al., 2000
),
suggesting that Atr2 has a role in DRPLA disease development or progression.
DRPLA is one of a family of nine polyglutamine diseases that includes
Huntington's disease, Spinal and Bulbar Muscular Atrophy, and several
spinocerebellar ataxias. Although the proteins encoding polyglutamine tracts
are distinct in each disease, there is evidence for a similar underlying
pathogenic mechanism involving disruption of gene regulation in the nucleus
(McCampbell et al., 2001
;
Nucifora et al., 2001
;
Ross, 2002
). The normal
function of vertebrate Atr1 is not understood but, as it binds to Eto1, a
component of nuclear co-repressor complexes, a role in transcriptional
repression has been proposed (Wood et al.,
2000
).
Whereas mouse and human genomes contain two distinct atrophin genes, the
Drosophila genome contains only one, known as Atro or
Grunge. Mutation of this gene causes a variety of defects, including
disruption of embryonic patterning (Erkner
et al., 2002; Fanto et al.,
2003
; Zhang et al.,
2002
). Atro binds to the Even skipped (Eve) and Huckebein
transcription factors, and acts as a co-repressor for Eve
(Zhang et al., 2002
). The
C. elegans genome also has a single atrophin ortholog,
egl-27, which is also required for the development of embryonic
pattern (Ch'ng and Kenyon,
1999
; Herman et al.,
1999
; Solari et al.,
1999
). Atr2, Atro and EGL-27, but not Atr1, each has N-terminal
homology to metastasis-associated protein 2 (Mta2), a core component of the
NuRD (Nucleosome Remodeling and Deacetylase) complex
(Zhang et al., 1999
). NuRD and
other histone deacetylase complexes silence genes by altering chromatin
structure such that the DNA becomes inaccessible to transcriptional activators
(Ng and Bird, 2000
). Because
of their homology to Mta2, it has been proposed that the vertebrate, fly and
worm atrophins may regulate transcription by acting in conjunction with
histone deacetylase complexes (Solari and
Ahringer, 2000
; Solari et al.,
1999
; Zhang et al.,
2002
). However, no biochemical evidence showing association of an
atrophin with such a complex has yet been reported.
Here, we report that Atr2-mutant embryos exhibit diverse developmental defects, and we focus on the role of Atr2 in patterning the anterior neural tube. We provide evidence that anterior neural defects are caused by disruption of two signaling centers, namely the anterior embryonic midline and the ANR. We show that the N terminus of Atr2 is sufficient to recruit histone deacetylase 1 (Hdac1), but not other NuRD core subunits, suggesting that Atr2 is part of a novel histone deacetylase complex. We propose that the molecular mechanism underlying om defects involves loss of function of this complex, and specific disregulation of Shh and Fgf8.
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Materials and methods |
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The full-length mouse Atr2 cDNA was cloned by RT-PCR and sequences verified by comparing with consensus sequences derived from NCBI and celera databases. 3x flag tags were added to the N termini of Atr2 proteins using the 3xFlag-CMV7 expression vector (Sigma). Fragments of Atr2 were created by dividing the cDNA into N-terminal (N-Atr2) and C-terminal (C-Atr2) coding portions using a unique EcoRI site between the SANT- and GATA-coding sequences.
Labeling of embryos, histology and 5' RACE
Embryos of desired ages were obtained by carrying out timed matings between
genotyped carriers. Whole-mount in situ hybridization was performed as
described (Zoltewicz et al.,
1999). The Atr2/ß-galactosidase fusion protein in PT026
embryos was visualized with X-gal substrate using standard procedures. Some
embryos were embedded in paraplast and sectioned at 7 µm using standard
procedures; others were embedded in agarose and vibratome-sliced. 5'
RACE was performed using the First Choice RLM-RACE kit from Ambion.
Co-immunoprecipitation
Flag-tagged Atr2 constructs were transiently transfected into 293 cells
using Lipofectamine Plus (Invitrogen). Transfected cells were harvested after
24 hours in culture and lysed in Ripa buffer [150 mM NaCl, 50 mM Tris (pH
7.5), 0.5% sodium deoxycholate, 0.5% NP-40, 0.1% SDS, complete protease
inhibitors (Roche)]. Nucleic acids were degraded by benzonase nuclease
(Novagen) treatment. An equal volume of buffer, similar to Ripa but lacking
denaturing detergents, was added and the insoluble debris pelleted. Soluble
extracts were added to M2 anti-flag beads (Sigma) pre-blocked with BSA, and
extracts plus beads were nutated for one hour at 4°C. Beads were washed 5
times, then bound were proteins eluted, resolved on 4-12% Bis-Tris gels
(Fig. 5B) or 3-8% Tris-Acetate
gels (Fig. 5C,D), and
transferred to PVDF membrane (Amersham) using the XCell II system
(Invitrogen). Blots were probed with anti-flag M5 (Sigma), anti-Hdac1,
anti-RbAp46, anti-RbAp48 (Affinity Bioreagents) and anti-Atr1 (Santa Cruz).
The 286-1 antibody was created by immunizing rabbits with a peptide matching
the C terminus of Atr1 (Synpep). It was affinity purified and DSS-crosslinked
(Pierce) to protein G sepharose.
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Results |
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PT026 carriers were also intercrossed to produce PT026 homozygous embryos. These mutants ranged from severely affected (identical to openmind homozygotes) to apparently normal, suggesting the PT026 allele might be hypomorphic. RT-PCR analysis confirmed the existence of the wild-type transcript in PT026 mutants (not shown), indicating that PT026 is indeed a hypomorphic allele. Even though some homozygous embryos looked normal at E9.5, no homozygous pups were ever recovered, indicating that the quantity of wild-type message is not sufficient to rescue viability. However, this report is not intended to be a detailed characterization of the PT026 mutant phenotype. Here, we used PT026 for two limited purposes: to confirm correct identification of the mutated gene and to define the wild-type expression pattern of Atr2.
A close examination of Atr2 genomic sequences revealed the existence of a second CpG island in the 12th intron (Fig. 2A). To determine whether this island represented an internal promoter producing another Atr2 transcript, 5' Race was performed. This analysis revealed a transcript that initiates within the second CpG island, and it was named Atr2S for Atr2 short form. Seven identical clones representing the 5' most end of the Atr2S mRNA were isolated, revealing that the Atr2S transcript contains about 100 bases of unique 5' GC-rich leader sequence that splice into exon 13 (data not shown). Because Atr2S initiates far downstream of the openmind point mutation, and because the om mutation affects expression of the full-length Atr2 only (Fig. 5D), Atr2S does not contribute to the om phenotype and will be characterized elsewhere.
Atr2 is widely expressed during embryonic development
The expression of Atr2 in the developing embryo was examined using
ß-galactosidase (ßgal) staining to detect the PT026 fusion protein,
and whole-mount in situ hybridization to detect the endogenous mRNA. In order
to visualize only full-length Atr2, a 5' specific RNA probe
having no sequences in common with Atr2S was used for the in situ
hybridization. The expression patterns visualized by these two methods were
similar. Atr2 expression was examined in detail from E7.5 through E11.5, using
PT026 heterozygotes. Atr2 expression was observed in every cell of
the embryo at all these stages. Some regions, such as the heart and dorsal
neural tissues, downregulate expression but do not lose it entirely. Atr2 was
expressed uniformly at E7.5 throughout the embryo in all three germ layers
(not shown). At early headfold, expression is mostly uniform but begins to be
upregulated in the anterior portion of the embryonic midline
(Fig. 3A). By E8.25, expression
is strongly elevated in the anterior midline, but remains at a uniform level
posteriorly (Fig.
3B,b,b'). By E8.5, expression is upregulated along the
entire notochord (Fig.
3C,c,c'). At E8.75, expression remains high in the
notochord, and begins to elevate ventrally in the anterior CNS
(Fig. 3D,d,d'). At E9.5,
additional sites of elevated expression appear, including the apical
ectodermal ridge, the isthmus, the ventral diencephalon and ventral neurons
(Fig. 3E,K). At E10.5, neurons
in the spinal cord and brain strongly upregulate expression
(Fig. 3F). The mRNA pattern at
E9.75 is similar to that of the fusion protein at E10.5
(Fig. 3G). Both the mRNA and
the fusion protein show elevation in the AER at E9.5
(Fig. 3H). Expression of Atr2
in the AER is required for normal development, as the AER does not form
properly in mutant embryos (Fig.
3I,J). E11.5 embryos express Atr2 in a pattern similar to that
seen at E10.5, showing an even greater upregulated expression in neurons
throughout the neural tube (Fig.
3L,M). We have not analyzed functions of Atr2 in the AER or
neurons because mutant embryos die from heart failure shortly after E9.5.
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The anterior midline is defective in om embryos
The expansion of Pax6 and Emx2 into the anterior neural
midline, and the reduction of Nkx2.1 ventrally, suggested a loss of
ventralizing signals. To discover whether such signals were present in
om embryos, we examined Shh expression. Shh is a
ventralizing signal produced by the prechordal plate, notochord and
prospective floorplate during day 8
(Epstein et al., 1999).
Interestingly, in om embryos at E8.75, Shh expression was
normal posterior to the hindbrain, but was almost completely absent from its
anterior domain (Fig. 4G;
om left, wild type right). Only a small spot of Shh
expression remained in om (arrowhead), which was likely responsible
for inducing the residual Nkx2.1 expression
(Fig. 4B), and possibly for
keeping the central neural plate clear of dorsal-specific transcripts. To
determine whether the anterior-specific loss of Shh at E8.75 is due
to a failure to initiate or a failure to maintain expression, Shh
expression was examined in very early headfold embryos. In mutants at E8.0,
Shh is missing from the anteriormost portion of the midline, but is
expressed normally in the posterior midline
(Fig. 4I). Because the onset of
Shh expression is very close to E8.0, the lack of transcripts
observed at this stage strongly suggests that Shh fails to initiate
expression in its anterior domain in om mutants.
We looked further, for the expression of genes regulated by Shh
Gli1 and Gli3 (Ruiz i
Altaba, 1998). At E8.75 the normal Gli1 expression
pattern is similar to that of Shh, but is wider ventrally
(Fig. 4J; wild type right),
whereas Gli3 exhibits a complementary dorsal-restricted pattern
(Fig. 4K). Gli1 was
significantly reduced anteriorly in om mutants
(Fig. 4J; left), whereas
Gli3 was expanded towards the ventral midline
(Fig. 4L), consistent with
reduced Shh function. Gli3 encroached on the midline from the
forebrain through the posterior hindbrain. These mutant expression patterns
illustrate that Atr2 is required for Shh production by midline cells that
underlie the developing brain. However, by E9.5, om mutants appear to
recover normal patterns of Shh, Gli1 and Gli3 expression
(not shown), indicating that the onset of anterior Shh expression is
significantly delayed in om mutants, rather than abolished. We could
not explore the effects of this delay on subsequent brain development in any
detail because by E9.5, mutants are unhealthy owing to imminent cardiac
failure.
Even though no Shh deficit was detected in the posterior of
mutants during day 8, mutants exhibit incomplete floorplate formation in the
spinal cord at E9.5. At E9.5 in normal embryos, Shh is expressed in
the notochord and neural floorplate; furthermore, the ventral spinal cord
midline is thin relative to the lateral edge, reflecting floorplate
development (Fig. 4M). In
om mutants of the same stage, although Shh expression is
normal, the ventral midline of the spinal cord is not thin, and the notochord
is abnormally large (Fig. 4N).
Embryos were also stained for Hnf3b (Foxa2 Mouse
Genome Informatics), a floorplate marker and transcription factor involved in
inducing Shh through direct binding to Shh promoter elements
(Epstein et al., 1999). At
E9.5, mutants express Hnf3b normally in the ventral spinal cord, but
they show morphological floorplate and notochord abnormalities
(Fig. 4P). These results
indicate that Atr2 is not required for the expression of floorplate-specific
genes in the spinal cord, but that it is necessary for the development of
floorplate morphology, and for normal convergence/extension of the
notochord.
To determine whether the anterior decrease in Shh during day 8
reflected a loss of anterior cells or a specific failure of these cells to
express Shh, younger mutants were stained for Hnf3b and for
brachyury (Wilkinson et al.,
1990), both markers of the developing notochord. Mutant embryos
show normal and robust expression patterns of both Hnf3b at E8.5
(Fig. 4R), and brachyury at
E8.25 (Fig. 4T). E8.5 embryos
were also stained for goosecoid (gsc), a marker of the prechordal
plate and anterior ventral neural plate
(Blum et al., 1992
). No
difference in gsc expression was detected between wild type
(Fig. 4U) and mutants
(Fig. 4V). These results
strongly suggest that anterior midline cells are indeed present in om
mutants, but that they lack the ability to express Shh.
Fgf8 expression is abnormal in the anterior neural ridge (ANR) of om embryos
The anterior neural ridge (ANR) is a signaling center involved in
patterning the vertebrate telencephalon
(Rubenstein et al., 1998;
Shimamura and Rubenstein,
1997
). The patterning function of the ANR is meditated at least in
part by Fgf8, a potent signaling molecule required for telencephalic
development (Meyers et al.,
1998
). At E8.5, the ANR is located at the anterior border of the
neural plate, at the junction of the ectoderm and neurectoderm. Fgf8
expression is normally tightly localized within the ANR at this stage
(Fig. 4W; red circle)
(Crossley and Martin,
1995
).
To determine whether ANR signals are produced in om mutants, the
expression of Fgf8 was examined. Expression of Fgf8 in the
vicinity of the ANR in mutant embryos is reduced in intensity and delocalized
relative to wild type (Fig.
4X,Y). In mutants, Fgf8 expression is no longer limited
to the border between the neurectoderm and the epidermal ectoderm, but spreads
abnormally into the epidermal ectoderm and the neuroepithelium. The
delocalization of Fgf8 in mutant embryos indicates that Atr2 is
necessary to limit Fgf8 to the ANR. As Fgf8 can repress Emx2
(Crossley et al., 2001),
reduced Fgf8 signaling may contribute to the expansion of Emx2
expression observed in the om mutant neural plate
(Fig. 4F). Hesx1 is a
transcription factor required for telencephalic development, and for normal
levels of Fgf8 expression in the ANR
(Martinez-Barbera and Beddington,
2001
). In om embryos, Hesx1 is maintained in the
medial neural plate, but is absent from the ANR and the anterolateral neural
plate (Fig. 4Z').
Reduction of Hesx1 in the mutant ANR may contribute to the observed
decrease in Fgf8 expression.
Atr2 associates with Hdac1 but not other NuRD components
The Drosophila genome encodes a single atrophin-related protein
(Atro) that functions as a co-repressor for eve, and, given the
complexity of patterning defects in mutant embryos, probably for other
transcription factors as well (Erkner et
al., 2002; Zhang et al.,
2002
). Although the molecular co-repression mechanism for Atro and
Eve has not been elucidated, Atro and Atr2 (but not Atr1) show significant
N-terminal protein homology to vertebrate metastasis associated factor 2
(Mta2) (Fig. 2A), suggesting
they may have a similar function. Mta2 is a core subunit of NuRD, a histone
deacetylase complex with transcriptional repressive activity
(Zhang et al., 1999
). Other
NuRD core subunits include Mbd3b, Hdac1, Hdac2, RbAp46, and RbAp48.
To determine whether Atr2 associates with NuRD components in vivo,
flag-tagged Mta2 (a gift of D. Reinberg) and full-length flag-tagged Atr2
(Fig. 5A) were transiently
overexpressed individually in 293 cells. Soluble extracts were incubated with
anti-flag beads to immunoprecipitate transfected and associated proteins.
Immunoprecipitated proteins were examined by western blotting for the presence
of the transfected protein, and then for NuRD complex proteins using
antibodies specific for Hdac1, RbAp48 and RbAp46
(Fig. 5B). Flag-Mta2 yielded
the expected band at about 80 kDa, and pulled down Hdac1, RbAp46 and RbAp48,
as previously demonstrated (Zhang et al.,
1999). Flag-Atr2 migrated at 250 kDa and pulled down Hdac1, but
not RbAp46 or RbAp48 (Fig.
5B).
To determine whether the association of Atr2 with Hdac1 was mediated by its Mta2-homologous domains or by another region, the protein was divided into two fragments (Fig. 5A). The N-terminal fragment (N-Atr2) encodes the BAH, ELM2 and SANT domains and the C-terminal fragment (C-Atr2) contains the remainder of the protein, from the GATA domain through the Atr1-homologous region. These fragments were flag-tagged and examined in 293 cells, as described above. These experiments show that Hdac1 immunoprecipitates with N-Atr2, but not with C-Atr2 (Fig. 5B), indicating that sequences through the SANT domain are sufficient for Hdac1 recruitment.
In order to look at the binding partners of Atr2 in mouse embryos, a
polyclonal antibody recognizing the C terminus of Atr2 was produced, called
286-1. This antibody was crosslinked to protein G sepharose beads and used to
immunoprecipitate endogenous Atr2 from wild-type E9.5 embryo extracts
(Fig. 5C). 286-1 beads pulled
down both full-length Atr2, and the160 kDa short form of Atr2, Atr2S.
Endogenous Atr2 was specifically associated with Atr1 and Hdac1, but not with
RbAp46. Thus, Atr2 associates with Hdac1 in 293 cells and in the mouse embryo,
but not with the core NuRD subunit RbAp46, suggesting that it exists in a
complex distinct from NuRD.
286-1 beads were also used to determine whether any full-length Atr2 is made by om cells. Because it was difficult to isolate E9.5 om mutants that were both healthy and similar in size to their unaffected littermates, fibroblasts were isolated from E8.75 wild-type and om mutant embryos, and grown in culture. Extracts were made from these mouse embryo fibroblasts (MEFs) and endogenous Atr2 immunoprecipitated. Although Atr2S was present in om MEFs, no full-length Atr2S could be detected (Fig. 5D). Atr1 was co-immunoprecipitated in wild-type and mutant cells (Fig. 5D), but no robust association with Hdac1 could be found in either cell line (not shown).
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Discussion |
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Atr2 is required for Shh expression in the anterior midline during day 8 of development
Regulation of the Shh expression pattern during mouse development
is complex. Multiple enhancers have been discovered, in the both mouse and
zebrafish Shh promoters, that are responsible for directing
Shh expression to distinct subdomains of its overall pattern
(Epstein et al., 1999;
Muller et al., 1999
). Along
similar lines, we have found that Atr2 is required for Shh to be
expressed in its anterior subdomain. Atr2 is necessary for initiation of
Shh in the anterior midline at E8.0, but not in the posterior midline
(Fig. 4I). Interestingly, the
same anterior midline cells that lose Shh in E8.0 mutants, normally
upregulate Atr2 at E8.0-E8.25 (Fig.
2A,B), consistent with the idea that a high level of Atr2 is
required for Shh to be transcribed anteriorly. One of the
transcriptional activators of Shh is Hnf3ß
(Ang and Rossant, 1994
;
Weinstein et al., 1994
).
Hnf3b is expressed in om mutants at E8.5
(Fig. 4R), but it is not
sufficient to activate Shh in the anterior midline. Anterior midline
cells are clearly present in om mutants, being marked by expression
of Hnf3b, gsc, and brachyury (Fig.
4Q-V). Taken together, these data demonstrate that even though
anterior midline cells are present and express a known inducer of
Shh, they fail to activate Shh expression.
This in turn suggests that an Atr2-regulated repressor of Shh is
expressed in the anterior midline. An apparent candidate for this repressor is
Gli3, because it is capable of downregulating Shh
(Ruiz i Altaba, 1998) and is
expanded in om. However, it is unlikely that the hypothetical
Atr2-regulated repressor is Gli3 for the following reasons. First,
Gli3 is still excluded from the ventral-most midline of the brain in
om mutants (Fig. 4L), where Atr2 is elevated (Fig.
3d). Second, if depression of Gli3 were a direct event in
om mutants, reducing the dosage of Gli3 by creating compound mutants
between om and Gli3 extra-toes
(Hui and Joyner, 1993
) should,
at least to some extent, rescue ventral development. Instead, double mutants
exhibit a more severe phenotype (J.S.Z. and A.S.P., unpublished). Therefore it
is more likely that Atr2 silences an as yet unidentified repressor of
Shh during day 8. Because om mutants appear to recover a
normal Shh expression pattern by E9.5 (not shown), the window of
activity of this repressor or the competence to respond to it appears to be
limited to day 8 of development. Alternatively, Atr2 may act as an activator
of Shh.
Atr2 is required to localize Fgf8 to the ANR
Atr2 is also necessary for correct restriction of Fgf8 to the ANR
signaling center, because om mutants show reduced and disorganized
Fgf8 expression at and around the anterior neural margin
(Fig. 4X,Y). Fgf8 signals from
the ANR normally contribute to patterning the telencephalic primordia
(Eagleson and Dempewolf, 2002;
Martinez-Barbera and Beddington,
2001
; Rubenstein et al.,
1998
; Shanmugalingam et al.,
2000
; Shimamura and
Rubenstein, 1997
). The mechanisms that ordinarily restrict Fgf8 to
the border between the neurectoderm and the epidermal ectoderm are not known.
For the midbrainhindbrain isthmic organizer, another signaling center using
Fgf8, Fgf8 restriction involves complex positive and negative
regulatory mechanisms; at the boundary between the Otx2 and Gbx2 expression
domains, for example, Otx2 represses Fgf8 while Gbx2 maintains
Fgf8 expression (Wurst and
Bally-Cuif, 2001
). By analogy, positioning of Fgf8 at the
ANR is also likely to involve positive and negative influences. In this
context, Atr2 appears to act as a requisite component of a transcriptional
repressor that is needed both for full-level expression of Fgf8 from
the ANR, and for limiting Fgf8 expression to the ANR. Hesx1 is a
transcriptional regulator that functions in the anterior neurectoderm
(Martinez-Barbera et al.,
2000
). Hesx1 is necessary for full expression of Fgf8
from the ANR-derived commissural plate
(Martinez-Barbera and Beddington,
2001
). Atr2 in turn is necessary for the expression of
Hesx1 in the ANR at E8.5 (Fig.
4Z'). Thus a normal role of Atr2 may be to silence a
repressor of Hesx1 in the ANR so that this factor is available to
support normal Fgf8 expression levels. However, this same mechanism
of Fgf8 regulation can clearly not be operating in the presumptive
telencephalic neurectoderm or the adjacent epidermal ectoderm. We hypothesize
the existence of additional repressors, both in the neurectoderm and in the
epidermal ectoderm, that cooperate with Atr2 to silence Fgf8 in these
tissues. Although there is a mild upregulation of Atr2 in the isthmus at E9.5
(Fig. 3E), om mutants
show correct restriction of Fgf8 transcripts to the isthmus at E9.0
(not shown).
Atr2 is required for normal AER formation
The AER is a specialization of the ectoderm that lies at the dorsoventral
boundary of the developing limb bud and that is involved in controlling limb
pattern (Capdevila and Izpisua Belmonte,
2001). Atr2 is expressed at high levels in the AER at E9.5
(Fig. 3H-J). When Atr2 function
is reduced, AER precursors aggregate abnormally in the center of the limb bud
instead of lining up along the boundary
(Fig. 3J). Although death of
mutants prior to limb bud outgrowth prevented studying mutant limb phenotypes,
Atr2 appears to be required for the correct initial setup of this limb bud
organizer. The generation of conditional alleles of Atr2 will allow us to
examine the effects of loss of Atr2 on limb patterning.
Does Atr2 play a role in neural degeneration in DRPLA?
Atr2 is strongly expressed in developing neurons
(Fig. 3K-M), suggesting it may
interact with Atr1 in regulating neuronal development and/or function. A
growing body of data points towards dysregulation of transcription as an
important pathogenic mechanism in polyglutamine diseases
(Freiman and Tjian, 2002).
Polyglutamine-expanded proteins can sequester transcriptional regulators like
CBP and thereby disrupt transcriptional control
(Nucifora et al., 2001
). Atr1
and Atr2 bind directly to each other, and their binding is stimulated by
expanded glutamine in Atr1 (Yanagisawa et
al., 2000
), suggesting that the neural degeneration in DRPLA
involves depletion of Atr2 and derepression of Atr2 target genes. Dissecting
the role of Atr2 in neurons, alone and in conjunction with Atr1, is likely to
provide insight into the normal cellular functions of these proteins, and by
extension help clarify the molecular mechanisms of neurodegeneration in DRPLA.
The study of the neuronal functions of Atr2 will require construction of
conditional alleles to allow survival of mutant animals beyond E9.5, which is
currently in process.
Atr2 may function as a co-repressor
Although mammalian genomes contain two atrophin genes, the
Drosophila and C. elegans genomes contains only one each,
Atro and egl-27, respectively
(Ch'ng and Kenyon, 1999;
Erkner et al., 2002
;
Herman et al., 1999
;
Solari and Ahringer, 2000
;
Solari et al., 1999
;
Zhang et al., 2002
). Atro
functions during early development as a co-repressor for the Eve transcription
factor (Zhang et al., 2002
),
and later regulates transcription in the planar polarity pathway
(Fanto et al., 2003
). The
widespread expression pattern of Atro, and the diverse developmental
defects in Atro and egl-27 mutants, strongly suggest that
other sequence specific transcription factors also use atrophins as
co-repressors. Interaction between Atro and Eve occurs through the C-terminal
domain of Atro. An analogous interaction in mammals has also been detected;
yeast two-hybrid screens with a mammalian transcriptional repressor have
pulled the C-terminal portion of both Atr1 and Atr2 from libraries (V. J.
Bardwell and M. W. Murphy, personal communication). Presumably then, the
co-repressor function of atrophins is conserved in vertebrates.
Further supporting the co-repressor hypothesis, is the conservation of
N-terminal sequences between Atro, EGL-27, Atr2 and Mta2. We have shown that
the N-terminal, Mta2-homologous region of Atr2, excluding the GATA domain, is
required and sufficient for recruitment of Hdac1 in 293 cells
(Fig. 5B). We have also shown
that endogenous Atr2 recruits Hdac1 in the mouse embryo
(Fig. 5C). Association of Atr2
with a histone deacetylase in vivo is significant, because such proteins alter
the structure of chromatin and silence transcription
(Ng and Bird, 2000),
supporting the idea that Atr2 functions as a transcriptional repressor during
development. The fact that Atr2 associates with Hdac1 but not with RbAp46 or
RbAp48 suggests that Atr2 acts in a protein context distinct from NuRD.
Combining all the available data, we hypothesize that the mechanism of gene
silencing by Atr2 involves binding directly to sequence-specific transcription
factors, which brings associated histone deacetylases to bear on specific
promoters.
We have also observed an association between endogenous Atr2 and Atr1 in mouse embryos and MEFs, providing evidence that these two proteins form heterodimers during embryonic development. As full-length Atr2 was absent from om MEFs but an association with Atr1 was still observed, it is clear that Atr2S is able to associate with Atr1. It is unclear why an association between Atr2 and Hdac1 was not seen in MEFs. One possibility is that Atr1 blocks the ability of Atr2 to associate with Hdac1. Additional binding studies are necessary to determine whether Atr2, Atr2S, Atr1 and Hdac1 all exist in a complex together, or if some components are mutually exclusive.
In summary, we have analyzed the phenotype of mutant alleles of Atr2, and we have found that Atr2 is required for normal embryonic patterning and for the specific regulation of Shh in its anterior domain. We have shown that Atr2 is required for establishment of proper dorsoventral pattern in the anterior neural plate. We have provided evidence that mutant anterior neural pattern results from the disruption of two signaling centers, the anterior midline and the ANR. Finally, in accord with a proposed role for atrophin family members as transcriptional co-repressors, we have presented biochemical evidence that Atr2 can recruit histone deacetylase in vivo. Taken together, these data suggest that the embryonic defects observed in Atr2 mutants are caused by the loss of a novel histone deacetylase complex and the subsequent derepression of developmentally important genes.
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ACKNOWLEDGMENTS |
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