Department of Molecular and Cell Biology, University of California, Berkeley, CA 94720, USA
Author for correspondence (e-mail:
harland{at}socrates.berkeley.edu)
Accepted 23 December 2004
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SUMMARY |
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Key words: SRp38, Neurogenesis, Notch/Delta, Neurogenin, NeuroD, Ribosomal RNA, Xenopus
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Introduction |
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In mammals, translational regulation has been shown to be important for the
development of the nervous system. Disruption of the basal splicing machinery
has been implicated in two common human diseases, retinitis pigmentosa and
spinal muscular atrophy (reviewed by
Faustino and Cooper, 2003). A
number of human neurological diseases have also been linked to misregulation
of splicing. For example, nucleotide mutations within the gene MAPT,
which encodes the neuronal microtubule associated tau, result in the
selection of an alternative splice site and these mutations have been
implicated in Parkinson's disease, frontotemporal dementia and others
(Dumanchin et al., 1998
). When
NOVA-1, a KH domain-containing RNA-binding protein, is mutated in mice,
splicing of two neuronal receptors, GlyR
2 and GABAA, is
defective (Dumanchin et al.,
1998
; Jensen et al.,
2000
). However, despite these studies, our knowledge of the
developmental regulation and biological functions of vertebrate RNA binding
proteins is limited.
Until recently, SR proteins had been suggested to have two distinct roles
during pre-mRNA processing. First, during selection of alternative splice
sites, increasing the amount of SR proteins both in vivo and in vitro biases
splice site selection towards sub-optimal upstream splice acceptors
(Ge and Manley, 1990;
Krainer et al., 1990
;
Sun et al., 1993
;
Tian and Maniatis, 1993
).
Second, SR proteins select exonic splicing enhancers in a process requiring
sequence specific binding of the RNA recognition motif (RRM)
(Graveley, 2000
). Recent work
suggests that SR proteins may also be important in regulation of
mRNA-processing events through translation
(Sanford et al., 2004
). In
general, SR proteins contain two functionally separable domains: one or more
RNA-recognition motifs (RRMs); and a serine-arginine rich motif (RS)
(Caceres and Krainer, 1993
;
Tacke and Manley, 1995
;
Zuo and Manley, 1993
). The RRM
binds to target RNAs in a sequence-specific manner, while the SR domain
interacts with partner proteins, presumably for the recruitment of other
splicing machinery (reviewed by Graveley,
2000
). Transcription of SR proteins has been shown to be tissue
specific and developmentally regulated
(Hanamura et al., 1998
;
Tian and Maniatis, 1993
). The
proteins themselves are regulated by phosphorylation
(Colwill et al., 1996
;
Wang et al., 1998
;
Xiao and Manley, 1998
). Thus,
the activity of the SR proteins can be controlled rapidly by intracellular and
extracellular signals (Du et al.,
1998
).
The identification of SRp38 (also known as NSSR-1, TASR-2,
fusip1 and SRrp40) uncovers a surprising new role for SR
proteins in splicing regulation (Clinton
et al., 2002; Komatsu et al.,
1999
; Shin and Manley,
2002
; Yang et al.,
2000
; Yang et al.,
1998
). Unlike the other SR proteins identified, SRp38
cannot activate splicing; in fact, it is a potent repressor of splicing when
dephosphorylated. SRp38 is specifically dephosphorylated during mitosis and
heat shock and its activity is required for both mitotic and stress-related
splicing repression (Shin et al.,
2004
; Shin and Manley,
2002
).
SRp38 was first identified in a yeast two-hybrid screen for
proteins interacting with TLS/FUS
(Yang et al., 2000;
Yang et al., 1998
).
TLS/FUS (translocated in liposarcoma/fusion protein) is a
DNA/RNA-binding protein implicated in the most common chromosomal
translocation in liposarcomas. It has been shown to bind both RNA pol II and
splicing factors, suggesting both a link between transcription and splicing
and a potential role in aberrant splicing during carcinogenesis. Yang and
colleagues later showed that the two TASR isoforms were generated by
alternative splicing (Clinton et al.,
2002
). Mouse NSSR-1 (long) and -2 (short)
(Fusip1 Mouse Genome Informatics) were discovered in a search
for SR proteins in a neural specific cDNA library
(Komatsu et al., 1999
). A
third group found SRrp40 in a database search for SR proteins
(Cowper et al., 2001
).
Finally, Shin and Manley identified SRp38 in a yeast two-hybrid
screen with the human splicing regulators Tra2
and
Tra2ß (Shin and Manley,
2002
). All of these groups had observed the inhibitory splicing
abilities of SRp38 but Shin and Manley found that SRp38 is
specifically activated by dephosphorylation during mitosis and heat shock
(Shin et al., 2004
). They also
observed that SRp38 is required for the inhibition of pre-mRNA splicing that
occurs in mitotic cell extracts. Dephosphorylation of SRp38 might result in
weakened interactions with other SR proteins; however, SRp38 is likely to also
bind target RNAs in a sequence-specific fashion, as Shin and Manley were able
to identify a high-affinity target sequence by SELEX and use this sequence to
deplete SRp38 from mitotic extracts.
Despite this progress in understanding the biochemical regulation and
activity of SRp38, many questions remain. In this study, we characterize a
novel mechanism underlying the control of neurogenesis. Primary neuronal
differentiation in Xenopus is governed by a sequential cascade of
basic helix-loop-helix (bHLH) transcription factors beginning with
neurogenin, a vertebrate homolog of Drosophila atonal.
neurogenin activates transcription of a series of bHLH factors including
neuroD. This proneural pathway results in the expression of markers
of neuronal differentiation (Kintner,
2002; Ma et al.,
1996
). Using an expression screening approach, we identified
Xenopus SRp38 as a modulator of neurogenesis
(Grammer et al., 2000
). We
found that SRp38 is expressed in the neural plate of the
Xenopus embryo at the time of primary neurogenesis and is itself
induced by neuroD. SRp38 regulates neurogenesis at a crucial step
downstream of neurogenin activity and this regulation is Notch dependent.
Depletion of SRp38 activity results in a context-dependent increase in
neurogenesis, which suggests that SRp38 is a negative feedback regulator that
is induced by neuroD during neuronal differentiation. Finally, SRp38
interacts with a 289 nucleotide sequence in domain V of the 28s rRNA, which
includes the peptidyltransferase domain of 28S ribosomal RNA. This suggests
that SRp38 may act by regulation of ribosome biogenesis or function in the
developing nervous system.
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Materials and methods |
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Cloning of SRp38
Xenopus SRp38 (clone #19A5) was originally identified in an
expression screen from a Xenopus laevis neurula stage (stage 19-22)
cDNA library (Grammer et al.,
2000; Mariani and Harland,
1998
). Single amino acid substitutions, noted in text, were made
using DPN mutagenesis (Braman et al.,
1996
).
Synthesis and injection of mRNA
Synthetic capped mRNA was generated using the mMessage mMachine Kit
(Ambion). RNA was precipitated first with one half volume of 6M LiCl, washed
in 80% ethanol, resuspended in 100 µl DEPC-treated water, reprecipitated
with ammonium acetate and washed again in 80% ethanol to ensure removal of all
LiCl. All synthetic mRNA was quantified using incorporation of trace amounts
of 32P-UTP and master stocks were stored at a concentration
of 1 µg/µl at 80° until further dilution. Each mRNA (1 µg)
was analyzed by agarose gel electrophoresis for quality and quantity
assurance. In general, RNAs were then diluted in DEPC-treated water and
injected in 5 nl volumes at the one- to four-cell stages.
Ectodermal `animal cap' explants and RT-PCR
For ectodermal explants, mRNA was injected into the animal hemisphere at
the one-cell stage. Embryos were then aged to blastula stages (stage 9) either
at 25°C (5 hours) or at 12°C (overnight). Animal caps (400 µm)
were cut from devitellinized embryos using eyebrow knives (courtesy of Dale
Frank) or from non-devitellinized embryos using the Gastromaster (Xenotek
Engineering). Explants and untreated stage control embryos were then cultured
in 75% NAM (+gentamycin 500 µg/ml) until indicated stages and harvested for
reverse transcriptase-polymerase chain reaction (RT-PCR) or mRNA in situ
hybridization (described below). For RT-PCR analysis, RNA was extracted and
RT-PCR performed as described (Wilson and
Melton, 1994
). All PCRs were performed at 25 cycles, except
EF1
and MA at 21 cycles, Sox3 at 23 cycles and S11 at 18 cycles. The
following RT-PCR primer pairs were used.
Morpholino oligonucleotides
SRp38 was BLASTed against the NCBI expressed sequence tag (EST)
database. ESTs were downloaded and aligned in ClustalX to determine sequence
similarity, in particular in the 5'UTR and at the start codon. This
allowed us to identify paralogous genes (owing to the presumed
pseudotetraploidy of X. laevis) and design oligonucleotides targeted
towards one or both copies of the genes. The following sequences were used to
block translation of SRp38: AMO1, 5'-GCG GCC TTG AAT AGC GAG ACA
TCC T-3'; AMO2, 5'-CAA GCG CCA CAC TTC GAC AAC AAT
A-3'. The control oligonucleotide is 5'-CCT CTT ACC TCA GTT ACA
ATT TAT A-3'. All morpholino oligonucleotides were ordered from
GeneTools, and resuspended at 1 mM concentrations in DEPC-treated
0.1xMR. AMOs were further diluted in DEPC-treated water at the
concentrations indicated. Both AMO1 and AMO2 inhibited in vitro translation of
the original SRp38 cDNA but not that of a Myc-tagged SRp38
that lacked the 5'UTR. Both AMOs also inhibited activity of SRp38 when
co-injected in vivo.
Whole-mount RNA in situ hybridization and immunohistochemistry
Embryos were fixed for 1 hour in MEMFA, dehydrated in methanol and stored
at 20°C until further processing. In some cases, the vitelline
membrane was removed before fixation using watchmaker's forceps.
RNA in situ hybridization was performed using a multibasket technique
previously described (Sive et al.,
2000) with the following modifications. In situs were developed
using BM Purple (Boehringer Mannheim). For certain probes (neuroD),
in situs were developed using 0.45 µl NBT (stock 75 mg/ml in 70%DMF) and
3.5 µl BCIP (stock 50 mg/ml in 100% DMF) per ml of AP buffer
(Lee et al., 1995
). All in
situs were postfixed in Bouin's Fix. Embryos were rinsed in
1xPBS-0.1%Tween or TE-buffered 70% ethanol then bleached in
0.5xSSC, 5% formamide and 1%H2O2.
For antibody staining, embryos were aged to the indicated stages, dissected out of their vitelline membranes, fixed for 45 minutes in 1x MEMFA and dehydrated in methanol. Embryos were rehydrated in a stepwise fashion and subsequently washed in 1x PBS, 1xPBS + 0.1% Triton X-100 (PBS-Tr) and preblocked for 1 hour in 1xPBSTr+10% heat inactivated goat serum. Primary antibodies were then added at the indicated dilutions and incubated overnight at 4°C. Embryos were washed several times in PBS-Tr and then incubated with secondary antibodies, again as indicated for 1 hour at room temperature. Samples were then washed in PBS-Tr repeatedly for at least 5 hours at room temperature. HRP activity was revealed using H2O2 and diaminobenzidine (DAB) as the histochemical substrate.
TUNEL staining
To assess the amount of apoptosis in the embryo, we used the TdT-mediated
dUTP-digoxigenin nick end labeling (TUNEL) technique to label dying cells in
situ. Embryos were analyzed essentially as described by Hensey and Gautier
(Hensey and Gautier, 1997).
The anti-digoxigenin antibody incubation was carried out in 2% BM Block
(Boehringer Mannheim) in 1x MAB and all subsequent steps were performed
essentially as those for in situ hybridization (above).
RNA immunoprecipitation
In an initial pilot experiment, 300 embryos each were untreated or injected
with 250 pg of synthetic flag-SRp38 mRNA or flag-SRp38* at the one- to
two-cell stage and aged to neurula stages. Co-immunoprecipitated RNAs from the
pilot experiment were radiolabelled using T4 RNA ligase for analysis (see
below). Subsequently, in two independent experiments, 1000 embryos in each
group were treated as above. In each experiment, all three sets of embryos
were lysed in 10 µl per embryo of homogenization buffer (HB: 15 mM HEPES pH
7.6, 10 mM KCl, 1.5 mM MgCl2, 0.5 mM EGTA, 44 mM sucrose) with the
addition of 3 mM Na2VO4, 1 mM DTT, 20 µg/ml
aprotinin, 10 µg/ml leupeptin, 4 µg/ml pepstatin, 0.75 mM PMSF and
1x vanadyl ribonucleoside complex (Gibco). Lysates were spun for 10
minutes at 4°C, 20,817 g and supernatants removed in order
to separate yolk. Anti-Flag antibody M2 (Sigma) was then added to a final
concentration of 2.8 ng/µl and incubated for 1 hour at 4°C. Protein A
Sepharose beads which had been equilibrated in HB (300 µl each sample) were
then added and again incubated for one hour at 4°C. Samples were
centrifuged for 15 minutes at 106 g at 4°C and washed
three times in HB. To elute the bound protein, beads were incubated three
times for 1 hour each in 1 ml of HB containing 30 µl of 5 µg/µl Flag
peptide (Sigma). A final elution was carried out overnight at 4°C. All the
supernatants were phenol:chloroform extracted and remaining nucleic acids were
precipitated in the presence of isopropanol, NH4OAc and glycogen.
Aliquots were removed at each step and saved for protein gel analysis. RNAs
co-immunoprecipitated in these experiments were cloned for subtractive
hybridization.
cDNA synthesis and subtraction of immunoprecipitated RNAs
RNAs were immunoprecipitated in two independent experiments using the SMART
PCR cDNA synthesis kit from Clontech. Efficiency of first and second strand
synthesis of cDNA was monitored by 32P-dCTP according to
standard protocols. Subtractive hybridization was performed essentially as
described by Diatchenko et al. (Diatchenko
et al., 1999
) using Flag-SRp38* (mutant RNP) as driver and
Flag-SRp38 as tester. The product of the subtractive hybridization was then
amplified by suppression PCR (Clontech).
Luciferase assays and 35S-methionine incorporation
At the one-cell stage, embryos were uninjected, injected with SRp38 or
mutant SRp38. Embryos were allowed to divide and each set of embryos was
subsequently injected with luciferase DNA. Embryos were then cultured and
harvested for luciferase readings at stage 21.
For 35S-methionine incorporation, embryos were injected at one cell stage with 250 pg luciferase, SRp38 or SRp38* mRNA. At stage 17, 35S-methionine was added to the culture media. Embryos were then lysed, proteins were precipitated and incorporated 35S-methionine was counted.
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Results |
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Sequencing of the clone 19A5 revealed an open reading frame of 239 amino acids (Fig. 1A). Analyses of the sequence by BLAST search showed that the RNA-binding domain of 19A5 is 87% identical to that of mouse SRp38. The serine-arginine rich (RS) domain is a region of low complexity and identity drops off considerably (Fig. 1B).
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SRp38 can inhibit differentiation during early germ layer formation
SRp38 was identified from two effects on development: induction of
ectopic pigmentation and disruption of the general neural marker nrp1
(Fig. 2A)
(Grammer et al., 2000). Closer
analysis revealed that differences in injection site could account for these
different activities: overexpression of SRp38 in the animal hemisphere at an
early stage (targeting the neurectoderm) resulted in the loss of nrp1
expression at neurula stages (Fig.
2A). Injection into the marginal zone (targeting presumptive
mesoderm) resulted in patches of ectopic pigmentation
(Grammer et al., 2000
).
Morphologically, the ectopic pigmentation appeared to result from an
accumulation of cells in the mesodermal tissue, rather than excess melanin
synthesis. We also found that mouse and human SRp38 recapitulates these
activities (data not shown).
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We then overexpressed SRp38 in the presence and absence of neuroD mRNA. SRp38 was unable to block the induction of synaptobrevin II by neuroD. Injection of neuroD mRNA induces ectopic expression of synaptobrevin II (Fig. 4C), while co-expression of SRp38 with neuroD does not inhibit neuroD induction of synaptobrevin II (Fig. 4C). Therefore, SRp38 was able to block the activity of neurogenin (Fig. 4A,B) but not that of neuroD (Fig. 4C), suggesting that neuroD is able to act downstream of SRp38.
Activation of Notch Signaling in tissues expressing SRp38
This activity seen above mimics activation of the Notch signaling pathway,
which also blocks the ability of neurogenin but not neuroD
to induce neuronal ß-tubulin (schematic in
Fig. 4D) (Ma et al., 1998;
Olson et al., 1998
). To test
whether SRp38 could be activating Notch signaling, we examined its effect on
lateral inhibition of the ciliated epidermis. Targeting the mRNA to the
ventral epidermis of the embryo resulted in fewer ciliated epidermal cells
(revealed by non-specific ß-tubulin expression;
Fig. 5A, right embryo), a
hallmark of increased Notch signaling
(Deblandre et al., 1999
).
Targeting the mRNA to the neural plate resulted in the loss of primary neurons
(marked by neural specific ß-tubulin,
Fig. 5A, center embryo), again
similar to ectopic Notch activation
(Chitnis et al., 1995
).
|
We also determined that Srp38 caused ectopic expression of Id3
mRNAs (Fig. 5E), and activation
of esr-1 transcription in animal caps (data not shown). Both
Id3 and esr1 are direct targets of Notch signaling and
expression of Id3 is often correlated with proliferating,
undifferentiated cell types
(Reynaud-Deonauth et al.,
2002; Wettstein et al.,
1997
). Id genes have been shown to inhibit differentiation by
binding to and inhibiting the transcriptional activities of bHLH proteins.
Id genes are spliced and Xenopus Id3 has been shown to be
cytoplasmically polyadenylated, suggesting that Id genes undergo
post-transcriptional regulation (Afouda et
al., 1999
). Using northern blotting, we looked for changes in the
mRNA of Id2, Id3 and Delta in samples of tissues
overexpressing SRp38. We analyzed total extracted RNA and polyA+ selected RNA
from whole embryos but found no change in the size or polyadenylation of these
genes (data not shown). We also examined the splicing status of Id2 and Id3 by
designing intron-spanning primers, but found no changes in the splicing of
these genes (Fig. 5F; data not
shown).
Inhibition of SRp38 translation using antisense morpholino oligonucleotides
To study the requirement for SRp38 activity during neurogenesis,
we used two different methods to inhibit SRp38 function. First, we
designed two antisense morpholino oligonucleotides (AMO) targeting the
SRp38 5'UTR and start codon (AMO1 and AMO2 see Materials and
methods). Antisense morpholino oligonucleotides bind to target RNAs in a
sequence-specific fashion and prevent translation
(Heasman et al., 2000). Both
AMOs inhibited the activity of SRp38 in vivo when co-injected with
exogenous mRNA and both AMOs inhibited translation of SRp38 in vitro
(data not shown). All loss-of-function experiments were initially performed
with AMO1 at a dose of 80 ng per embryo (control embryos were injected with a
control oligonucleotide, see Materials and methods). Results were then
independently confirmed using AMO2 (80 ng). Lower doses of the individual AMOs
had only subtle effects. We then found that a mixture of AMO1 and AMO2 (20 ng
to 40 ng each) was most effective. All experiments pictured used a combination
of the two AMOs.
We found that reduction of SRp38 translation in vivo using a
cocktail of AMOs (40 ng each) did not perturb expression of neuronal
ß-tubulin at early stages
(Fig. 6A, upper right). We
hypothesized that SRp38 acts in parallel with other mechanisms to
regulate the amount of neurogenesis in vivo and, thus, chose to examine loss
of SRp38 in sensitized assays. Expression of
Deltastu in the embryo results in increased neurogenesis
(Chitnis et al., 1995)
(Fig. 6A, lower left). When we
depleted SRp38 from embryos concurrently expressing
Deltastu we found that these embryos had an increase in
primary neurons above the levels induced by Deltastu alone
(Fig. 6A, lower right).
|
Inhibition of SRp38 activity using a function blocking consensus binding motif
Next, we took advantage of an SRp38 consensus binding motif (C3)
to inhibit SRp38 function in vivo. This sequence has been shown to
deplete SRp38 protein from cell extracts
(Shin and Manley, 2002). We
predicted that Xenopus SRp38 would also be able to bind specifically
to the consensus motif and that excess C3 might block SRp38 activity (mouse
and human SRp38 recapitulate the activity of Xenopus SRp38
in our assays, not shown). We found that co-injection of capped RNA from this
construct is sufficient to inhibit the differentiation-blocking activity of
injected SRp38 in vivo (Fig.
6B) in a dose-dependent fashion. Injection of SRp38
inhibits expression of the neural marker nrp1
(Fig. 6B;
Fig. 2B). When 5 ng of C3 was
co-injected with 250 pg of SRp38, nrp1 expression was rescued in 100%
(22/22) of the embryos (Fig.
6B). Lower doses of C3 gave less penetrant phenotypes [3.75 ng,
91% (20/22) rescue; 2.5 ng, 87% (20/23) rescue; 1.25 ng, 64% (16/25) rescue],
while injection of other capped mRNA sequences did not inhibit SRp38
activity (data not shown).
We then used the C3 sequence to analyze the requirement for endogenous SRp38 activity during neurogenesis. Using RT-PCR on ectodermal explants, we found that depletion of SRp38 activity in conjunction with ectopic expression of neurogenin resulted in an increase in neuronal ß-tubulin expression above that normally induced by neurogenin alone (compare lanes 5 and 4 in Fig. 6C). Conversely, we saw a comparable decrease in Id3 mRNA, marking an equivalent reduction in undifferentiated cells (again, compare lanes 5 and 4 in Fig. 6C). Thus, although reduction of SRp38 does not affect primary neurogenesis in vivo, there is a clear role for SRp38 in regulating levels of neurogenesis in response to changing levels of Deltastu or neurogenin. These data is consistent with a role for SRp38 as part of an inhibitory feedback mechanism during neuronal differentiation.
Effects of SRp38 on proliferation and cell death
Because SRp38 acts as a splicing repressor during mitosis
(Shin and Manley, 2002), we
considered the possibility that the phenotype we saw was due to selective
repression of splicing within the affected cells. This splicing repression
might prevent cells from exiting the cell cycle and differentiating;
alternatively, splicing repression might induce the block to differentiation
characteristic of mitotically active cells without triggering proliferation.
The process of neurogenesis is particularly sensitive to this effect, as it is
crucial that cells be able to exit the cell cycle in order to become neurons
(Vernon et al., 2003
).
Thus, cells expressing SRp38 might have several alternative fates. As these cells may not properly splice (and subsequently translate) appropriate genes, they might remain in an undifferentiated state or undergo apoptosis. Another possibility is that there would be an increase in mitotic cells, because splicing and translational silencing are characteristic of mitosis.
In order to determine whether SRp38-expressing cells undergo mitosis, we
used an anti-phosphorylated Histone H3 (anti-pH3) antibody that specifically
marks mitotic cells. We saw no change in nti-pH3 staining in
SRp38-injected embryos when compared with controls (data not shown). Because
perturbations in the cell cycle can often induce apoptosis
(Gartel and Tyner, 2002
), we
also analyzed injected embryos by TUNEL (TdT-mediated dUTP-digoxigenin nick
end labeling) to assess the amount of programmed cell death. There was a
clear, though variable, increase in the number of TUNEL-positive cells in
SRp38-injected embryos (see Fig.
6D); however, most cells expressing SRp38 were not TUNEL
positive (data not shown), suggesting that increased apoptosis in these
tissues might be secondary to the inability to differentiate. Finally, we saw
no change in the expression of the cell cycle regulators p27xic1
(Vernon et al., 2003
) or
Cyclin D1 (Ratineau et al.,
2002
) in neuralized tissues treated with C3
(Fig. 6C, compare lane 5 with
lane 4).
SRp38 is upregulated by neuroD
If SRp38 is indeed a component of a mechanism to limit neuronal
differentiation, we would expect it to be regulated by the neurogenic genes
(schematic, Fig. 7D).
Consistent with this, we found that increasing amounts of neuroD resulted in
increased amounts of SRp38 in the neural plate
(Fig. 7A,C) and in ectodermal
explants (Fig. 7B). Thus,
neuroD, a transcription factor that acts at the end of the neurogenic cascade,
induces expression of SRp38. SRp38 then inhibits upstream
transcription factors such as neurogenin
(Fig. 4), preventing excessive
neurogenesis. Similarly, neurogenin and neuroD induce
expression of Delta and this activation is also thought to limit the
amount of neurogenesis in the embryo (Ma
et al., 1998) (data not shown).
|
We then took a broader biochemical approach towards the identification of SRp38 targets. Flag-tagged wild-type SRp38 or a crippled RNA-binding mutant SRp38 were expressed in embryos and used to immunoprecipitate associated RNAs. In the SRp38 mutant (SRp38*), all four conserved phenylalanines and tyrosines in the RNA-binding domain were converted to alanines, changes that should abrogate sequence-specific binding (Fig. 8A). To identify targets, embryos were injected with Flag-SRp38 or Flag-SRp38* and allowed to develop to mid-neurula stages. Embryos were then lysed, Flag-SRp38 was immunoprecipitated with an anti-Flag antibody (Sigma, M2) and eluted with excess Flag epitope. The co-immunoprecipitated RNAs were then extracted, and the population precipitated with SRp38 was subtracted with the control population (SRp38*) prior to being cloned.
|
Specific binding of this sequence to SRp38 was confirmed by immunoprecipitation followed by gene-specific RT-PCR (Fig. 8B). Control, Flag-SRp38 or Flag-SRp38*-expressing embryos were lysed and immunoprecipitated with anti-Flag (M2) agarose beads. IP products were then subjected to RT-PCR for the S11 sequence. S11 was not immunoprecipitated in control uninjected embryos (lane 1) but was efficiently brought down in the wild-type SRp38 injected embryos (lane 2). A significantly smaller amount of S11 was also brought down with mutant SRp38*, and this may reflect interactions of S11 with SRp38 partner proteins. A proportion of the samples was also used for western blotting to determine efficiency of initial protein expression (input) and immunoprecipitation (IP). Thus, although SRp38 may inhibit splicing, it probably also interacts with the ribosome either during ribosome biogenesis or functionally during ribosome activity. SRp38 activity may halt ribosome function during mitosis, or it could be preserving partially processed pre-rRNA during mitosis.
Based on its interaction with the ribosome, we considered the possibility that SRp38 might influence translation in the embryo. We used two assays to test this possibility: first, we measured the rate of translation of a luciferase DNA reporter (Fig. 8F); and second, we determined the rate of 35S-methionine incorporation in treated embryos (Fig. 8G). Embryos injected with SRp38 show some decrease in the activity of the luciferase plasmid reporter when compared with mutant SRp38* (Fig. 8F). 35S-methionine incorporation was also mildly decreased in SRp38-injected embryos (Fig. 8G). From stage 17 to 21, there were no significant differences. By stage 28 (Fig. 8G, right) there was a small difference between SRp38- and SRp38*-injected embryos.
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Discussion |
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Our studies provide significant insights into the mechanism of SRp38
function, and control of neurogenesis. SRp38 had previously been shown to be a
neural-specific splicing repressor
(Komatsu et al., 1999) that is
activated in response to stress and cell cycle regulation
(Shin et al., 2004
). However,
these previous studies did not place SRp38 within a signaling cascade or
identify any of its biological targets. Our findings implicate SRp38 in the
regulation of vertebrate neurogenesis, linking it to signaling through both
proneural genes and to the Notch/Delta pathway. We show that SRp38 can inhibit
neural differentiation when overexpressed in the Xenopus embryo but
that SRp38 does not affect neural induction or competence
(Fig. 2). Furthermore, SRp38
inhibits the neurogenic activity of neurogenin but not that of neuroD
(Fig. 4). SRp38 inhibition of
primary neurogenesis requires active Notch signaling
(Fig. 6). Finally, neuroD
induces SRp38 expression, perhaps as a negative feedback mechanism to
limit neurogenesis in vivo (Fig.
7).
Notch is thought to maintain neural progenitor cells in an undifferentiated
state. Activation of Notch signaling during early neural patterning blocks
differentiation with a resulting decrease in neurogenesis, whereas in the
mouse, depletion of Notch1 in the mid-hindbrain boundary results in
the premature onset of neurogenesis (Ahmad
et al., 1997; Austin et al.,
1995
; Chitnis et al.,
1995
; Coffman et al.,
1993
; Lutolf et al.,
2002
). The findings that SRp38 acts in the context of Notch
signaling (Fig. 5) and that
active Notch signaling is required for inhibition of neurogenesis
(Fig. 6) suggest that a
combination of SRp38 and Notch signaling serve as a bridge between cell cycle
regulation and cell fates (Campos et al.,
2002
; Cereseto and Tsai,
2000
; Ohnuma et al.,
2002
; Ohnuma et al.,
1999
).
The control of proliferation and cell cycle progression is crucial for the
correct determination of the nervous system. Negative regulators of the cell
cycle, such as p27XIC1, have been shown to be required
precisely at the neurogenin to neuroD step during primary neurogenesis in the
Xenopus embryo (Vernon et al.,
2003). neuroD (also called Beta2) itself can induce cell cycle
withdrawal and neuroD/Beta2-null mice have an abnormal number of
proliferative cells in the small intestine
(Mutoh et al., 1998
). Thus, it
is possible that the regulation of SRp38 expression by neuroD is secondary to
neuroD control of the cell cycle. Conversely, positive regulators of the cell
cycle, such as cyclinD1, have been shown to repress the transcriptional
activity of neuroD in endocrine cells
(Ratineau et al., 2002
).
What is the requirement for SRp38 in the early embryo? The use of
high-affinity sequence-specific RNA binding had previously been used to
deplete SRp38 from mitotic and heat shocked cell extracts
(Shin et al., 2004). We used a
similar strategy to inhibit the activity of endogenous SRp38 in the early
embryo, as well as two different antisense morpholino oligonucleotides (AMOs)
to inhibit translation of embryonic mRNA
(Fig. 6). Depleting SRp38
activity when Delta activity is inhibited resulted in a synergistic increase
in neurogenesis (Fig. 6). There
was little effect on the development of whole embryos when they were treated
with C3 or the morpholino oligonucleotides, consistent with the idea that
SRp38 overlaps in function with other regulators of neurogenesis during
development. In the absence of SRp38 function, it may be that the amount of
neurogenesis continues to be limited by `redundant' mechanisms, including
appropriate spatial and temporal transcription of the proneural genes, cell
cycle regulation of transcription and translation, and coordination of exit
from the cell cycle. SRp38 activity would serve to reinforce these other
controls and thus only become obvious when the system is sensitized.
An illuminating result of these studies is the identification of the 28S
rRNA as a target of SRp38. SRp38 is capable of sequence specific binding and,
while an 11 nucleotide consensus binding sequence is known (C3), it provides
no insight into the RNA targets of SRp38. Using in vivo immunoprecipitation
and subtraction, we have identified a 289 nucleotide RNA containing the
peptidyltransferase domain of the 28S ribosomal RNA. This region has been
shown to be selectively bound by RNA-binding proteins, in particular by the
bacterial DEAD box proteins DbpA and YxiN
(Kossen et al., 2002;
Nicol and Fuller-Pace, 1995
).
Binding of SRp38 to this region of the ribosomal RNA suggests a function in
ribosome biogenesis or function. It is possible that other regulators of
splicing bind to this region of the ribosome or, this function may be unique
to the mechanism of SR splicing repressors. Importantly, this result also
provides a link between splicing regulation and translation.
Developmental heterogeneity of ribosome composition has been well
documented in the slime mold Dictyostelium discoideum
(Agarwal et al., 1999).
Ribosomes in Dictyostelium spores are quantitatively and
qualitatively different from those in the vegetative state and these
differences result from a range of changes, from transcription of ribosome
components to protein modifications (reviewed by
Ramagopal, 1992
). Additional
ribosome diversity is likely to result from interaction with cellular
proteins. For example, the fragile X mental retardation protein
(FMRP) has been shown to bind to polyribosomes and regulate translational
efficiency (Khandjian et al.,
2004
; Stefani et al.,
2004
). It is possible that SRp38 binds to 28s rRNA in a similar
mechanism for generating ribosome diversity. In this way, the composition of
the ribosome (and presumably, translational efficiency) as well as the
repertoire of transcription and signaling factors present would generate
specificity in specific cell types. Many unanswered questions remain, such as,
what factors dictate the mRNAs that escape from the transcriptional and
translational silencing imposed during mitosis? We suggest that SRp38
is likely to play a role in this process. This hypothesis is based on the fact
that SRp38 is required for mitotic splicing inhibition and this function is
required for limiting neuronal progenitors. It is also possible that
SRp38 plays a positive role in allowing specific sequences to be
processed during mitotic silencing.
It is likely that SRp38 or genes like SRp38 play similar roles in other developmental processes. For example, it is known that Notch signaling and cell cycle control are also important for myogenesis. Our data suggest that SRp38 may affect both mesoderm and endoderm development (Fig. 3; Table 1), however, because SRp38 expression in the Xenopus embryo was mostly detected in the neural plate (Fig. 1), we have primarily studied its function in that context. Supporting a general role in the development of animals, the SRp38 protein is found in Xenopus, mouse and man and the proteins are functionally interchangeable in our hands (data not shown). This study elucidates a novel mechanism for the control of neurogenesis, while revealing a developmental role for the unusual SR protein SRp38, thus providing an important link between transcriptional and translational regulation of neuronal cell fates.
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ACKNOWLEDGMENTS |
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Footnotes |
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