|
|
|
|
|
|
|
||||
| Home Help Feedback Subscriptions Archive Search Table of Contents | ||||
First published online 13 June 2007
doi: 10.1242/dev.002485
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
1 Department of Molecular Medicine and Pathology, School of Medical Sciences,
The University of Auckland, Private Bag 92019, Auckland, New Zealand.
2 Max Planck Institute für Entwicklungsbiologie, Spemannstrasse 35/III,
72076 Tübingen, Germany.
3 Cancer and Haematology Division, The Walter and Eliza Hall Institute of
Medical Research, 1G Royal Parade, Parkville, Victoria 3050, Australia.
Author for correspondence (e-mail:
ps.crosier{at}auckland.ac.nz)
Accepted 21 May 2007
 |
SUMMARY |
|---|
 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Key words: Runx1, Runx3, Rad21, Scc1, Cohesin
|
INTRODUCTION |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
subunits (Runx1, Runx2 or Runx3) plus a common
non-DNA-binding CBFß subunit. The Runx component specifies the biological
activity of CBF; Runx1 is an essential regulator of hematopoiesis, Runx2 is
involved in osteogenesis, and Runx3 is important in neurogenesis and gastric
epithelial cell growth control (Blyth et
al., 2005
; Ito,
2004). Deregulation of Runx function is commonly associated with
disease. For example, Runx1 is involved in several chromosomal translocations
underlying human leukemias. In addition, changes in the dose of Runx1
can contribute to neoplasia (Blyth et al.,
2005). An important route toward understanding the pathology of
Runx-mediated cancers is the elucidation of factors that control either the
expression of Runx genes, or the activity/stability of their protein
products.
In early zebrafish embryogenesis, runx1 is expressed in two
discrete hematopoietic regions; the anterior lateral-plate mesoderm (ALM),
which generates primitive myeloid cells, and the posterior lateral-plate
mesoderm (PLM) from which primitive erythroid cells develop
(Hsia and Zon, 2005). By
around 18 hours post-fertilization (h.p.f.), cells in the PLM have migrated
medially to form a central rod of hematopoietic precursors: the intermediate
cell mass (ICM). Hematopoietic expression of runx1 is downregulated
in all but the most posterior cells of the ICM at 21 h.p.f., and subsequently
reappears in definitive hematopoietic precursors in the ventral wall of the
dorsal aorta by 24 h.p.f. (Kalev-Zylinska
et al., 2002). runx1 is also expressed in Rohon-Beard
(RB) mechanosensory neurons and in specific neuronal cells
(Kalev-Zylinska et al.,
2002).
Studies in zebrafish (Burns et al.,
2005; Gering and Patient,
2005) and mice (Nakagawa et
al., 2006) have shown that Runx1 is a transcriptional
target of Notch signaling during definitive hematopoiesis. In zebrafish,
Hedgehog signaling is required for the migration of hematopoietic progenitors
to the midline, and for the subsequent formation of runx1+
definitive precursors (Gering and Patient,
2005). Early PLM runx1 expression appears to be
downstream of a Hox pathway regulated by caudal-related homeobox
genes cdx1a and cdx4
(Davidson et al., 2003;
Davidson and Zon, 2006). In
addition the timely initiation (but not maintenance) of runx1
expression depends on the transcription factors Scl and Lmo2
(Patterson et al., 2007;
Patterson et al., 2005). Other
factors known to contribute to Runx gene regulation include the BMP signaling
pathway (Pimanda et al.,
2007), and epigenetic modifications, such as promoter methylation
(Lau et al., 2006;
Mueller et al., 2007).
To search for potential regulators of runx1 expression in zebrafish, we conducted an in situ hybridization-based haploid genetic screen of F1 females carrying mutations generated by ethylnitrosourea (ENU). We isolated a mutant, termed nz171, which lacks some neuronal, and all hematopoietic runx1 expression in early embryogenesis. Through a positional cloning and candidate gene approach, we determined that the gene underlying nz171 was rad21, an integral subunit of mitotic cohesin.
Cohesin is a protein complex composed of four major subunits: SMC1, SMC3,
RAD21 and SA1 (or SA2), which interact to form a giant ring-like structure.
Mitotic cohesin acts as a `molecular glue' to hold replicated sister
chromatids together until the onset of anaphase
(Losada and Hirano, 2005;
Nasmyth and Haering, 2005).
Cohesin also has a DNA repair function
(Watrin and Peters, 2006).
Intriguingly, it seems that cohesin has additional non cell cycle-related
functions (Dorsett, 2007;
Hagstrom and Meyer, 2003). In
Drosophila, loss of Nipped-B, a protein that loads cohesin onto
chromosomes, affects expression of the cut gene
(Dorsett et al., 2005;
Rollins et al., 2004). The
human ortholog of Nipped-B/Scc2 is NIPBL, and mutations in the NIPBL
gene, or in the cohesin subunit SMC1, underlie the dominant
developmental syndrome, Cornelia de Lange (CdLS)
(Dorsett, 2007;
Musio et al., 2006;
Strachan, 2005). Although
cohesin clearly has a role in development as well as in the cell cycle, the
mechanisms underlying its developmental function are unknown.
Our study provides the first evidence of cohesin-dependent gene regulation in a vertebrate. We found that Runx gene expression depends on the function of the whole cohesin complex and that, as expected, loss of zebrafish cohesin subunits interferes with sister chromatid cohesion during mitosis. Differentiated blood cells were deficient in nz171 mutants, and we found that some of these cells could be rescued by microinjection of runx1. nz171 mutants also had neuronal defects and rad21 was robustly expressed in specific regions of the brain at 48 h.p.f., which might indicate a role in neuronal development. We observed a potent dosage effect of Rad21 on downstream gene expression, and determined that halving the dose of the rad21 gene reduces the expression of runx1 and proneural genes ascl1a and ascl1b. Our findings highlight a new role for cohesin in gene expression and development.
|
MATERIALS AND METHODS |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
). AB strain
males were mutagenized with four treatments of 2.5 mM ENU as described
previously (Pelegri, 2002),
and were bred with AB females to obtain F1 fish. Eggs from F1 females were
fertilized in vitro with UV-irradiated sperm to produce haploid progeny
(Pelegri, 2002), which were
subjected to in situ hybridization with a runx1 riboprobe. The nz171
F1 female carrier was crossed to the WIK strain for meiotic mapping.
Chromosome localization of the nz171 mutation was performed as described
previously (Geisler, 2002). `Z
marker' PCR primer sequences were obtained from the WashU Zebrafish Genome
Resources Project
(http://zfish.wustl.edu/)
and subsequent mapping was performed as described by Geisler
(Geisler, 2002). The genomic sequence of zebrafish rad21 is available in Ensembl (gene ID ENSDARG00000006092). For mutation analysis, rad21 was amplified in overlapping segments from cDNA made from 100 pooled nz171 homozygous mutant embryos and 100 wild-type AB embryos (48 h.p.f.). Primers used were Forward-1, 5'-TTCAATGAGAGGAAACGGTTGC-3'; Reverse-1, 5'-ATGATGTCACTGTAGGGCTCGG-3'; Forward-2, 5'-TTCAATGAGAGGAAACGGTTGC-3'; Reverse-2, 5'-GGAAGAGGCTGGTCAAAATCG-3'; Forward-3, 5'-CCCACTTCGTCCTCAGTAAACG-3'; Reverse-3, 5'-TGGTCTTGCTGTCCAACTCCTTC-3'; Forward-4, 5'-AGACAACCGTGAGGCAGCATAC-3'; Reverse-4, 5'-AGTGGAGTCAAGCAGCGAGTAAAC-3'; Forward-5, 5'-ATGTGGAAGGAGACTGGAGGTGTG-3'; Reverse-5 5'-TACAATGTGGAAGCGTGGTCCC-3'; Forward-6, 5'-TCAGGACCAAGAGGAGAGAAGGTG-3'; Reverse-6, 5'-CAACAAGTAGTGAAACTGCGGAGTC-3'. Amplified fragments were cloned and sequenced, and the coding region mutation at nucleotide 829 was represented by two independent overlapping clones and fourfold sequence coverage.
In situ hybridization and histology
In situ hybridization was performed as described previously
(Kalev-Zylinska et al., 2002).
Embryos to be sectioned were dehydrated in an ethanol series, embedded in JB-4
(Polysciences), cut to a thickness of 5 µm and stained with Giemsa
(BDH).
Cell cycle procedures
Bromodeoxyuridine (BrdU) incorporation was performed on whole embryos as
described previously (Shepard et al.,
2004). Mouse monoclonal anti-BrdU antibody (Zymed) was detected
using a Vectastain ABC Kit (Vector Laboratories). Phosphorylated histone H3
was detected as described previously
(Shepard et al., 2004). TUNEL
staining was performed using the ApopTag Kit from Chemicon as described
previously (Shepard et al.,
2004).
Microinjection
Full-length zebrafish and human rad21 clones (RZPD) were subcloned
into pCS2+. Amplified full-length rad21G277X
was subcloned using a ZeroBluntTOPO Kit (Invitrogen), sequenced entirely to
confirm the mutation, and subcloned into pCS2+. mRNAs for
microinjection were generated using a mMessage mMachine Kit (Ambion), and 100
pg of each was injected into nz171 mutant and sibling embryos at the 1-cell
stage. Full-length zebrafish runx1 in pCS2+
(Kalev-Zylinska et al., 2002)
was transcribed as above and 200 pg was injected as above. Morpholino
oligonucleotides were obtained from GeneTools LLC and diluted in water. For
microinjection, 2 nl of morpholino was injected into the yolk of wild-type
embryos from the 1- to 4-cell stages. Morpholino oligonucleotides targeting
rad21 were: rad21UTRMO,
5'-CACTACACCTGGAAGAAAACAG-3'; rad21ATGMO
5'-TCCTGCTTCACCCGCATTTTGTAAC-3' (start codon
underlined); rad21Splx3MO 5'-GATACAATACCTGGGCGGAAAG-3'
(targets 3' donor of exon 3). Morpholino oligonucleotides targeting
smc3 were smc3UTRMO,
5'-GCACAAAACACTCCTCAGAAAC-3'; smc3ATGMO,
5'-TGTACATGGCGGTTTATGC-3' (start codon underlined);
smc3Splx1MO, 5'-GTGAGTCGCATCTTACCTG-3' (targets 3'
donor of exon 1) and smc3Splx5MO,
5'-TTTCTTACTGGAAGTCTGTTGTCAG-3' (targets 3' donor of exon
5). All morpholinos were effective over the range of 0.5-3.0 pmol
injected.
Immunofluorescence and confocal microscopy
For immunofluorescence, embryos were fixed and stained with anti-Rad21
(Chemicon International) 1:100, anti--tubulin (Sigma-Aldrich) 1:500,
and DAPI as described previously (Shepard
et al., 2004). FITC- or TRITC-conjugated secondary antibodies
(Sigma, 1:500) were used. Flat-mounted samples were imaged using a Leica TCS
SP2 confocal microscope.
Quantitative immunoanalysis and quantitative RT-PCR
For immunoblotting, embryos were deyolked in Ringer's solution with EDTA
and PMSF, and 20 µg total protein was loaded per lane. For embryos under 12
h.p.f., entire single, or pools of up to 10 embryos were lysed in loading
buffer. Sample processing and immunoblotting was performed as described
previously (Westerfield, 1995)
using anti-Rad21 (Chemicon, 1:500), or anti--tubulin (Sigma, 1:2000).
Horseradish peroxidase-linked secondary antibodies (Sigma, 1:2000) and
enhanced chemiluminescence were used according to the manufacturer's
instructions (ECL Plus, Amersham Biosciences, Inc.). Signals were analyzed
using Fuji LAS-3000 imager and Fuji Image Gauge software. For quantitative
PCR, total RNA from pools of 30-100 embryos was extracted using Trizol
(Invitrogen), DNAse-treated, and used to synthesize random-primed cDNA
(Invitrogen, SuperScript III). SYBR green PCR Master Mix (Applied Biosystems)
was used to amplify cDNA, and relative start quantities were normalized to
ß-actin and wnt5a expression. Samples were analyzed
using an Applied Biosystems Sequence Detection System 7900HT. Primers for
quantitative PCR were designed using the Primer Express program (Applied
Biosystems). Sequences are: runx1 forward,
5'-AGACGTCTCCATCCTGGTCGTA-3', reverse,
5'-CCGTCAGCTCTGGACAGTGTAA-3'; rad21 forward,
5'-CAGCATACAATGCCATCACCTT-3', reverse,
5'-ATTCAGGGTGAACTGCTGTGCTA-3'; ascl1a forward,
5'-GGGCTCATACGACCCTCTGA-3', reverse,
5'-TCCCAAGCGAGTGCTGATATTT-3'; ascl1b forward,
5'-CCACATGGTTCGACAGATACGA-3', reverse,
5'-CAGCATGCAGCAAATCAAAGAC-3'; ß-actin
5'-CGAGCAGGAGATGGGAACC-3', reverse,
5'-CAACGGAAACGCTCATTGC-3'; wnt5a forward,
5'-GTTCGGCCGCGTCATG-3', reverse,
5'-TCGACTCACAGCATTCACAACA-3'.
|
RESULTS |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
)
(Fig. 1E). Strikingly, its
expression was completely absent in nz171 mutants at the 14-somite stage
(Fig. 1H). Although both
runx1 and runx3 are downregulated in nz171, their common
binding partner, cbfb was expressed normally in neuronal and
hematopoietic tissues in equivalent embryos
(Fig. 1I compared with 1F).
This indicates that the cell types that would normally express runx1
and runx3 are present, and that loss of runx1 and
runx3 expression was not due to gross developmental defects. Early
neurogenesis was abnormal in nz171 embryos; the TGG failed to develop properly
(although they are specified), and the number of RB neurons was reduced by
30-40% (data not shown). Furthermore, although genes that mark blood vessel
formation were expressed (see Fig. S1 in the supplementary material), no
visible circulation developed. From the 20-somite stage, nz171 embryos started
to exhibit developmental delay, which became increasingly apparent until
homozygotes arrested in development at around 35 h.p.f.
(Fig. 1B,C). nz171 embryos
develop no further, and die at around 3 days postfertilization.
|
nz171 embryos have a nonsense mutation in the rad21 gene
We mapped the nz171 mutation to chromosome 16 between simple sequence
length polymorphism (SSLP) markers z25049 and z51029
(Fig. 3A). The cohesin subunit
rad21 also maps to this region, and because of the observed mitotic
defect, it was a strong candidate gene for the mutation. rad21 cDNA
was isolated from nz171 homozygotes, sequenced, and a mutation identified in
the coding region of the gene (nucleotide 829, exon 8) changing codon 277 from
GGA, specifying glycine, to the stop codon TGA (G277X)
(Fig. 3B and see Fig. S2 in the
supplementary material). An antibody directed against the C-terminal region of
human RAD21 detected a specific band in wild-type, but not in mutant embryos
(Fig. 3C). An insertional
mutant previously mapped to the rad21 locus
(Amsterdam et al., 2004) has a
severe early embryonic phenotype (ZFIN ID: ZDB-LOCUS-041006-4) similar to that
of nz171, consistent with these defects affecting the same gene.
In wild-type embryos, rad21 mRNA was detected by RT-PCR at the
oocyte stage (Fig. 3D),
indicating that the transcript is maternally deposited. Whole-mount in situ
hybridization analysis showed that rad21 was expressed throughout the
embryo in early embryogenesis (Fig.
3E,F; data also available on ZFIN,
http://zfin.org/).
Expression in the brain and posterior tail regions at 26 h.p.f. was
particularly robust, most likely because these are areas of active cell
division. By contrast, rad21 transcription was dramatically
downregulated in nz171 mutants (Fig.
3E, Fig. 8D),
probably owing to nonsense-mediated mRNA decay
(Chang et al., 2007). At 48
h.p.f., rad21 was strongly expressed in discrete areas of the brain,
the mandibular cartilage and branchial arches, the otic vesicle and developing
pectoral fins (Fig. 3F).
Although some cells in these regions would be proliferating rapidly (e.g. the
pectoral fins) it is surprising that this expression pattern is so specific.
This might reflect a tissue-specific function for Rad21 that is not related to
the cell cycle.
To provide further evidence that we had found a mutation in zebrafish rad21, we designed antisense morpholino oligonucleotides (MOs) directed against the rad21 transcript (see Fig. S2 in the supplementary material). MO-injected embryos (termed `morphants') phenotypically resembled nz171 mutants, had excess pH3+ cells and showed dramatic reduction in hematopoietic runx1 expression (Fig. 4A-I,M). Furthermore, we were able to rescue nz171 mutants with rad21 mRNA: nz171 mutants approached wild-type morphology at 48 h.p.f. following injection of zebrafish rad21 mRNA (Fig. 4J,K). In addition, hematopoietic runx1 expression was rescued (Fig. 4N compared with L), as were mitoses (see Table S1 in the supplementary material; data not shown). By contrast, rad21 mRNA containing the G277X mutation (zrad21G277X) was unable to rescue nz171 mutants (see Table S1 in the supplementary material). Significantly, runx1 expression was also rescued by human RAD21 mRNA (Fig. 4O and see Table S1 in the supplementary material), indicating conservation of Rad21 function in both the cell cycle and gene expression through evolution. These results provide conclusive evidence that the nz171 mutation affects the rad21 locus. We have named our mutant allele rad21nz171 accordingly.
|
In early rad21nz171 embryos, we observed normal
expression of many other early transcription factors essential for
hematopoiesis, such as cbfb (Fig.
1I) scl (also known as tal1 - ZFIN) and
pu.1 (also known as spi1 - ZFIN)
(Fig. 6A,B), cmyb, cebpa,
drl and gata2 (see Fig. S1 in the supplementary material). The
only other early hematopoietic transcription factor affected in
rad21nz171 was gata1, which is reduced to about
half its normal expression in 10-somite embryos
(Fig. 6A). A time-course
analysis of gata1 expression revealed that its onset is slightly
delayed in rad21-compromised embryos, with levels consistently
reduced during early embryogenesis (see Fig. S3 in the supplementary
material). This might, at least in part, be due to the positive autoregulation
of gata1 (Kobayashi et al.,
2001) by a transcriptional regulatory complex that also contains
Runx1 (Elagib et al., 2003;
Waltzer et al., 2003).
Normal expression of early blood markers in rad21nz171 embryos occurred at the same stage that runx1 expression is lost. Therefore, loss of runx1 is not due to a developmental delay, or loss of hematopoietic precursors in the regions where runx1 is normally expressed.
Markers of differentiated blood cells are reduced or absent in rad21nz171
In contrast to markers of early hematopoiesis, we determined that later
expressed markers, such as cebpg, lyz, hbbe3
(Fig. 6C,D), lcp1 and
mpx (data not shown) were severely reduced or entirely absent in 24
h.p.f. rad21nz171 mutants. A summary of the blood defects
present in rad21nz171 is shown in Fig. S1 (see Fig. S1 in
the supplementary material). Injection of 200 pg runx1 mRNA into
rad21nz171 homozygotes at the 1-cell stage rescued
expression of lyz (Fig.
6D, n=21/23), demonstrating an ability for Runx1 to exert
function in early myeloid cells (see Discussion). It was not possible to
analyze the impact of early runx1 loss on definitive hematopoiesis in
rad21nz171 because of the developmental arrest by 35
h.p.f.
|
).
Intact cohesin is necessary for correct regulation of Runx gene expression
We next asked if Rad21 is operating as part of the cohesin complex in the
regulation of Runx gene expression. We used MOs to knock down the function of
Smc3, another integral subunit of the cohesin complex (see Fig. S4 in the
supplementary material). smc3 morphants appeared morphologically
similar to rad21 morphants and the rad21nz171
mutant (see Fig. S4 in the supplementary material). We observed mitotic cells
in smc3 morphants that had condensed chromosomes spread throughout
the cell, similar to cells lacking Rad21
(Fig. 7E), indicating a similar
role in chromosome cohesion. Lagging chromosomes and ectopic location of
chromosomes at the poles were frequently observed. Furthermore, smc3
morphants lacked early runx1 and runx3 expression
(Fig. 7F-I). By contrast, a
hydroxyurea and aphidicolin S-phase block of the cell cycle had no effect on
runx1 expression (data not shown). Since cells in
rad21nz171 homozygotes successfully complete mitosis at
the 14-somite stage, it is unlikely that loss of Runx gene expression at this
time is due to a cell cycle block or the activation of cell cycle checkpoints.
We therefore believe that the loss of Runx gene expression in
rad21nz171 homozygotes is directly related to a reduction
in cohesin. Taken together, our data indicate that the cohesin complex is
necessary for normal expression of Runx genes, in addition to its function in
sister chromatid cohesion.
|
), a
maternal complement of rad21 mRNA or Rad21 protein is likely to
sustain proliferation until cells in rad21nz171 embryos
eventually arrest in mitosis on depletion of Rad21 protein. In support of this
notion, we found that rad21 mRNA is maternally inherited
(Fig. 3D). Furthermore, Rad21
protein was detected by immunoblotting in 10-somite and 14-somite
rad21ATGMO morphants, and in 18-somite rad21nz171
homozygotes (Fig. 8A,B).
|
rad21nz171 homozygotes have defects in neuronal
development during early embryogenesis (J.A.H. and J.K-H.H., unpublished) and
rad21 is specifically expressed in restricted regions of the brain at
48 h.p.f. (Fig. 3F). To
determine if changes in the dose of rad21 can affect the expression
of neuronal genes in older embryos, we used quantitative RT-PCR to monitor
expression of selected neuronal genes in 48 h.p.f. embryos. We found that
expression of proneural genes ascl1a and ascl1b, which are
strongly expressed in the brain during embryogenesis
(Allende and Weinberg, 1994),
was severely reduced in rad21nz171 mutants and
significantly reduced in siblings (Fig.
8D, lower graphs). Our data indicate that although
rad21nz171 siblings appear to grow and develop normally,
they are unable to express wild-type levels of downstream genes. This finding
points to a regulatory function for cohesin that can be separated from its
cell cycle role.
Cohesin contributes to a novel regulatory mechanism for early runx1 expression
To understand how Rad21/cohesin might influence Runx gene expression, we
investigated whether loss of rad21 affects pathways known to be
upstream of runx1. The cdx (hox) pathway specifies
blood development from mesoderm, and is essential for runx1
expression in embryogenesis (Davidson et
al., 2003; Davidson and Zon,
2006). We determined that the expression of cdx4 and its
downstream hox targets (hoxa9a, hoxb4, hoxb6b, hoxb7a) were
unaffected in 10-somite rad21nz171 homozygotes (data not
shown). The Notch signaling pathway is upstream of definitive runx1
expression in zebrafish (Burns et al.,
2005). Interestingly, we found that several genes induced by Notch
signaling are downregulated in rad21nz171 embryos (J.A.H.
and S.H.A., unpublished results). However, consistent with previous data
(Burns et al., 2005;
Gering and Patient, 2005), we
found that early expression of runx1 is not Notch dependent (data not
shown), thereby eliminating this pathway as an intermediate between cohesin
and early runx1 expression. In summary, two previously characterized
pathways shown to contribute to runx1 expression do not mediate the
cohesin regulatory function. Therefore cohesin appears to have a novel role in
the regulation of early runx1 expression.
|
|
DISCUSSION |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
).
Therefore, rad21nz171 appears to be the first mutant that
affects early expression of runx1 alone. Although most other early
hematopoietic markers are expressed normally in
rad21nz171, early myelopoiesis is deficient and
hbbe3 expression is reduced. By 24 h.p.f., the cell cycle is severely
affected in rad21nz171
(Fig. 2) and therefore
hematopoietic progenitors might be unable to divide to form differentiated
progeny. In addition, cell death in the ICM almost certainly contributes to a
reduction in differentiated cells (Fig.
2L). However, live hematopoietic progenitors remain in the ICM of
rad21nz171 mutants, as indicated by robust scl
and gata2 expression at 24 h.p.f.; furthermore, inappropriate
maintenance of scl expression in the ICM at 48 h.p.f. points to the
persistence of immature precursors that cannot differentiate (see Fig. S1 in
the supplementary material). Loss of differentiated cells in
rad21nz171 might be due in part to loss of runx1
and reduction in gata1 mRNA. Rescue of lyz+ cells
by runx1 mRNA indicates that either, (1) runx1 is necessary
for early myelopoiesis and its restoration rescues differentiated cells, or
(2) expression of runx1 is able to overcome a developmental block in
the myeloid precursors of rad21nz171 by driving more cells
toward a myeloid fate at an earlier time. In support of the latter, we
observed an increase in the number of lyz+ cells in
siblings of runx1-injected rad21nz171 crosses
(n=41/41, data not shown). Unfortunately, the cell cycle block in
rad21nz171 prevented an analysis of effects on definitive
hematopoiesis.
Our results add to increasing evidence that chromatin-modifying proteins
can have specific roles in hematopoiesis. In previous studies, a nucleosome
assembly protein NAP1L was shown to operate upstream of scl in
Xenopus hematopoiesis (Abu-Daya et
al., 2005). Furthermore, Brg1 (a SWI/SNF subunit) appears to have
a distinct role in the activation of the ß-globin locus in erythropoiesis
(Bultman et al., 2005). The
involvement of trithorax (Ernst et al.,
2004a; Ernst et al.,
2004b) and polycomb (Lessard
and Sauvageau, 2003; Lessard
et al., 1999) group members as epigenetic regulators of
hematopoietic stem cell development is relatively well characterized.
A role for cohesin in vertebrate gene regulation
We report the first direct example of cohesin-dependent gene regulation in
a vertebrate. Previous studies in Drosophila implicated cohesin
loading protein, Nipped-B, and cohesin subunits in regulation of the
cut and Ultrabithorax loci
(Dorsett et al., 2005;
Rollins et al., 1999). In
C. elegans, the cohesin-loading factor MAU-2 (the Scc4 ortholog) is
essential for axon migration during development
(Seitan et al., 2006). Our
observations of neuronal abnormalities and altered neuronal gene expression in
rad21nz171 embryos, together with the C. elegans
data, are consistent with the idea that chromosome cohesion proteins have
specific roles in neuronal development.
|
|
Linking cohesin function and Runx gene regulation
It is tempting to speculate that loss of early runx1 and
runx3 expression in rad21nz171 is causally
related to the connection between the Runx family and the cell cycle. Runx
protein levels are dynamically regulated during the cell cycle; e.g.
expression of Runx2 oscillates during the cell cycle of MC3T3
osteoblasts (Galindo et al.,
2005). In cell lines, Runx1 levels were also shown to oscillate in
a cell cycle-dependent manner: Runx1 protein levels increase during S phase
and G2 (Bernardin-Fried et al.,
2004) and at the G2-M phase transition, Runx1 is degraded by the
anaphase-promoting complex (Biggs et al.,
2006; Wang et al.,
2006). Cohesin becomes stably associated with chromatin during S
and G2 (Gerlich et al., 2006),
and perhaps this is necessary for runx1 expression in a particular
developmental context. Runx1 degradation is also concomitant with cohesin
cleavage at G2-M.
Runx proteins appear to have cell cycle-specific functions. Overexpression
of Runx1 causes a shortening of G1 phase
(Strom et al., 2000) and Runx1
physically interacts with cyclin D3 to repress its own transcription
(Peterson et al., 2005).
During mitosis, Runx2 selectively regulates specific target genes
(Young et al., 2007b), and
represses transcription of ribosomal RNA genes
(Young et al., 2007a). These
authors also reported similar unpublished observations for Runx1. Therefore,
Runx proteins might regulate cell growth through control of ribosome
biogenesis, and it was proposed that by this mechanism Runx proteins might
coordinate cell proliferation and differentiation. Clearly, Runx proteins
provide a mechanistic link between the cell cycle and development. Therefore a
mechanism by which Runx gene expression is coordinated with the cell cycle
would make sense. Cohesin is an integral part of the cell cycle machinery, and
is therefore a good candidate to participate in cell cycle-dependent gene
regulation.
Cohesin-dependent gene regulation and the implications for development
There are a number of human developmental disorders associated with loss of
sister chromatid cohesion, including CdLS
(Krantz et al., 2004;
Musio et al., 2006;
Strachan, 2005). This raises
the interesting possibility that such developmental disorders might be
contributed to by a reduction in Runx gene expression in embryogenesis, as a
result of reduction in cohesin function. CdLS patients present with
neurodevelopmental, gastrointestinal and skeletal abnormalities
(Strachan, 2005). The
development of each of these systems depends on the proper regulation of Runx
proteins, which are themselves dose-sensitive in function
(Blyth et al., 2005). In
summary, our findings provide strong evidence for a novel developmental
function for cohesin. The next challenge will be to determine exactly how
cohesin contributes to regulation of gene expression and developmental
pathways.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/134/14/2639/DC1
|
ACKNOWLEDGMENTS |
|---|
|
Footnotes |
|---|
Present address: Department of Genetics, Harvard Medical School, 77 Avenue
Louis Pasteur, Boston, MA 02115, USA
|
REFERENCES |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Abu-Daya, A., Steer, W. M., Trollope, A. F., Friedeberg, C. E.,
Patient, R. K., Thorne, A. W. and Guille, M. J. (2005).
Zygotic nucleosome assembly protein-like 1 has a specific, non-cell autonomous
role in hematopoiesis. Blood
106,514
-520.
Allende, M. L. and Weinberg, E. S. (1994). The expression pattern of two zebrafish achaete-scute homolog (ash) genes is altered in the embryonic brain of the cyclops mutant. Dev. Biol. 166,509 -530.[CrossRef][Medline]
Amsterdam, A., Nissen, R. M., Sun, Z., Swindell, E. C.,
Farrington, S. and Hopkins, N. (2004). Identification of 315
genes essential for early zebrafish development. Proc. Natl. Acad.
Sci. USA 101,12792
-12797.
Bernardin-Fried, F., Kummalue, T., Leijen, S., Collector, M. I.,
Ravid, K. and Friedman, A. D. (2004). AML1/RUNX1 increases
during G1 to S cell cycle progression independent of cytokine-dependent
phosphorylation and induces cyclin D3 gene expression. J. Biol.
Chem. 279,15678
-15687.
Biggs, J. R., Peterson, L. F., Zhang, Y., Kraft, A. S. and
Zhang, D. E. (2006). AML1/RUNX1 phosphorylation by
cyclin-dependent kinases regulates the degradation of AML1/RUNX1 by the
anaphase-promoting complex. Mol. Cell. Biol.
26,7420
-7429.
Blyth, K., Cameron, E. R. and Neil, J. C. (2005). The RUNX genes: gain or loss of function in cancer. Nat. Rev. Cancer 5,376 -387.[CrossRef][Medline]
Bultman, S. J., Gebuhr, T. C. and Magnuson, T.
(2005). A Brg1 mutation that uncouples ATPase activity from
chromatin remodeling reveals an essential role for SWI/SNF-related complexes
in beta-globin expression and erythroid development. Genes
Dev. 19,2849
-2861.
Burns, C. E., Traver, D., Mayhall, E., Shepard, J. L. and Zon,
L. I. (2005). Hematopoietic stem cell fate is established by
the Notch-Runx pathway. Genes Dev.
19,2331
-2342.
Chang, Y. F., Imam, J. S. and Wilkinson, M. F. (2007). The nonsense-mediated decay RNA surveillance pathway. Annu. Rev. Biochem. 76 (in press).
Davidson, A. J. and Zon, L. I. (2006). The caudal-related homeobox genes cdx1a and cdx4 act redundantly to regulate hox gene expression and the formation of putative hematopoietic stem cells during zebrafish embryogenesis. Dev. Biol. 292,506 -518.[CrossRef][Medline]
Davidson, A. J., Ernst, P., Wang, Y., Dekens, M. P., Kingsley, P. D., Palis, J., Korsmeyer, S. J., Daley, G. Q. and Zon, L. I. (2003). cdx4 mutants fail to specify blood progenitors and can be rescued by multiple hox genes. Nature 425,300 -306.[CrossRef][Medline]
Dorsett, D. (2007). Roles of the sister chromatid cohesion apparatus in gene expression, development, and human syndromes. Chromosoma 116, 1-13.[CrossRef][Medline]
Dorsett, D., Eissenberg, J. C., Misulovin, Z., Martens, A.,
Redding, B. and McKim, K. (2005). Effects of sister chromatid
cohesion proteins on cut gene expression during wing development in
Drosophila. Development
132,4743
-4753.
Elagib, K. E., Racke, F. K., Mogass, M., Khetawat, R.,
Delehanty, L. L. and Goldfarb, A. N. (2003). RUNX1 and GATA-1
coexpression and cooperation in megakaryocytic differentiation.
Blood 101,4333
-4341.
Ernst, P., Fisher, J. K., Avery, W., Wade, S., Foy, D. and Korsmeyer, S. J. (2004a). Definitive hematopoiesis requires the mixed-lineage leukemia gene. Dev. Cell 6, 437-443.[CrossRef][Medline]
Ernst, P., Mabon, M., Davidson, A. J., Zon, L. I. and Korsmeyer, S. J. (2004b). An Mll-dependent Hox program drives hematopoietic progenitor expansion. Curr. Biol. 14,2063 -2069.[CrossRef][Medline]
Galindo, M., Pratap, J., Young, D. W., Hovhannisyan, H., Im, H.
J., Choi, J. Y., Lian, J. B., Stein, J. L., Stein, G. S. and van Wijnen, A.
J. (2005). The bone-specific expression of Runx2 oscillates
during the cell cycle to support a G1-related antiproliferative function in
osteoblasts. J. Biol. Chem.
280,20274
-20285.
Geisler, R. (2002). Mapping and cloning. In Zebrafish, A Practical Approach. Vol.261 (ed. C. Nusslein-Volhard and R. Dahm), pp.175 -212. Oxford: Oxford University Press.
Gering, M. and Patient, R. (2005). Hedgehog signaling is required for adult blood stem cell formation in zebrafish embryos. Dev. Cell 8,389 -400.[CrossRef][Medline]
Gerlich, D., Koch, B., Dupeux, F., Peters, J.-M. and Ellenberg, J. (2006). Live-cell imaging reveals a stable cohesin-chromatin interaction after but not before DNA replication. Curr. Biol. 16,1571 -1578.[CrossRef][Medline]
Hagstrom, K. A. and Meyer, B. J. (2003). Condensin and cohesin: more than chromosome compactor and glue. Nat. Rev. Genet. 4,520 -534.[CrossRef][Medline]
Hsia, N. and Zon, L. I. (2005). Transcriptional regulation of hematopoietic stem cell development in zebrafish. Exp. Hematol. 33,1007 -1014.[CrossRef][Medline]
Ito, Y. (2004). Oncogenic potential of the RUNX gene family: `overview'. Oncogene 23,4198 -4208.[CrossRef][Medline]
Kalev-Zylinska, M. L., Horsfield, J. A., Flores, M. V., Postlethwait, J. H., Vitas, M. R., Baas, A. M., Crosier, P. S. and Crosier, K. E. (2002). Runx1 is required for zebrafish blood and vessel development and expression of a human RUNX1-CBF2T1 transgene advances a model for studies of leukemogenesis. Development 129,2015 -2030.[Medline]
Kalev-Zylinska, M. L., Horsfield, J., Flores, M. V., Postlethwait, J., Chau, J. Y. M., Cattin, P. M., Vitas, M. R., Crosier, P. S. and Crosier, K. E. (2003). Runx3 is required for hematopoietic development in zebrafish. Dev. Dyn. 228,323 -336.[CrossRef][Medline]
Kobayashi, M., Nishikawa, K. and Yamamoto, M. (2001). Hematopoietic regulatory domain of gata1 gene is positively regulated by GATA1 protein in zebrafish embryos. Development 128,2341 -2350.[Medline]
Krantz, I. D., McCallum, J., DeScipio, C., Kaur, M., Gillis, L. A., Yaeger, D., Jukofsky, L., Wasserman, N., Bottani, A., Morris, C. A. et al. (2004). Cornelia de Lange syndrome is caused by mutations in NIPBL, the human homolog of Drosophila melanogaster Nipped-B. Nat. Genet. 36,631 -635.[CrossRef][Medline]
Lau, Q. C., Raja, E., Salto-Tellez, M., Liu, Q., Ito, K., Inoue,
M., Putti, T. C., Loh, M., Ko, T. K., Huang, C. et al.
(2006). RUNX3 is frequently inactivated by dual mechanisms of
protein mislocalization and promoter hypermethylation in breast cancer.
Cancer Res. 66,6512
-6520.
Lessard, J. and Sauvageau, G. (2003). Polycomb group genes as epigenetic regulators of normal and leukemic hemopoiesis. Exp. Hematol. 31,567 -585.[CrossRef][Medline]
Lessard, J., Schumacher, A., Thorsteinsdottir, U., van Lohuizen,
M., Magnuson, T. and Sauvageau, G. (1999). Functional
antagonism of the Polycomb-Group genes eed and Bmi1 in hemopoietic cell
proliferation. Genes Dev.
13,2691
-2703.
Losada, A. and Hirano, T. (2005). Dynamic
molecular linkers of the genome: the first decade of SMC proteins.
Genes Dev. 19,1269
-1287.
Mueller, W., Nutt, C. L., Ehrich, M., Riemenschneider, M. J., von Deimling, A., van den Boom, D. and Louis, D. N. (2007). Downregulation of RUNX3 and TES by hypermethylation in glioblastoma. Oncogene 26,583 -593.[CrossRef][Medline]
Musio, A., Selicorni, A., Focarelli, M. L., Gervasini, C., Milani, D., Russo, S., Vezzoni, P. and Larizza, L. (2006). X-linked Cornelia de Lange syndrome owing to SMC1L1 mutations. Nat. Genet. 38,528 -530.[CrossRef][Medline]
Nakagawa, M., Ichikawa, M., Kumano, K., Goyama, S., Kawazu, M.,
Asai, T., Ogawa, S., Kurokawa, M. and Chiba, S. (2006).
AML1/Runx1 rescues Notch1-Null mutation-induced deficiency of para-aortic
splanchnopleural hematopoiesis. Blood
108,3329
-3334.
Nasmyth, K. and Haering, C. H. (2005). The structure and function of SMC and kleisin complexes. Annu. Rev. Biochem. 74,595 -648.[CrossRef][Medline]
Patterson, L. J., Gering, M. and Patient, R.
(2005). Scl is required for dorsal aorta as well as blood
formation in zebrafish embryos. Blood
105,3502
-3511.
Patterson, L. J., Gering, M., Eckfeldt, C. E., Green, A. R.,
Verfaillie, C. M., Ekker, S. C. and Patient, R. (2007). The
transcription factors Scl and Lmo2 act together during development of the
hemangioblast in zebrafish. Blood
109,2389
-2398.
Pelegri, F. (2002). Mutagenesis. In Zebrafish, A Practical Approach. Vol.261 (ed. C. Nusslein-Volhard and R. Dahm), pp.145 -174. Oxford: Oxford University Press.
Peterson, L. F., Boyapati, A., Ranganathan, V., Iwama, A.,
Tenen, D. G., Tsai, S. and Zhang, D. E. (2005). The
hematopoietic transcription factor AML1 (RUNX1) is negatively regulated by the
cell cycle protein cyclin D3. Mol. Cell. Biol.
25,10205
-10219.
Pimanda, J. E., Donaldson, I. J., de Bruijn, M. F., Kinston, S.,
Knezevic, K., Huckle, L., Piltz, S., Landry, J. R., Green, A. R., Tannahill,
D. et al. (2007). The SCL transcriptional network and BMP
signaling pathway interact to regulate RUNX1 activity. Proc. Natl.
Acad. Sci. USA 104,840
-845.
Rollins, R. A., Morcillo, P. and Dorsett, D.
(1999). Nipped-B, a Drosophila homologue of chromosomal adherins,
participates in activation by remote enhancers in the cut and Ultrabithorax
genes. Genetics 152,577
-593.
Rollins, R. A., Korom, M., Aulner, N., Martens, A. and Dorsett,
D. (2004). Drosophila nipped-B protein supports sister
chromatid cohesion and opposes the stromalin/Scc3 cohesion factor to
facilitate long-range activation of the cut gene. Mol. Cell.
Biol. 24,3100
-3111.
Seitan, V. C., Banks, P., Laval, S., Majid, N. A., Dorsett, D., Rana, A., Smith, J., Bateman, A., Krpic, S., Hostert, A. et al. (2006). Metazoan Scc4 homologs link sister chromatid cohesion to cell and axon migration guidance. PLoS Biol. 4, e242.[CrossRef][Medline]
Shepard, J., Stern, H. M., Pfaff, K. L. and Amatruda, J. F. (2004). Analysis of the cell cycle in Zebrafish embryos. In The Zebrafish: Cellular and Developmental Biology. Vol. 76, 2nd edn (ed. H. W. Detrich, 3rd, M. Westerfield and L. Zon), pp. 109-125. San Diego: Elsevier.[CrossRef]
Sonoda, E., Matsusaka, T., Morrison, C., Vagnarelli, P., Hoshi, O., Ushiki, T., Nojima, K., Fukagawa, T., Waizenegger, I. C., Peters, J. M. et al. (2001). Scc1/Rad21/Mcd1 is required for sister chromatid cohesion and kinetochore function in vertebrate cells. Dev. Cell 1,759 -770.[CrossRef][Medline]
Strachan, T. (2005). Cornelia de Lange Syndrome and the link between chromosomal function, DNA repair and developmental gene regulation. Curr. Opin. Genet. Dev. 15,258 -264.[CrossRef][Medline]
Strom, D. K., Nip, J., Westendorf, J. J., Linggi, B.,
Lutterbach, B., Downing, J. R., Lenny, N. and Hiebert, S. W.
(2000). Expression of the AML-1 oncogene shortens the G(1) phase
of the cell cycle. J. Biol. Chem.
275,3438
-3445.
Waltzer, L., Ferjoux, G., Bataille, L. and Haenlin, M. (2003). Cooperation between the GATA and RUNX factors Serpent and Lozenge during Drosophila hematopoiesis. EMBO J. 22,6516 -6525.[CrossRef][Medline]
Wang, S., Zhang, Y., Soosairajah, J. and Kraft, A. S. (2006). Regulation of RUNX1/AML1 during the G2/M transition. Leuk. Res. doi:10.1016/j.leukres.2006.08.016 .
Watrin, E. and Peters, J. M. (2006). Cohesin and DNA damage repair. Exp. Cell Res. 312,2687 -2693.[CrossRef][Medline]
Westerfield, M. (1995). The Zebrafish Book. A Guide for the Laboratory Use of Zebrafish (Danio rerio). Eugene, OR: University of Oregon Press.
Wo
