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Molecular and Cellular Biology, June 2000, p. 4169-4180, Vol. 20, No. 11
0270-7306/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
S-Phase Progression Mediates Activation of a
Silenced Gene in Synthetic Nuclei
Alison J.
Crowe,
Julie L.
Piechan,
Ling
Sang, and
Michelle C.
Barton*
Department of Molecular Genetics,
Biochemistry and Microbiology, University of Cincinnati, Cincinnati,
Ohio
Received 21 January 2000/Returned for modification 19 February
2000/Accepted 28 February 2000
 |
ABSTRACT |
Aberrant expression of developmentally silenced genes,
characteristic of tumor cells and regenerating tissue, is highly
correlated with increased cell proliferation. By modeling this process
in vitro in synthetic nuclei, we find that DNA replication leads to
deregulation of established developmental expression patterns. Chromatin assembly in the presence of adult mouse liver nuclear extract
mediates developmental stage-specific silencing of the tumor marker
gene alpha-fetoprotein (AFP). Replication of silenced AFP chromatin in
synthetic nuclei depletes sequence-specific transcription repressors,
thereby disrupting developmentally regulated repression. Hepatoma-derived factors can target partial derepression of AFP, but
full transcription activation requires DNA replication. Thus, unscheduled entry into S phase directly mediates activation of a
developmentally silenced gene by (i) depleting developmental stage-specific transcription repressors and (ii) facilitating binding
of transactivators.
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INTRODUCTION |
Cellular commitment,
differentiation, and specificity are determined primarily by the
interaction of protein complexes with chromatin DNA. This
gene-regulatory programming is challenged during each cell cycle by
passage of the replisome during DNA synthesis. DNA replication may
facilitate competition between transiently disrupted histones and
transacting factors at the replication fork, potentially changing
established chromatin structure and gene expression patterns (2,
7, 20, 29, 59). During tumorigenesis, terminally differentiated
cells that have withdrawn from the cell cycle are induced to resume
cycling, often through functional inactivation of tumor suppressor
genes such as Rb and INK4a (30, 33). This cyclic disruption
of chromatin structure provides a fertile environment, similar to that
which exists during fetal development, for altering gene expression
patterns. Aberrant adult activation of genes normally expressed only in
the fetus is characteristic of many tumors (reviewed in references
31 and 39). For example, a
hepatoma marker gene, alpha-fetoprotein (AFP), is transcriptionally
repressed shortly after birth (reviewed in reference
52) and is reexpressed in mature hepatocytes only when they leave G0 quiescence and begin cycling following
partial hepatectomy or during hepatocellular carcinoma (HCC)
(42; reviewed in reference 52).
Expression of AFP is therefore closely linked to cell cycle
progression; the renewal of DNA replication following transition from
quiescence to S phase may play an epigenetic role in modulating gene expression.
The relative local concentration of transacting factors at the time of
replication can determine whether a given chromatin structure is
maintained or converted to an alternate conformation (17,
58). Modulating the balance of repressors and activators present
during replication could be achieved through a variety of mechanisms,
including functional inactivation of repressors and activation of
transcription factors. Alternatively, the availability of
transactivators could be controlled by shifting the replication timing
of a given gene. Highly expressed genes, the majority of which
replicate early in S phase, may bind limiting transcription factors,
thereby depleting available pools for late-replicating silent or
repressed genes (reviewed in reference 46).
To identify a potential role for DNA replication in renewed expression
of a developmentally silenced gene, such as AFP, we have mimicked the
reentry of differentiated G0 cells into the cell cycle by
synthetic nucleus assembly and S-phase progression in vitro using
Xenopus egg extracts (25, 40, 41). We assessed the functional consequences of S-phase progression and DNA replication uncoupled from protein synthesis through in vitro transcription analysis of AFP expression. Using this approach, we have recapitulated the switch in AFP gene expression that occurs during HCC and liver regeneration from developmentally silenced to transcriptionally active.
Our data provide the first direct evidence that deregulation of an
established developmental expression pattern requires DNA replication.
S-phase progression/DNA replication mediates activation of AFP in two
ways: (i) by depleting a developmentally staged transcription
repressor(s), thereby inhibiting the reestablishment of developmentally
silenced templates postreplication, and (ii) by facilitating the
binding of trans-acting factors to their sites in
nucleosomal DNA.
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MATERIALS AND METHODS |
DNA templates.
The AFP(3.8)-lacZ plasmid
(51) contains 3.8 kb of sequence upstream of the AFP
transcriptional start site, encompassing both enhancer I and the
previously described developmental repressor domain (reviewed in
reference 11). The 11-kb AFP(3.8)-lacZ
plasmid was digested with EcoRI and ClaI enzymes
to generate a 9.0-kb linear fragment encompassing the AFP enhancer I
and promoter sequences fused to the lacZ gene. Biotin
end-labeling reactions contained DNA (50 µg of DNA per 100-µl
reaction volume), Klenow enzyme (NEB; 5 U per 100-µl reaction
volume), 20 µM biotin 21-dUTP (Clontech, Palo Alto, Calif.), and dATP
(20 µM) and were incubated for 30 min at 37°C to generate
biotin-labeled EcoRI sites. Unincorporated deoxynucleotides,
biotinylated deoxynucleotides, and small fragments of digested DNA are
removed by gel filtration (Chromaspin 1000; Clontech).
Purified biotinylated DNA was coupled to streptavidin-coated M-280
Dynal paramagnetic beads in Kilobase Binding Buffer (Dynal Corporation,
Oslo, Norway). Beads were washed twice in an equal volume of binding
buffer. Biotinylated DNA (50 µg) was added to the resuspended beads
to give a 1:1 final dilution. Coupling reaction mixtures were incubated
at room temperature overnight on a rotating wheel. Coupled DNA-beads
(solid-phase DNA templates) were washed three times with wash buffer (2 M NaCl, 10 mM Tris-HCl [pH 8.0], 1 mM EDTA) using a magnetic
concentrator (Dynal Corporation) and stored in wash buffer at 4°C
(12, 13).
The 13.5-kb pAlbN albumin construct is a modification of pAN3-42
(generous gift of K. Zaret, Brown University). pAlbN contains
8.8 kb of
albumin sequence including the albumin enhancer (
12.0
to
4.0 kb)
fused at
800 bp of the albumin promoter relative
to the transcription
start site. The albumin sequences were linked
to the
neo
coding region in a pUC18 backbone. Unique biotinylation
and coupling
restriction sites were generated by insertion of
a polylinker
(
XmaI/
AscI/
PacI) at the
SmaI site (+1837) of pAlbN.
The DNA was digested with
KpnI and
XmaI, biotinylated, and coupled
to
paramagnetic beads as described
above.
PCR-mediated mutagenesis of AFP templates was conducted as previously
described by Lee et al. (
35) to generate templates
deleted
between
1000 and
541 or
209 of the AFP upstream regulatory
region. Deletions between
1000 and
850,
765, or
586 were made
using the Erase-A-Base system of mutagenesis, exactly as described
by
the manufacturer (Promega Corporation, Madison, Wis.). Endpoints
of
deletion and integrity of PCR-generated templates were determined
by
DNA
sequencing.
Protein extracts and in vitro transcription.
Adult mouse
liver extract was prepared by the method of Gorski et al.
(23). Hepatocarcinoma cell extracts were prepared from human
HepG2 (ATCC catalog no. HB-8065) and mouse Hepa 1-6 (ATCC catalog no.
CRL-1830) cells as described by Dignam et al. (14) with
minor modifications (35). HeLa nuclear extract was prepared
by the method of Workman and Abmayr (5). All extracts contained total proteins in concentrations of 5 to 10 mg/ml.
Xenopus egg extracts were prepared as described previously
(8). Fractionation by centrifugation of Xenopus
egg clarified extract (LSS) yields a vesicular membrane fraction and a
soluble high-speed supernatant (HSS) in Xenopus laevis (Xl)
buffer (100 mM KCl, 4 mM MgCl2, 10 mM potassium HEPES [pH
7.2], 100 mM sucrose, 0.1 mM EGTA). Generally, chromatin was assembled
in membrane-depleted HSS prior to synthetic nucleus formation in crude,
clarified LSS. Where noted, HSS plus membrane vesicles have been used
in transcription and replication assays as described previously
(7). Replication assays were also performed in LSS as
described below.
In vitro chromatin assembly and transcription reactions were performed
as described previously (
12). Cellular extracts were
supplemented with nuclear dialysis buffer (NDB: 20 mM HEPES [pH
7.9],
50 mM KCl, 0.5 mM EDTA, 20% glycerol, 2 mM dithiothreitol
[DTT]) to
an equal volume in all reactions (a total of 25 µl).
These were added
to 500 ng of solid-phase DNA templates during
a 20-min room temperature
preincubation or as indicated by the
experimental design (final protein
concentration during preincubation,
8 µg/µl). Following
preincubation, HSS supplemented with ATP to
6 mM, poly(dI:dC) to 4 ng/µl, and 1 M MgCl
2 to a 10 mM final concentration
was
added to give a total of 50 µl (usually 600 to 800 µg of protein)
and incubated for 1 h at 22°C. In order to inhibit any
background
DNA replication during the chromatin assembly period, the
DNA
polymerase inhibitor aphidicolin was added to each reaction at
a
40-ng/µl final concentration. Chromatin-assembled solid-phase
DNA was
washed three times by placement on a magnetic concentrator
(magnetic
concentration) and removal of unbound extract and proteins,
followed by
addition of 200 µl of NDB (containing 0.01% NP-40)
to each reaction.
Chromatin can be transcribed immediately or
assembled into synthetic
nuclei prior to in vitro transcription.
Washed chromatin templates were
assembled into synthetic nuclei
by addition of approximately 1.5 mg of
total protein of LSS (supplemented
with ATP and MgCl
2 to
give final concentrations in the assembly
reaction mixture of 3 mM ATP
and 5 mM MgCl
2), plus cellular extracts
or NDB in a total
volume of 50 µl. Reaction mixtures were incubated
for 2 h at
22°C. Templates were washed as described above prior
to in vitro
transcription under standard transcription conditions
(
12).
Synthetic nucleus analysis.
Nuclear transport studies were
performed by the method of Gorlich et al. (22). Briefly, 1 µg of AFP(3.8)-lacZ solid-phase bead-DNA was incubated in
100 µl of unfractionated Xenopus egg extract (LSS)
containing 1 mM ATP, 0.2 mM GTP, 10 mM creatine phosphate (Boehringer
Mannheim), and 9.5 U of creatine kinase (Boehringer Mannheim) per ml to
form solid-phase synthetic nuclei. After 1.5 h, 150 ng of the
indicated recombinant fusion protein (LEF-1:eGFP or GST:eGFP) stored in
25 mM Tris (pH 8.0)-50 mM ammonium sulfate-5% glycerol-1 mM EDTA-1
mM DTT was added and further incubated in transport buffer (final
concentrations: 4 mM HEPES [pH 7.8], 22 mM potassium acetate, 1.0 mM
sodium acetate, 0.4 mM magnesium acetate, 0.2 mM EGTA, 0.4 mM DTT).
After 30 min, 2-µl aliquots were removed and mixed with 2 µl of the
DNA-specific stain propidium idodide (Oncor, Washington, D.C.). Nuclei
were analyzed by fluorescent and phase-contrast microscopy at a
magnification of ×400.
DNA replication was assayed by steady-state "continuous labeling"
of newly synthesized DNA with 1 µCi of [
-
32P]dATP
(3,000 Ci/mmol; ICN) present during the entire incubation
period with
Xenopus egg extract (
50). DNA replication
reactions
were performed with 500 ng of bead-DNA in (as noted in the
figures
or text) 25 µl of LSS alone, 25 µl of HSS plus membrane
vesicles
(
7), or sequential HSS incubation, washes, and LSS
incubation
conditions described for in vitro transcription (see above),
plus
NDB or cellular extracts (100 to 120 µg) in a total volume of
50 µl. Replication assays were terminated by addition of 2.5%
sodium
dodecyl sulfate-60 mM EDTA. DNA was purified by overnight
digestion
with proteinase K (0.8 mg/ml) at 37°C followed by phenol-chloroform
(1:1) and chloroform extractions and ethanol precipitation prior
to gel
electrophoresis on a Tris-borate-EDTA-0.8% agarose gel.
The
proteinase digestion step also severs the biotin-streptavidin
linkage
between the biotinylated template and the paramagnetic
bead support.
Replicated DNA was visualized by autoradiography
of the dried
gel.
Relative levels of full-length DNA replication within solid-phase
nuclei were quantified by ImageQuant analysis of scanned
autoradiograms
comparing full-length replicated DNA products under
each of the
incubation conditions noted. Nucleosome assembly of
the
32P-labeled, replicated AFP-bead DNA was assessed by
micrococcal
nuclease (MNase; Boehringer Mannheim) digestion after a 2-h
incubation
with LSS containing 3 mM ATP, 5 mM MgCl
2 and 1 µCi of [
-
32P]dATP (3,000 Ci/mmol; ICN). Samples were
digested and analyzed
as described previously (
8). MNase
protection patterns were
identical whether samples were incubated in
LSS or HSS plus membrane
vesicles (
8).
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RESULTS |
Hepatoma-specific derepression of chromatin-assembled AFP
templates.
The tumor marker gene AFP provides an ideal model for
studying aberrant gene activation due to its strict tissue-specific and
developmental stage-specific regulation in vivo. AFP is highly expressed during fetal development in endoderm-derived tissues, including the yolk sac, gut, and liver. Developmental stage-specific silencing occurs shortly after birth, and under normal circumstances, AFP expression remains repressed in the differentiated hepatocyte throughout adult life (reviewed in references 11 and
52). However, if hepatocytes are induced to resume
cycling, as occurs both during HCC and after liver damage, the silenced
AFP gene is reactivated. This is in contrast to the evolutionarily
related albumin gene, which lies directly upstream of AFP. Albumin
levels remain relatively constant throughout adult life, and these high levels are maintained during both liver regeneration and HCC
(36).
To identify the molecular mechanisms responsible for the aberrant
reactivation of AFP, we have established in vitro chromatin
and
synthetic nucleus transcription systems for the AFP gene.
Based on a
design for transcriptional analysis of the
Drosophila hsp70
promoter (
47,
55), we have coupled the AFP template
to
streptavidin-coated paramagnetic beads. The generation of immobilized
templates facilitates washing of chromatin-assembled templates
and
removal of any unbound proteins present in cellular or
Xenopus assembly extracts prior to in vitro transcription.
The details
of biotin end labeling, DNA coupling, and manipulation of
the
bead-DNA (solid phase) templates in
Xenopus chromatin
and synthetic
nucleus assembly systems have been described elsewhere
(
12,
13). As diagrammed in Fig.
1A, solid-phase AFP templates containing
3.8 kb of upstream AFP regulatory sequence including enhancer
I and the
distal and proximal promoter elements are assembled
into chromatin by
incubation with fractionated
Xenopus egg extract
(HSS)
(
41,
45).
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FIG. 1.
Reconstitution of in vivo expression patterns in vitro.
(A) Diagram of coupled chromatin assembly-transcription system. (B)
Hepatoma-specific derepression of AFP and albumin chromatin templates.
Immobilized AFP (lanes 1 to 3) or albumin (lanes 4 to 7) templates were
preincubated (preinc.) in NDB (lanes 1 and 5), adult mouse liver (ML)
extract (lanes 2 and 7), HepG2 (lane 3), or Hepa 1-6 extract (lane 6)
prior to chromatin assembly in HSS. Chromatin templates were washed and
in vitro transcribed in HeLa extract. A nucleosome-free albumin
template (lane 4) was in vitro transcribed in HeLa extract (lane 4).
(C) Differential regulation of AFP and albumin in simultaneous
transcription reactions. Immobilized AFP and albumin templates were
coincubated with either NDB (lanes 1, 2, and 5), HepG2 (lanes 3 and 6),
or ML (lanes 4 and 7) extract prior to chromatin assembly in HSS (lanes
2 to 4 and 6 to 7) or incubation in Xl buffer in the absence of
chromatin assembly (lanes 1 and 5). All templates were washed and in
vitro transcribed in HeLa extract. After RNA isolation, each reaction
mixture was divided in two, and primer extension analysis was performed
with either an AFP-specific primer (lanes 1 to 4) or an
albumin-specific primer (lanes 5 to 7). The extension products for AFP
(84 bp) and albumin (ALB; 105 bp) are indicated. Radiolabeled  X174
DNA digested with HaeIII was run as a molecular size
standard (lane MW).
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To program the AFP gene in either its developmentally silenced or
tumorigenically active state, we have incubated the gene
with tissue or
cellular extracts prior to chromatin assembly.
We have used the AFP-
and albumin-expressing HCC cell lines human
HepG2 and mouse Hepa 1-6 as
sources of hepatoma
trans-acting factors.
Adult mouse liver
(ML) extract was used to provide proteins present
in a differentiated
tissue where AFP is developmentally silenced
and albumin is
constitutively transcribed. As shown in Fig.
1B,
preincubation of the
AFP gene with HepG2 extract established a
transcriptionally active AFP
chromatin template (lane 3), resulting
in a 27-fold increase in
expression over a chromatin-repressed
template (lane 1). Preincubation
with ML extract (lane 2) resulted
in only very low levels of AFP
transcription (1.8-fold over a
chromatin-repressed template). This
developmentally silenced state
of AFP, established by chromatin
assembly in adult ML extract,
is resistant to magnetic concentration,
isoosmotic washing, and
transcription. In contrast, both ML extract and
Hepa 1-6 extract
established transcriptionally active albumin chromatin
(lanes
6 and
7).
Similar results were obtained when AFP and albumin templates were
present in the same chromatin assembly and transcription
reaction
mixtures and then divided in half for separate primer
extension
analysis (Fig.
1C). Differential expression of repressed
AFP (lane 4)
and activated albumin (lane 7) by chromatin assembly
in ML extract
recapitulated regulated expression in differentiated
hepatic cells.
Preincubation with HepG2 extract established transcriptionally
active
albumin (lane 6) and AFP (lane 3) templates. HepG2 extract
was used as
a source of hepatoma factors in all of the remaining
experiments. The
only observed difference in expression of AFP
and albumin templates in
vitro between mixed templates and those
transcribed in parallel is an
apparent loss of albumin transcription
start site integrity (lane 5).
Whether the two mixed templates
in naked DNA transcriptions compete for
factors that maintain
the start site of transcription is unknown at
this
time.
Together, these data indicate that solid-phase chromatin transcription
accurately reflects the expression patterns observed
in HCC and
differentiated liver. Thus, the ML extract contains
all the necessary
transactivators to activate a chromatin-assembled
gene, such as
albumin, that is normally expressed in adult liver.
There is likely a
balance of transactivators and repressors present
in the adult liver
that directly bind both AFP and albumin genes
to generate the silenced
and active states,
respectively.
Solid-phase synthetic nuclei support nuclear protein transport and
DNA replication.
In vivo activation of the AFP gene in adults
occurs only under conditions of rapid cellular proliferation such as
exist during liver regeneration and HCC. We reasoned that derepression
of AFP after chromatin assembly in vitro may require on-going DNA
replication. To test this hypothesis directly, we needed to assemble
functionally competent synthetic nuclei with AFP DNA coupled to
streptavidin beads. Addition of the vesicular fraction of
Xenopus egg extract or unfractionated Xenopus egg
cytoplasm (LSS) to chromatin DNA results in the formation of a
nucleus-like structure around cloned DNA, which enables nuclear
functions of protein transport and semiconservative DNA replication
(41).
Nuclei that form around DNA-coupled 2.4-µm paramagnetic beads
(solid-phase nuclei) incubated in interphase LSS (
41) are
competent for both nuclear transport and DNA replication (Fig.
2)
(
27). As shown in Fig.
2A,
solid-phase nuclei transport proteins
in a nuclear localization signal
(NLS)-dependent manner. Recombinant
transcription factor LEF-1 protein,
which contains a highly basic
B-box NLS, fused to an enhanced green
fluorescent protein (eGFP)
tag (a generous gift from M. Prieve and M. Waterman) is efficiently
localized by nuclear transport within
preformed solid-phase nuclei
(Fig.
2A, panel b). Transported LEF-eGFP
appears as a green, fluorescent
ring (b) around the DNA-coupled
paramagnetic beads (c) encapsulated
by a membrane (a). No concentrated
green fluorescence is detected
when LEF-eGFP is added to DNA-coupled
beads in the absence of
nucleus formation (d), or when a glutathione
S-transferase (GST)-eGFP
fusion protein lacking an NLS is
added to the preformed nuclei
(Fig.
2A, panel e), demonstrating the
specificity of the observed
transport. The yellow fluorescence observed
after long exposure
times in these latter two cases is due to intrinsic
fluorescence
of the beads themselves. These data show that solid-phase
nuclei
selectively transport proteins in an NLS-dependent manner,
confirming
the functional integrity of the nuclear membrane.
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FIG. 2.
Solid-phase synthetic nuclei. (A) Solid-phase synthetic
nuclei support nuclear transport. Immobilized AFP templates were
assembled for 1.5 h in LSS to generate solid-phase synthetic
nuclei (a to c and e), or in Xl buffer alone (d). Assembly reaction
mixtures were further incubated for 30 min with approximately 150 ng of
either LEF-1:eGFP (a to d) or GST:eGFP (e). After addition of propidium
iodide, samples were visualized by phase-contrast microscopy (a) or by
fluorescence microscopy (b to e) with either a GFP wide-band filter (b,
d, and e) or a rhodamine filter (c). In the absence of a nuclear
membrane (d) or an NLS (e), no concentrated GFP fluorescence was observed. (B) Steady-state DNA
replication. Immobilized AFP templates were assembled into synthetic
nuclei in the presence of [-32P]dATP and incubated for
0 (lane 1), 1 (lane 2), 2 (lane 3), or 4 (lane 4 and 5) hours in LSS.
Aphidicolin (40 ng/µl) was added to lane 5. Purified DNA was
subjected to electrophoresis and visualized by autoradiography of the
dried gel. Labeled arrows point to replicated full-length DNA,
unresolved replication intermediates or concatemers, and the location
of the well. Radiolabeled lambda DNA digested with
HindIII was run as molecular size standards (lane MW).
(C) DNA replication under transcription conditions. Immobilized AFP
templates were assembled into synthetic nuclei by preincubation
(preinc.) in HSS plus membranes (MEMB) (lanes 1 to 10) or in LSS (lanes
11 and 12) or by preassembly in HSS followed by nucleus assembly in LSS
(lanes 13 to 15). Templates were incubated with either NDB (lanes  )
or adult ML (ml), or HepG2 (hep) extract at the indicated times during
assembly. Templates in lanes 13 to 15 were preincubated with ML extract
prior to assembly in HSS, and these silenced templates were then
assembled into nuclei in the presence of the indicated extracts.
Incubations and assembly reactions were performed exactly as diagrammed
in Fig. 3 and 4. DNA replication was monitored by inclusion of
[-32P]dATP during synthetic nucleus assembly. Purified
DNA was subjected to electrophoresis and visualized by autoradiography
of the dried gel. Labeled arrows point to replicated full-length DNA
and unresolved replication intermediates. Relative replication levels
(fold) in comparison to buffer controls are indicated below the lanes.
(D) Nucleosome assembly on replicated DNA. Immobilized templates were
assembled into synthetic nuclei in LSS in the presence of
[-32P]dATP for 2 h. DNA was then digested with
MNase, and aliquots were withdrawn for analysis at 0, 2.5, 5, 10, 20, 40, 60, and 80 min. Nucleosome spacing was estimated by comparison to a
123-bp DNA ladder (Gibco-BRL) and radiolabeled  HindIII fragments (lane MW). Arrows point to
MNase-protected fragments.
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DNA replication within solid-phase nuclei formed by incubation in
Xenopus egg extract LSS was measured by steady-state levels
of [
-
32P]dATP-labeled DNA accumulated over time (Fig.
2B). Radioactive
incorporation is specifically blocked by addition of a
DNA polymerase
inhibitor, aphidicolin, indicating that radiolabel
incorporation
is polymerase specific (Fig.
2B, lane 5).
Slower-migrating, unresolved
replication intermediates or intertwined
daughter molecules are
often apparent (Fig.
2B and C). We have
previously confirmed that
DNA radiolabeled under steady-state
conditions is the end product
of semiconservative DNA replication and
not gap repair by methylation
state analysis with the isoschizomers
DpnI and
Sau3AI (
12).
Relative
replication levels were assessed under a number of incubation
conditions that were later used in transcription analyses (Fig.
2C):
preincubation in buffer only, adult ML, or HepG2 hepatoma
cell extract,
followed by assembly in HSS plus membrane vesicles,
LSS only, or
sequential HSS, washes, and LSS incubation, as indicated.
Under each
incubation condition, a baseline was established by
addition of buffer
only, followed by nucleus assembly and replication
in the presence of
[
-
32P]dATP, DNA purification, and separation by gel
electrophoresis.
Comparison of replicated, full-length DNA reveals a
slight stimulation
of replication in ML extract of 1.4-fold when
preincubated (Fig.
2C, lane 2) but not when added during nucleus
assembly (lane 3)
in HSS plus membrane vesicles. Similarly, HepG2
extract enhanced
replication when preincubated with the template
(1.6-fold; Fig.
2C, lane 6), but not when added during assembly (lane
8). Aphidicolin
addition to HSS and membrane vesicles inhibited
replication by
87 and 97% in the presence of ML extract and hepatoma
extract,
respectively (Fig.
2C, lanes 4 and 10), and by 93% when added
to LSS assembly reactions (lane 12). Addition of ML extract to
preassembled developmentally repressed chromatin templates during
nucleus assembly in LSS (Fig.
2C, lane 15) again had a stimulatory
effect (1.8-fold) on DNA replication. These results indicate that
ML
factors may slightly increase the efficiency of DNA replication
in
synthetic nuclei. No increase was detected when hepatoma extract
was
added during nucleus assembly of preassembled chromatin templates
(Fig.
2C, lane
14).
To assess the chromatin structure of newly replicated DNA, solid-phase
AFP templates assembled into synthetic nuclei in LSS
were replicated in
the presence of [
-
32P]dATP followed by digestion with
MNase (Fig.
2D). A time course
of digestion with MNase revealed
extensive assembly of nucleosomes
on the newly replicated DNA.
Nucleosomal protection of DNA occurred
with a spacing of approximately
160 to 180 bp, identical to that
observed with uncoupled DNA assembled
into chromatin by
Xenopus egg extract (data not shown)
(
9).
DNA replication mediates hepatoma-specific activation of
chromatin-repressed AFP.
After establishing that solid-phase
synthetic nuclei supported both nuclear transport and DNA replication,
we directly tested whether DNA replication was required for
hepatoma-induced reprogramming of AFP gene expression. To recreate the
proliferative environment present in hepatoma cells, AFP templates were
assembled into synthetic nuclei by incubation in fractionated
Xenopus egg extract (HSS) plus membrane vesicles. These
conditions have been shown previously to direct stepwise chromatin
assembly and synthetic nucleus formation (41, 49), capable
of derepressing the 
-globin promoter in the presence of
transactivating red blood cell extracts (7).
We assayed the ability of hepatoma factors to activate
chromatin-repressed AFP templates under replicating and nonreplicating
conditions, as illustrated in Fig.
3A.
Complete derepression of
nucleosome-assembled templates occurred only
when hepatoma extract
was provided either prior to chromatin
assembly/replication (Fig.
3B, lane 2) or during DNA replication (1 to
1.5 h after
Xenopus extract addition; lane 5). As the
majority of nuclei are already
assembled after a 1-h incubation in HSS
plus membranes (
41),
proteins added after this point (Fig.
3B, lanes 4 to 9) must likely
be actively transported across the
nuclear membrane in order to
influence gene regulation. However, even
under these stringent
conditions, transcription levels measured on
chromatin-repressed
templates (Fig.
3B, lane 1) replicated in the
presence of hepatoma
extract (10.2-fold derepression; lane 5) were 91%
of those obtained
by hepatoma preincubation (11.3-fold derepression;
lane 2), demonstrating
a nearly complete reversal of the
chromatin-repressed state during
replication.
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FIG. 3.
Hepatoma-specific activation of AFP during DNA
replication. (A) Diagram of solid-phase nucleus assembly and
transcription system. (B) In vitro transcription. Immobilized AFP
templates were incubated with NDB (lanes 1 and 4 to 9) or HepG2 (lane
2) or ML (lane 3) extract. Templates were assembled into synthetic
nuclei by incubation in HSS plus membranes. Synthetic nuclei were
further incubated for an additional 30 min in the presence of NDB
(lanes 1 to 4 and 7) or HepG2 (lanes 5 and 8) or ML (lanes 6 and 9)
extract. Aphidicolin (lanes 7 to 9) was added concomitantly with the
HSS plus membranes. All samples were washed and in vitro transcribed in
HeLa extract. Radiolabeled X174 DNA digested with HaeIII
was run as molecular size standards (lane MW).
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Chromatin-mediated repression of AFP gene expression was maintained
during replication in the absence of cellular extract
(Fig.
3B, lane
4), indicating that the physical process of DNA
synthesis alone is
insufficient to activate AFP transcription.
Addition of ML extract
during DNA replication (Fig.
3B, lane 6)
resulted in maintenance of the
repressed state; it is unclear
whether this repression is propagated
through passive nucleosome
assembly, as in lane 4, or reflects the
AFP-silenced, differentiated
hepatic state imposed by ML extract (lane
3), or both. These two
models of ML-mediated repression during DNA
replication have been
further dissected, as described
below.
Hepatoma-specific proteins activated a low level of transcription
(25%, comparing lane 8 with lane 5 in Fig.
3B) when added
to
solid-phase nuclei in the presence of a replication inhibitor
(aphidicolin). As we have seen that aphidicolin inhibits DNA
replication
97% under these conditions (Fig.
2C, lane 10), the
hepatoma-specific
activation observed in lane 8 is most likely not
mediated by residual
DNA replication. These data suggest either a
limited ability of
hepatoma factors to bind their sites in nucleosomal
DNA or the
presence of chromatin-remodeling activity, or both. ML
extract
was unable to derepress chromatin during this postassembly
incubation
(Fig.
3B, lane 9). Importantly, chromatin remodeling factors
present
in the
Xenopus egg extract itself are insufficient
to activate
a chromatin-repressed AFP gene (Fig.
3B, lane
7).
DNA replication mediates activation of developmentally
silenced AFP in vitro.
These results suggest that DNA
replication facilitates binding of hepatoma activators to a
chromatin-repressed template, resulting in derepression and
transactivation of the chromatin template. In the above experiments,
chromatin-repressed AFP templates were used as the starting material to
examine the effects of DNA replication on AFP gene expression. However,
in the adult liver, repression of AFP occurs through an active process
involving developmental stage-specific repressor proteins (35,
54). To more accurately model the developmentally silenced state
of AFP, we employed a two-stage chromatin-synthetic nucleus assembly
system (38) coupled to in vitro transcription (Fig.
4A). In the first stage, developmentally silenced AFP templates were generated by preincubation with ML extract
and chromatin assembly in HSS, followed by an isoosmotic wash step to
remove all protein not interacting with DNA or chromatin. Silenced
templates were then exposed to one of two pathways in the second stage:
(i) stable maintenance of chromatin templates by incubation in buffer
only, or (ii) formation of synthetic nuclei and semiconservative
replication of chromatin-assembled templates by incubation in
unfractionated Xenopus egg extract (LSS). Using this
approach (sequential HSS-LSS incubations), the functional consequences
of synchronized DNA replication (38) within solid-phase nuclei (pathway 2) can be assessed independently of nuclear transport and under conditions of limited or excess protein or extract
concentration.
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FIG. 4.
DNA replication alone is sufficient to activate a
developmentally silenced AFP template. (A) Diagram of two-step
solid-phase nucleus replication and transcription system. (B) In vitro
transcription. Immobilized AFP templates were incubated with ML extract
(lanes 1 to 6) prior to chromatin assembly. Chromatin templates were
washed and either incubated in Xl buffer (lanes 1, 3, and 5) or further
assembled into synthetic nuclei by incubation with LSS (lanes 2, 4, and
6). NDB (lanes 1 and 2), HepG2 extract (lanes 3 and 4), or ML extract
(lanes 5 and 6) was included during the 2-h postchromatin/nucleus
assembly period. All samples were washed and in vitro transcribed in
HeLa extract. Radiolabeled X174 DNA digested with HaeIII
was run as molecular size standards (lane MW).
|
|
After passage of a replication fork,
trans-acting factors
must bind their respective sites on the newly replicated DNA in
order
to reestablish a given gene expression pattern. The local
concentration
of factors during chromatin reassembly would therefore
be expected to
influence the outcome of this process. To determine
whether the
concentration of ML proteins present during replication
might affect
the reassembly of developmentally silenced templates
postreplication,
we replicated the silenced templates in buffer
only. As shown in Fig.
4B, we found that, unlike our results with
chromatin-repressed AFP
templates, DNA replication of developmentally
silenced AFP
chromatin templates in the absence of any additional
cellular extracts
resulted in transcription activation (5.3-fold;
Fig.
4B, lane 2).
Limiting the concentration of adult ML extract,
through washing away of
unbound proteins, activated AFP transcription
in a DNA
replication-dependent manner (Fig.
4B, compare lanes
1 and 2).
Therefore, DNA replication may mediate the loss of sufficient
DNA-bound
transcription repressors to inhibit reformation of silencing
complexes
on the replicated
DNA.
Addition of hepatoma extract during DNA replication of developmentally
silenced templates increased transcription only 1.2-
to 2-fold compared
with the effect of replication alone (Fig.
4B, compare lanes 4 and 2).
However, as shown in Fig.
3B, both
HepG2 extract and DNA replication
are required to activate a chromatin-repressed
template. These data
suggest that further supplementation with
hepatoma factors does not
significantly amplify replication-mediated
activation of
developmentally silenced AFP. Additionally, under
these "washed
chromatin" conditions, HepG2 extract added in the
absence of
replication (Fig.
4B, lane 3) was unable to alter the
repressed state
of AFP, whereas replication-independent activation
by HepG2 was
observed on the "unwashed" chromatin templates (Fig.
3B, lane 8).
Chromatin remodeling complexes present in the
Xenopus egg
extract may therefore participate in the replication-independent
activation.
Depletion of developmental repressor(s) plays a role in AFP
activation.
Replication-mediated depletion of key repressors
should occur only under limiting protein conditions. The paramagnetic
bead concentration and isoosmotic washes of chromatin performed prior to solid-phase nucleus formation imposed these conditions (Fig. 4A).
Based on this assumption, replication of the silenced template in the
presence of excess ML extract should negate the repressor titration
effect as efficiently as inhibition of DNA replication does. As shown
in Fig. 4B, addition of ML extract during replication resulted in
maintenance of the silenced state (lane 6). Analysis of the DNA
replication levels under these conditions (Fig. 2C, lane 15) indicated
that the presence of ML extract during replication led to a 1.8-fold
increase in replication. Thus, even under enhanced replication
conditions, excess ML extract was sufficient to maintain the silenced state.
The activation observed upon replication of the ML-silenced AFP
template suggests that duplication of the target DNA may result
in
titration of critical, developmental stage-specific repressors,
allowing ML activators to reassemble a transcriptionally competent
template. We reasoned that if increasing the DNA concentration
by DNA
replication is sufficient to deplete a repressor(s) under
limiting
adult ML extract conditions, then exogenous addition
of increasing
amounts of AFP DNA during replication in excess
adult ML extract should
mimic this effect and activate transcription.
To test this possibility
directly, we replicated ML-silenced AFP
chromatin in excess ML extract
plus various concentrations of
supercoiled AFP DNA (Fig.
5). Addition of AFP DNA blocked excess
adult ML-mediated repression (Fig.
5B, compare lanes 3 and 4 to
lane
2). Derepression of AFP transcription was also detected following
addition of competitor AFP DNA that lacked the
lacZ coding
region
(data not shown). Titration of repressors by AFP DNA required
the physical process of replication, as inclusion of aphidicolin
maintained the repressed state (Fig.
5B, compare lanes 9 and 10).
Together, these results strongly support our hypothesis that DNA
replication mediates tumor marker gene activation both by facilitating
binding of transactivators and through depletion of repressor
proteins.
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FIG. 5.
Sequence specificity and replication dependence of
repressor depletion. (A) Diagram of two-step solid-phase nucleus
replication and transcription system. (B) In vitro transcription.
Immobilized AFP templates were incubated with ML extract prior to
chromatin assembly in HSS (lanes 1 to 10). Chromatin templates were
washed and further incubated in the presence (lanes 2 to 6 and 8 to 10)
or absence (lanes 1 and 7) of LSS to assemble synthetic nuclei.
Aphidicolin was added during synthetic nucleus formation to block DNA
replication in lane 10. ML extract was added during synthetic nucleus
formation (lanes 2 to 6 and 8 to 10). The indicated amount of
supercoiled AFP(3.8)-lacZ DNA (lanes 3, 4, 9, and 10) or
supercoiled pUC DNA (lanes 5 and 6) was added as a competitor during
synthetic nucleus formation. All samples were washed and in vitro
transcribed in nuclear HeLa extract. Radiolabeled X174 DNA digested
with HaeIII was run as molecular size standards (lane MW).
|
|
To determine whether depletion of a repressor(s) was due to competition
in
trans by sequence-specific binding, we compared
the
ability of nonspecific pUC DNA versus AFP DNA to compete for
repressor
binding. Addition of equivalent amounts of pUC DNA (Fig.
5B, lanes 5 and 6) had no effect on AFP expression, indicating
that depletion of a
repressor(s) requires the presence of AFP-specific
sequences.
Importantly, the sequence specificity of the activation
rules out the
formal possibility that excess DNA nonspecifically
depleted histones
from the LSS, reducing the capacity for chromatin-mediated
repression.
Studies of factors bound to competitor DNA and the
chromatin structure
of the competitors after replication-mediated
depletion are ongoing.
These data indicate that sequence-specific
repressors can be depleted
from the local environment during DNA
replication, thereby inhibiting
the reformation of a transcriptionally
silenced
template.
Repression of AFP transcription is sequence specific.
To begin
characterization of the developmental repressors depleted by DNA
replication, we transcribed AFP templates deleted within the
developmental repressor region, a broadly defined 750-bp region (1000
to 250) that mediates postnatal repression of AFP expression in
transgenic analysis (54). AFP DNA was deleted by
PCR-mediated mutagenesis from 1000 to 541 (removing approximately one-half of the developmental repressor region) and from 1000 to
209 (removing the entire developmental repressor region). RNA
transcribed from deletion templates is distinguished from full-length
primer extension products by PCR-engineered removal of a 13-nucleotide
polylinker connecting AFP to lacZ (Fig. 6A and
B). The activities of these templates
assembled into chromatin in the presence of ML extract (Fig. 6A, lanes
2, 5, and 7) were corrected for effects on basal transcription (lanes
1, 3, and 6) prior to comparison with each other. Removal of
nucleotides between 1000 and 541 led to a partial loss of silencing
by ML extract (lane 5), a 2.3-fold derepression between this template and the full-length construct. Complete removal of the developmental repressor region to 209 led to a threefold derepression (lane 7).
Comparison of these deletion templates as chromatin-free bead-DNA transcribed in HeLa and ML extracts (Fig. 6B) revealed a similar pattern of derepression (compare lanes 5 and 6 to lane 4). In vitro
transcription of the deletion templates in HeLa extract (Fig. 6B, lanes
1 to 3) again showed the same slight increase in basal transcription
(1.2-fold). Loss of silencing on chromatin-free templates was 5.8-fold
upon deletion to 541 and 7.4-fold upon deletion to 209. We
therefore focused our analysis of developmental repressor protein
interaction on the region between 541 and 1000 of AFP DNA.
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FIG. 6.
Developmental repression is mediated through
sequence-specific binding. (A) Derepression of developmentally silenced
AFP chromatin. Immobilized full-length AFP templates (AFP) and AFP
templates containing deletions from 1000 to 541 (AFP 541) and
1000 to 209 (AFP209) were preincubated (preinc.) in either NDB
(lanes 1, 3, 4, and 6) or adult ML extract (lanes 2, 5, and 7).
Templates were then either assembled into chromatin in HSS (lanes 2, 4, 5, and 7) or incubated in Xl buffer (lanes 1, 3, and 6). All samples
were washed and in vitro transcribed in HeLa extract. The primer
extension products obtained from both the full-length template and the
deletion templates (13-bp smaller) are indicated by arrows. (B)
Developmental repression of AFP is not mediated by chromatin.
Immobilized full-length AFP, AFP541, and AFP209 were in vitro
transcribed in either HeLa extract (lanes 1 to 3) or adult ML extract
(lanes 4 to 6). The primer extension products are indicated by arrows.
(C) Deletion of 150 bp is sufficient to derepress AFP. Immobilized
full-length AFP (AFP) and AFP templates containing deletions from
1000 to 850 (850), 1000 to 765 (765), and 1000 to 586
(586) were in vitro transcribed in either HeLa extract (lanes 1 to
4) or ML extract (lanes 5 to 8). These deletion mutants were designed
to yield a primer extension product identical to that of the
full-length AFP transcript, and this product is indicated by a single
arrow. Lanes 5 to 8 were overexposed relative to lanes 1 to 4 in order
to visualize the ML primer extension products.
|
|
Further deletions were constructed by exonuclease digestion from
1000
to
850, to
765, and to
586. Analysis was performed
by
transcribing the deletion templates in both HeLa and ML extracts
in the
absence of chromatin assembly (Fig.
6C). Removal of 150
bp to
850
augmented basal transcription slightly relative to
the full-length
template (1.3-fold; Fig.
6C, compare lanes 1 and
2) and derepressed
transcription in ML extracts 7.3-fold (compare
lanes 5 and 6). Further
deletions to
586 did not result in any
further effects on either
basal (1.3-fold; Fig.
6C, compare lanes
1 and 4) or derepressed
(7.0-fold; compare lanes 5 and 8) transcription.
As above, derepression
was calculated after correction for effects
on basal transcription
activity. These analyses show that a developmental
repressor
DNA-binding element between
1000 and
850 mediates
a majority of
developmental silencing independently of chromatin
assembly. Other
trans-acting proteins may modulate AFP expression,
but
clearly a regulatory element for silencing lies within this
region.
One candidate repressor protein that binds within this region is the
p53 tumor suppressor protein. In vitro analysis revealed
that p53
protein binds and excludes hepatocyte nuclear factor
3 activator from
an overlapping site between
860 and
833 of
AFP. Postnatal
repression of AFP expression correlated with induction
of p53 protein
in 2-week and adult liver nuclear extracts. By
transient-transfection
analysis with hepatoma and fibroblast cells,
we have previously found
that p53-mediated repression of AFP expression
is tissue specific
(
35). Studies are under way to identify tissue-specific
and
ubiquitously expressed repressor proteins that interact with
p53
protein and/or bind within the identified 150-bp repressor
element to
silence AFP expression during
development.
| |
DISCUSSION |
By establishing a hepatoma-like environment in solid-phase,
synthetic nuclei, we have modeled the aberrant activation of AFP gene
expression that occurs in vivo during liver tumor formation. We present
evidence that replication-mediated activation occurs through a bimodal
mechanism: (i) depleting the local concentration of developmental
repressor proteins and (ii) facilitating binding of transcription
activators to their sites on nucleosomal DNA. These studies indicate
that DNA replication, when uncoupled from normal, cellular protein
biosynthesis, acts as a driving force in activation of developmentally
silenced genes and may amplify tumor marker gene expression during
proliferation of hepatoma cells.
DNA replication facilitates activator binding.
Reorganization
of existing chromatin structure can occur by both
replication-dependent and -independent mechanisms (e.g., nucleosome remodeling, DNA methylation, and histone
modification). It is likely that many layers of control exist to
modulate chromatin structure, particularly on genes that must respond
rapidly to hormonal fluctuations or environmental challenges (4,
6, 44, 48, 53, 61). Our studies with the AFP gene show that while
low levels of replication-independent activation can be observed, DNA
replication greatly facilitates derepression of AFP.
DNA replication allows both activators and repressors to gain access to
sites normally occluded by nucleosomes (
29; reviewed
in reference
58). As the majority of transactivators
bind with
reduced affinity to their sites on nucleosomal DNA versus
free
DNA (reviewed in references
1 and
60), DNA replication may
enable newly synthesized or
activated proteins to bind and change
gene expression profiles. During
tumorigenesis, established gene
expression patterns are often
disrupted, frequently resulting
in reversion to those representative of
early development (reviewed
in reference
39). The
unchecked cell cycling characteristic
of tumor cells may facilitate
this reprogramming of gene expression
patterns.
Parallels for gene regulatory shifts during tumorigenesis are found in
regulation at silent mating type loci and telomeres
in yeast cells.
Relief of telomeric silencing in cases of position
effect variegation
requires both cell cycle progression and transactivator
expression
(
3), with assembly of silent chromatin occurring
as a
default. At mating type loci, establishment of the silenced
state is an
active process that requires passage through S phase
(reviewed in
references
18 and
37). Thus,
although different
mechanisms may be involved, reversal of
developmental silencing
in tumor cells and gene switching at both
mating type loci and
telomeric regions appear to require progression
through the cell
cycle.
DNA replication mediates depletion of developmental repressor
proteins.
Replication of developmentally silenced AFP under
limiting ML conditions caused transcription activation. Importantly,
replication of chromatin-repressed AFP (no ML present) did not activate
AFP expression, demonstrating that the physical process of replication alone is insufficient to activate AFP. We have depicted the repressor complex as a brick wall of multiple component factors which together inhibit activation of the AFP promoter even in the presence of upstream
activator proteins (Fig. 7). As would be
predicted from this model, repressor (brick) depletion during S phase,
at a limiting repressor(s) concentration, may result in incomplete
reassembly of repressor complexes (brick walls) on newly replicated
DNA. We have shown that repression is recovered by providing additional developmental repressors (bricks) in the form of ML extract during replication. Repression of AFP expression did not occur indirectly by
inhibition of DNA replication in solid-phase nuclei, as addition of ML
extract does not decrease the relative amount of DNA replication.
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FIG. 7.
Model for DNA replication-mediated activation of a
developmentally silenced gene. The developmental repressor region of
the AFP gene may be bound by a collection of repressor proteins
(symbolized by bricks) during postnatal repression. This brick wall
prohibits expression of the gene, perhaps by disrupting an upstream
activator-promoter communication of an upstream activator(s) with the
promoter. Duplication of repressor-binding sites, through either DNA
replication or addition of exogenous AFP DNA, effectively depletes the
developmental repressors, resulting in incomplete reassembly of the
brick wall on the newly replicated DNA. In the presence of an
incomplete complex of repressors, activators can now interact with
proximal promoter-bound factors in order to initiate transcription.
|
|
Replication timing is linked to cell memory and gene
activation.
During cell cycle S phase, all genetic material must
be faithfully replicated. Numerous studies have demonstrated that
chromatin structure is stably propagated from generation to generation
(reviewed in reference 56). Passage of the
replication fork results in transient disruption of 1 to 2 nucleosomes
as well as a majority of DNA-associated trans-acting factors
(19, 34, 43), implicating a mechanism for reestablishing
protein-DNA contacts postreplication. Models proposed to explain cell
memory must consider how expression states, whether active or silent,
are both established and maintained. In one such model with particular
relevance to our studies, DNA replication timing determines the
expression state of a given gene (21). Comparison of
numerous tissue-specific and developmental stage-specific genes,
including AFP and albumin, with housekeeping genes has revealed a
correlation between DNA replication timing and gene transcription, with
highly expressed genes replicated early and repressed genes replicated
late in the cell cycle (reviewed in references 15
and 26).
Timing of DNA replication has been implicated in silencing at mating
type loci and yeast telomeres, in X-chromosome inactivation
(reviewed
in reference
16), and in tissue-specific expression
of certain genes, e.g., globin and
Xenopus 5S genes
(reviewed
in references
24 and
28). Mutations in
Saccharomyces
cerevisiae which cause defective silencing at the HMR locus can be
suppressed
functionally by second-site mutations that increase the time
between
cell cycle phases, slowing progression through G
1,
S, or G
2/M
(reviewed in reference
18).
One implication of these studies
is that a critical concentration of a
multicomponent silencing
complex of proteins is required, and by
lengthening the timing
of cell cycle progression, a limiting protein(s)
can increase
in functional concentration by biosynthesis or
posttranslational
modification. Similarly, transactivator-mediated
derepression
of a telomeric
URA3 gene displays strict
concentration dependence,
suggesting that competition occurs during
each cell cycle between
the establishment of active and silent states
(
3).
The present study, as well as dissection of AFP gene regulation in
transgenic mouse models, reveals that
trans-acting factors
are required to establish developmental stage-specific silencing
(
54). Transition between the active fetal state and the
silenced
postnatal one may rely on competition between repressors and
transactivators
during DNA replication. Indirect support for this model
lies in
the brief lag time between postnatal repression of AFP
expression
and a nearly complete cessation of DNA replication in
differentiated
hepatocytes (
52). In response to liver damage
and/or carcinogenesis,
differentiated hepatocytes revert in many ways
to a fetal phenotype,
dividing rapidly and expressing early
developmental markers such
as AFP. One intriguing possibility is that
hepatocyte replication
timing is shifted to an early-replicating phase,
similar to the
developmental switch in replication timing and
expression of
-globin
genes (
32). When challenged by DNA
replication early in S phase,
the concentration of AFP repressor
proteins may be too low to
maintain a silenced state, disrupting the
balance of differentiation.
Using the synthetic nucleus system, it will
be possible to introduce
various concentrations of a wide spectrum of
candidate activators
and repressors during DNA replication, with the
ultimate goal
of reconstituting AFP activation in a defined
environment.
The results presented here emphasize the importance of examining gene
activation under a range of protein concentrations not
only within the
context of physiological chromatin, but also during
on-going cellular
processes such as DNA replication. DNA replication
and protein
synthesis are strictly regulated in eukaryotic cells
(
10,
57). By uncoupling these two processes in synthetic nuclei,
we
have demonstrated that replication in the absence of protein
synthesis
can have severe effects on the maintenance of established
transcription
states. Rapid cell cycling, during tumorigenesis
or liver regeneration,
may therefore upset the balance of transacting
factors required for
faithful, epigenetic propagation of developmentally
regulated
expression
patterns.
| |
ACKNOWLEDGMENTS |
We are grateful to K. Wernke-Dollries for excellent technical
assistance with the DNA replication assays. We thank M. Prieve and M. Waterman (UC Irvine), B. Spear (U. Kentucky) and K. Zaret (Brown Univ.)
for generously providing reagents and K. Steigerwald for assistance
with fluorescence microscopy. We are also grateful to P. Brindle, J. Ma, K. Tepperman, K. Lee, and Y. Sanchez for insightful comments and/or
critical reading.
A.J.C. is the recipient of an NIH postdoctoral fellowship (CA73083).
This work was supported in part by a grant from NIH (GM53683) to M.C.B.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Dept. of
Molecular Genetics, Biochemistry and Microbiology, University of
Cincinnati, Cincinnati, OH 45267-0524. Phone: (513) 558-5541. Fax:
(513) 558-8474. E-mail: Michelle.Barton{at}UC.edu.
| |
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Molecular and Cellular Biology, June 2000, p. 4169-4180, Vol. 20, No. 11
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Abstract
Introduction
