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INTRODUCTION |
Faithful transmission of genetic material from one cell generation
to the next requires that DNA be precisely duplicated each cell cycle.
Consequently, cells possess mechanisms that trigger DNA replication at
a discrete point in the cell cycle and delay the onset of mitosis until
replication is complete. Cells also possess mechanisms that coordinate
the firing of multiple discrete replication origins with respect to one
another during S phase and inhibit re-replication before mitosis. Some
of these regulatory pathways directly affect the assembly or activation
of the multisubunit replication initiation complexes that form with
replication origins. Replication origins in the budding yeast
Saccharomyces cerevisiae were first identified by their
ability to promote replication of episomal DNA (1). Many of these
autonomously replicating sequences
(ARSs)1 were shown
subsequently to act as replicators in their native chromosomal loci
(1), and recent replication initiation mapping studies indicate that
DNA synthesis begins at one or a few discrete sites within ARSs (2,
3).
Early analyses indicated that ARSs are composed of an essential A
domain containing the conserved ARS consensus sequence (ACS; Ref. 4)
and a B domain. The archetypal ARS1 also contains a C domain (5, 6),
which is packaged in a uniquely positioned nucleosome. This nucleosome
may reduce the probability that the adjacent ACS will be packaged into
a nucleosome capable of suppressing ARS activity (cf. Ref.
7). Marahrens and Stillman (8) found later that the B domain of the
ARS1 could be subdivided into a B3 element that is bound by the
transcription factor Abf1p (cf. Ref. 9), a B2 element
thought to serve as a DNA-unwinding element (10, 11), and a B1 element.
The B1 element acts in conjunction with the ACS to bind a
six-polypeptide origin recognition complex (ORC; Ref. 12). ORC is
required for the initiation of replication (13, 14) and possesses an
ATPase activity that is modulated by origin binding (15). Although the
functional significance of this ATPase activity is unknown, one role of
ORC is to help recruit additional components of the replication
initiation complex. Specifically, ORC-ARS complexes recruit Cdc6p
during late mitosis or early G1 phase; this is followed by
recruitment of a 6-subunit complex consisting of MCM proteins 2 through
7 (16). These proteins make up the "pre-replicative complex," which
protects a large segment of ARS DNA from DNase I (17). Another
essential factor, Cdc45p, joins the already-formed pre-replicative
complex later in G1 phase to form the "pre-initiation
complex" (18, 19). The single-stranded DNA-binding protein RPA, DNA
polymerase 
, and presumably other replicative enzymes as well join
the pre-initiation complex at about the time of origin firing (18, 20)
to form the replicative complex. Origin firing appears to involve loss of the MCM2-7 complex and Cdc45p from the pre-initiation complex and
redistribution of Cdc45p and some of the MCM proteins to the nascent
replication forks (18). This last finding together with the discovery
that an MCM subcomplex possesses helicase activity in vitro
suggested that an MCM subcomplex serves as the replicative helicase in
yeast (21). The recent discovery that an MCM-like protein in
Methanobacterium thermoautotrophicum not only possesses helicase activity but also assembles into a double hexamer (22) suggests that MCM complexes are highly functionally conserved and serve
as replicative helicases in all eukaryotes. After origin firing, a
"post-replicative complex" remains associated with ARS DNA through
the remainder of S phase and into the G2 and M phases; the
post-replicative complex has an altered DNase I profile and may consist
solely of ORC and Abf1p bound to ARS1 DNA (17).
The sequence of events described above is tightly regulated by Cdc28p,
a cyclin-dependent kinase (CDK) protein kinase required for commitment
of cells to a program of replication and cell division (reviewed in
Ref. 23) and by cyclin-dependent kinase (CDK)-like protein kinases,
notably Cdc7p in association with its cyclin-like partner Dbf4p
(24-27). During late G1 phase, the Cdc28p kinase in
association with the Cln cyclins phosphorylates the B-type (Clb) cyclin
inhibitor Sic1p, which renders it susceptible to ubiquitin-mediated
proteolysis (28, 29). Cyclin-dependent kinase phosphorylation of Cdc6p
during late G1 renders it susceptible to proteolysis as
well (30, 31). Loss of Cdc6p and phosphorylation of MCM proteins
prevents their re-association with ARS-ORC complexes until late in
mitosis, when the Clb cyclins are degraded. This sequence of
phosphorylation and proteolysis events makes activation of replication
complexes a unidirectional event and helps prevent re-replication
before mitosis. The discovery that the Cdc7p-Dbf4p kinase complex
physically associates with individual replication initiation complexes
(32, 33) and acts throughout S phase to trigger the firing of
individual origins (34, 35) suggests that Cdc7p directly modifies one
or more factors in the replication initiation complex. This inference
is supported by in vitro evidence that the Cdc7p-Dbf4p
kinase modifies Mcm 2, 3, 4, 6, and 7 as well as the largest subunit of
DNA polymerase 
-primase (33, 36, 37). Cdc7p-Dbf4p homologs in
Schizosaccharomyces pombe, Xenopus laevis, and
humans appear to have similar in vitro substrate preferences
(38-43). It is noteworthy, however, that a specific Cdc7p bypass
mutation in S. cerevisiae, mcm5/cdc46-bob1,
resides in Mcm5p (44), a subunit of the MCM complex that is not
efficiently phosphorylated in vitro by Cdc7p (33). Thus, it
is not entirely clear if the in vitro targets of Cdc7p are
identical to its in vivo targets. Additionally, the
consequences of the action of Cdc7p at replication origins are still
unclear. What is clear is that Cdc7p plays a critical role in
activation of replication, and in this paper we have investigated
structural changes at replication origins that can be attributed to the
action of Cdc7p.
Previous in vivo studies of cell cycle-regulated events at
ARSs have used cdc mutants, the pheromone -factor, the
replication elongation inhibitor hydroxyurea (HU), and the
microtubule-destabilizing agent nocodazole to block cells at selected
points in the cell cycle (e.g. Refs. 17 and 45). Cells
blocked with -factor or at the Cdc7p execution point exhibit a
pre-replicative complex footprint, whereas cells blocked with HU or
nocodazole exhibit a post-replicative complex footprint (17). To
further dissect events that occur after the Cdc7p execution point but
before the HU-sensitive step, we investigated the use of additional
cdc mutants whose execution points have been mapped to
within the Cdc7p-HU interval. One such mutant is cdc8-1,
which encodes a thymidylate kinase required for DNA synthesis (46, 47).
cdc8-1 mutants display a quick stop replication arrest when
shifted to the restrictive temperature and resume DNA synthesis after
their return to the permissive temperature. This rapid and reversible
phenotype and the fact that Cdc8p is required shortly after Cdc7p (48)
suggested that the study of cdc8-1-blocked cells might
reveal structural events that could be attributed directly to the
action of Cdc7p. The ARS1 footprints and DNA topology in
cdc8-1-blocked cells proved to be distinct from those in
cdc7-1-blocked cells and suggest that localized DNA
unwinding occurs before the CDC8 execution point. The
cdc8-1 block is fully reversible, suggesting that the structure evident in cdc8-1-blocked cells reflects a
bona fide replication intermediate. The DNA topology as well
as micrococcal nuclease (MNase) cleavage and KMnO4
modification patterns characteristic of cdc8-1-blocked cells
were also evident in both mcm5/cdc46-bob1 single mutants and
cdc7
mcm5/cdc46-bob1 double mutants that had been blocked in G1 phase by -factor. Since
mcm5/cdc46-bob1 is a CDC7- and
DBF4-specific bypass mutant, this result strongly suggests
that the structural changes evident in cdc8-1-blocked cells
are normally induced by the Cdc7p kinase and involve changes in the MCM complex.
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EXPERIMENTAL PROCEDURES |
Plasmids and Cell Strains--
Plasmid pJL347 (provided by J. Li, University of California at San Francisco) was used in episomal
footprinting and topoisomer studies and contains a centromere
(CEN6), a selectable marker (URA3), and the A and
B domains of ARS1. Plasmid pMD-2 was used as a footprinting control and
was constructed by inserting into the NheI site of pJL347 a
second copy of ARS1 rendered non-functional by a G to T substitution in
the ACS. Plasmid pKWD50N (provided by M. Gartenberg, University of
Medicine and Dentistry of New Jersey) contains the 2-mµ ARS,
URA3, and an excision cassette that consists of a 2480-bp
BglII-XhoI fragment from the yeast LYS2 gene flanked by direct repeats of a 58-bp
Zygosaccharomyces rouxii recombinase recognition sequence
(49). Co-transfection of pKWD50N and the recombinase expression vector
pHM153 (50) followed by induction of recombinase expression leads to
excision and recircularization of the excision cassette to form an
ARS-less plasmid that was used as a control in the topoisomer studies. Plasmids were transfected by the method of Ito et al. (51)
into S. cerevisiae strain TCK1 (W3031A;
bar1::HIS3) (provided by J. Kurjan, University of
Vermont), strains Y236 (MATa bar1 his6 ura3 trp1 leu2
cdc7-1), Y728 (Mata bar1 his6 trp1-289 leu2-3, 112 ura3-52 bob1-1 his31 lys2), and Y312 (MATa bar1 his6
ura3 trp1 leu2 can1 cdc8-1) (provided by R. Sclafani, University
of Colorado Health Science Center), and strain MVY19 (MATa lys2
ura3-52 his31 leu2-3, 112 cyh2
cdc71::HIS3 mcm5/cdc46-bob1 gal1
bar1::LEU2). MVY19 was derived from strain P211 (from R. Sclafani) by disruption of the BAR1 gene.
Genomic Footprinting--
The procedures used for genomic
footprinting were shown previously to faithfully reflect protein-DNA
interactions in cells (52-54), and the results for each cell strain or
cell cycle point examined in this paper are based on at least three
independent experiments. Briefly, cells were grown to early to mid-log
phase (1-2 × 107cells/ml) in synthetic complete
medium that lacked nutrients required for plasmid selection and
contained 25 mM phthalate (pH 5.5 with NH4OH).
Nuclei were isolated and treated with varying amounts of MNase, and DNA
was isolated. The overall extent of digestion of chromatin and naked
DNA control samples was estimated by agarose gel electrophoresis (not
shown). Samples digested to approximately equal extents were selected
for footprint analyses and cleaved with NdeI and
EcoRV. The fraction of template molecules that support primer extension to either the NdeI or EcoRV
sites provided a direct measure of the extent of nuclease cleavage
within the ARS1-DNA-containing region. Cleavage of templates with
NdeI and EcoRV also provided an internal control
for the specificity of the primers used for mapping cleavage sites.
Nuclease cleavage sites in plasmid chromatin and naked DNA control
samples were then mapped by polymerase chain reaction-mediated,
reiterative extension of end-labeled oligonucleotide primers
(5'-GCAATTTAACTGTGATAAACTACCGCATTAAAGCTTATCG-3',
5'-AAATGATGAATTGAATTGAAAAGCTGTGGCGGCCG-3', 5'-CGACAGCATCGCCAGTCACTATGGCGTGC-3', and
5'-GCGCATAGAAATTGCATCAACGCATATAGC-3'), as described (55). Genomic
footprints of ARS1 in chromosome IV were mapped using
chromosome-specific primers 5'-AAATGGCGTTATTGGTGTTGATGTAAGCGGAGGT-3' and 5'-TTGCGGTGAAATGGTAAAAGTCAACCCCCTGCGATG-3'. Extension
products were resolved by electrophoresis on sequencing gels and
visualized by autoradiography with Kodak X-Omat film. Cleavage sites
were mapped by comparison with sequence ladders generated in parallel polymerase chain reactions using the same end-labeled primers and
linearized plasmid DNA templates.
For cell cycle experiments, strains were grown at 23 °C to ~1 × 107 cells/ml and blocked in G1 phase by the
addition of -factor (Sigma) to a concentration of 20 nM.
After 3 h, unbudded cells were collected by centrifugation and
resuspended in fresh 37 °C media containing 0.1-1 mg/ml protease
(Sigma). After 1 h at 37 °C, at least 90% of cells had reached
their characteristic terminal phenotype, as judged by microscopy (56,
57) and were harvested for footprint analyses as above, except that
elevated temperatures were used to sustain the blocked state until the
moment of cell lysis.
KMnO4 footprinting was carried out essentially as described
by Park et al. (58). Briefly, 50-ml aliquots of yeast cells, grown as described above, were treated with 2-8 mM
KMnO4 (Sigma) for 1 min at 42 °C. The reaction was
terminated by the addition of an equal volume (50 ml) of ice-cold STE
buffer (100 mM NaCl, 10 mM Tris-HCl, pH 8.0, and 1 mM EDTA) containing freshly added 5 mM
dithiothreitol. The resulting mixture was chilled on ice, and cells
were pelleted at 3600 rpm in RT6000B centrifuge (Sorvall) at 4 °C
for 4-6 min, washed with 1 ml of 5 mM Tris-HCl, pH 8.0, and resuspended in 700 µl of cold 25× ETM (25 mM EDTA,
100 mM Tris-HCl, pH 8.0) containing freshly added
-mercaptoethanol (Sigma). Cells were lysed using glass beads, and
DNA was isolated and cleaved with EcoRV. Samples were then
treated with a final concentration of 0.1 M NaOH for 5 min
at room temperature and neutralized by the addition of HCl to a final
concentration of 0.1 M. Modification sites in chromatin and
naked DNA control samples were visualized and mapped by reiterative
extension of end-labeled primers, as described above.
Topoisomer Analyses--
Initial studies of pJL347 and TRP1ARS1
topoisomers were performed using DNA isolated from cells grown in
glucose-containing media and blocked at selected points in the cell
cycle, as described above. However, since galactose is required to
induce expression of the Z. rouxii recombinase (50), we also
examined pJL347 topoisomers isolated from cells that had been shifted
to galactose, as described below, and found that the different carbon
sources had no detectable effect on the topoisomer distributions (data
not shown). To examine the topology of plasmids lacking an ARS, cells
containing both pKWD50N and pHM153 were grown to mid-log phase at
23 °C in synthetic medium containing 2% raffinose. Cells were
blocked in G1 phase by the addition of -factor to 20 nM. After 2-3 h in -factor, recombinase expression was
induced by the addition of galactose to a final concentration of 2%.
After 1 h in the presence of both galactose and -factor,
-factor was removed, and half the culture was shifted to 37 °C,
whereas the other half was maintained at 23 °C. At appropriate
times, cells were lysed with glass beads in the presence of 10 mM N-ethylmaleimide (Sigma), and plasmid DNA was
isolated. Plasmid topoisomers were resolved by gel electrophoresis for
15 h at 2 V/cm in 0.9-1.4% agarose gels containing chloroquine diphosphate (Sigma). Chloroquine concentrations were varied in trial
experiments to ensure that amounts used were sufficient to induce a
positive writhe in the topoisomers to be examined (59). After transfer
to Protran nitrocellulose membranes (Schleicher & Schuell), the
excision cassette, pJL347, pKWD50N, and TRP1ARS1 topoisomers were
visualized by hybridization with 32P-labeled LYS2, pBR322,
and TRP1ARS1 probes, respectively. For each plasmid, individual
topoisomers were assigned an integral value, and the amount of each
topoisomer was quantified (and corrected for background) using a
Bio-Rad phosphoimager. The topoisomer amounts were summed, and the
number average distribution was calculated. The number average linking
number for plasmids isolated from -factor-blocked cells at 23 °C
was used as a reference value. Thus, the Lk reported in Table I for
plasmids in other cell types or at other points in the cell cycle
refers to the linking number change relative to the linking number in
-factor-blocked cells. Each Lk reported is based on the average
value of at least three separate experiments.
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RESULTS |
Alterations in the ARS1 Genomic Footprint during the Cell
Cycle--
As outlined in the Introduction, DNA replication requires
the assembly and activation of initiation complexes followed by DNA
unwinding by helicases, stabilization of unwound DNA by single-stranded DNA-binding proteins, and synthesis of an RNA primer by DNA primase. To
link these events to factors that control cell cycle progression, we
examined the chromatin structure of ARS1 in both its native location in
chromosome IV and in an episomal plasmid, pJL347, at specific points in
the cell cycle. Cells were blocked either in G1 phase with
the mating pheromone -factor or in S phase with hydroxyurea (HU). We
also blocked cells at selected points in the cell cycle using cell
strains containing temperature-sensitive mutations in either Cdc28p,
Cdc7p, or Cdc8p. To ensure maximal viability of mutants used in this
study and thereby avoid possible artifacts associated with cell death,
cells were first blocked in G1 phase with -factor and
then released to the restrictive temperature for just 1 h. The
viability of both cdc7-1 and cdc8-1 mutants
subjected to this regimen exceeded 85%. We also compared nuclease
cleavage patterns in wild type cells arrested with -factor at both
permissive and restrictive temperatures (23 °C and 37 °C,
respectively). No temperature-dependent differences were
evident (data not shown). Thus, any alterations in the chromatin
structure evident in cdc7-1 and cdc8-1-blocked
cells could be ascribed to loss of Cdc7p or Cdc8p function rather than
to exposure of cells to elevated temperatures before isolation of
nuclei. Finally, the cleavage pattern in wild type cells arrested in
-factor was also evident in cdc28 mutants blocked at
START and in cdc7-1 and cdc8-1 mutants arrested
in G1 phase with -factor (not shown). These observations
ruled out the possibility that structural changes in cdc7-1
and cdc8-1-blocked cells described below are due to defects
evident at earlier points in the cell cycle.
Earlier studies of ARS1 chromatin using DNase I as a probe indicated
that the characteristic pre-replicative complex footprint in
-factor-blocked cells is also evident in cdc7-1-blocked
cells (17), and this proved true for MNase as well (compare cleavages in lanes 5-7 with those in lanes 8-9 in Fig.
1A). Consequently, the
footprinting results for these two points in the cell cycle were used
as a reference point for later comparisons and are described together
here. As expected, comparison of MNase cleavages in nuclei (Fig.
1A, lanes 5-9) with those in naked DNA controls
(Fig. 1A, lane 4) revealed evidence of previously
documented protein-DNA interactions. On the A-rich strand, for example,
the B3 element is flanked by hypersensitive cleavage sites (Fig.
1A, filled circles, lanes 5-9),
which, along with the suppression of cleavages within the B3 element,
probably reflect the binding of Abf1p. On the same strand, two closely
spaced naked DNA cleavage sites within the B2 element also occur in
chromatin, but a third is suppressed; there is also a
chromatin-specific cleavage site within B2 (Fig. 1A,
filled circle, lanes 5-9). The
chromatin-specific cleavages in B2 DNA might reflect interactions with
Orc2p, as have been observed in vitro (60), or an altered
DNA conformation in vivo, reflecting the probable role of B2
as a DNA-unwinding element. Finally, we observed enhanced cleavage of
phosphodiester bonds between the B1 element and the ACS on the A-rich
strand (Fig. 1A, filled circles, lanes
5-9) as well as partial suppression of cleavages within the ACS
and B1 elements. In the T-rich strand, certain phosphodiester bonds
that are sensitive to cleavage in naked DNA are also protected in
nuclei, whereas two sites within the ACS and two others just 3' of the
B1 element show enhanced sensitivity to cleavage in nuclei (Fig.
2A, compare lanes
7-9 with lanes 5-6). Control experiments indicate
that mutations that reduce B1 function or abolish ACS function alter
the corresponding B1 and ACS cleavage patterns. For example, two
prominent chromatin-specific cleavage sites within and near the wild
type ACS (denoted by filled circles in Fig. 1A,
lanes 5-9) are protected in an ACS mutant (open
circles, Fig. 1B), suggesting that the cleavage
patterns observed in the B1 and ACS region of the wild type ARS1
reflect the binding of ORC.
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Fig. 1.
Genomic footprinting of the A-rich strand of
ARS1 at selected points in the cell cycle. A, nuclei
containing pJL347 were isolated from wild type cells blocked in
G1 phase with -factor or in S phase with HU, from
cdc7-1 and cdc8-1 mutants blocked with -factor
and then released to the restrictive temperature, and from
cdc7 mcm5-bob1 mutants blocked with
-factor. Isolated nuclei were treated with varying amounts of MNase,
and DNA was isolated. Single-strand cleavage sites in the A-rich strand
in chromatin and in naked DNA controls (lane N) were mapped
by polymerase chain reaction-mediated primer extension, as described
under "Experimental Procedures." Cleavage sites were mapped using
sequencing ladders (lane T). Primer extension reactions
performed with intact and mock-digested DNA templates (lanes
N0 and C0) showed only minimal pausing by
Taq polymerase. In the diagram to the right of each
panel, open rectangles refer to the ACS, B1, B2,
and B3 elements in ARS1. Filled circles mark a subset of the
phosphodiester bonds that are hypersensitive to cleavage in nuclei.
Stars mark chromatin-specific cleavage sites in
cdc8-1- and HU-blocked cells and in -factor-blocked
cdc7 mcm5-bob1 cells that differ from those in
cdc7-1- and -factor-blocked cells. B, cleavage
sites in a version of ARS1 rendered non-functional by a G to T mutation
in the ACS (asterisk) were mapped by genomic footprinting and compared to cleavage sites in naked DNA controls
(lane N1). Filled circles mark four cleavage
sites identical to those marked in Fig. 1A; open
circles mark two cleavage sites that are prominent in the wild
type ARS1 but absent in the acs mutant. C, cells
were collected at selected points in the cell cycle and prepared for
FACS analysis as described (83).
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Fig. 2.
Genomic footprinting of the T-rich strand of
ARS1 at selected points in the cell cycle. A,
single-strand MNase cleavage sites in the T-rich strand of ARS1 in
pJL347 were mapped as described in Fig. 1. In the diagram to the right
of each panel, open rectangles refer to the ACS,
B1, and B2 elements. Filled circles mark a subset of
cleavage sites that are hypersensitive in nuclei as compared with naked
DNA. Stars indicate DNA cleavage sites that displayed
increased MNase sensitivity in cdc8-1-arrested cells,
HU-blocked cells, and -factor-blocked cdc7
mcm5-bob1 cells as compared with that observed in
cdc7-1-arrested cells (nuclease sensitivity was estimated by
comparison with a site at the edge of the B2 element, marked with a
filled circle: cf. "Results"). A
vertical bar marks a segment of DNA that also is cleaved
differently at different points in the cell cycle. B, MNase
cleavage sites of the T-rich strand in the chromosomal copy of ARS1
were mapped as described above. Filled circles and
stars mark the same cleavage sites marked in
A.
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We next compared MNase cleavages of both the A-rich and T-rich strands
of ARS1 in cdc8-1-blocked cells with those in
cdc7-1-blocked cells. The cleavage pattern on the A-rich
strand changed only slightly as cells progressed from the
CDC7 to the CDC8 execution point (Fig.
1A, compare lanes 8-9 with lanes
10-11). These changes included the suppression of cleavage at a
site within the ACS and enhanced cleavage at a site near the edge of B1
(denoted by stars in Fig. 1A, lanes
10-11). On the T-rich strand of ARS1, the frequency of cleavage
at sites within the B2 element on the T-rich strand (indicated by
stars adjacent to lanes 12, 15, and 17 in Fig. 2A) increased relative to cleavage at
a site at the edge of the B2 element (indicated by a filled
circle adjacent to lanes 9, 12,
15, and 17 in Fig. 2A) as cells
progressed from the CDC7 to the CDC8 execution
point. Modest changes in the pattern and relative intensity of
cleavages flanking the B1 element also were evident (compare bands
adjacent to the vertical bar in lanes 7-9 with
those in lanes 10-12 in Fig. 2A). Together,
these changes suggested an alteration either in protein-DNA contacts or
in DNA structure, particularly at the B2 element, following the action of Cdc7p.
To directly relate the footprinting results to the DNA topology
analyses described below, most of the footprint analyses in this paper
were done using low copy number plasmids. However, differences similar
to those described above for the episomal ARS1 also occurred at ARS1 in
its native locus in chromosome IV (Fig. 2B). This result
strongly suggests that the alterations in the ARS1 footprints following
the CDC7 execution point reflect bona fide
structural changes associated with the initiation of replication.
Although previous studies have also documented changes in ARS1
chromatin following the CDC7 execution point, those studies typically compared cleavage patterns seen in cdc7-1-blocked
cells with those in HU-blocked cells (e.g. 17). The
transition from the CDC8 execution point to the HU-sensitive
step led to a pronounced asymmetry between the A- and T-rich strands of
ARS1 in their sensitivity to MNase. This can be seen by comparing the
intensity of "parental" bands at the tops of lane 13 in
Fig. 1A and lane 15 in Fig. 2A (parental bands reflect the fraction of template molecules that have
escaped MNase cleavage within the region of interest and can thus be
used to estimate local nuclease cleavage rates. The same DNA sample was
used in these two lanes to visualize cleavage sites in, respectively,
the A- and T-rich strands of ARS1, yet at this point in digestion,
virtually all of the T-rich strand molecules have been cleaved, whereas
a large fraction of the A-rich strands remain intact. This asymmetry in
MNase sensitivity can be attributed to an increase in sensitivity of
the T-rich strand (as opposed to a decrease in sensitivity of the
A-rich strand) as cells progress from the CDC8 execution
point to the HU-sensitive step. Specifically, the amounts of nuclease
used to produce the samples for lanes 13, 14, and
15 in Fig. 2A (50, 100, and 200 units × min/ml, respectively) were equal to the amounts used for samples in
lanes 10, 11, and 12 in Fig.
2A. Despite this, the amount of undigested template in
lanes 14 and 15 is severalfold lower than in
lanes 11 and 12. There also were more specific
differences in the cleavage patterns in cdc8-1 and
HU-blocked cells, particularly in the T-rich strand of ARS1. For
example, nuclease-sensitive bonds in the B2 element in
cdc8-1-blocked cells appeared even more sensitive in
HU-blocked cells, and additional cleavage sites became evident as well
(cf. starred bands in Fig. 2A,
lanes 13-15). Additional cleavage sites near the ACS and B1
elements also appeared. Thus, the cleavage pattern observed for
cdc8-1 blocked cells not only differs from that in
-factor and cdc7-1-arrested cells but also from that in
HU-arrested cells.
Evidence That DNA Unwinding Occurs after the Cdc7p Execution
Point--
Fig. 3 summarizes the
previously identified DNA synthesis start sites (2, 3) as well as sites
sensitive to DNase I (12, 17), MNase (this study), and
KMnO4 (this study, see below). Interestingly, the major
changes in the cleavage pattern of ARS1 DNA after the CDC7
execution point involve the T-rich bottom strand, whereas only minor
differences are evident in the A-rich top strand (Fig. 1A
and 2A). One possible explanation for this asymmetric change
in cleavage pattern is suggested by the finding that the T-rich bottom
strand in this segment of ARS1 serves as a template for lagging strand
synthesis, whereas the A-rich top strand serves as a template for
leading strand synthesis (2, 3). At an early step in replication, the
lagging strand template may have more single-strand character than the
leading strand, and unlike DNase I, MNase cleaves single-strand DNA
efficiently (61). This line of reasoning suggested that Cdc7p might
trigger local DNA unwinding at ARSs.
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Fig. 3.
Summary of nuclease-sensitive and DNA
synthesis start sites in ARS1. Open rectangles refer to
the ACS, B1, B2, and B3 elements. Open arrowheads mark DNA
synthesis start sites in the chromosomal ARS1, and filled
arrowheads mark DNA synthesis start sites in episomes containing
ARS1 (2, 3). Open diamonds mark DNase I hypersensitive sites
(12, 17). Chromatin-specific MNase hypersensitive sites marked in Fig.
1 are indicated by filled circles. Stars denote
cleavage sites that displayed increased cleavage frequency in
cdc8-1-arrested cells, HU-blocked cells, and
-factor-blocked cdc7 mcm5-bob1 cells as
compared with that observed in cdc7-1-arrested cells
(cf. Fig. 2 legend). The asterisk denotes the
KMnO4-modified site evident in the ACS of
cdc8-1- and HU-blocked cells, in -factor-blocked
cdc7 mcm5-bob1 cells, and in
-factor-blocked mcm5-bob1 cells (Fig. 4).
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To directly test whether DNA unwinding occurs after the CDC7
execution point, we used KMnO4 to probe for single-stranded
regions within and near ARS1 at selected points in the cell cycle.
KMnO4 preferentially modifies unpaired thymidine (T)
residues and, at higher concentrations, unpaired adenine (A) residues.
Certain distortions in double-stranded DNA can also lead to enhanced
KMnO4 sensitivity (62, 63). In either case, modification
sites can be mapped by treating modified residues with NaOH, which
creates a lesion that cannot be bypassed by Taq DNA
polymerase (64). In exponentially growing, wild type cells, ARS1 DNA
was only minimally reactive to KMnO4 (data not shown). This
was as expected, since only a small fraction of ARS1 molecules in an
asynchronous population of cells is undergoing replication during the
1-min exposure of cells to KMnO4. Likewise, in cells
blocked with -factor in G1 phase and in
cdc7-1 mutants blocked at the CDC7 execution
point, ARS1 DNA displayed little or no KMnO4 sensitivity
(Fig. 4A, lanes 5-9). By contrast, thymidine residues within and flanking ARS1 DNA in HU-blocked wild type cells showed an increased level of KMnO4 sensitivity, most likely due to DNA unwinding (Fig.
4A, lanes 13-14). Sequences within and near ARS1
in cdc8-1-blocked cells were similarly
KMnO4-sensitive (Fig. 4A, lanes
10-12). Modified residues in both cdc8-1- and
HU-blocked cells included a thymidine that lies within the ORC binding
site on the T-rich strand (Fig. 4A, asterisk).
KMnO4 modification of thymidine residues outside ARS1 in
cdc8-1- and HU-blocked cells suggest fairly extensive DNA
unwinding at both of these points in the cell cycle. The
KMnO4 modification pattern outside ARS1 in
cdc8-1-blocked cells differed somewhat from that in
HU-blocked cells (compare lanes 10-12 with 13-14, Fig. 4A), and we were able to generate
the HU pattern by returning cdc8-1-blocked cells to the
permissive temperature in the presence of HU (data not shown). This
result indicated that the cdc8-1 block is fully reversible
and occurs before the HU-sensitive step in DNA replication.
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Fig. 4.
Genomic footprinting of ARS1 with
KMnO4 indicates that cdc8-1-blocked cells
contain single-stranded DNA. A, cells harboring
pJL347 were treated with varying amounts of KMnO4 at
selected stages of the cell cycle. DNA was isolated by glass bead
lysis, and single-strand modification sites in the T-rich strand were
mapped by polymerase chain reaction-mediated primer extension, as
described under "Experimental Procedures." Extension reactions
performed with intact DNA (lane C0) show only minimal
pausing by Taq DNA polymerase. Lane M contains
molecular size markers. Lanes A and T denote A-
and T-specific sequencing ladders. An asterisk marks a
thymidine residue in the ACS that is modified in cdc8-1- and
HU-blocked cells, in -factor-blocked cdc7
mcm5-bob1 cells, and in -factor-blocked
mcm5-bob1 cells but not in either -factor or
cdc7-1-blocked cells. B, KMnO4
modification sites within and near a version of ARS1 rendered
non-functional by a G to T mutation in the ACS (asterisk)
were mapped by genomic footprinting. Filled circles mark
thymidine-specific modification sites in HU-blocked cells that reside
in sequences near a neighboring, functional ARS.
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To determine if the enhanced sensitivity of ARS1 DNA to
KMnO4 following the CDC7 execution point depends
on a functional ARS, we conducted parallel studies with a version of
ARS1 rendered non-functional by a point mutation in the ACS (Fig.
1B) (65). In all cell cycle stages tested, including
cdc8-1- and HU-arrested cells, this mutation abolished the
ARS-specific sensitivity to KMnO4. Sequences 3' of the
non-functional ARS still exhibited moderate KMnO4
sensitivity in HU-arrested cells (Fig. 4B, lanes 12-13), most likely due to a stalled replication intermediate initiated from a nearby, functional ARS. Together, these observations strongly suggest that DNA unwinding at replication origins is CDC7-dependent and requires a functional
ARS.
Analysis of DNA Topoisomers at Selected Points in the Cell
Cycle--
Unwinding of circular DNA produces compensatory positive
superhelical stress. This stress may be relieved by the action of DNA
topoisomerases, resulting in a change in linking number or Lk (Ref.
66 and references therein). However, particularly if the DNA unwinding
in cdc8-1-blocked cells is localized and limited in extent,
the resulting superhelical stress might be relieved through changes in
local protein-DNA interactions, with little or no Lk. Loss of a
single nucleosome, for example, would release enough negative
superhelical density to compensate for the unwinding of ~10 bp DNA
(52, 67-69). To determine if the CDC7-dependent structural
changes documented above are accompanied by a linking number change, we
examined the distribution of pJL347 topoisomers at selected stages in
the cell cycle (Fig. 5 and Table I). DNA topoisomers were resolved by gel
electrophoresis in buffer containing chloroquine diphosphate such that
the least negatively supercoiled topoisomers migrated most rapidly. We
first examined the topoisomer distribution in wild type cells and in
cdc7-1 and cdc8-1 mutants blocked with -factor
at both 23 °C and 37 °C. Cells were monitored to ensure that the
block was efficient at both temperatures. The DNA linking number
distribution in cdc7-1 and cdc8-1 mutants blocked
with -factor was virtually identical to that in -factor blocked
wild type cells. This result is in accord with our footprinting
results, which suggested that mutations in CDC7 and
CDC8 have no effect on replication initiation complexes prior to START. To block cells at the CDC7 and
CDC8 execution points, it was necessary to shift mutant
cells from 23 °C to 37 °C. Because DNA untwists with increasing
temperature (70), DNA isolated from cells incubated at 37 °C will be
more negatively supercoiled than DNA isolated from cells grown at
23 °C. The magnitude of this change for DNA in yeast chromatin is
approximately 
0.008 degree twist/°C/bp (71). Therefore, the
predicted in vivo Lk for the 6027-bp pJL347 following a
shift from 23 °C to 37 °C is 0.008 × (37 °C 23 °C = 14 °C) × 6027/360° = 1.80 turns. The measured Lk associated with a shift from 23 °C to 37 °C in
-factor-blocked cells was 1.68, 1.69, and 1.81 for wild type
cells, cdc7-1 mutants, and cdc8-1 mutants,
respectively (Table I). These results indicated that cdc7-1
and cdc8-1 mutants respond to temperature-induced superhelical stress in the same fashion as wild type cells. Thus, any
Lk that accompanied the transition from START to the CDC7 and CDC8 execution points could be attributed to structural
changes that occur at those points in the cell cycle.
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Fig. 5.
ARS-dependent changes in linking
number at selected points in the cell cycle. Wild type
(WT) cells containing either pJL347 or a non-replicating
ARS-less plasmid control (Excision cassette; see
"Experimental Procedures") were blocked either in G1
phase with -factor, in S phase with HU, or in G2/M with
nocodazole, and DNA was isolated by glass bead lysis. DNA also was
isolated (at the restrictive temperature) from cells blocked at the
CDC7 and CDC8 execution points. The topoisomerase
inhibitor N-ethylmaleimide was included in all cases to
inhibit changes in DNA topology during the isolation. Topoisomers were
resolved by electrophoresis in chloroquine-containing buffers such that
less negatively supercoiled topoisomers migrated most quickly and were
visualized by Southern blotting. After electrophoresis, DNA was
subjected to partial depurination and cleavage to ensure uniform
transfer of topoisomers. Topoisomers were quantified by
phosphorimagery, and the number average distribution was calculated as
described under "Experimental Procedures" (cf. Table
I).
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Table I
Shift in average linking number with changes in the cell cycle
Linking number changes evident in cdc7-1-blocked cells were
calculated relative to the average linking number for all three cell
types (i.e. W303-1A and the cdc7-1 and
cdc8-1 mutants) in -factor-blocked cells after
incubation at 37 °C; this eliminated the need for temperature
corrections. The linking number change for the transition from the
Cdc8p step to the HU-dependent step is equal to the
(Lk-HU Lk-cdc8-1] + [Lk · 37 °C Lk · 23 °C]. For pIL347, this is [4.68 (1.11)] + [1.88 (0.15)] = 4.06. WT, wild type; ND, not determined;
NA, not applicable.
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Between START and the Cdc7p execution point, there was a small but
statistically significant decrease in negative superhelicity of
approximately +1.6 (Fig. 5A and Table I). KMNO4
footprint analyses (Fig. 4A) showed no evidence of DNA
unwinding during this interval and indeed, had unwinding occurred, we
might expect the Lk to remain the same or become more negative. If the
observed Lk were due to pre-replicative events at ARS1, the Lk
for different sized plasmids might be similar. To test this prediction,
we measured Lk values at the same points in the cell cycle for
TRP1ARS1, a multicopy plasmid that is just one-quarter the size of
pJL347, and for pKWD50N, a larger plasmid (8278 bp) used in studies to be described below. The TRP1ARS1 and pKWD50N topoisomer distributions both changed between the -factor and Cdc7p steps in the same manner
as for pJL347, but the change was proportional to plasmid size (Table
I). This result suggested that the observed Lk might be the
consequence of cell cycle-specific alterations in chromatin structure
rather than structural changes associated specifically with ARS DNA.
The transition from the CDC7 to the CDC8
execution point was accompanied by a small, negative Lk for pJL347
(Fig. 5A, Table I). This result coupled with evidence of DNA
unwinding from the KMNO4 studies suggested that the helical
stress associated with initial DNA unwinding is either largely
inaccessible to topoisomerase or balanced by changes in protein-DNA
architecture. By contrast, there was a substantial Lk of +4.1 during
the transition between the cdc8-1-and HU-sensitive steps,
(Fig. 5, lane 7, Table I). The change in DNA topology
evident in HU-blocked cells was also seen in cdc7-1 and
cdc8-1 mutants that first were released from an -factor
block and shifted to the restrictive temperature for 1 h and then
returned to 23 °C in the presence of HU (data not shown). These
results further demonstrated that the cdc7-1- and cdc8-1-blocked states are fully reversible. The increased
linking number in HU-blocked cells was surprising since the predicted further DNA unwinding would be expected to lead to a reduction in
linking number (66). Events that may underlie this Lk are addressed
under "Discussion." Here, we note only that the substantial Lk
associated with the transition from the CDC8 execution point to the HU-sensitive step clearly demonstrates that our methods for
isolating DNA are capable of capturing changes in DNA topology and that
the post-replicative complex defined by the HU block differs from the
cdc8-1-blocked state.
Changes in DNA Topology in Late G1 and Early S Phase
Are ARS-dependent--
It was possible that the observed
cell cycle-specific changes in DNA topology are unrelated to DNA
replication. To address this possibility, we transfected cells with a
plasmid encoding the Z. rouxii R site-specific recombinase
under control of the GAL promoter (50) together with a
second plasmid containing recombinase target sites. Expression of the
recombinase during G1 phase results in the formation of a
circular, ARS-less plasmid (Ref. 49 and "Experimental Procedures").
cdc7-1 and cdc8-1 mutants and wild type cells
containing these plasmids were grown to mid-log phase and blocked in
G1 phase by the addition of -factor. Recombinase expression was then induced for 1 h in the presence of -factor, after which -factor was removed, and cells were either shifted to
the restrictive temperature or blocked by addition of HU. Topological analyses of ARS-less circular DNA revealed no changes in linking number
between the -factor-, Cdc7p-, Cdc8p-, and HU-sensitive steps (Fig.
5B, Table I). These results indicate that the topological changes we observed in ARS-containing plasmids are in fact
ARS-dependent.
The Structural Attributes in ARS1 DNA Evident in cdc8-1-blocked
Cells Are CDC7-dependent--
As described in the
Introduction, Cdc7p appears to act directly on origin complexes to
trigger the initiation of DNA replication. Thus, we wanted to determine
if the structural and topological changes evident in
cdc8-1-blocked cells were due to the direct action of Cdc7p.
Because CDC7 is essential, we took advantage of a strain
that can bypass the requirement for Cdc7p by virtue of a mutation in
MCM5 known as mcm5/cdc46-bob1 (25, 44). We reasoned that if Cdc7p triggers a structural change by modifying a
component of the pre-initiation complex, that change may be evident
earlier in the cell cycle in a cdc7 mcm5/cdc46-bob1
bar1 mutant (strain MVY19). To test this prediction, mid-log phase MVY19 cells were arrested in G1 phase with -factor at
both 23 °C (Figs. 1A and 2A) and 37 °C (not
shown), and ARS1 chromatin structure was examined using genomic
footprinting and DNA topology assays. Even though FACS analyses show no
indication of premature replication in the -factor-arrested
cdc7 mcm5/cdc46-bob1 cells (Fig.
1C; Refs. 33 and 44), the MNase-derived cleavage pattern for
ARS1 was distinct from that evident in -factor-blocked wild type
cells and in cdc7-1-blocked cells and strikingly similar to
the pattern seen in cdc8-1-blocked cells (Fig.
1A, compare lanes 14-16 with 10-11
and Fig. 2A, compare lanes 16-17 with
10-12). Thus, the cleavage pattern normally evident only in
early S phase is already present in G1 phase
cdc7mcm5/cdc46-bob1 cells.
As with ARS1 DNA in cdc8-1- and HU-blocked cells, ARS1 DNA
in -factor-arrested, cdc7 mcm5/cdc46-bob1
cells was also sensitive to KMnO4 (Fig. 4A,
compare lanes 15-16 with lanes 5-6). In
particular, the same thymidine residue within the ACS that is sensitive
to KMnO4 in cdc8-1-and HU-blocked cells but not
in -factor-blocked wild type cells or in cdc7-1-blocked
cells was KMnO4-sensitive in -factor-blocked
cdc7 mcm5/cdc46-bob1 cells (Fig.
4A, asterisk). It was important as well to ensure
that the premature KMnO4 sensitivity of ARS1 and
surrounding sequences in MVY19 cells was due to the mcm5/cdc46-bob1 mutation and not the absence of Cdc7p. We
therefore treated -factor-blocked mcm5/cdc46-bob1 cells
with KMnO4 and observed the same pattern of modification
within and surrounding ARS1 as before (Fig. 4A). As before,
the KMnO4 sensitivity was dependent on a functional ARS
(Fig. 4B). These results suggested that the increased
sensitivity to KMnO4 modification reflects a change in the
structure of the MCM complex that ordinarily is induced by Cdc7p.
Because the MNase and KMnO4 profiles in cdc7
mcm5/cdc46-bob1 cells arrested in G1 phase with
-factor were similar to those in cdc8-1-blocked cells, we
predicted that cdc7 mcm5/cdc46-bob1 mutants
might also exhibit premature changes in DNA topology. Although it must
be emphasized that the magnitude of the predicted Lk was small,
pJL347 DNA isolated from cdc7 mcm5/cdc46-bob1 cells arrested with -factor had a linking number similar to that seen in cdc8-1-blocked cells (Table I). pJL347 DNA isolated
from cdc7 mcm5/cdc46-bob1 cells that were
released from -factor and subsequently arrested with HU showed a
change in linking number comparable with that in wild type,
cdc7-1, and cdc8-1 mutant cells arrested with HU
(Table I). Taken together, these results argue that both the DNA
structural and topological changes evident in cdc8-1-blocked
cells are due to the action of the Cdc7p protein kinase at replication origins.
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DISCUSSION |
Structure of Replication Complexes in cdc8-1-blocked Cells--
In
this paper, we have investigated structural events that occur at ARS1
at discrete points in the cell cycle. Following the CDC7
execution point, we observed increased MNase sensitivity of
phosphodiester bonds within the probable DNA-unwinding element (B2) and
increased sensitivity of ARS1 and surrounding DNA to modification by
permanganate (Figs. 1, 2, and 4). Given that permanganate preferentially modifies unpaired thymidine residues and that MNase efficiently cleaves single-strand DNA, our results strongly suggest that ARS1 DNA unwinds following the action of Cdc7p. This inference is
consistent with cross-linking experiments that indicate that replication protein A, the eukaryotic replicative single-strand DNA-binding protein, becomes associated with ARSs soon after the CDC7 execution point (20). As cells progress from the
CDC8 execution point to the HU-sensitive step, ARS1 DNA
becomes substantially more MNase-sensitive, due principally to an
increased rate of cleavage of the presumptive template for lagging
strand synthesis (2, 3). These results might reflect arrest before the
onset of lagging strand synthesis or at a point during replication when the lagging strand has more single-strand character than does the
leading strand. Alternatively, the asymmetric nuclease sensitivity might reflect an asymmetric distribution of initiation factors or
replicative enzymes.
The pattern of nuclease sensitivity and distribution of topoisomers in
cdc8-1-blocked cells differs from that in both
cdc7-1- and HU-blocked cells. Additionally, the topoisomer
distribution and nuclease sensitivity characteristic of HU-blocked
cells becomes evident after releasing cdc8-1-blocked cells
into HU-containing media. These observations suggest that the
CDC8-dependent step in replication precedes the
HU-sensitive step, an inference consistent with the temporal ordering
of the CDC8 execution point by Hartwell (48). It is unclear,
however, if the ARS1 structure in cdc8-1-arrested cells
reflects a distinct, previously uncharacterized replication intermediate. Specifically, the cdc8-1-mediated arrest
appears stringent, whereas HU-arrested cells eventually overcome the HU block (i.e. adapt) and resume replication. The observed
structural differences between HU- and cdc8-1-arrested cells
might therefore reflect more extensive DNA synthesis in HU-treated cells.
The known function of Cdc8p is to supply cells with dTTP. Thus, it is
not immediately clear why cdc8-1 mutants would arrest at
what appears to be a very early step in replication. One possible explanation is that Cdc8p is also important for the structural integrity of the multienzyme replication apparatus. Such a role could
help explain the "quick stop" replication arrest that
cdc8-1 mutants display when shifted to the restrictive
temperature (46) as well as the limited Lk that accompanies the
transition from the CDC7 to the CDC8 execution
point (Fig. 5 and Table I). Although there do not appear to be any
genetic data that support the hypothesis that Cdc8p contributes to the
structural integrity of the multiprotein replication complex
(cf. Ref. 9), several other observations are at least
consistent with this idea. For example, DNA synthesis in permeabilized
cdc8-1 mutants halts when they are shifted to the
restrictive temperature, despite the presence of exogenous dTTP and
other dNTPs (72, 73). Additionally, a multienzyme complex of DNA
precursor-synthesizing enzymes, which includes thymidylate kinase,
forms in both T4 phage and Chinese hamster embryo model replication
systems and appears to directly participate in DNA replication (74,
75). Finally, kinases responsible for the sequential phosphorylation of
thymidine to dTTP appear to be localized at discrete sites in nuclei
(76).
Role of Cdc7p in Replication Initiation--
The permanganate
sensitivity of ARS1 and surrounding sequences that we observed in
cdc8-1- and HU-blocked cells was also evident in
mcm5/cdc46-bob1 and cdc7
mcm5/cdc46-bob1 mutants arrested in G1 phase
with -factor. The fact that a replication intermediate that normally
forms in early S phase is also evident in G1 phase in an
mcm mutant that bypasses the requirement for Cdc7p strongly suggests that the action of Cdc7p leads to structural changes in the
MCM complex. The Cdc7p·Dbf4p complex physically associates with
individual replication initiation complexes and acts throughout S phase
to trigger the firing of individual origins. Cdc7p-Dbf4p modifies
Mcm2p, -3p, -4p, -6p, -7p in vitro (33, 36), and a specific
Cdc7p bypass mutation, mcm5/cdc46-bob1, resides in Mcm5p
(44). This strongly suggests that one target of Cdc7p-Dbf4p in
vivo is the MCM2-7 complex. Possibly, phosphorylation of Mcm proteins and the bob1 lesion in Mcm5p both alter the MCM2-7
complex in the same manner. This alteration might then activate the MCM helicase (if in fact an MCM subcomplex serves as the replicative helicase). Although this scenario is speculative, it could account for
the apparent unwinding of ARS DNA in both -factor-blocked mcm5/cdc46-bob1 mutants and cdc8-1 cells blocked
in early S phase. Despite what appears to be premature DNA unwinding in
these mutants, there is no evidence of premature DNA synthesis (Fig.
1C; Refs. 33 and 44). Thus, although the MCM2-7 complex in
mcm5/cdc46-bob1 mutants may exist in a replication-competent
state, this is not sufficient to trigger DNA replication. As noted in
the Introduction, DNA replication also requires Cdc28-Clb kinase, an
activity that is absent in early G1 phase and in
-factor-arrested cells.
Changes in DNA Topology during the Initiation of DNA Replication in
Yeast--
Our study has revealed small, ARS-dependent
changes in DNA topology between START and the G1/S phase
boundary. To interpret these results, it is useful to consider how
events associated with replication affect DNA topology. Replication of
SV40, for example, begins with the binding of the SV40 T-antigen, which serves as a replicative helicase. Upon binding, T-antigen reduces the
DNA twist independent of its helicase activity (77). If the helicase
used during chromosomal replication resides in an MCM subcomplex (18,
21), association of the MCM complex with Cdc6p-ORC-ARS complexes early
in G1 phase might also alter the local DNA twist but before
rather than after START. The Lk that occurs later, between START and
the G1/S phase boundary, varied with plasmid size (Table
I), suggesting that it is the result of a change in chromatin structure
rather than a change in the pre-replicative complex. The observed Lk
corresponds to an average decrease in negative superhelical density of
approximately +0.003 or a Lk of roughly +0.04 to +0.05 per
nucleosome. Thus, the Lk could be due entirely to changes in histone
acetylation, which have been found to alter the Lk associated with
nucleosome formation by up to ~0.2 (78). These calculations and the
fact that the observed Lks during late G1 and the
following S phase are ARS-dependent lead us to speculate
that replication is preceded by ARS-directed changes in chromatin
structure that facilitate the subsequent passage of replication forks
through chromatin templates.
There was virtually no Lk associated with the transition from the
CDC7 to the CDC8 execution point despite the
apparent DNA unwinding that occurs during this interval. This result
suggests that the positive superhelical stress resulting from initial
DNA unwinding is either balanced by changes in chromatin structure, as
described above, or inaccessible to topoisomerases. In this latter
case, positive superhelical stress may "diffuse" into the unwound
region (66); right-handed wrapping of the presumptive DNA template
strands on a "virtual" surface within this region might even
facilitate interactions between enzyme complexes that form on leading
and lagging strands (cf. Refs. 79 and 80).
The transition from the CDC8 execution point to the
HU-sensitive step was accompanied by a Lk of +4.1 in pJL347 (Table
I). This result was surprising since the linking number of replicating plasmid DNA must eventually go to zero (cf. Ref. 66). There are two possible explanations for this result. First, release of cells
from an -factor block directly into HU-containing media, as was done
in our experiments, may activate a checkpoint that inhibits ARS1 firing
or permits only limited DNA synthesis from ARS1. In this case, the
positive Lk would most likely reflect pre-replicative chromatin
remodeling events. This explanation would seem to be ruled out by the
finding that other early-firing ARSs support synthesis of long nascent
DNA strands in the presence of HU (81), although such synthesis might
have occurred in only a small fraction of cells or from early firing
ARSs that escaped checkpoint inhibition. A second explanation for the
positive Lk observed in HU-blocked cells is that the plasmids used
in this study are fully replicated during the 1-h HU treatment but that chromatin maturation is incomplete. The packaging of newly replicated DNA into nucleosomes not only requires specific acetylated forms of
histone H3 and H4 but also assembly factors that couple deposition of
H3-H4 tetramers to DNA replication (Ref. 82 and references therein). HU
treatment might slow the final steps in nucleosome assembly and thereby
delay restoration of a linking number close to that in
-factor-blocked cells. The available data do not distinguish between
these two explanations, but each explanation makes specific, testable predictions.
§,
, and
TOP
ABSTRACT
INTRODUCTION
