Blackwell Science, LtdOxford, UKMMIMolecular Microbiology0950-382XBlackwell Science, 200246Original ArticleSupercoiling, knotting and replicationL. Olavarrieta et al.
Molecular Microbiology (2002) 46(3), 699–707
Knotting dynamics during DNA replication
L. Olavarrieta, M. L. Martínez-Robles, P. Hernández,
D. B. Krimer and J. B. Schvartzman*
Departamento de Biología Celular y del Desarrollo,
Centro de Investigaciones Biológicas (CSIC), Velázquez
144, 28006 Madrid, Spain.
Summary
The topology of plasmid DNA changes continuously
as replication progresses. But the dynamics of the
process remains to be fully understood. Knotted bubbles form when topo IV knots the daughter duplexes
behind the fork in response to their degree of intertwining. Here, we show that knotted bubbles can form
during unimpaired DNA replication, but they become
more evident in partially replicated intermediates containing a stalled fork. To learn more about the dynamics of knot formation as replication advances, we
used two-dimensional agarose gel electrophoresis to
identify knotted bubbles in partially replicated molecules in which the replication fork stalled at different
stages of the process. The number and complexity of
knotted bubbles rose as a function of bubble size,
suggesting that knotting is affected by both precatenane density and bubble size.
Introduction
Segregation of the products of plasmid DNA replication is
a rather complex topological problem. Nature designed a
special kind of enzyme, DNA topoisomerases, specifically
to deal with this problem. As replication proceeds, unwinding of the parental strands by DNA helicase leads to the
formation of (+) DLk ahead of the fork, which must be
removed for the fork to keep progressing (Kornberg and
Baker, 1992). This (+) DLk diffuses across the replication
fork so that it is redistributed both ahead of and behind
the fork (Champoux and Been, 1980; Peter et al., 1998).
In bacteria, two specific type II DNA topoisomerases act
in different ways and at different places to reduce (+) DLk
during replication: DNA gyrase introduces (-) DLk ahead
of the fork; and topoisomerase IV (topo IV) removes pre-
Accepted 15 August, 2002. *For correspondence. E-mail
[email protected]; Tel. (+34) 91 564 4562 ext. 4233, (+34) 91
561 1800 ext. 4233; Fax (+34) 91 564 8749.
© 2002 Blackwell Publishing Ltd
catenanes in the replicated region (Ullsperger et al., 1995;
Peter et al., 1998; Alexandrov et al., 1999; Lucas et al.,
2001). Negative supercoiling assists any process that
requires double helix opening (Kanaar and Cozzarelli,
1992). Moreover (+) DLk can lead to replication fork reversal through the formation of Holliday-like junctions, at least
in vitro (Postow et al., 2001; Olavarrieta et al., 2002).
These observations suggest that plasmids should remain
negatively supercoiled throughout the process for replication to go on, but there is no experimental evidence indicative of the latter. As DLk distributes both ahead of and
behind the fork (Champoux and Been, 1980; Peter et al.,
1998), and the replicated portion enlarges as replication
advances, this (-) DLk would switch progressively from
(-) supercoils to negatively twisted precatenanes. In other
words, the number of precatenanes is expected to rise as
a function of bubble size. But measuring the number of
precatenanes in vivo as replication advances is not an
easy task. However, there are ways of inferring changes
in the number of precatenanes as replication progresses.
Knotted bubbles form when a type II topoisomerase
crosses two successive precatenanes (Postow et al.,
1999; Sogo et al., 1999). For this reason, they reflect the
number and pattern of DNA crossings trapped between
the two segments that participated in the strand passage
event. As knotted bubbles occur in the replicated portion
of partially replicated plasmids, the two segments involved
are the two daughter duplexes. Plasmids containing a
stalled fork, however, might have an excess of (-) DLk,
because of a possible residual action of DNA gyrase when
DNA helicase has already stopped (Olavarrieta et al.,
2002). This is unlikely, however, at least in the case of
plasmids containing forks stalled at a Ter–TUS complex.
In this case, the newly replicated nascent leading strands
go as far as 3–5 bp behind the TUS binding site (Mohanty
et al., 1998), leaving no physical space for DNA helicase
and DNA gyrase to remain bound to the unreplicated
parental duplex, at least if DNA helicase and gyrase are
physically linked and act in a co-ordinate manner (Duguet,
1997). Interestingly, in partially replicated ColE1 plasmids
containing a stalled fork, most of the nodes of knotted
bubbles have a positive sign (Sogo et al., 1999). Knots
generated by type II DNA topoisomerases reflect not only
the number but also the pattern of DNA crossings trapped
within a single topological domain. For this reason, positive supercoiling leads to DNA knots with negative nodes,
700 L. Olavarrieta et al.
whereas negative supercoiling leads to knots with positive
nodes (Krasnow et al., 1983; Wasserman and Cozzarelli,
1991). The observation that, in partially replicated molecules containing a stalled fork, most of the nodes of
knotted bubbles have a positive sign (Sogo et al., 1999)
indicates that in vivo these molecules had precatenanes
that were negatively twisted (Postow et al., 1999). This
would be valid only for molecules containing a stalled fork
if the putative residual action of DNA gyrase proved to be
key, but it could also be true for partially replicated molecules during unimpaired DNA replication if the latter is
negligible (see above).
To find out whether the number of knotted bubbles
changes at different stages of replication, we constructed
three different plasmids in which knotted bubbles could be
readily examined after 25%, 52% and 81% of replication.
Owing to DNA stiffness, it is conceivable that small
bubbles will not be able to accommodate complex knots.
DNA curvature increases for knots with large numbers of
nodes. For this reason, only large bubbles would have the
flexibility required to accommodate them. To deal with this
potential problem, we constructed rather large plasmids
(ª 10 kb), where the bubble of the smallest 25% replicated
was 2.5 kb. Although this does not eliminate the possibility
that bubble size could influence knotting, especially in the
case of complex knots, it was shown previously that bubbles of this size could easily accommodate knots with up
to 16 nodes (Olavarrieta et al., 2002). The results obtained
indicated that the number and complexity of knotted bubbles rose as a function of bubble size.
Results
Knotted bubbles are readily detected by two-dimensional
(2D) agarose gel electrophoresis in partially replicated
plasmids containing a stalled fork after digestion with an
enzyme that cuts plasmid DNA only in the unreplicated
portion (Viguera et al., 1996; Sogo et al., 1999;
Santamaría et al., 2000; Olavarrieta et al., 2002). After
digestion with PstI, 2D gel analysis of pBR322 DNA
enriched for replication intermediates (RIs) corresponding to monomeric forms led to a very peculiar pattern
(Fig. 1C). The pattern expected, as predicted by the 2D
gel model (Viguera et al., 1998), was a bubble arc that
should switch to a double-Y arc as soon as the replication
fork reaches the PstI site (Fig. 1B). However, not one but
several bubble arcs were clearly identified in the autoradiogram (Fig. 1C and D). All these bubble arcs started at
the 1.0¥ linear form and vanished abruptly at the position
expected for the fork to reach the PstI site. These bubble
arcs showed increasing electrophoretic mobility during the
first dimension and decreasing mobility during the second
dimension of the 2D gel system. Such behaviour was
already observed for linear molecules analysed in 2D gels
Fig. 1. Unknotted and knotted bubble arcs formed during unimpaired DNA replication as visualized by 2D agarose gel electrophoresis.
A. Genetic map of pBR322 showing the relative position of its most relevant features: the ColE1 unidirectional origin, the ampicillin and tetracycline
resistance genes, the copy number control rop gene and the recognition site for PstI.
B. The RIs of pBR322 after digestion with PstI and the corresponding 2D gel pattern as predicted by the 2D gel computer model (Viguera et al.,
1998).
C. Autoradiogram of the 2D gel.
D. Diagrammatic interpretation of the signals observed in the autoradiogram. KnB, knotted bubbles; UnknB, unknotted bubbles; DY, double-Ys;
SY, simple-Ys; Xr, X-shaped recombinants; BrB, broken bubbles.
© 2002 Blackwell Publishing Ltd, Molecular Microbiology, 46, 699–707
Supercoiling, knotting and replication 701
(Bell and Byers, 1983) as well as for knotted bubbles
of relatively high molecular weight (Viguera et al., 1996;
Sogo et al., 1999; Santamaría et al., 2000; Olavarrieta
et al., 2002). Also, a complete simple-Y arc was absent in
the autoradiogram, as expected for the RIs corresponding
to monomeric forms (Martín-Parras et al., 1991; 1992).
The series of bubble arcs showed no continuation after
the fork had passed the PstI site, as a single double-Y arc
was detected. We believe this series of additional bubble
arcs corresponded to RIs containing a knotted bubble
with increasing numbers of nodes. Knotted bubbles are
detected only in linear forms containing an internal bubble
(Viguera et al., 1996; Santamaría et al., 1998; Sogo et al.,
1999; Olavarrieta et al., 2002) and are expected to dissolve as soon as a restriction enzyme introduces a
double-stranded break within the bubble to generate a
double-Y.
pTerE25, pTerE52 and pTerE81 are all derivatives of
pBR10 (Fig. 2), in which the Escherichia coli polar
replication terminator TerE (Hill et al., 1988; Bastia and
Mohanty, 1996) was cloned in its active orientation at
three different positions. Progression of the replication
fork halts as it encounters a TerE–TUS complex in the
proper orientation on account of its contrahelicase action
(Khatri et al., 1989). We designed these three plasmids
so that the replication fork would stop either after 25%,
52% or 81% of replication. To confirm the accumulation of
RIs replicated to different extents, plasmid DNAs were
digested with AlwNI and ScaI. Comparative amounts of
the three digested plasmids were mixed and analysed in
Fig. 2. Genetic maps of pBR10, pTerE25, pTerE52 and pTerE81
showing the relative position of their most relevant features: the ColE1
unidirectional origin, the E. coli terminator TerE, the ampicillin resistance and copy number control rop genes and the recognition sites
for a number of restriction endonucleases.
© 2002 Blackwell Publishing Ltd, Molecular Microbiology, 46, 699–707
a single 2D gel. Finally, the DNA in the gel was Southern
blotted and hybridized with an appropriate probe. We used
the 2D gel computer model (Viguera et al., 1998) to predict the shape and 2D gel patterns expected (Fig. 3). In
all three cases, blockage of the replication fork at TerE
would lead to the accumulation of linear molecules
containing an internal bubble that would be 1.25¥ the
mass of non-replicating molecules for pTerE25, 1.52¥ for
pTerE52 and 1.81¥ for pTerE81. The spots corresponding
to these molecules were expected to occur on top of the
bubble arc (Brewer and Fangman, 1987; Martín-Parras
et al., 1991; Friedman and Brewer, 1995) at three different
positions depending on their corresponding masses
(Fig. 3). Figure 4 shows a photograph of the autoradiogram obtained together with a diagrammatic interpretation
to the right. Unknotted and knotted bubbles were readily
detected for all three different plasmids, and their relative
positions fitted the pattern expected (see Fig. 3). The electrophoretic mobility of the ‘beads-on-a-string’ signal corresponding to knotted bubbles, however, differed for the
three plasmids. The signal extended downwards and
steeply to the right for pTerE25, in a smoother manner for
pTerE52 and upwards to the right for pTerE81 (Fig. 4).
This observation indicated that, during the first dimension
of the 2D gel system that was run at low voltage in a lowpercentage agarose gel, all knotted bubbles increased
their electrophoretic mobility as DNA knots became more
and more complex. But, during the second dimension,
which was run at high voltage in a high-percentage agarose gel, this correlation changed depending on the mass
of the molecules. It still held true for pTerE25, but not for
pTerE52 and pTerE81. In the latter case and at least for
the left half of the ‘beads-on-a-string’ signal, the electrophoretic mobility actually decreased as node number
increased. To compare the number of knotted bubbles
between plasmids better, their corresponding DNAs were
analysed in independent 2D gels. Comparative autoradiogram exposures corresponding to the three plasmids
are shown in Fig. 5. A densitometric profile of each autoradiogram is shown below. To make comparison more
reliable, densitometric profiles were adjusted so that the
area corresponding to the signal responsible for unknotted
bubbles was made equal for all three different profiles.
Then we proceeded to measure the area corresponding
to knotted bubbles in each case. Although evident even
by the naked eye, densitometric measurements showed
clearly that the number and complexity of knotted bubbles
increased as a function of bubble size. Only 35% of the
bubbles were knotted for pTerE25, in which knotted bubbles with up to nine nodes could be identified. In the case
of pTerE52, 45% of the bubbles were knotted, and this
number jumped to 66% for pTerE81. Knotted bubbles with
up to 14 and 22 nodes could be identified for pTerE52 and
pTerE81 respectively.
702 L. Olavarrieta et al.
Fig. 3. The RIs of pTerE25, pTerE52 and pTerE81 as predicted by the 2D gel computer model (Viguera et al., 1998). The linear map of each
plasmid after digestion with AlwNI and ScaI is shown on top. The shapes of a selected number of RIs with their corresponding masses appear
below. The dashed lines point to the location of the Ter site in each case. The expected 2D gel pattern for a mixture of the three plasmids is
shown below. The grey spots on top of the bubble arc indicate the signals expected for RIs containing an unknotted bubble in each case.
Another very interesting observation that was particularly evident for the autoradiograms corresponding to
pTerE52 and pTerE81 was the presence of at least two
different and independent families of accumulated intermediates in each case. A higher magnification of the
autoradiogram corresponding to pTerE52 together with a
diagrammatic interpretation indicating these two families
of stereoisomers is shown in Fig. 6. During the seconddimension electrophoresis, members of the less abundant
family (depicted in grey in the diagram) migrated faster
than the most abundant family (depicted in black). Two
possible interpretations for these complex signals are
discussed below.
Discussion
Knotted bubbles can form during unimpaired
DNA replication
RIs containing knotted bubbles were originally identified
in bacterial plasmids containing two inversely oriented
unidirectional origins (Viguera et al., 1996). The advancing replication fork halts as soon as it runs on a second
ColE1 origin in the opposite orientation because of the
inability of the DnaB helicase to disrupt the RNA 3¢ end
of a DNA–RNA hybrid (Santamaría et al., 1998). This
blockage leads to the accumulation of partially replicated
molecules containing an internal bubble. Analysis of these
RIs in 2D gels after digestion with enzymes that cut outside the bubble revealed the presence of a ‘beads-on-astring’ signal that was assigned to knotted bubbles by
neutral-alkaline 2D gels (Viguera et al., 1996) and electron microscopy of RecA-coated molecules (Sogo et al.,
1999). It was subsequently shown that knotted bubbles
could also be readily detected if replication forks were
blocked at a Ter–TUS complex (Santamaría et al., 2000;
Olavarrieta et al., 2002). The electrophoretic mobility of
knotted molecules increases as a function of node number (Stasiak et al., 1996). As the trefoil knot – the simplest
knot – has three nodes, there is always a gap between
© 2002 Blackwell Publishing Ltd, Molecular Microbiology, 46, 699–707
Supercoiling, knotting and replication 703
Fig. 4. 2D gel autoradiogram corresponding to a mixture of comparative amounts of the three plasmids (pTerE25, pTerE52 and pTerE81)
after digestion with AlwNI and ScaI. The diagram to the right highlights the signals corresponding to unknotted bubbles (on top of the
bubble arc) and knotted bubbles.
extent of replication. In other words, all the plasmid’s DLk
occurs as supercoils at the beginning of replication,
whereas most of it takes the form of precatenanes as
completion of replication approaches (Peter et al., 1998).
Knotted bubbles form when topo IV knots the two daughter duplexes behind the fork (Postow et al., 1999; Sogo
et al., 1999). Interestingly, in partially replicated molecules
containing a stalled fork, most of the nodes of knotted
bubbles have a positive sign (Sogo et al., 1999), indicating
that precatenanes were negatively twisted at the time of
knot formation (Postow et al., 1999). Experimental evidence showing that the number of knotted bubbles rises
as a function of bubble size during unimpaired DNA replication is still missing. But, assuming that the topological
effects of fork stalling could be negligible (see Introduction), our finding that, in partially replicated molecules, the
number of knotted bubbles rose as a function of bubble
size would favour the notion that plasmids keep their
(–) DLk from the beginning to the end of the replication
process.
the electrophoretic mobility of the unknotted and the
first knotted form (Stasiak et al., 1996; Viguera et al.,
1996; Vologodskii et al., 1998; Santamaría et al., 2000;
Olavarrieta et al., 2002). This gap becomes evident when
comparing the electrophoretic mobility of molecules with
no, three, four, five and more nodes. The series of bubble
arcs observed in Fig. 1C complied with this pattern. This
observation, together with the fact that the series of bubble arcs showed no continuation in the form of double-Ys,
strongly favours the hypothesis that they correspond to
RIs containing a knot with an increasing number of nodes.
Knotting has potentially devastating effects (Pieranski
et al., 2001), but cells are able to remove DNA knots
efficiently. It was shown recently that topo IV alone is
responsible for unknotting DNA in E. coli (Deibler et al.,
2001).
The number and complexity of knotted bubbles increase
as a function of bubble size
Maintenance of (–) supercoiling is essential for the
promotion of DNA double helix opening (Crisona et al.,
2000). This explains why progression of the replication
fork is impeded when both gyrase and topo IV are mutated
or inhibited (Hiasa and Marians, 1994; 1996; Levine et al.,
1998; Khodursky et al., 2000). As DLk diffuses across
the replication fork (Champoux and Been, 1980; Peter
et al., 1998), its distribution between the unreplicated
and replicated portions of the plasmid is a function of the
© 2002 Blackwell Publishing Ltd, Molecular Microbiology, 46, 699–707
Fig. 5. Autoradiograms of the 2D gel area in which the unknotted and
knotted bubbles corresponding to pTerE25 (top), pTerE52 (middle)
and pTerE81 (bottom) migrated after digestion with AlwNI and ScaI.
Note the difference in the number and complexity of knotted bubbles.
To help visualization of this difference, a densitometric profile of
unknotted and knotted bubbles (made using version 1.61 of NIH
IMAGE) is shown below each autoradiogram. The densitometric profiles were normalized so that the areas corresponding to the signal
responsible for unknotted bubbles were made equal.
704 L. Olavarrieta et al.
Fig. 6. Higher magnification of the area of the
2D gel autoradiogram in which pTerE52 unknotted and knotted bubbles migrated. Note the
presence of two independent series of ‘beadson-a-string’ signals. The diagram to the right
highlights these two families. The most abundant family is depicted in black and the less
abundant in grey. The stippled triangular spot
(marked with a star) could correspond to
unknotted bubbles containing a hemiprecatenane or to the first element of the family of
prime knotted bubbles (see text for details).
On the nature of two different populations of
knotted bubbles
Two different families of knotted bubbles were clearly identified for pTerE52 and pTerE81. A similar observation was
made for DNA knots made in vitro (Trigueros et al., 2001).
The less abundant family that we observed for both plasmids could be interpreted in two different ways. They
might correspond to unknotted and knotted bubbles carrying a hemiprecatenane. In such a case, the stippled spot
marked with a star in the diagrammatic interpretation in
Fig. 6 would correspond to unknotted bubbles containing
a hemiprecatenane. The term hemicatenane was originally used to define a structure arising after homologous
pairing of a single-stranded DNA molecule with a duplex,
promoted by the RecA protein of E. coli (DasGupta et al.,
1980; Cunningham et al., 1981; Bianchi et al., 1983) and
was later extended to account for linked duplexes in which
one strand of a duplex is wound around one strand of
another duplex (Kmiec and Holloman, 1986; Sogo et al.,
1986; Laurie et al., 1998). Evidence for the occurrence of
hemicatenanes in vivo was obtained by electron microscopy in psoralen cross-linked replicating SV40 minichromosomes (Sogo et al., 1986; Laurie et al., 1998) and in
vitro in plasmid DNA from Ustilago maydis (Kmiec and
Holloman, 1986). Evidence in vivo was also obtained by
2D agarose gel electrophoresis after inactivation of topoisomerases in SV40 and the yeast 2 mm plasmid (Levac
and Moss, 1996) as well as in pBR322 DNA replicating in
Xenopus egg extracts (Lucas and Hyrien, 2000). In all
these cases, however, it was not possible to determine
whether or not the molecules involved were fully replicated. In our case, it would seem more appropriate to call
them hemiprecatenanes, as the arms involved were
the daughter duplexes of partially replicated molecules
(Fig. 7). Alternatively, the less abundant family of knotted
bubbles observed could correspond to composite knotted
bubbles, molecules containing two independent knots. In
this case, the stippled spot marked with a star in the
diagrammatic interpretation in Fig. 6 would correspond to
Fig. 7. Cartoons illustrating a portion of an RI displaying a relaxed
bubble, a precatenane, a hemiprecatenane, a prime knotted bubble,
a prime knotted bubble with a hemiprecatenane and a composite
knotted bubble. Parental strands are depicted in green and blue,
whereas nascent strands in red. For the sake of simplicity, no attempt
was made to keep the length of the daughter duplexes equal. An
arrowhead points to a hemicatenane, and the insert depicts a hemiprecatenane at a higher magnification.
© 2002 Blackwell Publishing Ltd, Molecular Microbiology, 46, 699–707
Supercoiling, knotting and replication 705
the first element of the most abundant family displaying
the simplest trefoil knotted bubble. During the first dimension of the 2D gel system that was run at low voltage in
a low-percentage agarose gel, the first element of a family
of composite knotted bubbles formed by two independent
trefoil knots should migrate as a six-noded prime knotted
bubble, and this was precisely the case. The observation
that a similar case was reported for DNA knots made in
vitro (Trigueros et al., 2001) further supports this latter
interpretation.
at 0.45 V cm-1 at room temperature for 69 h. The lane containing the lambda DNA/HindIII marker sizes was excised,
stained with 0.3 mg ml-1 ethidium bromide and photographed.
In the meantime, the lanes containing the DNA problem were
kept in the dark. The second dimension was in a 0.58%
agarose gel in TBE containing 0.3 mg ml-1 ethidium bromide
at a 90∞ angle with respect to the first dimension. The dissolved agarose was poured around the excised lane from the
first dimension, and electrophoresis was at 0.89 V cm-1 also
at room temperature and for 95 h. Southern transfer was
performed as described previously (Viguera et al., 1996;
Martín-Parras et al., 1998; Santamaría et al., 1998;
Olavarrieta et al., 2002).
Experimental procedures
Bacterial strains and culture medium
Non-radioactive hybridization
The E. coli strain used was DH5aF¢. Competent cells were
transformed with monomeric forms of the plasmids as
described previously (Viguera et al., 1996; Santamaría et al.,
1998; Olavarrieta et al., 2002). Cells were grown at 37∞C in
LB medium containing 75 mg ml-1 ampicillin.
Probes were labelled using the Random Primer Fluorescein
kit (NEN Life Sciences Products). Membranes were
prehybridized in a 20 ml prehybridization solution (2¥
SSPE, 0.5% Blotto, 1% SDS, 10% dextran sulphate and
0.5 mg ml-1 sonicated and denatured salmon sperm DNA) at
65∞C for 4–6 h. Labelled DNA was added, and hybridization
lasted 12–16 h. Then, membranes were washed successively with 2¥ SSC and 0.1% SDS, 0.5¥ SSC and 0.1% SDS,
0.1¥ SSC and 0.1% SDS for 15 min each at room temperature except for the last wash, which occurred at 65∞C. Detection was performed with an antifluorescein-AP conjugate and
CDP-Star (NEN) according to the instructions provided by the
manufacturer.
Plasmid construction
pBR322 and pBR10 derivatives were used. pBR10 is a
derivative of pBR18 (Santamaría et al., 2000), in which the
5.8 kb EcoRI fragment of human rDNA was inserted at the
unique AvaI site of pBR18. The resulting 10.2 kb plasmid
contains a ColE1 unidirectional origin and confers resistance
only to ampicillin (Fig. 1). To construct pTerE25, two oligos,
5¢-CGCGTCTTAGTTACAACATACTTTAAAGAGCT-3¢ and 5¢CTTTAAAGTATGTTGTAACTAAGA-3¢, containing the 23 bp
that constitutes the E. coli TerE terminator (Hill et al., 1988;
Bastia and Mohanty, 1996) with a 3¢ SacI and a 5¢ MluI tails
were annealed to each other and inserted between the
unique SacI and MluI sites of pBR10. To construct pTerE52,
another pair of oligos, 5¢-TCTTAGTTACAACATACTTTAAAT
GCA-3¢ and 5¢-TTTAAAGTATGTTGTAACTAAGATGCA-3¢,
including TerE with two NsiI tails were annealed to each other
and inserted at the unique NsiI site of pBR10. Finally, pTer81
was constructed using a third pair of oligos, 5¢-AATTCGGCT
TAGTTACAACATACTT TAAA-3¢ and 5¢-AGCTTTTAAAGTAT
GTTGTAACTAAGCCG-3¢, containing TerE with a 3¢ EcoRI tail
and a 5¢ HindIII tail. These oligos were annealed to each
other and inserted between the unique EcoRI and HindIII
sites of pBR10.
Isolation of plasmid DNA
The isolation of plasmid DNA was performed as described
previously (Viguera et al., 1996; Martín-Parras et al., 1998;
Santamaría et al., 1998; Olavarrieta et al., 2002) with two
minor modifications: Triton X-100 was used instead of Brij-58
and sodium deoxycholate, and DNA precipitation was performed using isopropanol instead of 95% ethanol.
Two-dimensional gel electrophoresis and
Southern transfer
The first dimension was in a 0.28% agarose gel in TBE buffer
© 2002 Blackwell Publishing Ltd, Molecular Microbiology, 46, 699–707
Densitometry
Autoradiograms were scanned using a GS-800 Bio-Rad
calibrated densitometer and analysed using the Bio-Rad
QUANTITY ONE program, version 4.2.2. and NIH IMAGE, version
1.61.
Acknowledgements
We thank all the other permanent and transient members of
the laboratory (Alberto Benguría, Sylwia Baranska, Ma José
Fernández, Ana García, Agurtzane Irizar, Tatiana Ivanenko,
Leonor Rodríguez, Alicia Sánchez, Nora Soberón, Raúl
Torres and Pilar Robles) for their help and constant support.
We also acknowledge Andrzej Stasiak and José Manuel
Sogo for their suggestions and critical reading of the manuscript. This work was partially financed by grants PM97-0138,
PB98-048 and SAF2001-1740 from the Spanish Ministerio
de Ciencia y Tecnología (MCyT), and by grant 99/0850 from
the Spanish Fondo de Investigación Sanitaria (FIS).
References
Alexandrov, A.I., Cozzarelli, N.R., Holmes, V.F., Khodursky,
A.B., Peter, B.J., Postow, L., et al. (1999) Mechanisms of
separation of the complementary strands of DNA during
replication. Genetica 106: 131–140.
Bastia, D., and Mohanty, B.K. (1996) Mechanisms for completing DNA replication. In DNA Replication in Eukaryotic
706 L. Olavarrieta et al.
Cells. DePamphilis, M.L. (ed.). New York: Cold Spring Harbor Laboratory Press, pp. 177–215.
Bell, L., and Byers, B. (1983) Separation of branched from
linear DNA by two-dimensional gel electrophoresis. Anal
Biochem 130: 527–535.
Bianchi, M., DasGupta, C., and Radding, C.M. (1983) Synapsis and the formation of paranemic joints by E. coli RecA
protein. Cell 34: 931–939.
Brewer, B.J., and Fangman, W.L. (1987) The localization of
replication origins on ARS plasmids in S. cerevisiae. Cell
51: 463–471.
Champoux, J.J., and Been, M.D. (1980) Topoisomerases and
the swivel problem. In Mechanistic Studies of DNA Replication and Genetic Recombination. Alberts, B. (ed.). New
York: Academic Press, pp. 809–815.
Crisona, N.J., Strick, T.R., Bensimon, D., Croquette, V., and
Cozzarelli, N.R. (2000) Preferential relaxation of positively
supercoiled DNA by E. coli topoisomerase IV in singlemolecule and ensemble measurements. Genes Dev 14:
2881–2892.
Cunningham, R.P., Wu, A.M., Shibata, T., DasGupta, C.,
and Radding, C.M. (1981) Homologous pairing and
topological linkage of DNA molecules by combined action
of E. coli RecA protein and topoisomerase I. Cell 24: 213–
223.
DasGupta, C., Shibata, T., Cunningham, R.P., and Radding,
C.M. (1980) The topology of homologous pairing promoted
by RecA protein. Cell 22: 437–446.
Deibler, R.W., Rahmati, S., and Zechiedrich, E.L. (2001)
Topoisomerase IV, alone, unknots DNA in E. coli. Gene
Dev 15: 748–761.
Duguet, M. (1997) When helicase and topoisomerase meet!
J Cell Sci 110: 1345–1350.
Friedman, K.L., and Brewer, B.J. (1995) Analysis of replication intermediates by two-dimensional agarose gel electrophoresis. In DNA Replication, Vol. 262. Campbell, J.L.
(ed.). San Diego, CA: Academic Press, pp. 613–627.
Hiasa, H., and Marians, K.J. (1994) Topoisomerase IV can
support oriC DNA replication in vitro. J Biol Chem 269:
16371–16375.
Hiasa, H., and Marians, K.J. (1996) Two distinct modes of
strand unlinking during theta-type DNA replication. J Biol
Chem 271: 21529–21535.
Hill, T.M., Pelletier, A.J., Tecklenburg, M.L., and Kuempel,
P.L. (1988) Identification of the DNA sequence from E. coli
terminus region that halts replication forks. Cell 55: 459–
466.
Kanaar, R., and Cozzarelli, N.R. (1992) Roles of supercoiled
DNA structure in DNA transactions. Curr Opin Struct Biol
2: 369–379.
Khatri, G.S., MacAllister, T., Sista, P.R., and Bastia, D.
(1989) The replication terminator protein of E. coli is a
sequence-specific contra-helicase. Cell 59: 667–674.
Khodursky, A.B., Peter, B.J., Schmidt, M.B., DeRisi, J.,
Botstein, D., Brown, P.O., and Cozzarelli, N.R. (2000)
Analysis of topoisomerase function in bacterial replication
fork movement: use of DNA microarrays. Proc Nat Acad
Sci USA 97: 9419–9424.
Kmiec, E.B., and Holloman, W.K. (1986) Homologous pairing
of DNA molecules by Ustilago Rec1 protein is promoted
by sequences of Z-DNA. Cell 44: 545–554.
Kornberg, A., and Baker, T.A. (1992) DNA Replication, 2nd
edn. New York: W.H. Freeman
Krasnow, M.A., Stasiak, A., Spengler, S.J., Dean, F., Koller,
T., and Cozzarelli, N.R. (1983) Determination of the absolute handedness of knots and catenanes of DNA. Nature
304: 559–560.
Laurie, B., Katritch, V., Sogo, J., Koller, T., Dubochet, J., and
Stasiak, A. (1998) Geometry and physics of catenanes
applied to the study of DNA replication. Biophys J 74:
2815–2822.
Levac, P., and Moss, T. (1996) Inactivation of topoisomerase
I or II may lead to recombination or to aberrant replication
termination on both SV40 and yeast 2 mm DNA. Chromosoma 105: 250–260.
Levine, C., Hiasa, H., and Marians, K.J. (1998) DNA gyrase
and topoisomerase IV: biochemical activities, physiological
roles during chromosome replication, and drug sensitivities. BBA Gene Struct Express 1400: 29–43.
Lucas, I., and Hyrien, O. (2000) Hemicatenanes form upon
inhibition of DNA replication. Nucleic Acids Res 28: 2187–
2193.
Lucas, I., Germe, T., Chevrier-Miller, M., and Hyrien, O.
(2001) Topoisomerase II can unlink replicating DNA by
precatenane removal. EMBO J 20: 6509–6519.
Martín-Parras, L., Hernández, P., Martínez-Robles, M.L., and
Schvartzman, J.B. (1991) Unidirectional replication as
visualized by two-dimensional agarose gel electrophoresis.
J Mol Biol 220: 843–853.
Martín-Parras, L., Hernández, P., Martínez-Robles, M.L.,
and Schvartzman, J.B. (1992) Initiation of DNA replication
in ColE1 plasmids containing multiple potential origins of
replication. J Biol Chem 267: 22496–22505.
Martín-Parras, L., Lucas, I., Martínez-Robles, M.L.,
Hernández, P., Krimer, D.B., Hyrien, O., and Schvartzman,
J.B. (1998) Topological complexity of different populations
of pBR322 as visualized by two-dimensional agarose gel
electrophoresis. Nucleic Acids Res 26: 3424–3432.
Mohanty, B.K., Sahoo, T., and Bastia, D. (1998) Mechanistic
studies on the impact of transcription on sequence-specific
termination of DNA replication and vice versa. J Biol Chem
273: 3051–3059.
Olavarrieta, L., MartinezRobles, M.L., Sogo, J.M., Stasiak,
A., Hernandez, P., Krimer, D.B., and Schvartzman, J.B.
(2002) Supercoiling, knotting and replication fork reversal
in partially replicated plasmids. Nucleic Acid Res 30: 656–
666.
Peter, B.J., Ullsperger, C., Hiasa, H., Marians, K.J., and
Cozzarelli, N.R. (1998) The structure of supercoiled intermediates in DNA replication. Cell 94: 819–827.
Pieranski, P., Kasas, S., Dietler, G., Dubochet, J., and
Stasiak, A. (2001) Localization of breakage points in knotted strings. New J Physics 3: 10.1–10.10.13.
Postow, L., Peter, B.J., and Cozzarelli, N.B. (1999) Knot what
we thought before: the twisted story of replication. Bioessays 21: 805–808.
Postow, L., Ullsperger, C., Keller, R.W., Bustamante, C.,
Vologodskii, A.V., and Cozzarelli, N.R. (2001) Positive torsional strain causes the formation of a four-way junction at
replication forks. J Biol Chem 276: 2790–2796.
Santamaría, D., delaCueva, G., Martínez-Robles, M.L.,
Krimer, D.B., Hernández, P., and Schvartzman, J.B.
© 2002 Blackwell Publishing Ltd, Molecular Microbiology, 46, 699–707
Supercoiling, knotting and replication 707
(1998) DnaB helicase is unable to dissociate RNA–DNA
hybrids – its implication in the polar pausing of replication
forks at ColE1 origins. J Biol Chem 273: 33386–33396.
Santamaría, D., Hernández, P., Martínez-Robles, M.L.,
Krimer, D.B., and Schvartzman, J.B. (2000) Premature
termination of DNA replication in plasmids carrying two
inversely oriented ColE1 origins. J Mol Biol 300: 75–82.
Sogo, J.M., Stahl, H., Koller, T., and Knippers, R. (1986)
Structure of replicating Simian virus 40 minichromosomes.
J Mol Biol 189: 189–204.
Sogo, J.M., Stasiak, A., Martínez-Robles, M.L., Krimer, D.B.,
Hernández, P., and Schvartzman, J.B. (1999) Formation
of knots in partially replicated DNA molecules. J Mol Biol
286: 637–643.
Stasiak, A., Katrich, V., Bednar, J., Michoud, D., and
Dubochet, J. (1996) Electrophoretic mobility of DNA knots.
Nature 384: 122.
Trigueros, S., Arsuaga, J., Vazquez, M.E., Sumners, D.W.,
and Roca, J. (2001) Novel display of knotted DNA molecules by two-dimensional gel electrophoresis. Nucleic
Acids Res 29, e67.
© 2002 Blackwell Publishing Ltd, Molecular Microbiology, 46, 699–707
Ullsperger, C., Vologodskii, A.A., and Cozzarelli, N.R. (1995)
Unlinking of DNA by topoisomerases during DNA replication. In Nucleic Acids and Molecular Biology. Lilley, D.M.J.,
and Eckstein, F. (eds). Berlin: Springer-Verlag, pp. 115–
142.
Viguera, E., Hernández, P., Krimer, D.B., Boistov, A.S., Lurz,
R., Alonso, J.C., and Schvartzman, J.B. (1996) The ColE1
unidirectional origin acts as a polar replication fork pausing
site. J Biol Chem 271: 22414–22421.
Viguera, E., Rodríguez, A., Hernández, P., Krimer, D.B.,
Trellez, O., and Schvartzman, J.B. (1998) A computer
model for the analysis of DNA replication intermediates by
two-dimensional agarose gel electrophoresis. Gene 217:
41–49.
Vologodskii, A.V., Crisona, N.J., Laurie, B., Pieranski, P.,
Katritch, V., Dubochet, J., and Stasiak, A. (1998) Sedimentation and electrophoretic migration of DNA knots and catenanes. J Mol Biol 278: 1–3.
Wasserman, S.A., and Cozzarelli, N.R. (1991) Supercoiled
DNA-directed knotting by T4 topoisomerase. J Biol Chem
266: 20567–20573.