Step 1. Replication origins are determined, at least in part, by
the binding of the six subunit origin recognition complex (ORC),
which has been conserved in evolution from yeast to humans. In
budding yeast, ORC binds specifically to an essential, bipartite
sequence element within replication origins. ORC binds ATP and this
binding is required for DNA binding. ORC can also hydrolyse ATP;
however, ATP hydrolysis is not required for DNA binding and is, in
fact, inhibited upon origin binding. ORC remains bound to origins
throughout the cell cycle in budding yeast, even through mitosis.
In multicellular eukaryotes some ORC subunits remain bound
throughout G1, S and G2 phases, although there is conflicting
evidence as to whether ORC remains bound to chromosomes during
mitosis.
Figure 1. Metaphase-to-anaphase transition in rat kangaroo PtK2
cells. -Tubulin, green; centrosomes, red; DNA stained with
4',6-diamidino-2-phenylindole, blue.
Figure 2. Structure of cohesin and a possible mechanism by which
it might hold sister chromatids together. (A) Smc1 (red) and Smc3
(blue) form intramolecular antiparallel coiled coils, which are
organized by hinge or junction domains (triangles). Smc1/3
heterodimers are formed through heterotypic interactions between
the Smc1 and Smc3 junction domains. The COOH terminus of Scc1
(green) binds to Smc1's ABC-like ATPase head, whereas its NH2
terminus binds to Smc3's head, creating a closed ring. Scc3
(yellow) binds to Scc1's COOH-terminal half and does not make any
direct stable contact with the Smc1/3 heterodimer. Scc1's separase
cleavage sites are marked by arrows. Cleavage at either site is
sufficient to destroy cohesion. By analogy with bacterial SMC
proteins, it is expected that ATP binds both the Smc1 and Smc3
heads, alters their conformation, and possibly brings them into
close proximity. By altering Scc1's association with Smc heads, ATP
binding and/or hydrolysis could have a role in opening and/or
closing cohesin's ring. (B) Cohesin could hold sister DNA molecules
together by trapping them both within the same ring. Cleavage of
Scc1 by separase would open the ring, destroy coentrapment of
sister DNAs, and cause dissociation of cohesin from chromatin. (C)
Smc-containing complexes other than cohesin could also function via
chromatid entrapment. Condensin, for example (black), could
organize mitotic chromosomes by trapping supercoils. It and/or
other related complexes could hold distant loci together (arrow)
and thereby facilitate the function of long-range enhancers and
silencers of transcription. Figure 3. The anaphase-promoting
complex induces amphitelically attached chromatids to segregate to
opposite poles by destroying both cyclin B and securin. The
Ipl1/Aurora B kinase both eliminates syntelically attached
chromatid pairs and promotes inhibition of APC/C when centromeres
fail to come under tension. Chromatin, blue; microtubules, green;
centromeres, open circles; cohesin, red.
Figure 4. Chromatid individualization as cells enter mitosis
involves dissociation of most cohesin (red symbols) from
chromosomes, which is regulated by PLK. This process coincides with
condensin's (green symbols) association with chromosomes and their
compaction. Cohesin remaining on chromosomes, largely at
centromeres, is then cleaved by separase at the
metaphase-to-anaphase transition.
Figure 1. Summary of S.cerevisiae DNA damage, replication, and
mitotic checkpoints. The different stages of the cell cycle are
indicated above the horizontal line. Below the line are listed a
subset of the proteins that function in the indicated checkpoint
branches. These proteins are thought to detect the "damage" that
triggers each checkpoint. The primary effect of activating each
checkpoint is shown below the proteins. Listed below the three
checkpoint branches that function in S-phase are many of the known
proteins that function in the downstream signal transduction
cascade or are targeted by this cascade. The box at the right lists
human homologs of yeast checkpoint genes that are mutated in human
cancer susceptibility syndromes.
Figure 2. Illustration of break-induced replication, telomere
maintenance, and de novo telomere addition reactions. (A)
Break-induced replication. In this example, the 3' end of a broken
DNA invades the intact DNA to form a D loop. The resulting
heteroduplex region is then extended and a Holliday junction is
formed. By appropriate resolution of the Holliday junction, a
structure is formed that is equivalent to a replication fork. (B)
Telomere maintenance functions. The diagram shows the end of a
chromosome containing a single-stranded TG repeat end and the known
S.cerevisiae proteins that bind to telomeres and extend the TG
repeat sequences. (C) De novo telomere addition reactions. The
diagram shows the end of a broken DNA molecule and the proteins
that have been implicated in de novo telomere addition.
Figure 3. Multiple pathways function to maintain genome
stability in S.cerevisiae. DNA replication errors activate S-phase
checkpoint sensors, whereas telomere damage likely activates DNA
damage checkpoint sensors. These sensors then activate downstream
responses, including DNA repair, that are required for suppression
of genome instability and possibly cell cycle arrest or delay.
Multiple pathways work to correct the damage; the primary
nonmutagenic repair pathway appears to be break-induced replication
(BIR), although double-strand break (DSB) repair and nonhomologous
end joining (NHEJ) may play minor roles. The major mutagenic
pathways are the translocation pathway(s), the de novo telomere
addition pathway suppressed by PIF1, and aberrant recombination
reactions that are suppressed by DNA mismatch repair (MMR) and
other functions. The number of translocation pathways and whether
there are pathways that specifically suppress translocations remain
to be determined. Figure 1. Replication region of bacteriophage
genome and assembly of the replication complex. Genes, promoters
and terminators present in the replication region are indicated.
Transcripts are represented by arrows. The scheme is not drawn to
scale. See text for details.
Figure 2. Two pathways of plasmid replication. Large circles
represent plasmid DNA molecules. Small filled (orange) circles
indicate heritable replication complexes. See text for details.
Figure 3. Effects of various factors on formation and stability
of the heritable replication complex. The complex is symbolized by
large orange circle. Positive regulators are marked in green, and
negative regulators are marked in blue. Stimulation processes are
shown as arrows and inhibitory actions are shown by blunt-ended
lines. See text for details.
Figure 1. Model for TLS by specialized polymerases.
High-fidelity semiconservative replication is arrested at sites of
base damage or structural distortions in the replication fork
(solid triangle). Multiple specialized polymerases are able to
support TLS across offending template lesions. During TLS, the
correct (error-free) (N) or the incorrect (error-prone) (M)
nucleotide is inserted, depending on the nucleotide preference of
the engaged specialized polymerase. In either case, persistence of
the damaged template base precludes the resumption of normal DNA
replication until the primer strand is extended for some distance
(bold red), often by a second specialized polymerase, such as
prokaryotic pol V and eukaryotic pol . The replicative machinery
then reengages to continue high-fidelity synthesis. Once the site
of base damage is cleared, it is subject to normal DNA repair. If
TLS was error-free, the DNA is restored to the normal sequence
(N:N). However, if TLS resulted in nucleotide misincorporation, a
mutation will be fixed (M:M). The product of TLS is in red. Fig. 1.
A, schematic showing the primers and template DNAs prepared for
this study. The two methods of thymine oxidation are indicated as
well as the position and sequence context of the lesion. B,
literature values for the putative stereochemical make-up of each
thymine glycol sample. Note that in the TgOsO4 template, the 5R
stereoisomers are in greater relative abundance.
Fig. 2. GST-pol bypasses thymine glycol in template TgBr2.
Radiolabeled primer-templates (5nM) were incubated with all four
deoxynucleotide triphosphates (100M total) and the indicated
quantities of each polymerase and the resulting primer extension
products resolved by DPAGE. GST-pol largely bypasses thymine glycol
as does Klenow fragment (exo ) of E.coli polymerase I.In contrast,
the calf thymus pol is arrested. Lanes 1, 3, 6, and 8, control
experiments with either control or TgBr2 template but no enzyme
added. Lane 2, undamaged template with 5nM GST-pol ; lanes 4and 5,
TgBr2 template with 5and 10nM GST-pol ; lane 7undamaged template
with 1nM Klenow (exo ); lane 9, TgBr2 template with 1nM Klenow (exo
); lanes 10-12, undamaged template with 0.1unit of calf thymus pol
and increasing concentrations of PCNA; lanes 13and 14, TgBr2 with
0.4and 0.8units of calf thymus pol .
Fig. 3. A, GST-pol exhibits identical bypass of an alternatively
prepared thymine glycol template, TgOsO4, which is constitutionally
identical to TgBr2 (Fig. 2) experiments, but stereochemically
contains a greater proportion of 5R thymine glycol stereoisomers.
Radiolabeled primer-templates (5nM) were incubated with all four
deoxynucleotide triphosphates (100M total) at the indicated
quantities of each polymerase, and the resulting primer extension
products were resolved by DPAGE. Lanes 1and 4, control experiments
with no enzyme added. Lanes 2and 3, undamaged template with 1and
5nM GST-pol . Lanes 5and 6, TgOsO4 template with 1and 5nM GST-pol .
B, the TLS pattern for Klenow fragment of E.coli Pol I (exo ) with
TgOsO4, is identical to that shown for TgBr2 in Fig. 2, and T4 DNA
polymerase shows characteristic replicative arrest at Tg in TgOsO4.
Experiments were performed analogously to those in A.Lanes 1, 3, 5,
and 7, control experiments with no enzyme added. Lane 2, undamaged
template with 1nM Klenow (exo ). Lane 4, TgOsO4 template with 1nM
Klenow (exo ). Lane 6, undamaged template with 5nM T4 DNA
polymerase. Lane 8, TgOsO4 template with 5nM T4 DNA polymerase. C,
standing start experiments (primer P5-ox-ss) indicate nearly
identical bypass of the two Tg templates by GST-pol . Experiments
were performed analogously to those in A.Lanes 1and 4, control
experiments with no enzyme added; lane 2, TgBr2 template with 1nM
GST-pol ; lane 3, TgBr2 template with 5nM GST-pol ; lane 5, TgOsO4
template with 1nM GST-pol ; lane 6TgOsO4 template with 5nM GST-pol
; lane 7, undamaged template with 5nM GST-pol added.
Fig. 4. GST-pol preferentially incorporates adenine across from
thymine glycol. Radiolabeled primer-templates (5nM) were incubated
with the indicated deoxynucleotide triphosphate(s) (100M total
[dNTP]) and GST-pol (5nM) and the resulting primer extension
products were resolved by DPAGE. Lanes 1-6 contained undamaged
template DNA; lanes 7-12 contained TgBr2 template. Lanes 1and 7,
control experiments with no enzyme added. Lanes 2and 8, only dATP;
lanes 3and 9, only dCTP; lanes 4and 10, only dGTP; lanes 5and 11,
only TTP. Lanes 6and 12contained a mixture of all four dNTPs.
Fig. 5. Representative results for steady state kinetics
analysis for incorporation of the four dNTPs opposite thymine
glycol by GST-pol . The nucleotide incorporated is shown
immediately above each gel, and the micromolar dNTP concentration
incubated in each reaction is shown immediately below the
corresponding lane of each gel. Experiments were performed using
the undamaged template (A), the TgBr2 template (B), or the TgOsO4
template (C). Raw data were analyzed as described under
"Experimental Procedures," and the resulting steady state kinetic
parameters are reported in Table I. Note that most of the panels
shown represent initial conditions with a broad dNTP range. dCTP
and TTP gels in B show representative results from more narrowly
focused working ranges used in subsequent experiments.
Fig. 6. Graphical representation of select kinetic parameters
from Table I. A, kcat/Km values for nucleotide insertion opposite
thymine glycol in the TgOsO4 template (black bars), the TgBr2
template (blue bars), and the undamaged template (yellow bars). The
left graph shows directly plotted kcat/Km values obtained for
incorporation of the bases A, C, G, or T, emphasizing the
difference in incorporation efficiency of A between the control and
Tg templates. The right graph is a y axis blown up version of the
same graph to emphasize that A is incorporated in great preference
to the other bases in all three of the templates and that GST-pol
may exhibit stereochemical preference during insertion of A
opposite of thymine glycols. B, comparison of the finc values
obtained for each template, measuring the degree of preference for
incorporation of the correct base. finc is defined as
(kcat/Km)incorrect/(kcat/Km)correct. The left graph is a direct
plot of the finc values for nucleotide insertion opposite thymine
glycol in the TgOsO4 template (black bars), the TgBr2 template
(blue bars), and the undamaged template (yellow bars), emphasizing
the high degree of preference for A incorporation in each template.
The graph on the right is a y axis blown up version of the same
graph to emphasize that the misincorporation frequency of G
increases in template TgOsO4, suggesting that thymine glycol
stereochemistry influences the ability of GST-pol to discriminate
between the purines during incorporation opposite the lesion.
Fig. 7. Schematic drawing of the most important conformers
expected from the most abundant stereoisomers of thymine glycol.
The 5R,6S form (left drawings, putatively in greater abundance in
TgOsO4) may give rise to G-T wobble base-pairing in the GST-pol
active site, resulting in higher levels of misincorporation of
G.The 5S,6R isomer (right drawings) may adopt a much different
conformation. dR, deoxyribose. As a result of different half-chair
conformations, the stereoisomers may each present the hydrogen
bonding surface of the base at different angles, giving rise to the
observed differences for misincorporation of G between the TgBr2
and TgOsO4 templates.
Figure 1. Model for the sequential assembly of the various
components of nucleotide excision repair. Global genome repair
involves the initial binding of the XPC-hHR23B and XPE binding
proteins followed by downloading of the TFIIH transcription and
helicase complex that remodels the damaged site. Transcription
coupled repair involves initial response to damage by stalling of
the RNA polymerase II apparatus, and coupling by the CSA and CSB
proteins. Subsequent steps proceed in common, consisting of loading
the XPG nuclease, the XPA-RPA DNA binding proteins, and the
ERCC1-XPF nuclease. After nuclease cleavage around the dimer site,
the excision complex departs and the site is resynthesized by
PCNA-polymerase delta and ligase I. Figure has been redrawn (37)
with addition of the transcription repair pathway.
Figure 2. Causative mutations reported in the XPD gene after
elimination of non-functional mutations. The DNA/DNA helicase boxes
are shown by clear boxes with the amino acid involved indicated by
numbers above the gene. DNA/RNA helicase boxes shown in lighter
shade below the gene with roman numerals for numbering. The
mutations are indicated by the number and the type of amino acid
change, with those causing XPD or XPD/CS above, and those causing
XPD/TTD below. The interaction domain between XPD and the p44
component of the TFIIH complex is shown. Reproduced from (117).
Figure 3. The mechanism of tumor formation in basal cell nevus
syndrome involving the hedgehog (Hh), patched (PTC) and smoothened
(SMO) signal transduction pathway. Top left: normal signal
transduction pathway. PTC inhibits SMO and no nuclear transcription
signal is induced. Top right: normal pathway after sonic Hh binds
to PTC, removes binding to SMO, and results in signal transduction
to nuclear transcription factor Gli that induces cPTC
over-expression. Bottom: three mechanisms of derangement of the
signal transduction pathways in tumors involving either PTC, SHh or
SMO. Redrawn from initial diagrams provided by M. Aszterbaum MD,
PhD.
Figure1.Schematic illustration of mechanisms and barriers in
liposome-mediated gene delivery to hepatocytes. Potential barriers
are highlighted with dark background. Lipoplexes formed from
specific ligand-labeled cationic liposomes and plasmid DNA are
administered through either peripheral vein or portal vein. In both
administration routes, the lipoplexes may interact with plasma
proteins, which results in large aggregates. The aggregates may be
large enough to be trapped in the lung circulation and only small
part of injected lipoplexes will reach the liver when the
lipoplexes are administrated intravenously. Lipoplexes are
endocytosed by hepatocytes when they have been recognized by the
cells. Endocytosed lipoplex components including plasmid DNA may
undergo degradation by catalytic enzymes in lysosomes or endosomes,
and only a small portion of DNA is able to enter the nucleus
through the nuclear pore complex. Episomal DNA in the nucleus will
be transcribed, and transgene expression can be detected in cell
lysates. ASGP-R = asialoglycoprotein receptor.
Figure 1. Structure of genetically dissected replication origins
in eukaryotes. Consensus ORC binding sites are indicated in red.
Additional sequences important for origin activity are shown in
brown. Transcription units and regulatory sequences are shown in
green. Sites of ORC binding, where known, are indicated. Sites of
initiation of replication are indicated with a bidirectional arrow
passing through a bubble.
Figure 2. Once-per-cell-cycle genome duplication is independent
of the positions or density of replication origins. (A) Duplication
of DNA exactly once-per-cell division is achieved with two mutually
exclusive periods of the cell cycle during which either pre-RCs can
be assembled but replication cannot initiate or replication can
initiate but pre-RCs cannot be assembled. Regardless of where
pre-RCs assemble, DNA is completely replicated and cannot be
re-replicated until after cell division. (B) The assembly of
extraneous pre-RCs can ensure that the entire segment of DNA is
replicated in a timely fashion, without the need for specific DNA
sequences to optimize origin spacing. Any pre-RCs that do not
initiate are destroyed by passage of the replication fork and so
cannot re-initiate on the replicated strands.
Figure 3. Exclusion of pre-RCs from specific regions could
create the need for origin focusing mechanisms. In this model,
passage of the transcription apparatus before replication depletes
transcription units (black boxes) of pre-RCs (yellow stars). (A)
Without transcription, there is no selective pressure to focus
initiation to specific sites as the assembly of pre-RCs at many
sites ensures timely genome replication. (B) When transcription
units are sparse, the assembly of multiple pre-RCs at many sites
within the intergenic regions is sufficient to accomplish genome
replication. (C) When intergenic regions are few and far between,
specific DNA sequence recognition elements are required to ensure
that at least one pre-RC is assembled at intervals appropriate to
accomplish the timely replication of the DNA segment.
Fig. 1. Pre-RCs are formed and activated on immobilized linear
DNA fragments. Equivalent amounts of magnetic beads (lane 1) or
magnetic beads coupled to a 3-kb DNA fragment of pBluescript (lanes
2-6) were incubated with egg cytosol for 20min, isolated, washed,
and analyzed by Western blotting. In lane 4, the egg cytosol
contained geminin. In lanes 5and 6, the incubation in egg cytosol
was followed by 15min of incubation in aphidicolin-supplemented NPE
that contained (lane 6) or lacked (lane 5) 1M p27Kip. Western blots
were probed with MCM3 (all panels), ORC2 (left and center panels),
or Cdc45 (right panel).
Fig. 2. ORC and MCM are efficiently recruited to an 82-bp DNA
fragment. A and B, a 251-bp PCR product spanning the pBluescript
polylinker was coupled to beads. The beads were divided into
equal-sized pools and mock-digested or digested with different
enzymes to generate 251(lane 1), 154(lane 2), 120(lane 3), 94(lane
4), and 67(lane 5)-bp DNA fragments. Lane 6contained beads lacking
DNA. After digestion, DNA attached to beads was analyzed on a 2%
agarose gel (A), or equal quantities of the various beads were
incubated with egg cytosol and the attached proteins analyzed by
Western blotting (B). C, a similar set of DNA beads as those used
in A were incubated with egg cytosol containing buffer (lanes 1-5)
or geminin (lanes 6-10), and the bound proteins were analyzed by
Western blotting. D, proteins bound to equivalent amounts of 94-bp
beads (lane 1) and 251-bp beads (lane 2) were analyzed by Western
blotting alongside 2-fold dilutions of a 1:1 molar ratio of
purified MCM3 and ORC2 (lanes 3-6). The amount of ORC2 and MCM3 in
lane 3is 16and 24.6ng, respectively. E, binding of proteins to
digested DNA beads as in B but with the addition of 82-(lane 3) and
54-bp (lane 5) fragments. As in B, all lanes contained equivalent
amounts of beads that were all derived by digestion from the 251-bp
parental DNA fragment.
Fig. 3. MCM complex binding is proportional to the length of DNA
fragments attached to beads, whereas ORC binding is independent of
DNA fragment length. A, a 6-kb PCR product was coupled to magnetic
beads and digested to generate 3-,1-,and 0.35-kb DNA fragments.
Binding of MCM3 and ORC2 to equal amounts of digested beads was
examined in egg cytosol in the presence (lanes 5-8) and absence
(lanes 1-4) of geminin. B, an immobilized 6-kb DNA fragment (lane
1) or sperm (lane 2) was incubated in egg cytosol, and binding of
MCM3 and ORC2 was examined. C, a 1-kb PCR product was attached to
beads and digested to generate a 100-bp DNA fragment. Equivalent
amounts of the 1-kb DNA beads (lanes 1and 4), 100-bp DNA beads
(lanes 2and 5), and beads lacking DNA (lane 3) were incubated in
egg cytosol (EC) containing (lanes 4and 5) or lacking (lanes 1-3)
geminin, and the attached proteins were analyzed by Western
blotting. D, proteins bound to an immobilized 1-kb DNA fragment
(lane 4) were analyzed on a Western blot alongside known quantities
of purified MCM3 and ORC2. Lanes 1-3, 0.5ng of ORC2; lane 1, 15.3ng
of MCM3; lane 2, 7.7ng of MCM3; lane 3, 1.5ng of MCM3.
Fig. 4. A, lateral MCM complexes are bound tightly to chromatin.
100-bp DNA beads derived by digestion from 251-bp DNA beads (lanes
1-4) or 6-kb DNA beads prepared separately (lanes 5-8) were
incubated with egg cytosol, diluted with buffer containing Triton
X-100 and increasing concentrations of salt as indicated, isolated,
washed in the same buffer, and probed with MCM3, MCM7, or ORC2
antibodies. B, sperm chromatin was incubated with egg cytosol,
diluted with buffer containing Triton X-100 and 0.1M KCl, isolated,
washed in buffer containing 0.1(lane 1) or 1M (lane 2) KCl, and
probed with ORC2 antibody (lower panel) or a mixture of MCM3 and
MCM7 antibodies (upper panel). C, Cdc45 binding is independent of
DNA length. Equal quantities of 6-kb DNA beads (lanes 1, 5, and 9),
1-kb DNA beads (lanes 2, 6, and 10), or 100-bp DNA beads derived
from the 1-kb DNA beads by digestion (lanes 3, 7, and 11), as well
as beads lacking DNA (4, 8, 12), were incubated with egg cytosol
(EC) (lanes 1-4), egg cytosol followed by NPE (lanes 5-8), or egg
cytosol followed by NPE containing 1M p27Kip (lanes 9-12). The NPE
contained aphidicolin. Chromatin-bound proteins were analyzed by
Western blotting using MCM3 antibody (upper panel) or a mixture of
ORC2 and Cdc45 antibodies (lower panel).
Fig. 5. Unlike binding of MCM2-7, chromatin binding of Cdc45 is
rate-limiting for DNA replication. A, sperm chromatin (10,000/l)
was incubated with egg cytosol containing (lane 3) or lacking
(lanes 1, 2, and 4) geminin and then supplemented with 2volumes of
NPE containing 50g/ml aphidicolin (lanes 2-4) and p27Kip (lane
4only). Immediately before (lane 1), or 20min after the addition of
NPE (lanes 2-4), sperm chromatin was isolated, and the equivalent
of 10,000sperm was analyzed on a Western blot using MCM3 antibodies
(top panel) or a mixture of ORC2 and Cdc45 antibodies (bottom
panel). The Western blot also contained purified proteins (lanes
5-12). Lane 5, 0.4ng of ORC2, 0.6ng of MCM3; lane 6, 0.4ng of ORC2,
1.8ng of MCM3: lane 7, 0.4ng of ORC2, 6ng of MCM3; lane 8, 0.4ng of
ORC2, 18ng of MCM3; lane 9, 0.4ng of Cdc45; lane 10, 1.2ng of
Cdc45; lane 11, 4ng of Cdc45; lane 12, 12ng of Cdc45. B, sperm
chromatin was incubated with 6l of Cdc45-depleted egg cytosol and
then supplemented with 12l of total of mock-depleted and
Cdc45-depleted NPE mixed in the following ratios: 1:0 (triangles),
1:4 (circles), 1:9 (diamonds), 1:19 (squares), 0:1 (filled
squares). 9l of each sample was mixed with [ -32P]dATP and
replication measured 20,40,60,and 80min after NPE addition (graph).
The other 9l of each sample was mixed with aphidicolin, and
chromatin binding of MCM, Cdc45, and RPA was measured 25min after
NPE addition. C, mock-depleted and MCM7-depleted egg cytosol were
mixed in 1:0 (lane 1), 1:1 (lane 2), 1:3 (lane 3), 1:7 (lane 5),
and 1:15 (lane 6) ratios and incubated with sperm chromatin
(10,000/l). Immediately before (top 3panels) or 15min after
addition of NPE (bottom panel), sperm chromatin was isolated, and
bound proteins were analyzed by Western blotting with antibodies
against MCM3, ORC2, or Cdc45. An aliquot of each reaction was
supplemented with [ -32P]dATP and replication measured 60min after
addition of NPE (bar graph).
Fig. 6. All chromatin-bound MCM complexes are phosphorylated in
a Cdc7-dependent fashion. Xenopus sperm chromatin was incubated
with egg cytosol (10,000/l) for 30min and then supplemented with
2volumes of NPE. Aliquots containing 40,000sperm were withdrawn
immediately before (lane 1) or at the indicated times after NPE
addition (lanes 2-7), isolated, and the bound proteins analyzed by
Western blotting using antibodies against MCM4 and MCM7 (top panel)
or ORC2 (bottom panel). Lane 6contained the same sample as lane 1,
and lane 9contained a 30:1 mixture of MCM3 (22.5ng) to ORC2
(0.5ng). Top panel, probed with MCM3 serum; bottom panel, probed
with ORC2 serum.
Fig. 7. Actinomycin D stimulates Cdc45 binding. A, sperm
chromatin was incubated with egg cytosol and then supplemented with
NPE containing buffer (lane 1), 10g/ml actinomycin D (lane 2), or
aphidicolin (lane 3). After 20min, chromatin was isolated and
probed with MCM3 and Cdc45 antibodies. B, sperm chromatin was
incubated with egg cytosol containing (lane 4) or lacking (lanes
1-3 and 5) geminin. Subsequently, NPE containing 10g/ml actinomycin
D (lanes 2, 4, and 5), aphidicolin (lane 3), and p27Kip (lane 5)
was added. After 20min, chromatin was isolated and probed by
Western blotting with antibodies against MCM3 and Cdc45 alongside
10ng of purified MCM3 and 0.66,2.2,and 6.6ng of purified his-Cdc45
(lanes 6-8).
Fig. 8. Model for the initiation of DNA replication. A, ORC,
Cdc6, and Cdt1 stimulate MCM2-7 binding to sites widely distributed
around ORC. B, at the G1/S transition, Cdc7 binds to and
phosphorylates many MCM complexes. C, Cdk2/cyclin E stimulates the
association of Cdc45 with a subset of these MCM complexes.
Activation of the first MCM complexes by Cdc45 may lead to
inactivation of neighboring MCM complexes, thereby restricting
initiation to defined intervals.
Figure 1. A) Schematic illustration of mature HSV-1 virion
showing major structural components. B) Diagram of HSV genome. The
unique long and short segments (UL, US) are shown flanked by repeat
sequences. Genes encoded are shown adjacent to their loci in the
schematic genome; those above the schematic are not essential for
replication in tissue culture. Gene names are color-coded according
to their functions: teal = regulatory; blue = capsid and DNA
packaging; red = envelope; orange = tegument; green = DNA
replication, and black = uncertain function.
Figure 2. Flow chart illustrating the cascade of regulatory
events that results in ordered sequential expression of genes
during HSV-1 lytic infection. As ICP4 and ICP27 are both absolutely
required for gene expression to proceed to the E or L stages, viral
replication can be blocked by deleting one or other of these
essential IE genes.
Figure 3. Schematic summary of the in vivo life cycle of HSV-1.
An epithelial surface, with its innervating sensory neuron is
shown. The nucleus of the neuron has been enlarged to depict the
intranuclear events occurring during lytic and latent
infection.
Figure 4. Schematic summary of the vectors discussed in the
text. The name of each virus is shown to the left of the schematic;
the viruses used in the studies reviewed here are referred to by
name throughout the text. The diagrammatic genomic map of each
vector is aligned with that of the HSV-1 genome in Figure 4A to
facilitate comparison between viruses. Each schematic depicts the
positions and types of foreign transgenes inserted into each
construct, and which subset of viral genes has been
inactivated.
Figure 5. Schematic summary of neurological applications for
HSV-1 vectors. For each region of the nervous system, the possible
disease applications are listed with types and specific examples of
transgenes whose delivery and expression may be beneficial. Data
pertaining to many of these are discussed in the text.
Figure 6. Some applications of HSV vectors in the peripheral
nervous system. A) The cell bodies and central terminals of
transduced afferent nerve fibers are depicted schematically.
Vector-mediated expression of pre-proenkephalin in these cells
results in appropriate processing, transport, and packaging, such
that enkephalin is found at central presynaptic terminals. Several
models indicate that painful stimuli cause enkephalin release,
resulting in the inhibition of pain neurotransmission. Nerve growth
factor (NGF) expression and release can also be demonstrated in
these cells following transduction with an appropriate vector; the
site of action of this agent is uncertain at present. B) Injection
of the rat footpad with formalin results in pain behavior that may
be scored. Following an initial transient nociceptive response,
pain behavior reappears and lasts for approximately 1 hour,
reflecting a more chronic pain mechanism. Pretreatment of rats with
a pre-proenkephalin expressing vector reduces the chronic pain
behavior in this model without affecting the initial nociceptive
response. The asterisks (*) denote results that show a
statistically significant difference between the experimental
(SHPE) and control (SHZ). C) Dorsal root ganglion cells may be
transduced with a replication-defective vector expressing NGF,
which protects the cells of the ganglion against the toxicity of a
hydrogen peroxide challenge. The neuroprotective effect is most
pronounced at 3 days post-transduction when using a construct in
which a viral immediate-early promoter drives NGF expression (SHN).
HSV LAP2-driven transgene expression increases through day 14;
neuroprotection is maximal at this later time point (SLN). The
protective effect is similar in magnitude to that seen when
recombinant NGF is applied to the culture, and is not seen when a
promoterless construct (SN) is used. Stable, chronic expression
from the LAP2 promoter may enable long-term therapeutic transgene
expression in sensory neurons; in other studies, reporter gene
expression has been detected over a year after the initial
transduction event.
Figure 7. Strategies for increasing bystander lysis in
antiglioma suicide gene therapy. In HSV transduced cells, the
pro-drug GCV is activated by viral thymidine kinase to form an
oncolytic monophosphate derivative. The active drug may enter and
kill surrounding cells; this may be enhanced by coexpression of
Cx43 to encourage gap junction formation between transduced cells
and their neighbors. In addition, TNF- secretion from transduced
cells makes the tumor more sensitive to radiation. Both of these
enhancements to TK-GCV suicide gene therapy are effective in
promoting the survival of animals in an in vivo model of malignant
glioma. The Kaplan-Meier survival curves on the right side of the
figure show the survival data from nude mice that have been
intracerebrally inoculated with U87 glioma cells and then treated
with the vectors and drugs shown in the chart legends.
Figure 8. Non-neurological applications of HSV vectors. We are
currently studying ex vivo transduction of stem cells for the
introduction of therapeutic genes or differentiation factors. In
vivo transduction of muscle may allow delivery of dystrophin or
other gene products that are absent in various muscle diseases.
"Depot" tissues, such as adipose and ligament, may be transduced
with genes encoding circulating proteins, with a variety of
potential applications.
Figure 1. A simplified model of the bacterial cell cycle. DNA
(dark gray lines), origin (oriC, gray circles), terminus (terC,
dark gray square), replisome (overlapping triangles, one for each
replication fork), cytokinetic ring (dashed line). In this model,
DNA replication initiates at or near mid cell. The sister origins
rapidly separate from each other and become anchored on opposite
halves of the cell. DNA replication continues followed closely by
refolding of newly replicated DNA until there are two complete and
separate chromosomes. Finally, the cell divides medially. The model
is simplified to ignore multifork replication.
Figure 2. A model of a circular chromosome that is undergoing
multifork replication in a rod-shaped bacterium. The cell contains
four copies of the origin region (gray circles). The initial round
of replication in the middle is duplicating the parental chromosome
(thin black) to generate the gray copies. A second round has begun
at the quarters, duplicating the daughters (gray) to generate the
granddaughters (thin black). The cell contains four copies of the
origin region (gray circles), located approximately at the one-,
three-, five-, and seven-eighth positions along the cell length,
and a single copy of the terminus region (dark gray square) located
near mid cell.
Figure 3. The extrusion-capture model for bacterial chromosome
partitioning. After the origin region is replicated, the two sister
origins (light gray circles) are extruded from (arrows pointing
toward cell poles) the centrally located replisome (overlapping
triangles) and captured on opposite halves of the cell at or near
the cell quarters. The terminus region (dark gray square) remains
at mid cell until it is duplicated.
Figure 4. Terminus-specific chromosome partitioning events. (A)
Chromosome decatenation. (B) If formed, a chromosome dimer is
resolved to two monomers by two site-specific recombinases acting
on the dif site in the terminus region. (C) When chromosomes (gray)
are trapped in an invaginating division septum (black vertical
lines), SpoIIIE (black ovals) pumps the chromosomes out of the way
(arrows above cell indicate direction each chromosome will
move).
Fig. 1. DNA replication vs. homologous recombination.
Chromosomes are shown as double lines. Parental strands are filled;
daughter strands are open. (A) A chromosome. (B) Chromosome
replication has been initiated. (C) Chromosome replication is
nearing completion. (D) Chromosome replication is complete. (E)
Strand degradation in preparation for homologous recombination has
started. (F) Strand degradation is nearing completion, whereas
annealing of the complementary strands is going on.
Fig. 2. The pathways of replication fork stalling/disintegration
with subsequent resetting/repair. DNA duplexes are shown as double
lines; a protein tightly bound to DNA is shown as a bricked circle.
For all Holliday junctions, one of the two possible resolution
directions is indicated by the small arrows. (A) A replication
fork. (B) The replication fork approaching a single-strand
interruption in template DNA. (C) The replication fork has
collapsed at the interruption. (D) Double-strand end invasion to
restore the replication fork structure. (E) A stalled replication
fork. (F) Regression of the stalled replication fork forms a
double-strand end and a Holliday junction. (G) Double-strand end
invasion to restore the replication fork structure. Resolution of
the Holliday junction in F leads to replication fork breakage (C).
Resolution of the Holliday junctions in D or G, or exonucleolytic
degradation of the linear tail in F leads to restoration of the
replication fork structure (A).
Fig. 3. Two ways to repair a replication fork by single-strand
annealing. DNA duplexes are shown as double lines; sister chromatid
cohesion is indicated by thin dumbbells. (A) A replication fork
approaching a single-strand interruption in template DNA. (B) The
replication fork has collapsed. (C) Rad52-promoted reannealing of
the detached end with the complementary single-strand gap on the
full-length chromatid. (D) Repair of the single-strand
interruption. (E) Filling-in the single-strand gap on the
full-length chromatid. Sister chromatid alignment is shown. The
thin arrows indicate a hypothetical signal from the Rad52-bound
double-strand end to the intact sister chromatid. (F)
Rad54-catalyzed unwinding of the intact chromatid in the vicinity
of the double-strand end. (G) Rad52-catalyzed annealing of the
double-strand end with the open sister duplex to generate a
replication fork structure.
Fig. 1. Three systems required for the precise duplication of
chromosomal DNA replication. Top panel, a small segment of DNA
carrying three replication origins passing through the cell cycle.
Red boxes denote ORC and a plus denotes a licensed origin. The
metaphase chromosome decondenses, assembles licensed origins, is
assembled into a nucleus, replicates, and then recondenses. Below
are the activities of the origin recognition system, the licensing
system and SPF during the course of the cell cycle. See text for
further details.
Fig. 2. Events occurring at a replication origin in the Xenopus
early embryo from late mitosis to late G1. (A)Assembly of ORC onto
origin DNA. (B)Binding of Cdc6 and RLF-B/Cdt1 onto ORC. (C)Multiple
Mcm(27) heterohexamers are assembled onto each origin to license
it. (D)Once licensing is complete Cdc6 is removed, whilst ORC binds
less tightly (unshaded ORC denotes weak binding to DNA). (E)Cdc7 is
recruited to the licensed origin and phosphorylates Mcm(27)
complexes (purple circles denote phosphorylation). (F)Following
nuclear assembly, CDK activity induces the assembly of Cdc45 onto
the origin.
Fig. 1. Model of SOS translesion replication by DNA polymerase
V.The two DNA strands are shown as green lines, and the
replication-blocking lesion is represented by the red rectangle.
The three major steps in TLR are pre-initiation (2), in which the
RecA nucleoprotein filaments assembles; initiation (3and 4), which
involves binding of pol V to the primer-template and loading of the
subunit clamp; and lesion bypass by pol V holoenzyme (5). SSB is
suggested to help in displacing RecA from DNA both at the
initiation and lesion bypass steps.
Fig. 1. Pathways for recombinational DNA repair of a stalled
replication fork. A pathway involving fork regression is shown for
gap repair (a-f), and a double-strand break repair path is shown
for the repair of a fork collapsed at the site of a DNA strand
break (g-l). The pathways shown are intended to be generic and do
not incorporate all current ideas for fork repair. The dashed line
represents a pathway in which the Holliday junction is converted to
a double-strand break by the action of RuvABC, as observed by
Michel and colleagues (180) and others. Arrowheads on DNA strands
denote 3' ends. If a DNA lesion (other than a strand break) is
responsible for halting the progress of a replication fork, note
that replication fork repair does not entail repair of the lesion
itself. Instead, the recombination and replication steps set up the
lesion for repair by providing an undamaged complementary DNA
strand. The degree to which excision repair and other DNA repair
processes are integrated with the recombinational repair pathways
is unknown.
Fig. 2. A replication intermediate from Drosophila embryos,
photographed by Inman (49). One of the apparently regressed forks
has a single-stranded tail (arrow). (Reprinted from Biochim.
Biophys. Acta, 783,Inman, R.B., "Methodology for the study of the
effect of drugs on development and DNA replication in Drosophila
melanogaster embryonic tissue," pp. 205-215, Copyright 1984,with
permission from Excerpta Medica, Inc.; ref. 49.)
Figure 1. Models for replication restart by recombination.
(Left) Holliday junction (HJ) formation by fork regression allows
bypass of unreplicatable DNA damage. (Right) A stalled fork can be
restored after breakage by D-loop formation and strand
invasion.
Fig. 1. Recombination repair of broken replication forks. (A)
Rescue of blocked replication forks (adapted from ref. 17). The
replication fork is blocked at the Ter site in the presence of Tus.
A DSB occurs in the lagging-strand template. The RecBCD enzyme
enters at the double-strand end and initiates homologous
recombination catalyzed by RecA. Completion of the recombination
reaction by resolution of the Holliday junction leads to
restoration of a replication fork. Binding of the primosome allows
loading of a new replisome to promote DNA replication restart. To
account for the viability of a strain carrying an ectopic Ter site
in a recombination-proficient background, one needs to assume that
the newly reconstituted replication fork is not arrested again, and
hence that Tus has been removed from Ter during the recombination
reaction. DSB on the lagging strand is shown, but a similar model
can apply to breakage and repair of the leading strand. (B)
Replication fork collapse (adapted from ref. 31). The progressing
fork encounters a single-strand interruption in the leading-strand
template, because of a defect in closure of the lagging strand at
the previous replication round. Reincorporation of the broken DNA
strand by homologous recombination and replication restart are
catalyzed by the same enzymes as on breakage of the fork. The full
lines represent the two DNA strands, the dashed lines represents
newly synthesized DNA strands. The arrowhead corresponds to DNA 3'
ends.
Fig. 2. RuvAB/RecBCD-mediated rescue of blocked replication
forks (adapted from ref. 44). In the first step (A) the replication
fork is blocked and the two newly synthesized strands anneal,
forming a Holliday junction (see Fig. 3 for the different pathways
proposed to promote this step). In a second step (B) the junction
is stabilized by RuvAB binding. (C) In recombination proficient
strains, RecBCD binds to the double-strand tail (C1); degradation
takes place until the first recognized CHI site (CHI is an
octameric sequence that switches RecBCD from an exonuclease to a
recombinase enzyme) and is followed by a genetic exchange mediated
by RecA (C2); RuvC resolves the first Holliday junction bound by
RuvAB (C3). In C2 and C3, the double-strand end is reincorporated
into the circular chromosome by homologous recombination and the
Holliday junction is resolved, which results in the reconstitution
of a replication fork. This pathway is presumably used in
recombination-proficient cells. (D) RecBCD-mediated degradation of
the tail progresses up to the RuvAB-bound Holliday junction.
Replication can restart when RecBCD has displaced the RuvAB
complex. D can take place in recombination-proficient strains if
RecBCD reaches RuvAB before encountering a CHI site; it is the only
pathway that leads to a viable chromosome in recA and ruvC mutants.
(E) RuvC resolves the RuvAB-bound Holliday junction. This pathway
is used in the absence of RecBCD and leads to the RuvABC-dependent
DSBs observed in recBC mutants. Continuous and discontinuous lines
represent the template and the newly synthesized strand of the
chromosome respectively; the arrowheads indicate the 3'end of the
growing strands.
Fig. 3. Models for formation of Holliday junctions at arrested
replication forks by RFR. (A) RecA binds to the single-stranded
region of the lagging-strand template, polymerizing in the 5' to 3'
direction. Pairing of the lagging-strand template with the
leading-strand template renders the leading strand free to anneal
with the 5' end of the lagging strand. This results in the
formation of a Holliday junction that can be bound directly by
RuvAB. (Adapted from ref. 64.) (B) RecG binds to the replication
fork and migrates toward the chromosomal replication origins,
displacing the 5' end of the lagging strand. RecG activity
ultimately creates a four-stranded junction. (Adapted with
modifications from ref. 76.) (C) (+) Topological stress that
accumulates downstream of the fork on arrest is relaxed by
unwinding of the two newly synthesized strands from the template
strands and their annealing. (Schematic representation based on
results in ref. 78.) Full and dashed lines represent the template
and the newly synthesized DNA, respectively; the arrowheads
indicate the 3' end of the growing strands.
Fig. 4. RFR in UV-irradiated cells (adapted from refs. 76, 80,
and 92). The replication fork is blocked by a UV photo-product
(black triangle) in the leading-strand template. RFR, proposed to
be catalyzed by RecG in E. coli (76), or by Rad51 (the yeast RecA
homologue) in S. pombe (80), renders the damaged DNA double
stranded and thereby allows direct repair by nucleotide excision
repair enzymes (A). If the lagging-strand polymerase has continued
synthesis past the lesion, leading-strand DNA synthesis using the
lagging strand as template followed by reverse branch migration
[proposed to be catalyzed by RecG in E. coli (76) and by Rqh in S.
pombe (80)] reconstitutes a fork on which the lesion has been
bypassed (B; ref. 94). Full and dashed lines represent the template
and the newly synthesized DNA, respectively; the arrowhead
indicates the 3' end of the growing strand.
Fig. 1. A scheme to illustrate stalled replication fork
rebuilding by replication fork regression, by using recombination
enzymes in the E. coli chromosome (adapted from refs. 17, 18, 24,
and 25). Pathways that lead to a single crossover (or an odd number
of crossovers) generate dimeric chromosomes, and those that act
without crossing over (or even numbers of crossovers) retain the
monomeric status of the chromosome. Pathways considered to be major
routes to retaining the monomeric chromosome status are overlaid
onto a beige background. Similarly, the arrows bounded by a bold
line are intended to indicate major pathways, with relative
contributions being indicated by arrow breadth. Those arrows in
dark green indicate pathways that would be expected to be
RecABCD-dependent, whereas those in light green indicate RuvC
cleavage from within a RuvABC complex. Arrows in light blue
indicate RuvAB helicase action, whereas that in pink indicates
similar action by RecG or RecQ helicases. Black lines are
unreplicated DNA, and red/pink lines newly replicated daughter
strands. (a) Reannealing of daughter strands at a stalled
replication fork (closed triangle indicates a nontemplate lesion on
the leading strand). Reannealing can be mediated by RecG (18) and
facilitated by positive supercoiling ahead of the fork (27). RuvAB
and RecA may also promote the growth of the reversed fork once fork
reversal has been initiated. Lagging strand synthesis is indicated
as proceeding beyond the lesion, thereby providing an opportunity
to replicate past the lesion by copying of the switched template in
a reaction that uses recombination proteins, but not recombination
(pink line; ref. 18). (b) Productive RuvAB branch migration to
extend the four-way HJ (RuvB is cartooned as a pair of cylinders on
opposed arms of the HJ). Note that RuvB binding to the other two
arms of the junction (step l) will lead to abortion of the four-way
junction by branch migration (step p). (c and m) Action of RuvABC
to cleave the strands 3' of the bound RuvB on the branch point
side, to generate broken forks (corresponding to single strand
lesions in the leading and lagging parental template strands
respectively; ref. 24). RecABCD-mediated reinvasion of the broken
ends leads to rebuilt replication forks that most readily yield
noncrossover (d-f) and crossover (n) chromosomes, respectively.
Note that after reinvasion, a further round of RuvABC action is
required; in each case the "productive" orientation of RuvB binding
gives the majority species shown (f and n). Binding of RuvB in the
abortive configuration either will act to reverse the invasion or
will lead to RuvC cleavage to give the minority products, dimers
and monomers, respectively (k, o). (g-i). RecABCD-mediated invasion
of the end created by fork reversal into its homologous region to
generate a molecule containing two HJs (or a HJ and a three-way
junction). Such an intermediate can be processed by crossover (i)
or noncrossover (h) pathways by several ways that involve the
simultaneous or sequential action of proteins at each of the branch
points; we predict dimers to predominate over monomers in these
events. (j) Processing of the reversed replication fork
intermediate by RecG or RecQ helicases (equivalent to step p,
promoted by RuvAB).
Fig. 2. Alignment of E. coli DnaX, YcaJ, and RuvB (CLUSTAL X
multiple sequence alignment program, v.
1.8,http://www.ebi.ac.uk/clustalw/). The sequences of YcaJ, RuvB,
and DnaX all contain well-conserved nucleotide-binding sites with
Walker A (GxxxxGKT/S) and Walker B (Dexx) motifs. The Zn-binding
motif of DnaX is absent in YcaJ, but the putative ATPase sensor
motifs (29) are present. Colors represent types of amino acids.
Fig. 3. (A) An outline of the Xer recombination reaction. XerCD
bind cooperatively at dif, psi, or cer recombination sites,
ensuring synapsis (with the help of accessory sequences and
proteins in the case of psi/cer) (i). XerC initiates catalysis (ii)
to form a HJ intermediate, which undergoes a conformational change
(iii) to provide a substrate for catalysis by XerD, which can then
complete the recombination reaction (iv). There is normally a
barrier to this conformational change, and XerC frequently
catalyzes the conversion of the HJ back to substrate (ii). In
recombination at psi, the proteins PepA and ArgA-P facilitate the
HJ conformational change, whereas in recombination at dif, FtsKc is
thought to facilitate this change (34, 35). (B, C) Species
specificity of FtsK action. FtsK cells (DS9041) were transformed
with pBAD expression vectors (48) carrying full-length FtsK
proteins (B) or the C-terminal domains (C) of different species. To
assay for Xer recombination, they were transformed with a plasmid
containing two dif sites and grown in conditions of repression ( ;
0.2% glucose) or induction (+; 0.2% arabinose) of the expression
vectors (34). Induction was checked by Western blot analysis by
using an antibody directed against a FLAG epitope fused to the N
termini of the constructs, after resolution of the protein extracts
on a 6% (B) or an 8% (C) SDS/PAGE. (D) Alignment of the C-terminal
domains of FtsK homologues. Identical residues are indicated by
stars, conservative substitutions by dots. Open boxes underline
regions predicted to adopt an -helix conformation by PREDICTPROTEIN
PHD software v.
1.96,http://www.embl-heidelberg.de/predictprotein/predictprotein.html
whereas black arrows underline those predicted to form sheets. Ec:
E. coli FtsK, Hi: H. influenzae FtsK and Bs: B. subtilis
SpoIIIE.
Fig. 4. The C-terminal domain of FtsK is randomly distributed
throughout the cytoplasm. FtsK cells (DS9041) were transformed with
pBAD expression vectors carrying an N-terminal fusion of the green
fluorescent protein (GFP) to full-length FtsK or to the C-terminal
domain of FtsK (FtsKc). Cells were grown to midexponential phase in
LB supplemented with 0.1% (full-length) or 0.2% (FtsKc) arabinose.
Nucleoids were stained by using 4',6-diamidino-2-phenylindole
(DAPI). Phase-contrast and fluorescent images were acquired by
using a cooled charge-coupled device camera (Princeton Instruments,
Trenton, NJ) and METAMORPH IMAGE ACQUISITION software (Universal
Imaging, Media, PA) from an Olympus BX50 (New Hyde Park, NY)
fluorescence microscope. Shown are overlays of the DAPI image in
red and the GFP image in green. Full-length FtsK frequently
localizes to the septum, whereas FtsKc is always distributed
throughout the cytosol and never found at the septum.
Fig. 5. A model for chromosome segregation in E. coli. In
contrast to the replication origins (large dark gray circles), the
replication terminus region, which contains the dif site (open
triangle), stays localized at midcell (49). As a consequence,
sister dif sites can be synapsed by the XerCD recombinases (small
white and light gray circles) whether the chromosomes form a dimer
or not. This leads to cycles of HJ formation and resolution by
XerC. If the chromosomes are monomeric, segregation will eventually
break the synaptic complex and move the sites away from midcell
before septum closure. If the chromosomes form a dimer, the
synaptic complex will stay trapped at midcell, which allows access
to FtsK. FtsK mediates the HJ conformation change needed to
activate catalysis by XerD, thus coordinating resolution of
chromosome dimers to cell division.
Fig. 1. Conversion of plectonemic supercoils into knot and
catenane nodes by Int site-specific recombination. (A) A ( )
supercoiled Int substrate (black line) is shown with the att
recombination sites represented by red and blue arrows. When the
att sites are in inverse (head-to-head) orientation in the primary
sequence, the recombination products are right-handed torus knots;
these knots can be drawn without crossings on the surface of a
torus- or doughnut-shaped object (Upper). When the sites are
directly (head-to-tail) repeated, the product is a right-handed
torus catenane (Lower). (B) An electronmicrograph of a 13-noded Int
knot produced in vitro. The DNA was coated with RecA protein to
help visualize the crossings. (Reprinted from Cell 276,Spergler,
S.J., Stasiak, A.&Cozzarelli, N.R., "The stereostructure of
knots and catenanes produced by phage lambda integrative
recombination: 1985,Implications for mechanism and DNA structure,"
325-334, 1985,with permission from Elsevier Science; ref. 15.)
Fig. 2. Monte Carlo simulations of catenanes between relaxed and
supercoiled DNA molecules. Simulation of a singly linked catenane
between two relaxed plasmids (A) or between a relaxed and
supercoiled plasmid (B). The yellow chain represents a 3.5-kb DNA,
and the red chain a 7-kb DNA. Simulations courtesy of Alexander
Vologodskii.
Fig. 3. Conformations of replicating DNA. (A) A replication fork
is depicted at Left with the parental strands in black and the
daughter strands in red. Red arrows denote 3' ends. During
replication, the denaturation of the parental duplex causes a (+)
Lk in the replicating molecule (Center). This (+) Lk can be
expressed either as (+) supercoiling of the parental duplex in
front of the replication fork or (+) precatenanes between the
replicated duplexes behind the fork. We have shown the (+) Lk in
the usual fashion, which assumes that it is initially ahead of the
fork and must diffuse past the replisome to generate precatenanes.
It is, however, possible that the converse is true and that (+)
precatenanes are the primary consequence of the (+) Lk from
replication. Upon replisome dissociation (Right), the ends of the
nascent strands will be free to base pair with each other, forming
a four-way junction at the replication fork, the chickenfoot, that
allows the replication fork to regress until the molecule is
relaxed. (B) Electron micrograph of an in vitro replication
intermediate with replication stalled by the Tus/ter complex. The
molecule displays both supercoils in the unreplicated region (thick
line) and precatenanes in the replicated region (thin line).
(Reprinted from ref. 47, with permission from Elsevier Science.)
(C) Scanning force microscopy of an in vivo replication
intermediate incubated in ethidium bromide. This molecule displays
the linear duplexes of the middle toe of the chickenfoot (white
arrows) emerging from both the unidirectional origin of replication
and the terminus. (Reprinted from ref. 60, with permission from the
American Society for Biochemistry and Molecular Biology.) (For B
and C, the scale bar is 100nm.)
Fig. 4. Physiological implications of the chickenfoot.
Replication forks are shown with parental strands in black and
daughter strands in red, and with the 3' end tipped with an
arrowhead. (A) Recombination-mediated replication restart. The
four-way junction of the chickenfoot can be cleaved by Holliday
junction resolving enzymes such as RuvC (white arrowheads) [1].
Cleavage will sever one of the replicated arms, which can then be
processed by the RecBCD complex (Pac-man) [2], allowing it to
become a substrate for homologous recombination [3]. Recombination
with the sister replicated arm re-forms the replication fork,
allowing replication restart [4]. (B) Bypass of a lesion. An
unpaired lesion (black rectangle) on the parental strand blocks
leading strand replication, but lagging strand replication can
continue for more than 500nt [1]. Upon replisome dissociation, the
two nascent strands can base pair, requiring the denaturation of
more than 500bp at the lagging strand [2]. Once a chickenfoot is
formed, the lagging strand becomes a template for the leading
strand, allowing leading strand replication to continue past the
lesion [3]. Reabsorption of the chickenfoot re-forms the
replication fork, allowing replication to continue and a second
opportunity to repair the lesion [4].
Fig. 5. Model for topology of the replicating chromosome. A
segment of chromosomal DNA is depicted, with black lines as
parental strands and red lines as nascent strands. In the bacterial
chromosome, domain barriers (yellow boxes) isolate the topology
around the fork from the rest of the chromosome, which is ( )
supercoiled by DNA gyrase, as shown in the domains on either side
of the replication domain. Replication creates a (+) Lk in the
replicating domain (Center), which can cause (+) supercoils ahead
of the fork and (+) precatenanes behind it. Thus, either gyrase or
topo IV could support replication by removing (+) supercoils in
front of the fork, and topo IV could also support replication by
removing precatenanes behind the replication fork.
Fig. 1. Resolution of Holliday junctions in early replication
intermediates can generate a number of different products. (i) The
ERI. (ii) An ERI in which one end (the origin-proximal end) of the
nascent DNA has regressed. This could happen with equal probability
at the other end as well. (iii) Cleavage of the Holliday junction
in ii generates an structure. (iv) An ERI in which both ends of the
nascent DNA have regressed. Because resolution of each Holliday
junction can occur in one of two ways, two sets of products are
generated: a nicked circle (form II) and a short duplex DNA
corresponding to the distance on the replicated portion of the
template that is between the sites of resolution (which will vary
depending on the amount of nascent DNA that has regressed) (v); and
a linear molecule that is longer than the original template by the
distance on the template that is between the sites of resolution
(vi). Red, the leading-strand template; blue, the lagging-strand
template; green, nascent DNA.
Fig. 2. Both RusA and RuvC can cleave ERIs. Standard oriC
replication reactions containing either RusA (lanes 2-5,
concentration varied by a factor of 10in each lane, left to right),
RuvC (lanes 7-10, concentration varied by a factor of 10in each
lane, left to right), or having no addition (lanes 1and 6) were
incubated at 37C for 5min. The reactions then were terminated by
the addition of EDTA, and the DNA products then were analyzed by
electrophoresis through either a neutral 1% agarose gel (A) or a
denaturing 1% alkaline agarose gel (B) as described in Materials
and Methods. , structure; FII, form II (nicked, circular DNA). The
mobility difference between the ERI and the structure is subtle and
can be seen more clearly in Figs. 5-7.
Fig. 3. RuvA inhibits cleavage of the ERI by RusA. Standard oriC
replication reactions containing either 10nM RusA (lane 2), 10nM
RusA and RuvA (lanes 3-5, RuvA concentration varied by a factor of
10in each lane from left to right), 100nM RuvA (lane 6), or having
no addition (lane 1) were incubated first for 2min at 37C after
RuvA addition and then for an additional 5min at 37C after RusA
addition. The reactions were terminated by the addition of EDTA,
and the DNA products then were analyzed by electrophoresis through
a neutral 1% agarose gel.
Fig. 4. Holliday junctions form in inactivated ERI. Inactivated
ERI was formed as follows. Nine standard reactions for ERI
formation as described in Materials and Methods were incubated at
37C for 10min. The reactions then were terminated by heating to 65C
for 5min. After cooling, the reactions were pooled together. Nine
and one-half microliters of the inactivated ERI pool was incubated
in new reaction mixtures (10l) containing either no RuvA or RusA
(lane 1), RusA (lanes 2-7, concentration in lanes 2-4 varied by a
factor of 10from left to right and was 100nM in lanes 5-7), and
RuvA (lanes 5-8, concentration varied by a factor of 10in lanes 5-7
from left to right and was 500nM in lane 8) and incubated for 2min
at 37C after RuvA addition and for an additional 5min at 37C after
RusA addition. The reactions then were terminated by the addition
of EDTA, and the DNA products were analyzed by neutral 1% agarose
gel electrophoresis. F III', form III' (see text for definition); F
III, form III (full length, linear form).
Fig. 5. Negative supercoiling inhibits RusA cleavage of the ERI.
Inactivated ERI was formed as in the legend to Fig. 4 except that
12reaction mixtures were pooled. Fresh ATP was added to this pool
to a concentration of 2mM. Nine microliters of the inactivated ERI
pool was incubated in new reaction mixtures (10l) containing either
no RusA (lanes 1,7,and 12) or 100nM RusA (all other lanes), and
either Topo IV (lanes 3-7, concentration varied by a factor of 5in
lanes 3to 6from left to right and was 25nM in lane 7) or DNA gyrase
(lanes 8-12, concentration varied by a factor of 5in lanes 7-11
from left to right and was 25nM in lane 12) for 5min at 37C. The
reactions were terminated by the addition of EDTA, and the DNA
products were analyzed by neutral 1% agarose gel electrophoresis.
The autoradiogram of the gel is shown in A and a photograph of the
ethidium bromide stain gel is shown in B. FI, negatively
supercoiled template DNA; F I', form I' (intact template DNA with
no supercoils); SC, supercoiled ERI.
Fig. 6. RecG stimulates RusA-catalyzed cleavage of the ERI.
Inactivated ERI was formed as in the legend to Fig. 4 except that
six reaction mixtures were pooled. Fresh ATP was added to this pool
to a concentration of 2mM. Six and three-tenths microliters of the
inactivated ERI pool was then incubated in new reaction mixtures
(7l) containing either no RecG or RusA (lane 1), RecG (lanes 2-4
and 6-8, concentration varied by a factor of 10in lanes 2-4 and
6-8, left to right), and 10nM RusA (lanes 5-8) for 5min at 37C. The
reactions were terminated by the addition of EDTA, and the DNA
products were analyzed by neutral 1% agarose gel
electrophoresis.
Fig. 7. RecG promotes nascent strand regression in negatively
supercoiled ERI. Inactivated ERI was formed as in the legend to
Fig. 4 except that 12reaction mixtures were pooled. Fresh ATP was
added to this pool to a concentration of 2mM. Twelve and
three-quarter microliters of the inactivated ERI pool was then
incubated in new reaction mixtures (15l) containing DNA gyrase
(25nM), RecG (10nM), and RusA (100nM) as indicated for 5min at 37C.
The reactions were then terminated by the addition of EDTA.
One-half of the reaction mixture was analyzed by neutral 1% agarose
gel electrophoresis [autoradiogram (A); photograph of the ethidium
bromide-stained gel (C)] and the other half was analyzed by
denaturing 1% alkaline agarose gel electrophoresis [autoradiogram
(B)]. The arrow on the right-hand side of B marks a faint band that
may correspond to the linear fragment formed by cleavage of the ERI
at both ends of the nascent DNA (see Fig. 1v).
Fig. 1. Replication of Mu by transposition. In the first stage,
the phage-encoded transposition proteins aided by the histone-like
protein HU promote transfer of 3'-OH ends of miniMu (red) to each
strand of target DNA (green). Two sites, 5bp apart on target DNA,
that will be subjected to a nucleophilic attack by each Mu end are
indicated by arrows. Strand exchange produces a fork at each Mu
end, the target providing 3'-OH ends (indicated by half arrows)
that can potentially serve a primers for leading strand synthesis.
MuA transposase, which has been assembled into an oligomeric
transpososome, remains tightly bound to both Mu ends in the strand
exchange product (strand transfer complex, STC1). Host factors then
initiate Mu DNA synthesis from one end to duplicate Mu and form the
final cointegrate product. The DNA synthesis phase was initially
reconstituted in an eight-protein system supplemented with
partially purified host factors (MRF), as described in the
text.
Fig. 2. Components of the MRF. MRF was originally identified as
host factors needed in addition to the eight-protein system to
convert STC1 to cointegrates. Resolution of MRF into enzyme
fractions distinguishable by function and into pure components
(ClpX, PriA, PriB, and DnaT) is indicated.
Fig. 3. Transition from transpososome to replisome. The
molecular chaperone ClpX converts STC1 (A) to STC2 (B), altering
the conformation of the transpososome. MRF 2 then displaces the
transpososome to assemble the prereplisome at the Mu forks, forming
STC3 (C). PriA binds to the forked DNA structure created by strand
exchange (D) and begins the process of assembling a replisome at
one Mu end. The mechanism that determines which Mu end is used to
initiate DNA synthesis is not yet clear. PriA assembles a
preprimosome complex by recruiting PriB, DnaT, and the DnaB-DnaC
complex (E). In this process, DnaB must be bound to single-stranded
lagging strand template. To create this binding site, PriA unwinds
duplex DNA by translocating 3' to 5' along this template. Once
bound to DNA, DnaB attracts primase to form a primosome, which
catalyzes primer synthesis for lagging strand synthesis, and DnaB
promotes binding of the DNA pol III holoenzyme to complete
replisome assembly.
Fig. 4. Fragile property of the STC2 transpososome. Formation of
STC1, its conversion to STC2, crosslinking of the transpososome
with DSS, and detection of crosslinked MuA by Western blot analysis
was conducted as previously described (63). Lane 1: As control, the
strand exchange reaction mixture was incubated without HU protein,
conditions which do not permit transpososome assembly, and then MuA
was subjected to crosslinking with DSS. Lanes 2-6: STC1 was
isolated free of unbound proteins by filtration through a Bio-Gel
(Bio-Rad) A-15 m column equilibrated with 25mM Hepes-KOH (pH 7.5),
12mM magnesium acetate, and 60mM KCl. Isolated complexes were
incubated at 37C for 30min in the presence of ATP, ClpX being
included for conversion to STC2. The reaction mixture was adjusted
to 300or 500mM NaCl, as indicated, and allowed to stand at room
temperature for 15min before addition of DSS. For lane 2,STC1 was
not subjected to DSS treatment as control.
Fig. 5. MRF 2 consists of at least two distinct components. The
reconstituted Mu DNA replication reaction (50l) with [ -32P]dNTPs
was assembled with ClpX, the 12-protein system, and the indicated
MRF 2 components, and products were resolved by alkaline agarose
electrophoresis as previously described (75). Crude MRF (fraction
II) (30) and MRF 2 (fraction III) (63) were prepared as described.
Resolution of MRF 2 into two components, MRF 2A and MRF 2B, will be
described in a future publication (V.D. and H.N., unpublished
work). Approximately 10units (63) of the indicated MRF 2 components
were added. Where indicated, MRF 2A and MRF 2B were heated at 65and
100C, respectively, for 10min. In the reaction catalyzed with crude
MRF, greater than 95% of STC1 was converted to cointegrate. When
MRF 2 was supplied as two components, typically 50-95% of STC1 was
converted to a cointegrate. CO, position of the cointegrate; STC,
position of the strand exchange product (not radiolabeled and
therefore not visible).
Fig. 1. Alternative BIR mechanisms. A broken chromosome end will
be resected by 5' to 3' exonucleases, allowing the 3' end to
interact with various recombination proteins to carry out strand
invasion. (A) The 3' end of the invading strand initiates DNA
replication, leading to a migrating D-loop "bubble" as described by
Formosa and Alberts (5). the displaced newly synthesized DNA strand
can then be made double-stranded. (B) Strand invasion sets up a
replication fork that will result in semiconservatively synthesized
molecules. A Holliday junction will be resolved at some point. (C)
Strand invasion sets up a replication fork in which branch
migration enzymes displace both newly synthesized DNA strands as
the replication structure migrates down the template.
Fig. 2. BIR-dependent formation of LEU2 recombinants. (A)
EcoRI-digested plasmid pWYL37 has homology only at one end to sites
in the yeast genome. (B) The "LEU" segment at one end of the DSB
may initiate new DNA synthesis, but the completion of the event
requires that the newly synthesized DNA is displaced from the
template and must rejoin to the other end of the DSB by a
nonhomologous end-joining event. (C) A
