Molecular Life Sciences
DOI 10.1007/978-1-4614-6436-5_132-1
# Springer Science+Business Media New York 2014
Cycling of the Lagging Strand Replicase During Okazaki Fragment
Synthesis
Charles McHenry*
Chemistry and Biochemistry, University of Colorado at Boulder, Boulder, CO, USA
Synopsis
The E. coli replicase has enormous processivity, sufficient to synthesize over 100,000 bases without
dissociation. Yet, on the lagging strand of the replication fork, Okazaki fragments of 1,000–2,000
bases are made every 1–2 s. This requires a mechanism to trigger the lagging strand polymerase to
cycle in far less than one second to new primers synthesized at the replication fork. Several
mechanisms have been proposed to explain how this might occur. The most prominent have been
the collision model where a direct collision with the 50 -end of the preceding Okazaki fragment
triggers cycling and the signaling model where synthesis of a new primer at the fork triggers
polymerase release and rebinding to the new primer. Kinetic studies indicate release after collision
is far too slow to support the rate of lagging strand synthesis. Experimental support for the signaling
model has been obtained on synthetic mini-circle templates that contain high asymmetry in GC
composition between the two strands, allowing the rate of lagging strand synthesis to be selectively
perturbed. Furthermore, cycling can be induced by addition of exogenous primers, indicating that it
is the presence of a new primer, not the action of primase, that provides the signal.
Introduction
The Pol III HE has the processivity required to replicate >150 kb (Mok and Marians 1987a, b) and
perhaps the entire E. coli chromosome, without dissociation, yet it must be able to efficiently cycle to
the next primer synthesized at the replication fork upon the completion of each Okazaki fragment at
a rate faster than Okazaki fragment production. The rate of replication fork progression in E. coli is
about 600 nt/s at 30 C (Breier et al. 2005), approximately the rate of replicase progression on singlestranded templates (Johanson and McHenry 1982). Thus, most of the time in an Okazaki fragment
cycle is spent on elongation and little time (0.1 s) remains for the holoenzyme to release, bind the
next primer, and begin synthesis. A processivity switch must be present to increase the off-rate of the
lagging strand polymerase by several orders of magnitude.
There are two competing but nonexclusive models for the signal that throw the processivity
switch. The first (signaling model) proposes that a signal is provided by synthesis of a new primer at
the replication fork that induces the lagging strand polymerase to dissociate, even if the Okazaki
fragment has not been completed (Wu et al. 1992b). The second (collision model) was originally
proposed for T4 (Alberts et al. 1983) and then extended to the E. coli system (Leu et al. 2003). The
collision model posits that the lagging strand polymerase replicates to the last nucleotide (Leu
et al. 2003) or until the Okazaki fragment is nearly complete (Georgescu et al. 2009).
A communication circuit that proceeds through the t subunit has been proposed to sense the
conversion of a gap to a nick, signaling release.
*Email: [email protected]
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# Springer Science+Business Media New York 2014
A third model for cycling has been proposed that attributed dissociation of the lagging strand
polymerase induced by the inability of the dimeric polymerase to rotate around the template strand
once per helical turn, resulting in highly supercoiled product (Kurth et al. 2013). Shortly after the
discovery that t dimerizes the leading and lagging strand polymerases, it was recognized that
negative superhelical torque could be an issue for the leading strand, but that rotation within
single-stranded DNA would rapidly dissipate superhelical tension in the lagging strand (McHenry
et al. 1988). However, Marians observed that coupled leading and lagging strand rolling circle
replication could generate leading strand products of approximately 150,000 bases without the
addition of topoisomerases (Mok and Marians 1987a, b). This suggested that if this torque was
generated, it could be relieved by mechanisms that did not involve topoisomerase action. Cozzarelli
and colleagues considered the issue in more detail (Ullsperger et al. 1995) and discounted the notion
of extensive precatenane interwinding of the leading and lagging strand product created by rotation
of the lagging strand polymerase about the leading strand to remove the torque. As one plausible
solution, they proposed the 30 -end of the growing leading strand could occasionally be released from
the active site of the polymerase to allow dissipation of superhelicity (Ullsperger et al. 1995). It
would appear that the ring-like structure of the b sliding clamp might allow rotation of the helix
through its central pore without dissociation, maintaining processive leading strand synthesis.
Another possible mechanism for torque release would be dissipation of leading strand torque by
release of the lagging strand polymerase during cycling to the primer for synthesis of the next
Okazaki fragment. This mechanism would appear cumbersome, but not impossible, because of the
large mass and radius of gyration of the Pol III HE in the viscous milieu of the cell. However, if
cycling involves DnaX as the sensor that initiates cycling (section Cycling of the Lagging Strand
Polymerase in E. coli is Directed Exclusively by a Modified Signaling Model), it would be attached
to the priming apparatus, precluding this mechanism.
Evidence that a modification of the signaling model is exclusively used to drive cycling in E. coli
is presented in the last section of this essay.
Please note the inconsistency in how an internal cross-reference was handled on page 2 and 4. Either include
the solid triangle from both or exclude in both. If it is author's choice, we suggest inclusion.
Cycling Models Provided by Bacteriophages T4 and T7
In the more fully characterized systems provided by the replication apparatus of bacteriophages T4
and T7, signaling through synthesis or the availability of a new primer appears to play an important
role, with the collision pathway playing a backup role (Hamdan et al. 2009). A handoff of the
nascent tetraribonucleotide primer and the T7 polymerase is effected by direct primase-polymerase
interaction (Kato et al. 2004). It takes time for synthesis of a new primer, release of the lagging strand
polymerase, and initiation of synthesis on a new primer. Differing views have been presented on
how T7 overcomes this delay on the lagging strand to allow leading and lagging strand replication to
remain coordinated (Lee et al. 2006; Pandey et al. 2009). In one model, it is proposed that the
helicase and thus leading strand synthesis is halted during slow primer synthesis (Lee et al. 2006). In
the other model, it is proposed that the lagging strand primer is synthesized before it is needed and
held in a priming loop, close to the replication fork, facilitating handoff to the DNA polymerase
(Pandey et al. 2009). This model, if generally applicable, could explain the double loops sometimes
observed at replication forks by electron microscopy (Chastain et al. 2003). In the latter model, it is
proposed that the reactions remain coordinated because the lagging strand polymerase elongates
faster than the leading strand polymerase (Pandey et al. 2009).
In T4, primer synthesis does not halt progression of the leading strand polymerase. Primer
synthesis occurs by two mechanisms: (1) dissociative, with primase releasing from its helicase
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association during primer synthesis, or (2) processive, whereby primase remains associated and a
second loop is formed on the lagging strand DNA (Manosas et al. 2009; Yang et al. 2006). The
sliding clamp and clamp loader increase the processive/looping mechanism and it was suggested
that gp32 (T4 SSB) might further increase the processive looping mechanism in the natural system
by facilitating handoff of the nascent pentaribonucleotide primer (Manosas et al. 2009). A proposal
was made that the clamp loader/clamp interaction with a new primer might be the key signal required
for release of the lagging strand polymerase (Yang et al. 2004). More recent work supports that
proposal (Chen et al. 2013). The size distribution of short Okazaki fragments that derive from
signaling-induced cycling are affected by concentrations of the clamp loader while independent of
polymerase concentration, as expected for a processive, recycled lagging strand polymerase.
Does the OB Fold Provide the Processivity Switch for Cycling?
In the structure of Pol III a complexed to DNA, an OB fold is located close to the primer terminus
(Wing et al. 2008). Because OB folds commonly bind to ssDNA, a proposal was made that it could
be part of the sensing network (Bailey et al. 2006; Wing et al. 2008). Consistent with this hypothesis,
the ssDNA binding portion of Pol III was localized to a C-terminal region of a that contains the OB
fold element (McCauley et al. 2008). A test of the importance of the OB fold motif was made using a
mutant in which three basic residues located in the b1-b2 loop were changed to serine (Georgescu
et al. 2009). No ssDNA binding was observed in the mutant, indicating diminution in affinity.
However, even the wild-type polymerase bound ssDNA extremely weakly, near the limit of
detection in the assays used (Kd 8 mM).
The processivity of the mutant polymerase was decreased by the b1-b2 loop mutations, an effect
that was rescued by the presence of the t-complex (Georgescu et al. 2009). The latter observation
would seem to suggest that although the OB fold contributes to ssDNA affinity and processivity, it is
not the processivity sensor or at least that the residues mutated are not the key interactors. The OB
fold might bind to the nick generated by completion of an Okazaki fragment, as seen in human ligase
1 (Pascal et al. 2004), inducing a non-processive conformation. Alternatively, the OB fold might act
in concert with other binding changes as part of a more complex signaling network.
It is possible that the entire polymerase active site is the processivity switch. Steitz and colleagues
(Wing et al. 2008) have elegantly demonstrated a conformational change in a induced by substrate
binding in which the movement of several elements places the b2 binding domain in a position where
it can productively interact with the b2 clamp on DNA. Follow-on studies with a Gram-positive
polymerase suggest this observation is general (Evans et al. 2008; Wing 2010). The geometry and
spatial constraints around the active site when the exiting template is double-stranded might make
insertion of the last nucleotide energetically unfavorable. Upon insertion, the product might lose
affinity for the active site, triggering a reversal of the conformational changes that occurred upon
primer-template and dNTP binding, causing the b2 binding domain to be pulled away, and switching
the polymerase to a low processivity mode. The presence of an unliganded polymerase domain
serves to decrease the affinity of the C-terminus of Pol III a for b2 (Kim and McHenry 1996),
consistent with this view. The genetic screen for mutants that led to a loss of the dominant-negative
phenotype of D403E a (▶ DNA Polymerase III Structure, ▶ C-terminal Domains, ▶ t-binding
Domain) revealed a mutant at a conserved alanine in the interior of the b2 binding domain (A887E)
that no longer binds b2 tightly. Future studies should use this mutant to address whether the
communication circuit that modulates b2 affinity flows through this region. Perhaps a bulky residue
changes the presentation of the b2 binding loop or b2 binding surface, decreasing affinity and, if
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DOI 10.1007/978-1-4614-6436-5_132-1
# Springer Science+Business Media New York 2014
relevant to cycling, the dissociation rate. This type of cycling, induced by collision with a downstream 50 -end, would be relevant for the release of Pol III HE from repaired DNA, but not at the
replication fork during Okazaki fragment synthesis, because the release is too slow (section ▶ The
Rate of Polylmerase Dissociation Upon Collision with the Preceding Primer Is Too Slow to Support
Okazaki Fragment Synthesis).
Please note the inconsistency in how an internal cross-reference was handled on page 2 and 4. Either
include the solid triangle from both or exclude in both. If it is author's choice, we suggest inclusion.
What Is t’s Role in Cycling?
It has been proposed that t acts as a sensor for conversion of a gap to a nick upon completion of
Okazaki fragment synthesis (Leu et al. 2003; Lopez de Saro et al. 2003a, b). When bound to ssDNA,
t was proposed to lose contact with the C-terminus of a, leaving the C-terminus free to contact b.
When the t-ssDNA contact is lost, t was proposed to bind the C-terminus of a, displacing b and
allowing the polymerase to cycle to the next Okazaki fragment (Leu et al. 2003; Lopez de Saro
et al. 2003a, b). This model did not consider earlier data that showed that t and b2 did not compete
significantly for polymerase binding and that the critical binding site for b2 was internal and not at
the extreme C-terminus of a (Dalrymple et al. 2001; Kim and McHenry 1996; Wijffels et al. 2004).
Follow-up work rigorously confirmed that the internal b2 site is the one required for processive
replication (Dohrmann and McHenry 2005). Interestingly, replacement of the internal b2 binding
site with the consensus sequence identified by informatics (Dalrymple et al. 2001) increased the
affinity 120-fold, while the same change at the C-terminus had no effect on b2 binding but caused a
2,700-fold decrease in t binding (Dohrmann and McHenry 2005). This suggests that either the
internal site provides the b-interaction sequence in a unique conformation or additional local
contacts dictate the specificity of binding.
While it is possible that the C-terminus of a could interact with b2 for some undiscovered ancillary
purpose, all current mutational effects can be attributed to defects in t interaction or minor structural
perturbations. Recent functional and structural studies reveal only participation of the internal
Pol IIIa b binding site and occlusion of the C-terminal site (Jergic et al. 2013; Liu et al. 2013;
Ozawa et al. 2013; Toste et al. 2013). Furthermore, a consensus b2 binding motif is not found at the
C-terminus of many replicative DnaEs, whereas the internal sequence is conserved (Kurth
et al. 2013).
The function of t in enabling rapid cycling remains unclear. The earliest observation pertaining to
a possible role of t in cycling was made by Marians and colleagues (Wu et al. 1992a). They saw a
t-dependent acceleration that was codependent upon primase and, along with an observed lessening
of the proportion of short Okazaki fragments, proposed t accelerated transit from an old Okazaki
fragment to a new one (Wu et al. 1992a). Proposals for t being the sensor for gap to nick conversion
were derived from equilibrium measurements in which t decreased the affinity for a nick relative to a
gap (Leu et al. 2003). Photo-cross-linking experiments performed using diazirine side chains at a
large number of positions in the template ahead of the polymerase and in positions within a duplex
with which the elongating polymerase collides only detected a cross-links – none to t (Dohrmann
et al. 2011). Because irradiation of diazirines generates carbenes that will insert into any amino acid,
one can interpret not seeing t cross-links with confidence, eliminating the possibility that t directly
senses gap to nick conversion.
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The Rate of Polymerase Dissociation Upon Collision with the Preceding
Primer Is Too Slow to Support Okazaki Fragment Synthesis
Most of the measurements assessing pathways to trigger polymerase release and cycling have been
performed using equilibrium measurements. While a decreased affinity might be consistent with a
role in accelerating release, the real issue is whether the lagging strand can be triggered to release in
0.1 s or less upon collision with the preceding Okazaki fragment’s 50 -end.
This issue was pursued using a surface plasmon resonance assay in which immobilized primers
are placed a set distance from an oligonucleotide that models the 50 -end of the preceding Okazaki
fragment. It was found that filling in a gap to the final nucleotide accelerates release, but the rate is far
too slow to support the physiological rate of Okazaki fragment synthesis (Dohrmann
et al. 2011) – approximately 2–3 min rather than <0.1 s as expected from the requirements for a
physiological rate of Okazaki fragment synthesis. No requirement for t for the acceleration of
release upon gap filling was observed (Dohrmann et al. 2011).
Primase-Pol III Handoff
change dash to colon
A three-point switch model has been proposed (Yuzhakov et al. 1999) whereby Pol III HE does not
have access to primers synthesized by primase unless the w subunit of the Pol III HE first contacts the
SSB subunit to which primase is bound, permitting displacement of primase from the nascent
primer. This model was based largely on the behavior of the protein encoded by the temperaturesensitive ssb-113 mutant. In the presence of SSB-113, w was necessary for synthesis on templates
only when primers were synthesized by primase – not templates where synthetic primers were
provided. The report generalizes the finding to the replication fork by showing that lagging strand
synthesis is lost without w, perhaps because of its inability to use primers synthesized by primase.
However, it does not demonstrate directly one obvious prediction of the model – that replication
proceeding from primase-synthesized primers on single-stranded templates in the presence of wildtype SSB requires w. The model needs to be reexamined to resolve an inconsistency with experimental observations: replication of primase-primed M13Gori does not require w, provided adequate
d-d’ is present to saturate DnaX and salt is maintained at moderate levels (Fig. 1 at low potassium
glutamate concentrations in reference (Glover and McHenry 1998)). (Salt concentrations are not
directly relevant to the primase issue, but wc becomes essential at high salt because wc-SSB
interactions stabilize holoenzyme-DNA complexes at high salt regardless of the source of the
primer.) The reported experimental observations with SSB-113 are undoubtedly valid, but some
of the effects may reflect a gain-of-function defect in SSB-113. That is, SSB-113 causes a problem
that requires w to overcome it.
Pertinent to this issue, the availability of mutant forms of w that specifically interrupted SSB
interaction permitted testing of the functional consequences in reconstituted replication fork assays
(Marceau et al. 2011). The mutations had no effect on lagging strand synthesis other than a slight
shortening of Okazaki fragment length. The most profound effect was a marked decrease in leading
strand synthesis, consistent with a role of a network of interactions that stabilizes the leading strand
polymerase through interaction with the SSB-coated lagging strand (Marceau et al. 2011).
So, the molecular interactions and the mechanism of primer transfer from DnaB-associated
primase to the Pol III HE at the replication fork remains a pressing, but open question. In T7, a
direct polymerase-primase interaction enables the handoff (Kato et al. 2004). In the T4 system, the
clamp loader and/or clamp appear to be key players (Manosas et al. 2009). The apparent dominant
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negative role of SSB-113 that requires w to overcome the in vitro phenotype likely provides an
additional hint on the mechanism in E. coli (Yuzhakov et al. 1999).
Cycling of the Lagging Strand Polymerase in E. coli Is Directed Exclusively
by a Modified Signaling Model
To distinguish collision and signaling mechanisms in E. coli, modifications of technology initially
developed by the Benkovic lab for studying T4 DNA replication (Yang et al. 2006) were employed
(Yuan and McHenry 2014). Templates with highly asymmetric GC distribution were used. Instead of
perturbing lagging strand synthesis by nucleotide depletion (as with T4), it was perturbed by either
poisoning lagging strand synthesis specifically by addition of ddGTP or slowing elongation with the
analog dGDPNP in place of dGTP. The signaling and collision models make distinct predictions. If
an Okazaki fragment was prematurely terminated by incorporation of ddGMP, leaving a gap after its
30 -end, collision would not occur, abruptly terminating DNA synthesis. However, if cycling
occurred through signaling, rapid cycling would take place. Shortened Okazaki fragments would
be observed, but the molar level of Okazaki fragments would remain constant. The latter result was
obtained, supporting the signaling model (Yuan and McHenry 2014).
In the presence of dGDPNP, which slowed elongation but left the DNA unmarked after nucleotide
analog incorporation, the prediction of the signaling model is much the same as explained in the
preceding paragraph for ddGMP incorporation. A shortened population of Okazaki fragments
would be obtained. If the collision model was operational, slowing Okazaki fragment synthesis
while leading strand synthesis continued at an unperturbed rate would increase the time required for
collision with the 50 -end of the preceding Okazaki fragment to occur. Thus, each successive Okazaki
fragment would become increasingly longer. Results indicated the signaling model was exclusively
used (Yuan and McHenry 2014). The drastically opposed length predictions allowed the collision
model to be completely eliminated for E. coli.
The same results were obtained when exogenous oligonucleotides were used to prime synthesis,
demonstrating that the availability of a new primer provides the signal for cycling, not the action of
primase, per se (Yuan and McHenry 2014). The system was set up with substoichiometric Pol III HE
that has a high processivity and affinity for the fork, precluding the possibility that significant
synthesis occurred elsewhere.
Results suggested that the lagging strand polymerase is significantly faster than the leading strand
polymerase (at least with the dGDPNP analog), explaining how a large population of complete
Okazaki fragments could be obtained, even if signaling is the only mechanism driving cycling (Yuan
and McHenry 2014). Partial evidence supported speculation that the clamp loader was the sensor for
cycling (Yuan and McHenry 2014), suggesting that the model put forward by Benkovic for T4
(Chen et al. 2013; Yang et al. 2004) might be general for bacterial chromosomal replication.
The torque model for cycling induced by superhelical energy built up in the leading strand product
(Kurth et al. 2013) appears to be inconsistent with the observation that Okazaki fragment size varies
with primase concentration and the frequency of priming (Wu et al. 1992b). If leading strand torque
induced dissociation of the lagging strand polymerase, the lagging strand product should be reduced
to approximately the same size regardless of priming frequency, because the force generated should
be the same per unit length of leading strand synthesized. If torque drives cycling, under conditions
of limiting polymerase (Wu et al. 1992b; Yuan and McHenry 2014) that precludes extension by
incomplete Okazaki fragments by exogenous polymerase, the gaps between Okazaki fragments
should increase with decreasing priming frequency, but not fragment length. Multiple reinitiations of
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DNA elongation by the lagging strand polymerase on the same Okazaki fragment does not appear
feasible because the rate of initiation complex formation is a partially rate-limiting step during
Okazaki fragment synthesis (Downey et al. 2011; Hayner and Bloom 2013). A more definitive test
would be provided by addition of topoisomerase I to rolling circle replicative reactions. This enzyme
(and not an inactive control created by mutation of a critical active site residue) should inhibit
cycling if superhelical tension is the driving force.
Cross-References
By Springer guidelines, are cross-references within the same
article included here? We prefer not, but if you chose to include,
you should include both of the internal cross-references made in
this article (page 2 and 4).
▶ Cycling of the Lagging Strand Replicase During Okazaki Fragment Synthesis
▶ DNA Polymerase III Structure
References
By Springer guidelines, should subsections be included in this list as we noted on page 3, last
paragraph, or do you just want to list the chapter in which the cross-reference occurs as you
have done here? If it is the author's choice, we prefer listing just the cross-referenced chapter
as you have done here. We just want a clarification before proofing our remaining chapters.
Alberts BM, Barry J, Bedinger P, Formosa T, Jongeneel CV, Kreuzer KN (1983) Studies on DNA
replication in the bacteriophage T4 in vitro system. Cold Spring Harb Symp Quant Biol
47(Pt 2):655–668
Bailey S, Wing RA, Steitz TA (2006) The structure of T. aquaticus DNA polymerase III is distinct
from eukaryotic replicative DNA polymerases. Cell 126:893–904
Breier AM, Weier HU, Cozzarelli NR (2005) Independence of replisomes in Escherichia coli
chromosomal replication. Proc Natl Acad Sci U S A 102:3942–3947
Chastain PD, Makhov AM, Nossal NG, Griffith J (2003) Architecture of the replication complex and
DNA loops at the fork generated by the bacteriophage T4 proteins. J Biol Chem
278:21276–21285
Chen D, Yue H, Spiering MM, Benkovic SJ (2013) Insights into Okazaki fragment synthesis by the
T4 replisome: the fate of lagging-strand holoenzyme components and their influence on Okazaki
fragment size. J Biol Chem 288:20807–20816
Dalrymple BP, Kongsuwan K, Wijffels G, Dixon NE, Jennings PA (2001) A universal proteinprotein interaction motif in the eubacterial DNA replication and repair systems. Proc Natl Acad
Sci U S A 98:11627–11632
Dohrmann PR, McHenry CS (2005) A bipartite polymerase-processivity factor interaction: only the
internal b binding site of the a subunit is required for processive replication by the DNA
polymerase III holoenzyme. J Mol Biol 350:228–239
Dohrmann PR, Manhart CM, Downey CD, McHenry CS (2011) The rate of polymerase release
upon filing the gap between Okazaki fragments is inadequate to support cycling during lagging
strand synthesis. J Mol Biol 414:15–27
Downey CD, Crooke E, McHenry CS (2011) Polymerase chaperoning and multiple ATPase sites
enable the E. coli DNA polymerase III holoenzyme to rapidly form initiation complexes. J Mol
Biol 412:340–353
Evans RJ, Davies DR, Bullard JM, Christensen J, Green LS, Guiles JW, Pata JD, Ribble WK,
Janjic N, Jarvis TC (2008) Structure of polC reveals unique DNA binding and fidelity determinants. Proc Natl Acad Sci U S A 105:20695–20700
Georgescu RE, Kurth I, Yao NY, Stewart J, Yurieva O, O’Donnell M (2009) Mechanism of
polymerase collision release from sliding clamps on the lagging strand. EMBO J 28:2981–2991
Page 7 of 9
Molecular Life Sciences
DOI 10.1007/978-1-4614-6436-5_132-1
# Springer Science+Business Media New York 2014
Glover BP, McHenry CS (1998) The wc subunits of DNA polymerase III holoenzyme bind to
single-stranded DNA-binding protein (SSB) and facilitate replication of a SSB-coated template.
J Biol Chem 273:23476–23484
Hamdan S, Loparo JJ, Takahashi M, Richardson CC, van Oijen AM (2009) Dynamics of DNA
replication loops reveal temporal control of lagging-strand synthesis. Nature 457:336–339
Hayner JN, Bloom LB (2013) The beta sliding clamp closes around DNA prior to release by the
Escherichia coli clamp loader gamma complex. J Biol Chem 288:1162–1170
Jergic S, Horan NP, Elshenawy MM, Mason CE, Urathamakul T, Ozawa K, Robinson A, Goudsmits
JM, Wang Y, Pan X, Beck JL, van Oijen AM, Huber T, Hamdan SM, Dixon NE (2013) A direct
proofreader-clamp interaction stabilizes the Pol III replicase in the polymerization mode. EMBO
J 32:1322–1333
Johanson KO, McHenry CS (1982) The b subunit of the DNA polymerase III holoenzyme becomes
inaccessible to antibody after formation of an initiation complex with primed DNA. J Biol Chem
257:12310–12315
Kato M, Ito T, Wagner G, Ellenberger T (2004) A molecular handoff between bacteriophage T7
DNA primase and T7 DNA polymerase initiates DNA synthesis. J Biol Chem 279:30554–30562
Kim DR, McHenry CS (1996) Identification of the b-binding domain of the a subunit of Escherichia
coli polymerase III holoenzyme. J Biol Chem 271:20699–20704
Kurth I, Georgescu RE, O’Donnell ME (2013) A solution to release twisted DNA during chromosome replication by coupled DNA polymerases. Nature 496:119–122
Lee JB, Hite RK, Hamdan SM, Xie XS, Richardson CC, van Oijen AM (2006) DNA primase acts as
a molecular brake in DNA replication. Nature 439:621–624
Leu FP, Georgescu R, O’Donnell ME (2003) Mechanism of the E. coli t processivity switch during
lagging-strand synthesis. Mol Cell 11:315–327
Liu B, Lin J, Steitz TA (2013) Structure of the Pol III a-t(c)-DNA complex suggests an atomic
model of the replisome. Structure 21:658–664
Lopez de Saro FJ, Georgescu RE, Goodman MF, O’Donnell ME (2003a) Competitive processivityclamp usage by DNA polymerases during DNA replication and repair. EMBO J 22:6408–6418
Lopez de Saro FJ, Georgescu RE, O’Donnell ME (2003b) A peptide switch regulates DNA
polymerase processivity. Proc Natl Acad Sci U S A 100:14689–14694
Manosas M, Spiering MM, Zhuang Z, Benkovic SJ, Croquette V (2009) Coupling DNA unwinding
activity with primer synthesis in the bacteriophage T4 primosome. Nat Chem Biol 5:904–912
Marceau AH, Bahng S, Massoni SC, George NP, Sandler SJ, Marians KJ, Keck JL (2011) Structure
of the SSB-DNA polymerase III interface and its role in DNA replication. EMBO J 30:4236–4247
McCauley MJ, Shokri L, Sefcikova J, Venclovas C, Beuning PJ, Williams MC (2008) Distinct
double- and single-stranded DNA binding of E. coli replicative DNA polymerase III a subunit.
ACS Chem Biol 3:577–587
McHenry CS, Tomasiewicz H, Griep MA, F€
urste JP, Flower AM (1988) DNA polymerase III
holoenzyme: mechanism and regulation of a true replicative complex. In: Moses R, Summers
W (eds) DNA replication and mutagenesis. American Society for Microbiology, Washington,
DC, pp 14–26
Mok M, Marians KJ (1987a) Formation of rolling-circle molecules during fΧ174 complementary
strand DNA replication. J Biol Chem 262:2304–2309
Mok M, Marians KJ (1987b) The Escherichia coli preprimosome and DNA B helicase can form
replication forks that move at the same rate. J Biol Chem 262:16644–16654
Ozawa K, Horan NP, Robinson A, Yagi H, Hill FR, Jergic S, Xu ZQ, Loscha KV, Li N, Tehei M,
Oakley AJ, Otting G, Huber T, Dixon NE (2013) Proofreading exonuclease on a tether: the
Page 8 of 9
Molecular Life Sciences
DOI 10.1007/978-1-4614-6436-5_132-1
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complex between the E. coli DNA polymerase III subunits alpha, epsilon, theta and beta reveals a
highly flexible arrangement of the proofreading domain. Nucleic Acids Res 41:5354–5367
Pandey M, Syed S, Donmez I, Patel G, Ha T, Patel SS (2009) Coordinating DNA replication by
means of priming loop and differential synthesis rate. Nature 462:940–943
Pascal JM, O’Brien PJ, Tomkinson AE, Ellenberger T (2004) Human DNA ligase I completely
encircles and partially unwinds nicked DNA. Nature 432:473–478
Toste RA, Holding AN, Kent H, Lamers MH (2013) Architecture of the Pol III-clamp-exonuclease
complex reveals key roles of the exonuclease subunit in processive DNA synthesis and repair.
EMBO J 32:1334–1343
Ullsperger CJ, Vologodskii AV, Cozzarelli NR (1995) Unlinking of DNA by topoisomerases during
DNA replication. In: Eckstein F, Lilley DMJ (eds) Nucleic acids and molecular biology. Springer,
Berlin, pp 115–142
Wijffels G, Dalrymple BP, Prosselkov P, Kongsuwan K, Epa VC, Lilley PE, Jergic S, Buchardt J,
Brown SE, Alewood PF, Jennings PA, Dixon NE (2004) Inhibition of protein interactions with the
b2 sliding clamp of Escherichia coli DNA polymerase III by peptides from b2-binding proteins.
Biochemistry 43:5661–5671
Wing RA (2010) Structural studies of the prokaryotic replisome. Thesis/Dissertation, Yale University, p 170
Wing RA, Bailey S, Steitz TA (2008) Insights into the replisome from the structure of a ternary
complex of the DNA polymerase III a-subunit. J Mol Biol 382:859–869
Wu CA, Zechner EL, Hughes AJ Jr, Franden MA, McHenry CS, Marians KJ (1992a) Coordinated
leading and lagging-strand synthesis at the Escherichia coli DNA replication fork.
IV. Reconstitution of an asymmetric, dimeric DNA polymerase III holoenzyme. J Biol Chem
267:4064–4073
Wu CA, Zechner EL, Reems JA, McHenry CS, Marians KJ (1992b) Coordinated leading- and
lagging-strand synthesis at the Escherichia coli DNA replication fork: V. Primase action regulates
the cycle of Okazaki fragment synthesis. J Biol Chem 267:4074–4083
Yang J, Zhuang Z, Roccasecca RM, Trakselis MA, Benkovic SJ (2004) The dynamic processivity of
the T4 DNA polymerase during replication. Proc Natl Acad Sci U S A 101:8289–8294
Yang J, Nelson SW, Benkovic SJ (2006) The control mechanism for lagging strand polymerase
recycling during bacteriophage T4 DNA replication. Mol Cell 21:153–164
Yuan Q, McHenry CS (2014) Cycling of the E. coli lagging strand polymerase is triggered
exclusively by the availability of a new primer at the replication fork. Nucleic Acids Res
42:1747–1756
Yuzhakov A, Kelman Z, O’Donnell ME (1999) Trading places on DNA–a three-point switch
underlies primer handoff from primase to the replicative DNA polymerase. Cell 96:153–163
Page 9 of 9