The Rate of Polymerase Release upon Filling the Gap ... · Okazaki fragment at a rate faster than that of Okazaki fragment production. The rate of replication fork progression in

doi:10.1016/j.jmb.2011.09.039 J. Mol. Biol. (2011) 414, 15–27

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Journal of Molecular Biologyj ourna l homepage: ht tp : / /ees .e lsev ie r.com. jmb

The Rate of Polymerase Release upon Filling the Gapbetween Okazaki Fragments Is Inadequate to SupportCycling during Lagging Strand Synthesis

Paul R. Dohrmann, Carol M. Manhart, Christopher D. Downeyand Charles S. McHenry⁎Department of Chemistry and Biochemistry, University of Colorado, Boulder, CO 80309, USA

Received 4 August 2011;received in revised form16 September 2011;accepted 24 September 2011Available online1 October 2011

Edited by J. Karn

Keywords:DNA replication;Okazaki fragment synthesis;cycling;collision model;surface plasmon resonance

*Corresponding author. E-mail [email protected] used: SPR, surface

ssDNA, single-stranded DNA; TCEPphosphine hydrochloride.

0022-2836/$ - see front matter © 2011 E

Upon completion of synthesis of an Okazaki fragment, the lagging strandreplicase must recycle to the next primer at the replication fork in under 0.1 sto sustain the physiological rate of DNA synthesis. We tested the collisionmodel that posits that cycling is triggered by the polymerase encounteringthe 5′-end of the preceding Okazaki fragment. Probing with surfaceplasmon resonance, DNA polymerase III holoenzyme initiation complexeswere formed on an immobilized gapped template. Initiation complexesexhibit a half-life of dissociation of approximately 15 min. Reduction in gapsize to 1 nt increased the rate of dissociation 2.5-fold, and complete filling ofthe gap increased the off-rate an additional 3-fold (t1/2 ∼2 min). Anexogenous primed template and ATP accelerated dissociation an additional4-fold in a reaction that required complete filling of the gap. Neither a5′-triphosphate nor a 5′-RNA terminated oligonucleotide downstream of thepolymerase accelerated dissociation further. Thus, the rate of polymeraserelease upon gap completion and collision with a downstream Okazakifragment is 1000-fold too slow to support an adequate rate of cycling andlikely provides a backup mechanism to enable polymerase release when theother cycling signals are absent. Kinetic measurements indicate that additionof the last nucleotide to fill the gap is not the rate-limiting step for polymeraserelease and cycling. Modest (approximately 7 nt) strand displacement isobserved after the gap between model Okazaki fragments is filled. Todetermine the identity of the protein that senses gap filling to modulateaffinity of the replicase for the template, we performed photo-cross-linkingexperiments with highly reactive and non-chemoselective diazirines. Onlythe α subunit cross-linked, indicating that it serves as the sensor.

© 2011 Elsevier Ltd. All rights reserved.

Introduction

Cellular chromosomal replicases from all branchesof life are tripartite. They contain a polymerase

ress:

plasmon resonance;, tris(2-carboxyethyl)

lsevier Ltd. All rights reserve

(Pol III in bacteria and Pol δ and ɛ in eukaryotes), asliding clamp processivity factor (β2 in bacteria andPCNA in eukaryotes), and a clamp loader (DnaXcomplex in bacteria and RFC in eukaryotes).1–3 TheEscherichia coli replicase, the DNA polymerase IIIholoenzyme (Pol III HE), has the processivityrequired to replicate N150 kb,4,5 and perhaps theentire E. coli chromosome, without dissociation. Yet,the lagging strand polymerase must be able toefficiently cycle to the next primer synthesized atthe replication fork upon the completion of each

d.

mailto:[email protected]

http://dx.doi.org/10.1016/j.jmb.2011.09.039

16 Test of Collision Model

Okazaki fragment at a rate faster than that ofOkazaki fragment production. The rate of replicationfork progression in E. coli is about 600 nt/s at 30 °C,2

approximately the rate of replicase progression onsingle-stranded DNA (ssDNA) templates.6 Thus,most of the time in an Okazaki fragment cycle isspent on elongation, and little time (b0.1 s) remainsfor the polymerase to release, bind the next primer,and begin synthesis. A processivity switch must bepresent to increase the off-rate of the lagging strandpolymerase by several orders of magnitude.Upon complete replication of primed single strand

DNA, Pol III⁎ (a complex of Pol III and DnaXcomplexes containing the τ form of DnaX that bindsPol III) dissociates from a duplex and leaves the β2sliding clamp behind.7 There are two competing butnonexclusive models for the signal that throws theprocessivity switch. The first, the signaling model,proposes that a signal is provided by synthesis of anew primer at the replication fork that inducesthe lagging strand polymerase to dissociate, evenif the Okazaki fragment has not been completed.8

The second, the collision model, originally proposedfor T49 and then extended to the E. coli system,10

posits that the lagging strand polymerase replicates tothe last nucleotide10 or until the Okazaki fragment isnearly complete, signaling polymerase dissociation.11

A communication circuit that proceeds through the τsubunit has been proposed to sense the conversionof a gap to a nick, signaling release.10,12,13 τ wasproposed to act by competing with a C-terminal siteon Pol III for binding to β. However, other work hasshown the essential site required for β interaction tobe internal within the Pol III α subunit.14–16

Experiments designed to test whether the signalingor collision models are dominant have yieldedequivocal results.17

In the more fully characterized systems providedby the replication apparatus of bacteriophages T4and T7, signaling through synthesis or the availabil-ity of a new primer appears to play an importantrole, with the collision pathway playing an apparentbackup function.18–20 In T4, handoff of the nascentpentaribonucleotide primer occurs in a reaction thatis facilitated by T4 ssDNA binding protein (T4 SSB)and the clamp and the clamp loader.21,22 A proposalwasmade that the T4 clamp loader/clamp interactionwith a new primer might be the key signal requiredfor release of the lagging strand polymerase.22

In this report, we perform a kinetic assessment ofthe collision model on templates constructed suchthat elongating Pol III HE encounters an oligonu-cleotide hybridized to the template that mimics the5′ terminus of the preceding Okazaki fragment onthe lagging strand of a replication fork. Wecompared the rate of Pol III⁎ release with the rateof addition of the last nucleotide. To determinecandidate holoenzyme subunits that could serve asa sensor for completion of lagging strand synthesis,

we identified the protein that contacts the templateimmediately ahead of the primer terminus and the5′-end of the preceding Okazaki fragment.

Results

Most measurements assessing pathways thattrigger Pol III⁎ release and cycling have beenperformed using equilibrium binding measure-ments. While a decreased affinity of Pol III⁎ for thetemplate might be consistent with a role in acceler-ating release, the relevant issue is whether thelagging strand is triggered to release in less than0.1 s upon collision with the preceding Okazakifragment. To provide a method for quantitativeassessment of the kinetics of Pol III⁎ release, we setup a surface plasmon resonance (SPR) assay inwhich primers were placed on a surface-immobilizedtemplate a set number of nucleotides from anoligonucleotide that models the 5′-end of thepreceding Okazaki fragment (Fig. 1a). Initiationcomplexes are mobile on DNA, and unless bulkyblocking agents flank the complex, it rapidly slidesoff the ends of the template.23 We placed biotin nearthe 3′-end of the template to permit attachment to astreptavidin-covered surface for SPR measurements.The other end of the construct was blocked bycovalently attaching the 3′ terminus of the blockingoligonucleotide to His6-tagged O6-alkylguanine-DNA-transferase (SNAP-tag).24 The sequence of thetemplate downstream of the primer was designed topermit controlled advancement of the elongating PolIII HE by a subset of dNTPs.We observed stabile association of Pol III HE with

each primer–template to form an initiation complexthat dissociated with a half-life of 10–15 min(Fig. 1c–e). Addition of the full complement ofdNTPs required for conversion of the gap to a nickgreatly increased the amount of the Pol III⁎ released(Fig. 1c and d) and modestly (3- to 8-fold) increasedthe dissociation rate (Table 1). However, the half-lifeof polymerase release (∼2–5 min) is approximately1000-fold slower than that expected to support thephysiological rate of cycling upon completion of anOkazaki fragment. We observed that addition ofany nucleotide, independent of whether the nucle-otides present were able to fill the gap completely,led to an increase in the amount of Pol III⁎ thatdissociates (amplitude column in Table 1), butcomplete gap filling is needed to achieve themaximal rate.Kinetic analysis was complicated by a second,

more slowly dissociating component that requiredanalysis using a double-exponential decay equation(Fig. 1f). The second component dissociated with ahalf-life of 1–5 h. This required each dissociationexperiment to be run for up to 3 days to acquire adata set that allowed for complete decay (10 half-

Fig. 1. Pol III⁎ does not release rapidly upon filling a gap. The dissociation of Pol III⁎ from primed templates wasmeasured using SPR. (a) Three primed templates containing 4-, 5- and 10-nt gaps were anchored to the surface of astreptavidin-coated sensor chip using biotin-labeled oligonucleotides. The 3′-end of the blocking oligonucleotide wascovalently linked to the SNAP protein to prevent β2 from sliding off the end. (b) Initiation complexes were assembled ontothe immobilized DNA primer–template by injecting Pol III⁎, β2, and ATP. A second injection containing either buffer or asolution containing 40 μM dCTP, dTTP, and ddGTP (dNTPs) was made at 540 s. (c–e) Injection of either buffer (red) ordNTPs (green) over the 4-nt gapped template, 5-nt gapped template, and 10-nt gapped template, respectively.Dissociation was allowed to continue for 3 days. The data shown are only for the first hour after injection. (f) Curve-fitanalysis is shown for the Pol III⁎ dissociation in the presence of dNTPs over a 4-nt gapped template: data (open circles),curve fit to single-exponential decay (red), curve fit to double-exponential decay (cyan).

17Test of Collision Model

lives) for an accurate curve fit. In the text of the mainpaper, we refer to the presumably biologicallyrelevant fast dissociating component and presentthe full data under Supplementary Tables S1–S3.Recognizing that simple conversion of a gap to a

nick did not provide an adequate rate of Pol III⁎release to achieve a physiologically relevant rate ofcycling for Okazaki fragment synthesis, we soughtexternal effectors that might stimulate dissociation.We found that the presence of ATP and anexogenous template–primer accelerates dissociation

approximately 4-fold (Fig. 2 and Table 2). Raisingthe exogenous primed template concentration in theanalyte solution did not further increase the rate.ATPγS will not substitute in this reaction, suggest-ing that ATP hydrolysis is required. The rateenhancement by these factors is only fully achievedwhen a gap has been filled. SSB and exogenous β2are not required to accelerate Pol III release (Table 2).The blocking oligonucleotide used in our assay to

mimic the preceding Okazaki fragment contained a5′-hydroxyl group. The preceding fragment in cells

Table 1. Filling a gap completely is required to achieve themaximal rate of Pol III⁎ release

Injection

4-nt gaptemplatet1/2 (s)

[amplitude]Gapsize


[amplitude]Gapsize

Buffer 860 [12%] 4 620 [10%] 5dCTP 390 [69%] 3 360 [70%] 3ddCTP 660 [25%] 3 380 [26%] 4dCTP, dTTP 210 [62%] 1 190 [63%] 1dCTP, ddTTP 370 [36%] 2 350 [37%] 2dCTP, dTTP, dGTP 290 [55%] 0 180 [57%] 0dCTP, dTTP, ddGTP 110 [64%] 0 110 [63%] 0

Table 2. Exogenous primed template and ATP stimulatePol III⁎ release from completed Okazaki fragments

Injection


[amplitude]


[amplitude]

ATP 990 [60%] 920 [62%]dNTPs 110 [64%] 110 [64%]dNTPs, 20/70-mer, ATP 30 [86%] 40 [83%]dNTPs, ATP 60 [75%] 60 [77%]20/70-mer, ATP 330 [52%] 340 [51%]dNTPs, 20/70-mer 60 [80%] 70 [77%]dNTPs, 20/70-mer, ATP, SSB4 50 [77%] 50 [73%]dNTPs, 20/70-mer, ATP, SSB4, β2 30 [64%] 30 [59%]dNTPs, 20/70-mer, ATPγS 130 [68%] 130 [67%]


would be expected to contain a 5′-triphosphate tobe composed of RNA for ca 10 nt at the 5′-end. Todetermine whether either of these factors led tomore rapid polymerase release, we had blockingoligonucleotides synthesized that contained eithera 5′-triphosphate or a mixed RNA/DNA oligonu-cleotide with 12 RNA nucleotides at the 5′-end anda single 5′-phosphate. Neither of these modifica-tions led to acceleration of release (Table 3). Theexperiments described so far were conducted usingPol III HE with the presumed physiological PolIII⁎ composition: Pol III2-τ2γδδ′χψ.

1 To ensurethat a dimeric polymerase was not the cause ofslow release, we also prepared an assembly with aPol III-τγ2δδ′χψ composition. Upon addition ofdNTPs on the 4- and 5-nt gapped templates,polymerase dissociated with half-lives of 140 and130 s, respectively—similar to the 110 s observedwith the dimeric polymerase assembly. We ob-served a similar dissociation rate (t1/2=140 s) upon

Fig. 2. An exogenous primed template and ATP acceleradissociation of Pol III⁎ from primed template was measureddCTP, dTTP, and ddGTP) alone or dNTPs plus additional co20/70-mer). Data shown are normalized response units versusare for dNTPs alone with a 4-nt gapped template (green) and+ATP with 4-nt gapped template (magenta) and 5-nt gappeATPγS decreases the off-rate of Pol III⁎ in the presence of dNTP+20/70-mer+ATP with a 4-nt gapped template (magenta) a+ATPγS with a 4-nt gapped template (brown) and 5-nt gappedgapped templates are coincident and not resolved at most tim

the addition of dNTPs to allow gap filling for PolIII HE with a Pol III3-τ3δδ′χψ composition.The rate of polymerase release that we observe is

much slower than the rate of less than or equal to(0.5 s−1; t1/2∼1.4 s) reported previously.10 However,reanalysis of the published data yielded a fit to asingle exponential with a half-life of 14 s (0.05 s−1),too slow to support the rate of in vivo Okazakifragment synthesis, albeit faster than the rates weobserve in this work (Supplementary Fig. S1).Our results are consistent with gap completion

providing a signal for cycling, but with releaseslower than would be expected for cycling duringOkazaki fragment synthesis. We wanted to deter-mine whether the slow rate of polymerase releasewas due to a slow process after Okazaki fragmentcompletion such as a conformational change orwhether the rate of incorporation of the finalnucleotide(s) to complete the fragment was slow.

te Pol III⁎ release from templates with filled gaps. Theusing SPR following a second injection of dNTPs (40 μMmponents (+1 mM ATP or ATPγS and 0.5 μM exogenoustime for the first 1000 s of dissociation. (a) Curves showna 5-nt gapped template (teal) or for a dNTPs+20/70-merd template (red). (b) The slowly hydrolyzed ATP analogs and exogenous 20/70-mer. Curves shown are for dNTPsnd a 5-nt gapped template (red) or dNTPs+20/70-mertemplate (dark orange). The curves for the 4-nt and 5- ntes shown.

image of Fig. 2

Table 3. RNA or a triphosphate on the 5′-terminus of thepreceding Okazaki fragment does not contribute to therate of Pol III⁎ release

Injection


[amplitude]


[amplitude]

5′-OH DNA39 blocking oligo 30 [86%] 40 [83%]5′ PO4 RNA12/DNA27 blocking oligo 60 [71%] 70 [68%]5′ Tri-PO4 DNA39 blocking oligo 80 [51%] 90 [44%]


To enable these measurements, we constructedtemplates similar to those used for the SPRexperiments with 5′-32P-labeled primers. This per-mitted monitoring of the rate of primer elongationand gap completion using high-resolution poly-acrylamide gel electrophoresis. After formation ofinitiation complexes with Pol III HE on the primer–template, elongation reactions were initiated byaddition of dNTPs. The majority (79%) of primer–templates are completely gap filled before the firstmanually sampled time point could be taken (5 s), a

tended 10-nt gap primer–template; (12) 45-mer annealed to textended by Pol III HE. The samples in lanes 1 and 4–6 were exlanes 8–10 and 13 were extended by wild-type Pol III HE.

timescale faster than Pol III⁎ dissociation (Fig. 3).Thus, dissociation is not limited by the rate of gapfilling. The nicked duplex represents 22% of allelongated products. Interestingly, most of theelongation product (57% for wild-type Pol III HE)corresponds to a modest level of displacement of theblocking oligonucleotide. The wild-type Pol III HEforms significant products up to 14 nt beyond theblocking oligonucleotide, and the average stranddisplacement is approximately 7 nt.Previous models have proposed roles for either τ

or α to serve as the sensor for conversion of a gap toa nick upon completion of Okazaki fragmentsynthesis, serving as a trigger for acceleratedpolymerase release.10,11,25,26 To determine the pro-teins that contact the template and blocking oligo-nucleotide immediately ahead of the primerterminus, we performed cross-linking experiments.A phenyldiazirine linked to the 5-position ofthymidylate was placed at a unique position in aseries of templates that mimicked those used in theSPR experiments (Fig. 4a). We chose diazirinesbecause, when irradiated, they generate carbenes

Fig. 3. Pol III HE rapidly fillsa 10-nt gap and partially displaces ablocking oligonucleotide. (a) 5′-32P-end-labeled primer 35-mer was hy-bridized to a template 81-mer with a10-nt gap in front of blocking oligo-nucleotide 34-mer. This primer–tem-plate was incubated with Pol III HEand ATP for 60 s to form initiationcomplexes, followed by addition ofdNTPs to initiate primer elongationfor the 32P-labeled complexes. (b)Denaturing polyacrylamide gel anal-ysis of the rate of nucleotide incorpo-ration. The gel laneswere loadedwith(1) unblocked primer–template fullyextended by Pol III HE; (2) 5′-32P-end-labeled 45-mer corresponding toa fully gap-filled product, annealedto the 81-mer template; (3) unex-tended 10-nt gap primer–template;(4) 10-nt gap primer–template after a5-s extension time; (5) 10-nt gapprimer–template after a 10-s exten-sion time; (6) 10-nt gap primer–template after a 20-s extension time;(7) 45-mer annealed to the 81-mertemplate (4-fold more sample thanloaded than in lanes 2 and12); (8) 10-nt gap primer–template after a 20-sextension time; (9) 10-nt gapprimer–template after a 10-s extension time;(10) 10-nt gap primer–template aftera 5-s extension time; (11) unex-

he 81-mer template; (13) unblocked primer–template fullytended by exonuclease-deficient Pol III HE. The samples in

image of Fig. 3

Fig. 4. The Pol III α subunit and not τ is positioned to serve as the sensor for the completion of Okazaki fragmentsynthesis. (a) DNA constructs used to determine Pol III HE contacts with model templates. The position of thephenyldiazirine photo-cross-linker is indicated by red lowercase t. The oligonucleotide containing the photo-cross-linkerwas labeled with 32P on the 5′-end prior to annealing. Template T+4 also served as Templates T+3 and T+1 by theaddition of 10 μM ddTTP or dTTP, respectively. (b) Denaturing polyacrylamide gel analysis of Pol III HE subunitstandards. Generation of lanes 1–3 is described in Supplementary Fig. S4. Pol III (0.62 μM final concentration) used togenerate lane 4 and τ (2 μM) used to generated lane 5. (c) Identification of protein contacts with model templates. Photo-cross-linking took place within initiation complexes formed with the designated model templates. The gel lanes wereloaded with approximately equal counts of radioactivity. Lanes 1, 2, 12, and 13 are identical with lanes 4, 5, 4 and 2,respectively, in (b).


that efficiently insert into C–H bonds of all aminoacids.27 This permits interpretation of negativeresults with certainty—if a protein does not cross-link, it is not bound to the template position towhich the diazirine is linked.

To enable assignment of cross-linked proteins bytheir electrophoretic mobility, we forced template–protein contacts with high concentrations of singleor simple subsets of Pol III HE subunits to generatestandards (Supplementary Fig. S4 and Fig. 4b).

image of Fig. 4


Control experiments were performed with thephenyldiazirine placed at the penultimate (−2)position at the 3′-end of the primer terminus andat the template position opposite the 3′-primerterminus. As expected from previous work,25,28

α was observed to cross-link. Cross-linking attemplate positions 1, 3, 4, and 8 nt ahead of theprimer and at the template position opposite the5′-nucleotide of the blocking oligonucleotide led tothe cross-linking of the α subunit of Pol III and notτ (Fig. 4c). We also observed a cross-link to single-stranded template positions that may correspondto either δ or δ′. In a control experiment, where thematerial in lane 8 of Fig. 4c was mixed with lane 2,the τ and δ/δ′ cross-links were clearly distinguish-able (data not shown). These species may repre-sent two binding states—one with Pol III bound tothe primer terminus and another with DnaXcomplex bound. Cross-links to α were alsoobserved to the nucleotide in the gap for templatescontaining a single nucleotide gap (Fig. 4c). Whenphenyldiazirine was placed at the penultimate5′-position of the blocking oligonucleotide, a verylow yield of cross-links to α was observed,consistent with the strand displacement observedupon completion of synthesis (Fig. 3). This resultsuggests that displaced blocking oligonucleotidedoes not maintain contact with Pol III. Thus, α andnot τ is the only protein positioned to function asthe sensor for gap completion.

Discussion

To permit a kinetic evaluation of the rate ofpolymerase release upon completing Okazaki frag-ment synthesis, we immobilizedmodel templates ona surface, assembled initiation complexes, andmonitored polymerase release upon nucleotideaddition by SPR. We found that conversion of agap to a nick accelerates release, but the rate is fartoo slow to support the physiological rate ofOkazaki fragment synthesis. The differences fromother published models likely arise from theirderivation from equilibrium measurements andours from kinetic determinations.In a search for factors that accelerate collision-

induced dissociation, we found that a mixture ofATP and an exogenous primed template acceleratesdissociation 4-fold. ATPγS will not substitute in thisreaction, suggesting that ATP hydrolysis is required.These findings partially reconcile previous modelsand observations pertaining to the requirementfor gap completion,10 the availability of a newtemplate–primer,8 and the requirement for τ.10 Aseries of switches may exist, all of which must beaccommodated for maximal cycling. Gap comple-tion may put the Pol III HE in a state in whicha signal can be received from the associated

τ-containing DnaX complex that involves bindingof a new primer–template and ATP hydrolysis.Additional acceleration might be derived fromother factors or their arrangement/interaction atthe fork, or the collision mechanism might only be abackup, with the primary signal provided by eventsassociated with new primer generation. The Pol IIIHE is involved in mismatch repair and other repairreactions where large stretches of DNA need to becopied.29 The collision mechanism might provide ameans for dissociating Pol III⁎ once it has completedits task in repair reactions. Previous reports describethat gaps need to be completely filled10 or onlymostly filled11 before polymerase release is trig-gered. These measurements were made primarilyby equilibrium binding measurements. Our kineticexperiments support the conclusion that gaps needto be completely filled before the maximal rate ofrelease is induced, but this rate is too slow tosupport chromosomal replication.Our experiments were performed on a DNA

substrate that mimicked the gap in the final stagesof Okazaki fragment synthesis. True replicationforks contain additional interactions, including twocoupled leading and lagging strand polymerasesthat are bound to the replicative helicase. It ispossible that the polymerase could switch to a statewhere it could more rapidly dissociate in thepresence of these additional potential allostericeffectors. Nevertheless, our substrates are similarto those used to initially formulate the collisionhypothesis.Measurement of the rate of gap filling indicates

that the reaction is complete before the first timepoint (5 s) was taken. Thus, the rate of gap filling isfast relative to the rate of polymerase dissociation.For a majority of primer–templates, filling of the gapis followed by a strand displacement of approxi-mately 7 nt on average. This value is less than theprocessivity of 300 nt observed in a strand displace-ment reaction that benefitted from a stabilizinginteraction of χ with SSB.30 Comparison of theseresults suggests that the χ–SSB interaction increasesthe processivity of strand displacement approxi-mately 40-fold. The fact that most of the productobserved involves strand displacement arguesagainst exact gap filling to form a nick providingthe signal for Pol III⁎ release. If the signaling modelis dominant, many Okazaki fragments may endbefore the gap between their terminus and theprimer, for the following Okazaki fragment, is filled.However, if the lagging strand polymerase issufficiently fast relative to the leading strandpolymerase, complete gap filling will occur. In thatcase, strand displacement is likely to occur. DNApolymerase I is recognized as an enzyme that fillsany remaining gaps between Okazaki fragmentsand removes RNA primers with a duplex-specific5′–3′ exonuclease activity, providing a nick that can


be sealed by DNA ligase.31,32 If there is a portion ofthe product that arises from strand displacement,another repair enzyme would be required to excisethe displaced single-stranded flap, analogous tothe function of the flap endonuclease FEN1 ineukaryotes.33

We applied diazirine-based photo-cross-linking toidentify which proteins contact a uniquely reactiveposition within model templates. The advantages ofthe method include high yields, irradiation atwavelengths (350 nm) far removed from chromo-phores present in proteins or nucleic acids, and theability of the carbene formed upon irradiation toinsert into all amino acids. The efficiency and lack ofchemical specificity permit negative results to beinterpreted with confidence. Using this method, wehave identified that α, and not τ, contacts thetemplate and blocking oligonucleotide in front of theprimer, making it the candidate for sensing conver-sion of a gap to a nick, presumably triggeringchange to a conformation that more rapidly releasesfrom DNA.In a structure of Pol III α complexed to DNA, an

OB fold was located close to the primer terminus.25

Because OB folds commonly bind to ssDNA, aproposal was made that the OB fold could be part ofthe sensing network.25,26 Consistent with thishypothesis, the ssDNA-binding portion of Pol IIIwas localized to a C-terminal region of α thatcontains the OB-fold element.34 A test of theimportance of the OB-fold motif was made using amutant in which three basic residues located in theβ1–β2 loop were changed to serine.11 No ssDNAbinding was observed in the mutant, indicatingdiminution in affinity. However, even the wild-typepolymerase bound ssDNA extremely weakly, nearthe limit of detection in the assays used (Kd ∼8 µM);thus, a modest decrease in affinity would appear tobe a null result. The processivity of the mutantpolymerase was decreased by the β1–β2 loopmutations, an effect that was rescued by thepresence of the τ complex.11 The latter observationwould seem to suggest that although the OB foldcontributes to ssDNA affinity and processivity, it isnot the processivity sensor, or at least the residuesmutated are not the key interactors. The OB foldmight act in concert with other binding changes aspart of a more complex signaling network.In our view, the entire polymerase active site is

likely the processivity switch. Wing et al. haveelegantly demonstrated a conformational changeinduced by substrate binding in which severalelements move, one of the results of which is toplace the β2 binding domain in a position where itcan productively interact with the β2 clamp onDNA.25 Follow-on studies with a Gram-positivepolymerase suggest that this observation may begeneral.35,36 The geometry and spatial constraintsaround the active site when the exiting template is

double stranded might make insertion of the lastnucleotide and ensuing strand displacement ener-getically unfavorable. Thus, these products mayhave decreased affinity for the active site, triggeringa reversal of the conformational changes thatoccurred upon primer–template and dNTP binding,causing the β2 binding domain to be pulled away,and switching the polymerase to a low-processivitymode. The presence of a polymerase domain that isnot bound to DNA serves to decrease the affinity ofthe C-terminal domains of Pol III α for β2,

14 which isconsistent with this model.Our results demonstrate that the collision of the

Pol III HE with the downstream elongated primer ofthe preceding Okazaki fragment is inadequate tosupport the rapid release required for cycling to thenext primer at the replication fork. The onlyalternative explanation in the literature is providedby the signaling model, originally proposed by KenMarians.8 We have shown that providing a com-peting primer in the presence of ATP only modestlystimulates dissociation. Thus, the function of pri-mase in the signaling model must be more than justproviding a competing primer. One possibility isthat primase, upon primer synthesis, sends a signalto the Pol III HE through the replicative helicase towhich both are bound. Another possibility is that thecompeting primer is provided in a special contextcreated by the molecular interactions at the replica-tion fork.Methodologies have been developed in the T4 and

T7 systems18–20 that might enable a more rigoroustest of the signaling mechanism in E. coli. If thesignaling model proves correct, an understanding ofthe specific reaction steps andmolecular interactionsthat throw the processivity switch and direct thereplicase to the next primer will be the nextchallenge. This remains a critical deficit in ourunderstanding of all DNA replication systems.

Materials and Methods

DNA templates

The templates diagrammed in Fig. 1 were assembledfrom the following gel-purified oligonucleotides: template81-mer, 5′ GCT AAT GAA TTC CCG GTT CTT GAC TACATT ACT CTT GAT CAA GGT CAG CCA GCC TATGCG CGT GAT CTG TAC ACC GTT C biotinT T (biotinTrepresents T with biotin attached, via a linker, to the C5position); primer 30-mer, 5′ GAA CGG TGT ACA GATCAC GCG CAT AGG CTG (to make 10-nt gappedtemplate); primer 35-mer, 5′ GAA CGG TGT ACA GATCAC GCG CAT AGG CTG GCT GA (to make 5-ntgapped template); primer 36-mer, 5′GAACGG TGTACAGAT CAC GCG CAT AGG CTG GCT GAC (to make 4-ntgapped template); blocking oligonucleotide 39-mer, 5′ATC AAG AGT AAT GTA GTC AAG AAC CGG GAATTC ATT AGC-SNAP; blocking oligonucleotide 39-mer


(5′-phosphate RNA12/DNA27), 5′-PO4rArUrC rArArGrArGrU rArArU GTA GTC AAG AAC CGG GAA TTCATT AGC-SNAP; and blocking oligonucleotide 39-mer(5′-triphosphate), 5′-triphosphate-ATC AAG AGT AATGTAGTCAAGAACCGGGAATTCATTAGCC-SNAP.The exogenous primed template used to stimulatepolymerase release (Table 3) was assembled by annealinga 20-mer primer (5′-TTG TTC AGA TGA AGG CGC AT)to a 70-mer template (5′-TAAAAAAAAAAAAAAAAAAAG GAT TAC TGG ATC CGA AGG TCA GCC AGCCTA TGC GCC TTC ATC TGA ACA A).For experiments reported in Fig. 3, in addition to

oligonucleotides described in this subsection, the followingoligonucleotides were synthesized: blocking oligonucleo-tide 34-mer, 5′GAG TAA TGT AGT CAAGAA CCGGGAATT CAT TAG C-biotin, and extended primer 45-mer, 5′GAA CGG TGT ACA GAT CAC GCG CAT AGG CTGGCT GAC CTT GAT CAA.

Preparation of the SNAP-conjugated blockingoligonucleotides

A method24 for attaching the Hexa-His-taggedO6-alkylguanine-DNA-Transferase (SNAP-tag; NewEngland Biolabs) to a DNA oligomer was adaptedfor our purposes. The blocking oligonucleotides(PAGE purified from Integrated DNA Technologies orBiosynthesis, Inc. for the 5′-triphosphate oligonucleotide)contained 3′-O-CH2-CH2-SH blocked by formation of adisulfide bond. Disulfide-blocked oligonucleotide(35 nmol) was reduced by incubation (1 h) at roomtemperature with 10 mM tris(2-carboxyethyl)phosphinehydrochloride (TCEP) (Pierce) in 500 μl TE buffer [10 mMTris–HCl and 1 mM ethylenediaminetetraacetic acid(pH 8.0)] (Supplementary Fig. S2). The solution wasapplied to a NAP5 (0.5 ml) gel-filtration column (GEHealthcare) with TE as the running buffer to removeTCEP and the 3-carbon thiol linker (SupplementaryFig. S2). The most concentrated fractions, determinedspectrophotometrically, were combined, giving a total of800 μl. O6-benzylguanine-maleimide (300 μl; NewEngland Biolabs) (2.5 mM in dimethyl formamide) wasadded to the 800 μl of reduced 39-mer in a total volumeof 1.1 ml, and the reaction mixture was allowed toproceed at room temperature (1 h) before applicationto a 2.5-ml NAP25 column to remove unreactedO6-benzylguanine-maleimide. To verify attachment ofthe O6-benzylguanine to the 3′end of the 39-mer, weanalyzed product (12,738 Da) on a 9% denaturingpolyacrylamide gel using the unmodified 39-mer(12,316 Da) for comparison. Greater than 90% of theoligonucleotide was modified (Supplementary Fig. S3).Unreacted/unmodified oligomer was removed in thenext step.The BG-modified-39-mer (33 nmol) was covalently

attached to SNAP by incubating 1.5-fold excess ofoligonucleotide over SNAP in [50 mM sodium phosphate(pH 8.0), 300 mMNaCl, 20% glycerol, 1 mM dithiothreitol,and 5 mM imidazole] for 1 h at room temperature withgentle rotation of the tube in a final volume of 2 ml. Theoligonucleotide-SNAP conjugate was applied to a 0. 75-mlNi2+-NTA column (Qiagen) pre-equilibrated with washbuffer [50 mM sodium phosphate (pH 8.0), 300 mMNaCl,20% glycerol, 1 mM dithiothreitol, and 5 mM imidazole].

All subsequent column procedures were performed at0–4 °C. The column was washed with 10 column volumesof wash buffer [50 mM sodium phosphate (pH 8.0),300 nM NaCl, 20% glycerol, 1 mM dithiothreitol, and5 mM imidazole] to remove unreacted oligonucleotide.Protein was eluted with elution buffer [50 mMsodium phosphate (pH 8.0), 300 nM NaCl, 20%glycerol, 1 mM dithiothreitol, and 400 mM imidazole].Eluted fractions containing the protein samples weredetected by the Bradford method (Pierce). Analysis ofthe SNAP-tag-39-mer, monitored on a 4–20% gradientSDS-PAGE gel from its molecular mass shift (unmodifiedSNAP-tag, 22.2 kDa; modified SNAP-tag-39-mer, 34.9 kDa),indicated that approximately 90% of the SNAP-tag proteinthat eluted off the column contained a covalently attached39-mer oligonucleotide (Supplementary Fig. S3). Theprotein/oligo conjugate was frozen in small aliquots byflash freezing in liquid nitrogen.

DNA polymerase III holoenzyme components

Protein components were purified as described for PolIII,37 ɛ,38 τ,39 γ,39 δ,40 δ′,40 χ,41 ψ,41 β2,

42 and SSB4.43 Four

different DnaX protein complexes with stoichiometries ofτ2γδδ′χψ, τγ2δδ′χψ, τ3δδ′χψ, and γ3δδ′χψ were purifiedas described previously.44,45 Unless otherwise noted, the ɛsubunit used in our experiments was mutated at D12Aand E14A to eliminate endogenous 3′–5′ exonucleaseactivity, which degrades the primer.46

For experiments using Pol III⁎ exhibiting a τ2γstoichiometry, complexes were assembled from purifiedsub-complexes of Pol III (αɛθ; 40 nM final) and DnaXcomplex (τ2γδδ′χψ; 20 nM final). For experiments usingPol III⁎ exhibiting γ3 stoichiometry, complexes wereassembled from purified sub-complexes of Pol III (αɛθ;20 nM final) and DnaX complex (γ3δδ′χψ; 20 nM final).For experiments using Pol III⁎ exhibiting a τγ2stoichiometry, the Pol III⁎ was isolated and purifiedfirst by cation-exchange chromatography over a Mono-Scolumn.37,45,47 DnaX complex (τγ2δδ′χψ; 2 nmol) wasincubated with Pol III (αɛ D12A E14A, θ; 10 nmol) for15 min at room temperature. The incubated mixture wasapplied to a NAP25 2. 5-ml (GE Healthcare) gel-filtrationcolumn to exchange the buffer [25 mM Hepes (pH 7.5),5% glycerol, and 25 mM NaCl] prior to running themixture over a 1-ml Mono-S column at 4 °C. The columnwas washed with 4 column volumes of wash buffer[25 mM Hepes (pH 7.5), 5% glycerol, and 25 mM NaCl].Fractions (0.5 ml) were collected after application of a150-mM NaCl step [in 25 mM Hepes (pH 7.5) and 5%glycerol]. The Pol III⁎ fractions eluted approximately5 column volumes after the 150-mM NaCl step. PurifiedPol III⁎ was injected over the Sensor chip SA at a finalconcentration of 50 nM.

Surface plasmon resonance

A BIAcore 3000 instrument was used to quantify andmeasure the release of Pol III from immobilized oligonu-cleotide templates. A flow rate of 5 μl/min in KCl-EDBbuffer [50 mM Hepes (pH 7.5), 125 mM KCl, 10 mMmagnesium acetate, 0.005% Tween-20, and 10 μM TCEP]at 25 °C was used for all reactions. All buffers were filtered


and degassed before use. The Sensor Chip SA (BIAcore), asensor chip pre-immobilized with streptavidin for captureof biotinylated ligands, was pre-conditioned with three1-min injections of 5 μl of 1 M NaCl/50 mM NaOHprior to attachment of DNA primer–templates. Threedifferent DNA primer–templates were immobilized. The81-mer template annealed to the 30-mer primer wasattached to flowcell 1 of the Sensor Chip SA. Thetemplate 81-mer annealed to the 35-mer and the template81-mer annealed to 36-mer were attached to flowcells 2and 3, respectively. Flowcell 4 was not derivatized andwas used as a control for background subtraction.DNA primer–templates were assembled in two steps.

First, duplex DNA containing the template oligonucleo-tide 81-mer (500 pmol) and the binding oligonucleotide30-mer (or 35-mer or 36-mer) (5 nmol) was annealedin vitro in [10 mM Tris–HCl (pH 8.0), 300 mM NaCl, and30 mM sodium citrate] in a volume of 250 μl. The samplewas heated to 95 °C for 10 min and allowed to coolslowly to 23 °C at 1 °C/min. The sample was diluted1000-fold to a concentration of 2 fmol/μl, with respect tothe 81-mer, in KCl-EDB buffer and aliquoted. To preparethe flowcell with the DNA primer–template, a total of ca50 μl of the annealed DNA primer–template was injectedover the surface of an individual flowcell on the sensorchip SA, which resulted typically in an increase of ca270–300 response units. Second, SNAP-tag-39-mer wasdiluted to a final concentration of 80 fmol/μl in KCl-EDBbuffer, and 300 μl was injected over all four flowcellssimultaneously. The SNAP-tag-39-mer does not bind tothe underivatized flowcell. Complete annealing to theDNA templates on the remaining flowcells was observedfor the SNAP-tag-39-mer by determining the bindingsignal after injection. Since the output obtained from theBIAcore is directly proportional to the mass bound,stoichiometries could be calculated. Complete annealingwas observed for the SNAP-tag-39-mer.Pol III⁎ complexes were injected over the DNA primer–

template at concentrations determined empirically for eachindividual assembly. To determine the optimum concen-tration for binding, we injected a series of differentconcentrations ranging from 5 nM to 80 nM Pol III⁎complex. Concentrations were chosen that gavemaximumstoichiometry of binding. In a typical experiment, Pol III⁎,β2 (300 nM), and ATP (1 mM) were injected over all fourflowcells of the SA sensor chip simultaneously in a 45-μlinjection. The “KINJECT” command was used for mea-suring dissociation of Pol III complexes in the presence ofbuffer alone. Data were collected for 3 days following theinjection to allow the dissociation of complexes to come tobaseline. The long dissociation timewas required to gatherdata for 10 half-lives for the slow, non-physiologicalsecond phase. Having a complete time course for theslow phase proved essential for a proper kinetic fit foraccurate determination of the fast phase. To measure thedissociation of Pol III⁎ in the presence of dNTPs and/orother additional components, we used the “COINJECT”command. The COINJECT command introduces a secondsample injection (45 μl was used) immediately uponcompletion of the first. The following components wereused either individually or in combination to determinethe requirements for full recycling: dNTPs (40 μM each,final), ddNTPs (40 μM final), ATP (1 mM final), ATPγS(1 mM final), β2 (0.5 μM final), SSB4 (0.5 μM final), and20-mer/70-mer (0.5 μM final). All were diluted in KCl-

EDB running buffer. Pol III⁎ failed to bind to DNAprimer–templates using control reactions that lacked β2or ATP in the first injection, demonstrating dependenceon both for binding.The rate of Pol III⁎/β2 dissociation from DNA primer–

templates was determined using nonlinear regressionanalysis using SigmaPlot software. The dissociation datawere then fit to several models for exponential decay. Allmodels tested included an offset value that defined thepoint at which the decay curve leveled out. To help assesswhether a term was necessary to generate a good fit, wedetermined a “dependency” value for each parameter inthe fitting equation. The simplest model that best fit ourdata was double-exponential decay.

Kinetics of filling the gap within model templates

A primer–template system similar to the SPR experi-ments was used, with an identical primer 35-mer andtemplate 81-mer. Blocking oligonucleotide 34-mer wasused to generate a 10-nt gap on the template. The primer35-mer was 5′-32P end-labeled using T4 polynucleotidekinase. The construct was annealed by combining200 nM 5′-32P-labeled primer 35-mer, 250 nM template81-mer, and 300 nM blocking oligonucleotide 34-mer(to ensure complete capture of the primer by a fullyblocked template) and heating to 95 °C for 10 min andcooling to 23 °C at 1 °C/min. An unblocked controlsample was prepared by the same procedure but omittingthe blocking oligonucleotide 34-mer. A marker for theexpected product resulting from primer extension by10 nt to fully fill the gap was prepared by annealing200 nM 5′-32P-extended primer 45-mer with 250 nMtemplate 81-mer.Primer elongation assays were conducted at ambient

temperature in a buffer containing 50 mMHepes (pH 7.5),100 mM potassium glutamate, 10 mMmagnesium acetate,0.20 mg/ml bovine serum albumin, 10 mM dithiothreitol,2.5% (vol/vol) glycerol, and 0.02% (vol/vol) Nonidet-P40detergent. Initiation complexes were generated by pre-incubating 10 nM 32P-labeled 10-nt gap containingprimed template with 30 nM streptavidin and thencombining this mixture with 25 nM τ2γ DnaX complex,25 nM Pol III, 100 nM β2, 200 μM ATP, and 1 μMhexokinase and incubating for 60 s. For experiments withwild-type Pol III, this initiation complex formationreaction included 20 μM dATP, enabling the polymeraseto replace the 3′-terminal nucleotide of the primer 35-merif digested by the 3′–5′ exonuclease. Initiation complexformation was terminated, and primer elongation wasinitiated by adding an equal volume of 0.25 mg/mlactivated calf thymus DNA (a cold trap for unassociatedPol III HE), 10 mM glucose (to activate the hexokinaseATPase activity), and 40 μMdNTPs.48 The total volume ofeach gap filling reaction was 40 μl. All concentrationsabove are the final values after initiating gap filling.Primer elongation reactions were quenched after varyingreaction times with 40 μl 50 mM ethylenediaminetetraa-cetic acid. To ensure a high yield of the full-length product,we conducted the primer elongation reaction for theunblocked primer–template for 30 s in the absence of calfthymus DNA and glucose. All other control DNA sampleswere not exposed to Pol III HE. All reactions and controlsamples were ethanol precipitated, and the air-dried


pellets were each dissolved in 40 μl 75% formamide and17 μl loaded onto 10% polyacrylamide/8.0 M urea/12%formamide sequencing gels to resolve the products. Thegels were visualized using a Storm 840 Phosphor Imagerand analyzed using ImageQuant 5.2.

Attachment of phenyldiazirine to oligonucleotides

In the absence of ambient light, 4-[3-(trifluoromethy)-diazirin-3-yl]benzoate N-hydroxysucinimide (customsynthesized by BioSynthesis, Inc.) was dissolved inanhydrous dimethyl formamide (0.5 M final concentra-tion). A 4.4-μmol sample was combined with 20 nmolof oligonucleotide (Integrated DNA Technologies) con-taining amino-modified C2 dT from Glen Research in a0.1 M sodium tetraborate buffer (pH 8.5) (final volume,50 μl). This reaction mixture was incubated in the darkat room temperature overnight. The derivatized productwas HPLC purified using a Waters XBridge OST C18column (4.6 mm×50 mm, pore size of 2.5 μm). Thesample was applied in 0.1 M triethylammonium acetate(pH 7.0) at a flow rate of 1.0 ml/min and eluted with agradient changing to acetonitrile at 1.54%/min. Thecolumn was run at 60 °C, and absorbance was monitoredat 265 nm and 350 nm. Approximately 15 nmol ofphotoreactive oligonucleotide was recovered. The pres-ence of photo-cross-linker in the oligonucleotide wasverified by observing peaks at 260 nm and 350 nm and bya shift in chromatographic mobility from the unreactedoligonucleotide.

Photo-cross-linking oligonucleotides

Oligonucleotides (Fig. 4a) were combined with pro-teins and nucleotides at room temperature in a buffercontaining 50 mM Hepes (pH 7.5), 10 mM magnesiumacetate, 10 mM dithiothreitol, 20% (vol/vol) glycerol,0.02% (vol/vol) Nonidet-P40 detergent, and 100 mMpotassium glutamate (unless otherwise indicated) insiliconized Pyrex tubes covered in Parafilm and irradiat-ed at 350 nm in a Rayonet photochemical reactor for 1 hat room temperature. The samples were quenched in 2×SDS-PAGE sample buffer [0.25 M Tris–HCl (pH 6.8), 5%SDS, 0.02% β-mercaptoethanol, and 0.2% glycerol].Samples were analyzed by 4–11% gradient SDS-PAGEcontaining 4 M urea.For experiments in Fig. 4c, initiation complexes were

generated by pre-incubating 4 nM 32P-labeled primedtemplate (Fig. 4a) with 40 nM streptavidin and thencombining with 16 nM τ3 complex, 16 nM Pol III, 200 nMβ2, and 0.8 mM ATP prior to irradiation.

Acknowledgements

This work was supported by National Institutes ofHealth grant R01 GM060273. We thank AnnaWieczorek for preparation of proteins and DianeHager for assistance in figure preparation.

Supplementary Data

Supplementary data to this article can be foundonline at doi:10.1016/j.jmb.2011.09.039

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