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J. Biol. Chem., Vol. 276, Issue 31, 29559-29566, August 3, 2001

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Conditional Coupling of Leading-strand and Lagging-strand DNA Synthesis at Bacteriophage T4 Replication Forks^*

Farid A. Kadyrov and John W. Drake

From the Laboratory of Molecular Genetics, NIEHS, National Institutes of Health, Research Triangle Park, North Carolina 27709-2233

Received for publication, February 12, 2001, and in revised form, June 1, 2001

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Eight proteins encoded by bacteriophage T4 are required for the replicative synthesis of the leading and lagging strands of T4 DNA. We show here that active T4 replication forks, which catalyze the coordinated synthesis of leading and lagging strands, remain stable in the face of dilution provided that the gp44/62 clamp loader, the gp45 sliding clamp, and the gp32 ssDNA-binding protein are present at sufficient levels after dilution. If any of these accessory proteins is omitted from the dilution mixture, uncoordinated DNA synthesis occurs, and/or large Okazaki fragments are formed. Thus, the accessory proteins must be recruited from solution for each round of initiation of lagging-strand synthesis. A modified bacteriophage T7 DNA polymerase (Sequenase) can replace the T4 DNA polymerase for leading-strand synthesis but not for well coordinated lagging-strand synthesis. Although T4 DNA polymerase has been reported to self-associate, gel-exclusion chromatography displays it as a monomer in solution in the absence of DNA. It forms no stable holoenzyme complex in solution with the accessory proteins or with the gp41-gp61 helicase-primase. Instead, template DNA is required for the assembly of the T4 replication complex, which then catalyzes coordinated synthesis of leading and lagging strands in a conditionally coupled manner.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Genetic and biochemical studies have identified eight T4 gene products required for T4 DNA replication. These are a DNA polymerase with an intrinsic 3'5' proofreading exonuclease activity (gp43),¹ a clamp loader (a 4:1 complex of gp44:gp62), a clamp (gp45), an ssDNA-binding protein (gp32), a replicative DNA helicase (gp41), a primase (gp61), and a helicase-loading protein (gp59) (1-4). Except for weakly viable gene 61 mutants, amber mutants of these genes are strongly defective in DNA synthesis.

Biochemical studies of the purified proteins and of DNA replication reconstituted in vitro have clarified many structural and mechanistic details of this complicated process. The gp61 primase binds to DNA, whereupon in the presence of gp59 and either ATP or GTP, the gp41 helicase interacts with the gp61-DNA complex to form a primosome consisting of DNA, a helicase hexamer, and a primase monomer (5-7). The only known function of gp59 is to load the helicase-primase complex, and rates of DNA synthesis in vitro are independent of the presence of gp59 (5). Upon binding a template for lagging-strand synthesis, the gp41/gp61 helicase-primase complex moves processively in the 5'3' direction (8). The helicase-DNA association at the T4 replication fork has an 11-min half-life (9). The primase synthesizes predominantly pppApCpNpNpN pentaribonucleotide primers for lagging-strand synthesis (10, 11). The T4 DNA polymerase holoenzyme, comprising the gp43 DNA polymerase, the gp44/62 clamp loader, and the gp45 clamp, catalyzes continuous leading-strand synthesis at a rate in vitro of about 400 nucleotides/s (5). This value is similar to the rate in vivo, where 5-6 min are required to replicate the 169-kb phage genome.

To account for the high efficiency of lagging-strand synthesis, which requires rapid and coordinated loading of a lagging-strand polymerase on the next primer terminus, Alberts et al. (12) suggested a model for T4 DNA replication. The key aspect of this model was that, once loaded onto a replication fork, a polymerase dimer thereafter catalyzes the synthesis of both strands. Thus, the same lagging-strand polymerase must be recycled during repetitive rounds of Okazaki-fragment synthesis. Alberts et al. (12) also suggested that the T4 DNA replication apparatus is an example of a "replicative machine" because in their model the polymerase dimer is a complex of two polymerase holoenzymes that accomplish replicative synthesis of an entire phage genome. In support of the model, they presented data showing that decreasing the polymerase concentration over a range of 34-0.4 nM did not increase the size of Okazaki fragments, as would have been expected if DNA synthesis were uncoupled. Recently, further support for this model was obtained using a synthetic 70-nucleotide circle as a template for DNA synthesis catalyzed by T4 proteins (13). Coordinated synthesis of leading and lagging strands was observed with 200 nM exonuclease-deficient (D219A) gp43. On the other hand, experiments involving the dilution of pre-formed replication complexes have not been conducted with the T4 system. Dilution of pre-formed replication complexes is a powerful method for differentiating between coupled and uncoupled modes of DNA replication because in uncoupled synthesis, lagging-strand synthesis depends on the concentration of DNA polymerase and is sensitive to dilution. Because both polymerase and additional replication proteins are involved in lagging-strand synthesis in all analyzed replication systems, dilution experiments also clarify whether these proteins, once loaded, remain bound within a replication complex or function distributively (i.e. are recruited from solution for each cycle) during repetitive cycles of Okazaki fragment synthesis.

In both the complicated Escherichia coli and the simpler phage T7 systems for replication in vitro, dilution experiments showed that leading-strand and lagging-strand DNA replication are coupled (14-17). However, such coupling is conditional in the E. coli system in the sense that the bacterial primase and clamp ( subunit) act in a distributive manner during repetitive cycles of Okazaki fragment synthesis (14-16). There are two underlying differences between the E. coli and phage T4 replication systems. First, the E. coli DNA polymerase III holoenzyme is a tightly associated complex of 14 subunits whose structure can be summarized as two core polymerases held together by a dimer of  and one -complex clamp loader (18). In contrast, neither isolation of a T4 DNA polymerase holoenzyme nor association of the purified components into a stable holoenzyme have been reported, although a gp43 affinity column retained gp43 from T4-infected cell extracts, and eluates from such columns also contained the gp45 sliding clamp (12). Second, there is no evidence for a special T4 subunit corresponding to E. coli , which physically couples two DNA polymerase III holoenzymes and an E. coli replicative helicase and thereby increases the rate of replication-fork movement from 30-35 to 500-700 nucleotides/s (19). The absence of specific strong binding between gp43 and the gp41 helicase is also probable because a gp43 affinity column does not retain gp41 from T4-infected cell extracts and vice versa (12) and because analytical ultracentrifugation also detected no gp41-gp43 interaction (20). However, direct interactions may occur between these two proteins within the replication fork, because a tryptic product of the gp41 helicase that lacks 17-20 amino acids from the COOH end has normal helicase activity but fails to function as a helicase in the T4 replication fork (21).

Here we describe coordinated synthesis of leading and lagging strands which resists extensive dilution in a reaction mixture that lacks additional T4 DNA polymerase, replicative helicase, primase, and helicase-loading protein. However, omitting the clamp loader, the sliding clamp or the ssDNA-binding protein from the dilution mixture results in uncoordinated DNA replication and/or formation of larger Okazaki fragments. These results indicate that, once loaded onto the template DNA, two DNA polymerase molecules plus the helicase-primase complex catalyze conditionally coupled replicative synthesis of both DNA strands, whereas the clamp loader, the clamp, and the ssDNA-binding protein function distributively in the synthesis of Okazaki fragments. The mechanism that couples two gp43s during T4 DNA replication appears to require DNA because the polymerase is a monomer in solution, and the only complex with other replicative proteins detected in solution is with the gp45 clamp as reported previously (22). We also observed that an unrelated DNA polymerase, Sequenase (a modified DNA polymerase of phage T7 bound to its processivity factor), can replace T4 gp43 for leading-strand synthesis but does so poorly for lagging-strand synthesis. This result further implies that specific protein-protein interactions are required at the T4 replication fork for the synthesis of Okazaki fragments.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Strains and Plasmids-- E. coli strain MV1190/pPST4Pol containing phage T4 gene 43 under the control of the tac promoter, strain MV1190/pPST4Pol(D219A) containing T4 gene 43D219A under the control of the tac promoter, strains OR1265/pDH518 and N4830/pDH911 harboring plasmids with cloned T4 genes 41 and 61, respectively, under the control of the thermosensitive phage- P_l promoter, and strain N4830 were obtained from Nancy Nossal (NIDDK, NIH, Bethesda, MD). E. coli topA strain DM800 was from James Wang (Harvard University, Cambridge, MA). Plasmid p44/62 containing T4 genes 44 and 62 under the control of the T7 RNA polymerase promoter was from Jim Karam (Tulane University Medical Center, New Orleans, LA). Plasmids p45F and pYS6 bearing T4 genes 45 and 32, respectively, under the control of the thermosensitive phage- P_l promoter, were from William Konigsberg (Yale University, New Haven, CT).

Phage T4 gene 32 was cloned under the control of the T7 RNA polymerase promoter into translation vector pET-21a. The gene 32 DNA was first polymerase chain reaction-amplified using the oligonucleotides 5'-TTGCATATGTTTAAACGTAAATCTACT-3' and 5'-TTGAGATCTAGGGTCCCCAATTAA-3'. Amplified fragments were cleaved by NdeI and BglII, purified from agarose gels, and cloned into the NdeI-BamHI sites of pET-21a, yielding plasmid p323-21a. The cloned gene 32 was confirmed by DNA sequencing.

DNA Sequencing-- Sequencing was performed using the ABI Prizm^TM dRhodamine Terminator Cycle Sequencing Ready Reaction kit and an ABI 377 DNA sequencer.

DNA Preparations-- M13mp2 phage was propagated in E. coli NR9099. Phage particles were precipitated in 4% polyethylene glycol 8000, 0.5 M NaCl, resuspended in 20 mM Tris-HCl, pH 8.0, 0.15 M NaCl, 1 mM EDTA, and centrifuged at 18,000 rpm in a Beckman JA-20 rotor for 30 min. The particles were digested with proteinase K (0.25 mg/ml) at 55 °C for 30 min. Viral DNA was precipitated in 0.5% hexadecyltrimethylammonium bromide, dissolved in TE buffer, and precipitated with ethanol. The pellet was re-dissolved in TE buffer, extracted five times with phenol/chloroform, and ethanol-precipitated.

Buffers-- TE buffer contained 10 mM Tris-HCl, pH 8.0, 1 mM EDTA. Buffer A contained 20 mM Tris-HCl, pH 7.5, 10% glycerol (w/v), 0.5 mM DTT, 0.5 mM benzamidine chloride, 0.1 mM phenylmethylsulfonyl fluoride. Buffer A_0.025 is buffer A containing 0.025 M NaCl. Buffer B contained 20 mM potassium phosphate, pH 6.8, 10% glycerol (w/v), 0.5 mM DTT, 0.5 mM benzamidine chloride, 0.1 mM phenylmethylsulfonyl fluoride.

Protein Purification-- Overproduction and purification of T4 gp43, gp43D219A, gp44/62, gp45, gp32, gp41, gp59, and gp61 were performed as described (Ref. 23 and references therein) with some modifications. Overproducing E. coli strains were grown in 2× YT broth. The AKTApurifier system (Amersham Pharmacia Biotech) was used to purify most proteins at the final step.

Because our preparations of gp32 from E. coli N4830/pYS6 contained traces of topoisomerase I activity, we used E. coli topA strain DM800 transformed with the plasmid p323-21a as a host and phage CE6 (24) as a source of T7 RNA polymerase to overexpress gene 32. The gp32 was purified by chromatography on DEAE-Sepharose, ssDNA-cellulose, and phenyl-Sepharose columns as described (23).

T4 gp41 was purified by DEAE-Sepharose chromatography. This was followed by two rounds of precipitation with 1.2 M (NH₄)₂SO₄ as described (23).

T4 gp43 and gp43D219A were first purified by phosphocellulose chromatography as described (23). The phosphocellulose product was desalted by gel filtration on PD-10 columns (Amersham Pharmacia Biotech) equilibrated with buffer A_0.025 and loaded onto an FPLC MonoQ HR 5/5 column (Amersham Pharmacia Biotech) equilibrated with the same buffer. The column was washed with 5 ml of buffer and developed with 24 ml of a linear gradient of NaCl from 25 to 150 mM; gp43 eluted at 100-140 mM NaCl.

The gp44/62 complex was first purified by phosphocellulose chromatography as described (23). The phosphocellulose product was desalted by gel filtration on PD-10 columns (Amersham Pharmacia Biotech) equilibrated with buffer A_0.075 and was passed through a ssDNA-cellulose column equilibrated with the same buffer; gp44/62 does not bind to the column under these conditions. The ssDNA-cellulose gp44/62 fraction was loaded onto a 2-ml CHT2-I ceramic hydroxyapatite column (Bio-Rad) equilibrated with buffer B. The column was washed with 5 ml of buffer B and developed with 24 ml of a linear gradient of potassium phosphate from 0.02 to 0.3 M; gp44/62 eluted at 0.2 M potassium phosphate. The hydroxyapatite gp44/62 fraction was desalted by gel filtration on a PD-10 column (Amersham Pharmacia Biotech) equilibrated with buffer A_0.025 and loaded onto an FPLC MonoS HR 5/5 column (Amersham Pharmacia Biotech) equilibrated with the same buffer; the protein passes through the column under these conditions.

T4 gp45 was first purified by DEAE-Sepharose chromatography as described (23). The DEAE-Sepharose gp45 fraction was loaded onto a 2-ml CHT2-I ceramic hydroxyapatite column (Bio-Rad) equilibrated with buffer B. The column was washed with 5 ml of buffer B and developed with a 24-ml linear gradient of potassium phosphate from 0.02 to 0.2 M; gp45 eluted at 0.1 M of potassium phosphate. The hydroxyapatite fraction of gp45 was desalted by gel filtration on PD-10 columns (Amersham Pharmacia Biotech) equilibrated with buffer A_0.025. It was then loaded onto an FPLC MonoQ HR 5/5 column (Amersham Pharmacia Biotech) equilibrated with the same buffer and developed with a 24-ml linear gradient of 0.1-0.4 M NaCl; the gp45 peak eluted at 0.23 M of NaCl.

T4 gp59 was first purified by phosphocellulose chromatography as described (23). The gp59 fraction was loaded onto a 2-ml CHT2-I ceramic hydroxyapatite column (Bio-Rad) equilibrated with buffer B. The column was washed with 5 ml of buffer B and developed with a 36-ml linear gradient of potassium phosphate from 0.02 to 0.3 M; gp59 eluted at 0.20-0.27 M potassium phosphate. The hydroxyapatite gp59 fraction was desalted by gel filtration on PD-10 columns (Amersham Pharmacia Biotech) equilibrated with buffer A_0.025. It was loaded onto an FPLC MonoS HR 5/5 column (Amersham Pharmacia Biotech) equilibrated with the same buffer and developed with a 24-ml 0.025-0.8 M NaCl linear gradient; gp59 eluted at 0.4 M NaCl.

T4 gp61 was first purified by phosphocellulose chromatography as described (23). The phosphocellulose gp61 fraction was loaded onto a 2-ml CHT2-I ceramic hydroxyapatite column (Bio-Rad) equilibrated with buffer B. The column was washed with 5 ml of buffer B and developed with a 36-ml 0.02-0.3 M linear gradient of potassium phosphate; gp61 eluted at 0.2 M potassium phosphate. The hydroxyapatite gp61 fraction was desalted by gel filtration on PD-10 columns (Amersham Pharmacia Biotech) equilibrated with buffer A_0.025. It was then loaded onto an FPLC MonoQ HR 5/5 column (Amersham Pharmacia Biotech) equilibrated with the same buffer; the protein passed through the column under these conditions. The MonoQ gp61 fraction was loaded onto an FPLC MonoS HR 5/5 column (Amersham Pharmacia Biotech) equilibrated with buffer A_0.025, and a 24-ml 0.025-0.8 M NaCl linear gradient was then applied; gp61 eluted at 0.3 M NaCl.

All proteins were free of contaminating exo- and endo-deoxyribonuclease activities. The final fractions of gp43, gp44/62, gp45, gp32, gp59, and gp61 obtained after the last chromatographic steps were dialyzed overnight against a buffer containing 20 mM Tris-HCl, pH 7.5, 50% glycerol (v/v), 0.1 M KCl, 0.5 mM DTT, 0.5 mM benzamidine chloride, 0.1 mM phenylmethylsulfonyl fluoride, 0.5 mM EDTA. In the case of the gp41 helicase, the buffer also contained 10 mM magnesium acetate. After dialysis, the fractions were subdivided and stored at 80 °C.

Protein concentrations were determined as described (25)² and are expressed in monomer molarities. In the case of the gp44/62 heteromultimer, the given molar concentration is for a complex of four subunits of gp44 and one of gp62.

Substrate for DNA Replication Experiments-- DNA annealing was performed in a final volume of 200 µl of a buffer containing 20 mM Tris acetate, pH 7.8, 8 mM magnesium acetate, 50 mM potassium glutamate, 5 mM DTT, 250 nM mp2 ssDNA (as circular chromosomes), and 360 nM 55-mer oligonucleotide (5'-GCGTACCATTTTCGATAAAAGCGCAGGCGCGAGCTGAAAAGGTGGCATCAATTCT-3') (whose 30 3' nucleotides are complimentary to the viral ssDNA) for 5 min at 40 °C. The annealed DNA was immediately used in a DNA synthesis reaction in a total volume of 1 ml of a buffer containing 20 mM Tris acetate, pH 7.8, 8 mM magnesium acetate, 50 mM potassium glutamate, 12.6 mM KCl, 5 mM DTT, 6.3% glycerol (v/v), 50 nM annealed ssDNA, 1 mM ATP, 0.2 mM dGTP, 0.2 mM dATP, 0.2 mM dTTP, 0.2 mM dCTP, 36 nM gp43, 158 nM gp44/62, 396 nM gp45, and 2.1 µM gp32 for 40 min at 37 °C. The DNA products were extracted twice with phenol-chloroform, precipitated with ethanol, dissolved in TE buffer, and centrifuged through micro-spin columns (Bio-Rad). The DNA concentration was measured spectrophotometrically by absorption at 260 nm. Neutral gel electrophoresis of 1 µg of this DNA revealed no band corresponding to ssDNA, indicating that more than 95% of the DNA was converted into a double-stranded form. To estimate the length of the 5' tails, the DNA was digested with BamHI. If no strand displacement had occurred, then 2.15-kb fragments would have appeared. Limited strand displacement was achieved by using a low ratio (42:1) of gp32 per ssDNA molecule. The observed fragments had an average size of 2.30 kb, so that the average size of the 5' tails was 150 nucleotides.

Rolling-circle Replication Assays-- DNA replication reactions catalyzed by T4 proteins were performed in a final volume of 40 µl of a standard replication mixture containing 20 mM Tris acetate, pH 7.8, 50 mM potassium glutamate, 17.5 mM KCl, 9 mM magnesium acetate, 5 mM DTT, 8.7% glycerol (v/v), 500 µg/ml bovine serum albumin, 0.2 mM dATP, 0.2 mM dTTP, 0.2 mM dGTP, 0.2 mM dCTP, 1.5 mM ATP, 1.5 mM GTP, 0.4 mM CTP, 0.4 mM UTP, 69 µCi/ml [-³²P]dGTP (3000 Ci/mmol), and 3 nM 5'-tailed mp2 double-stranded DNA. The reaction mixtures were supplemented with 9.4, 4.7, or 2.35 nM gp43, 16.5 nM gp44/62, 14.2 nM gp41 (as a hexamer), 103 nM gp45 (as a trimer), 600 nM gp32, 32 nM gp61, and 18 nM gp59. Reaction mixtures without template DNA but with all T4 proteins except gp43 and gp32 were first incubated at room temperature for 3 min. Then gp43 and gp32 were added, the mixtures were transferred to a 37 °C water bath for 1 min, pre-warmed template DNA was added at time 0, and reactions were run at 37 °C. Samples (6 µl) were withdrawn at the indicated times and mixed with 25 µl of 75 mM EDTA, 30 mM NaOH. Samples (10 µl) of the diluted reaction products were separated in 0.6% alkaline agarose gels in 30 mM NaOH, 2 mM EDTA. Reactions with the modified T7 DNA polymerase (Sequenase, Amersham Pharmacia Biotech) were carried out under the same conditions except that T4 DNA polymerase was replaced with 47.5 nM (1.1 units) Sequenase. Gels were dried, and data collection and quantification were performed using a Storm 850 PhosphorImager and the ImageQuaNT^TM program (Molecular Dynamics).

To calculate the fraction of DNA used as a replication substrate, rolling-circle replication reactions were run as above without [-³²P]dGTP but with the 3 nM DNA end-labeled with ³²P using T4 polynucleotide kinase. The fraction of DNA used as a replication substrate was calculated as the fraction (products that moved slower than the substrate band)/(sum of the products and the substrate).

To quantify dGMP incorporation, 1.5-µl samples were separated by thin-layer chromatography on polyethyleneimine plates (Merck) in 1.3 M LiCl, 1 M acetic acid. Two rectangles were drawn on each chromatogram, one surrounding a spot of incorporated dGMP, and the other surrounding the spots of unincorporated dGTP and excised dGMP (the latter was a product of the proofreading activity of the phage DNA polymerases). Backgrounds were subtracted from these values, and incorporated dGMP was calculated using the equation µM incorporated dGMP = k(value for incorporated dGMP)/(sum of values for total radioactivity), where k = 200, the final µM dGTP in the reaction buffer.

Dilution Experiments-- The standard dilution mixture contained 20 mM Tris acetate, pH 7.8, 50 mM potassium glutamate, 17.5 mM KCl, 9 mM magnesium acetate, 5 mM DTT, 8.7% glycerol (v/v), 500 µg/ml bovine serum albumin, 0.2 mM dATP, 0.2 mM dTTP, 0.2 mM dGTP, 0.2 mM dCTP, 1.5 mM ATP, 1.5 mM GTP, 0.4 mM CTP, 0.4 mM UTP, 69 µCi/ml [-³²P]dGTP (3000 Ci/mmol), 8.2 nM gp44/62, 103 nM gp45 (as a trimer), and 43.8 nM gp32. DNA replication reactions were started as described in the figure legends. After 45-60 s, 1-2 µl of the reactions were mixed, either with prewarmed dilution mixture or with a stopping solution containing 50 mM EDTA and 30 mM NaOH to achieve a final dilution of 64- or 128-fold. Diluted reaction aliquots stopped by the addition of EDTA-NaOH were used as controls to estimate levels of DNA synthesis immediately before diluting. Diluted reactions were further incubated for 5 min and stopped by adding EDTA to 50 mM and NaOH to 25 mM. These reaction samples were centrifuged through micro-spin columns (Bio-Rad) to remove unincorporated [-³²P]dGTP and were analyzed by electrophoresis in 0.6% alkaline agarose gels.

Southern Analysis-- DNA products of diluted reactions separated in 0.6% alkaline agarose gels were transferred to a Nytran nylon membrane (Schleicher & Schell) using Posiblot 30-30 Pressure Blotter (Stratagene) and hybridized to lagging-strand products with a ³²P-labeled probe according to the manufacturer's instructions. To generate the probe, a 100-µl reaction mixture containing 20 mM Tris acetate, pH 7.8, 50 mM potassium glutamate, 6 mM KCl, 8 mM magnesium acetate, 5 mM DTT, 4% glycerol, 1 mM ATP, 50 µM dATP, 50 µM dGTP, 50 µM dTTP, 50 µCi of [-³²P]dCTP (3000 Ci/mmol), 10 nM mp2 ssDNA annealed with the 55-mer oligonucleotide (5'-GCGTACCATTTTCGATAAAAGCGCAGGCGCGAGCTGAAAAGGTGGCATCAATTCT-3'), 7.5 nM exonuclease-deficient gp43D219A, 13.2 nM gp44/62, and 110 nM gp45 (as a trimer) was incubated for 5 min at 37 °C. The proteins were inactivated by heating at 75 °C for 15 min. To remove unincorporated [-³²P]dCTP, the reaction mixture was passed through micro-spin columns (Bio-Rad). The DNA was cleaved with HhaI and HaeIII, which generated a 271-base pair DNA fragment including part of the oligonucleotide. The fragment was purified through a 6% polyacrylamide gel and used to probe lagging-strand products.

Gel-exclusion Chromatography-- Gel-filtration experiments were performed using an AKTApurifier system (Amersham Pharmacia Biotech) connected with a Superdex 200 HR 10/30 column. Samples (100 µl) were loaded and separated at 0.5 ml/min. Blue dextran (2000 kDa), thyroglobulin (669 kDa), ferritin (440 kDa), aldolase (158 kDa), bovine serum albumin (67 kDa), and ovalbumin (43 kDa) were used to calibrate the column. The following buffers were used for gel-filtering 100-µl samples of 4-20 µM gp43: a high salt buffer (20 mM Tris acetate, pH 7.8, 150 mM potassium acetate, 5% glycerol, 0.5 mM DTT), the same buffer supplemented with 10 mM magnesium acetate, a medium salt buffer (20 mM Tris acetate, pH 7.8, 50 mM potassium acetate, 10 mM magnesium acetate, 5% glycerol, 1 mM DTT), and a low salt buffer (20 mM Tris acetate, pH 7.8, 5 mM magnesium acetate, 5% glycerol, 1 mM DTT).

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Effect of Dilution on Coordination of Leading-strand and Lagging-strand DNA Synthesis-- To inquire whether T4 replication forks catalyze coupled synthesis of leading and lagging strands, we performed dilutions of active T4 replication complexes. Once initiated, highly processive T4 leading-strand synthesis should be unaffected by dilution. If coupled to leading-strand synthesis, lagging-strand synthesis should also be resistant to dilution. In these experiments, we used the eight purified T4 replication proteins (Fig. 1A) together with M13mp2 double-stranded DNA with a ~150-nucleotide 5' tail to form active replication forks. The 5'-tailed substrates are preferable for assembling T4 replication complexes because the gp41/gp61 helicase-primase complex requires such tails to load efficiently (26).

View larger version (32K): [in this window] [in a new window]   Fig. 1.   Synthesis of leading and lagging strands by undiluted T4 replication complexes. A, the purified T4 replication proteins. Lanes 1-7, gp43, gp44/62, gp45, gp32, gp41, gp61, and gp59, respectively, obtained after the final purification steps are displayed using 12% SDS-PAGE and staining with Coomassie Brilliant Blue. The gel was loaded with 4 µg of gp44/62 and 2 µg of the other proteins. B, standard DNA replication reactions with 9.4 nM gp43 ± CTP + UTP and either 3 or 1.5 nM template DNA for 1, 2, and 4 min. Molecular mass markers are from a 32P-labeled HindIII digest of  DNA. C, PhosphorImager analysis of the size distributions of Okazaki fragments formed at 1 and 4 min as shown in B. a, the 32P-labeled HindIII digest of  DNA. b and c, Okazaki fragments formed at 1 and 4 min with 3 nM template DNA, respectively. d and e, Okazaki fragments formed at 1 and 4 min with 1.5 nM template DNA, respectively.

Under our standard conditions, these T4 replication proteins catalyze the efficient synthesis of long leading strands of >20 kb and short lagging strands of 0.6-7 kb when analyzed by denaturing agarose gel-electrophoresis (Fig. 1B, lanes 1-6). Electron microscopic analysis of DNA products synthesized by T4 proteins showed that they comprise duplex circles with linear multigenomic tails (27). In addition to the bands representing leading-strand and lagging-strand synthesis, a band of about 8 kb appears. This band represents limited strand-displacement synthesis by complexes that have not acquired the primosome, and the relative band intensity decreases when the concentration of template DNA is decreased (Fig. 1B). As expected, the synthesis of lagging strands depends strongly on the CTP and UTP used by the helicase-primase complex to synthesize pentaribonucleotide primers (Fig. 1B, lanes 7-9). Because C residues occur in a 1:1 ratio in the strands that template leading-strand and lagging-strand synthesis, we used radioactively labeled precursor [-³²P]dGTP to quantify DNA synthesis. Under these conditions, the synthesis of both strands is coordinated, that is, the same amounts of dGTP are incorporate into both leading and lagging strands.

The average size of Okazaki fragments depends on several factors including the reaction time and the concentration of template DNA (Fig. 1, B and C). Increasing the incubation time and/or the DNA concentration increases the average size of Okazaki fragments (Fig. 1C). Another important factor is the concentration of potassium glutamate; increasing its concentration decreases the average size of Okazaki fragments. The optimum concentration of potassium glutamate for DNA synthesis under undiluted conditions is 50-150 mM (data not shown).

We then performed dilution experiments. The dilution mixture was the standard replication mixture but without template DNA, gp43 polymerase, gp41 helicase, gp61 primase, or gp59 helicase-loading protein. In addition, the concentrations of gp32 ssDNA-binding protein and gp44/62 clamp loader were decreased 14- and 2-fold, respectively, compared with the standard replication mixture. The concentrations of these proteins were lowered to avoid an inhibition of DNA synthesis that was otherwise observed in diluted reactions (data not shown). Lane 3 of Fig. 2A shows that when clamp, clamp loader, and gp32 were present in the dilution buffer, vigorous lagging-strand synthesis of 0.6-8-kb fragments continued after dilution. These amounted to about 47% of total incorporation into both lagging and leading strands (Table I). When clamp, clamp loader, or gp32 were omitted from the dilution buffers (Fig. 2A, lanes 4-7, and Table I), the fraction of 0.6-8-kb Okazaki fragments decreased. Active E. coli replication complexes diluted in buffer lacking the cognate primase generated large Okazaki fragments, suggesting that the E. coli primase must also be recruited from solution for each initiation event (14). To determine whether larger Okazaki fragments were formed in the diluted reactions shown in Fig. 2A (lanes 4-7), we analyzed the DNA products by hybridizing with a probe to lagging-strand products (Fig. 2B). As seen with DNA replication assays conducted in the presence of -³²P-labeled dGTP and shown in Fig. 2A (lane 3), Southern hybridization revealed an efficient accumulation of 0.6-8-kb Okazaki fragments when preformed replication complexes were diluted in the standard dilution buffer (Fig. 2B, lane 3). Note that the peak size of Okazaki fragments in lane 3 of Fig. 2A is about 3.5-4 kb, whereas hybridization displays a peak size of 2-3 kb. This difference occurs simply because in the former case, more ³²P-labeled precursor molecules were incorporated into larger than into smaller Okazaki fragments, obscuring the position of the true peak. Besides the Okazaki fragments and the 7.2-kb band of M13mp2 ssDNA (Fig. 2B, lane 3), lane 4 also has a band with a mobility slightly less than that of template DNA. We suspect that this band represents the products of snap-back DNA synthesis. In those diluted reactions in which clamp loader, clamp, or gp32 were omitted, the probe revealed a substantial fraction of larger Okazaki fragments. Thus, the results of the dilution experiments indicate that the gp44/62 clamp loader, the gp45 sliding clamp, and the gp32 ssDNA-binding protein are recruited from solution for each round of synthesis of Okazaki fragments.

View larger version (21K): [in this window] [in a new window]   Fig. 2.   Coordinated synthesis of leading and lagging strands by T4 replication complexes resists dilution provided gp44/62, gp45, and gp32 are provided. A, standard DNA replication reactions were carried out with 9.4 nM gp43 and with template DNA at 3 nM. After 45 s, incubation reactions were diluted 64-fold with a stopping solution of 50 mM EDTA, 30 mM NaOH (lane 2), or with dilution mixture (lanes 3-7) and were then incubated for another 5 min. In lanes 4-7, gp44/62, gp45, or both gp44/62 and gp45 or gp32 were omitted from the dilution buffer, respectively. In lane 1, template DNA, gp43, gp32, gp41, gp61, and gp59 were prediluted to obtain final concentrations of the proteins in the reaction identical to those in the reaction diluted 64-fold (lane 3) and were then incubated in the standard dilution mixture for 6 min. B, Southern hybridization of the DNA products formed in dilution reactions with a probe specific to lagging-strand DNA. Dilution reactions were performed as in panel A but without [-32P]dGTP. Lane 1 and 2-6 show products formed before or after dilution, respectively. In lanes 3-6, gp44/62, gp45, or both gp44/62 and gp45 or gp32 were omitted from the dilution buffer, respectively. Products were analyzed by hybridization as described under "Materials and Methods." C, comparison of DNA synthesis in 64-fold diluted and undiluted reactions. Lane 1, the 64-fold diluted reaction conducted as in A, lane 3. Lane 2, the undiluted reaction with 9.4 nM gp43 diluted 64-fold after a 5.75-min incubation in standard replication buffer. D, the fraction of substrate DNA used in DNA replication. Experiments were carried out as described under "Materials and Methods."  and , reactions carried out with 9.4 and 2.35 nM T4 DNA polymerase, respectively. E, dilution reactions were carried out as in A, but the concentration of gp43 to start the reactions was 2.35 nM, and the dilution factor was 128-fold. Lane 1 shows the products of DNA synthesis formed during the first 45 s (before diluting) and then terminated by diluting into stopping solution. Lane 2 shows the products of DNA synthesis formed during the first 45 s plus after a 128-fold dilution into the standard dilution mixture during the next 5 min. Lane 3 shows the products of DNA synthesis formed with 2.35 nM gp43 diluted 128-fold after a 5.75-min incubation in standard replication buffer. F, potassium glutamate modulates the size of Okazaki fragments synthesized by T4 replication forks. Dilution experiments were carried out as in Fig. 2A. Lane 1 shows products formed before dilution. Lanes 2 and 3 show products formed after a 64-fold dilution in the standard dilution mixture containing either 50 or 150 mM potassium glutamate, respectively.

View this table: [in this window] [in a new window]   Table I Fraction of [-32P]dGTP incorporated into 0.6-8-kb lagging strands compared to overall incorporation after a 64-fold dilution Volume integration of DNA products longer than about 20 kb and shorter than the 8-kb band corresponding to circular duplex DNA were used to quantify leading-strand and lagging-strand synthesis, respectively, after a 64-fold dilution. The volume data of a particular diluted reaction were subtracted from those of a 45-s reaction, the time at which dilution was performed. Averages ± S.D. are from six experiments.

These dilution experiments included two controls. In the first, the DNA polymerase, the gp41 helicase, the gp61 primase, the gp59 helicase-loading protein, and the template DNA were prediluted and then incubated with the four remaining T4 proteins (gp32, gp44/62, and gp45) to determine whether T4 replication complexes would form at these low concentrations (Fig. 2A, lane 1). The resulting level of DNA synthesis was 15-20-fold lower than in the standard dilution reactions (Fig. 2A, lane 3), indicating that few T4 replication complexes could form. In the second control, reaction products formed by the time that dilutions were started were diluted into a stopping mixture containing 50 mM EDTA and 30 mM NaOH to estimate the level of DNA synthesis immediately before diluting (Fig. 2A, lane 2). Clearly, most of the DNA synthesis recorded in lane 3 occurred after dilution. Incorporation of [-³²P]dGMP into leading and lagging strands increased by 6.4 ± 0.6-fold after 5 min in the diluted reaction. Comparing DNA synthesis in the reaction diluted 64-fold in the standard dilution buffer with the undiluted reaction showed that about 3.6-fold less DNA was synthesized under diluted conditions (Fig. 2C). However, new replication complexes continue to load throughout the 5.75 min of the undiluted reaction, and as a result, 2.8-fold more template DNA is used to form replication complexes than by the 45s when the dilutions were made (Fig. 2D).

Dilution experiments demonstrating coupling between leading and lagging strands have also been performed with E. coli DNA polymerase III (14-16) and with T7 DNA polymerase (17). The final concentration of T4 DNA polymerase in our 64-fold dilution experiments was 2- and 30-fold lower, respectively, than in the E. coli and T7 experiments. Because average Okazaki fragment size in the reactions diluted 64-fold in the standard dilution buffer was about 2-fold greater than in the undiluted reaction (Fig. 2C), we sought to explore even greater dilutions to test as severely as possible the hypothesis that coordinated synthesis does not depend on the concentration of phage T4 DNA polymerase. This was done in two steps, first by decreasing the gp43 concentration in the standard replication mixture by 4-fold and then by diluting 128-fold instead of 64-fold in the standard dilution buffer (Fig. 2E). Such diluted reactions produced Okazaki fragments of sizes similar to those of undiluted reactions (Fig. 2E, lanes 2 and 3). Quantification of the resulting DNA synthesis revealed a ratio of 0.6-8-kb lagging-strand incorporation as a fraction of the sum of lagging-strand and leading-strand incorporation of 46.1 ± 3.5% with 5.1 ± 0.5-fold increased total incorporation into leading and lagging strands after dilution. DNA synthesis into leading and lagging strands in the reactions diluted 128-fold (Fig. 2E, lanes 2) was 4.7 times less than that of undiluted reactions (Fig. 2E, lanes 2), reflecting the fact that 3.8 times more template DNA was used to form additional replication complexes in the undiluted reactions. Thus, coordinated synthesis remains resistant to the highest dilution compatible with our conditions.

Because a higher concentration of potassium glutamate decreases the average size of Okazaki fragments in undiluted reactions, we asked whether this higher concentration produces the same effect in diluted reactions. The results of such an experiment are shown in Fig. 2F. The average size of Okazaki fragments formed at 150 mM potassium glutamate is clearly less than at 50 mM. Although synthesis remains coordinated at 150 mM, total incorporation in leading and lagging strands was 1.5-fold lower than at 50 mM, the concentration in our standard replication mixture.

Taken together, these dilution experiments indicate that the synthesis of leading and lagging DNA strands catalyzed by the eight T4 replication proteins is conditionally coupled, that is, coupled provided that the dilution buffer is supplemented with the gp44/62 clamp loader, the gp45 clamp, and the gp32 ssDNA-binding protein.

Sequenase Can Replace T4 DNA Polymerase for Leading-strand Synthesis but Not for Coordinated Lagging-strand Synthesis-- We wished to test whether specific protein-protein interactions are important for coordinated synthesis of leading and lagging strands at T4 replication forks. To this end, we asked whether an unrelated polymerase, Sequenase (a derivative of phage-T7 DNA polymerase), is able to replace T4 DNA polymerase in reactions carried out in the presence of the other T4 replication proteins. The results of such a test using undiluted reaction mixtures together with control reactions are shown in Fig. 3A. Sequenase polymerase activity was stimulated by gp32 (compare lanes 1-3 with lanes 4-6). When Sequenase was incubated with gp32, gp41, gp61, and gp59, synthesis of both leading and lagging strands was observed (lanes 7-9). Adding the gp44/62 clamp-loader and the gp45 clamp (lanes 10-12) had no major effect. Lagging-strand synthesis was abolished upon omitting CTP and UTP (lanes 13-15). Quantification of DNA synthesis showed that 0.6-8-kb lagging-strand synthesis as a fraction of leading-strand plus lagging-strand synthesis was 14.1 ± 4.8% at 2 min and 25.7 ± 1.9% at 4 min (Fig. 3A, lanes 8 and 9). Fig. 3B shows total dGMP incorporation in Sequenase reactions containing either 47.5 or 23.75 nM Sequenase compared with reactions containing either 4.7 or 2.35 nM T4 DNA polymerase. Total DNA synthesis with 47.5 nM Sequenase and 2.35 nM T4 polymerase was about the same, indicating that 20-fold more Sequenase than T4 DNA polymerase is required for similar rates of DNA synthesis.

View larger version (20K): [in this window] [in a new window]   Fig. 3.   Sequenase can replace T4 DNA polymerase for leading-strand synthesis. A, Sequenase at 47.5 nM was incubated alone (lanes 1-3) or in the presence of the indicated T4 proteins (lanes 4-15) for the indicated times in the standard replication mixture. CTP and UTP were omitted from the reactions shown in lanes 13-15. B, total dGMP incorporation in reactions with T4 DNA polymerase holoenzyme or Sequenase. T4 DNA polymerase at 4.7 nM () or 2.35 nM () was incubated with the other seven T4 proteins in the standard replication mixture. Reactions with Sequenase at 47.5 nM () or 23.75 nM () were carried out under the same conditions as with T4 DNA polymerase except that gp44/62 and gp45 were omitted. C, Sequenase was incubated with gp32, gp41, gp61, and gp59, and after 1 min, the replication mixture was diluted 64-fold with 50 mM EDTA, 30 mM NaOH (lane 2) or with the standard dilution mixture (which lacked gp32, gp44-gp62, and gp45) (lane 3) and incubated for 5 min. In lane 1, template DNA, Sequenase, gp32, gp41, gp61, and gp59 were prediluted to obtain final concentrations identical to those in the 64-fold diluted reactions and were incubated in the dilution mixture for 6 min. Lane 4, 64-fold diluted sample of undiluted reaction mixture was incubated for 6 min.

To test whether complexes formed with Sequenase plus the T4 gp41/61 primosome are resistant to dilution, we carried out the experiments shown in Fig. 3C. The standard dilution mixture was the same as the standard replication mixture but without template DNA, Sequenase, or any T4 proteins. Surprisingly, these experiments showed that total DNA synthesis by Sequenase plus T4 primosome was resistant to dilution (lane 3 versus lane 2), with 3.2 ± 0.3-fold increased incorporation in leading and 0.6-8-kb lagging strands after dilution and with lagging-strand synthesis comprising 25.0 ± 2.7% of total synthesis. DNA synthesis into leading and lagging strands in the 64-fold diluted reactions (Fig. 3C, lanes 3) was 5.1 times less then that in undiluted reactions (Fig. 3C, lanes 4). Because Sequenase can replace T4 DNA polymerase for leading-strand synthesis but less well for lagging-strand synthesis, specific protein-protein interactions at T4 replication forks are likely to be more important for lagging-strand synthesis than for leading-strand synthesis.

T4 DNA Polymerase Is a Monomer in Solution-- Because coordinated T4 DNA replication continues under conditions of high dilution, we tested whether gp43 alone can form a dimer detectable by gel-exclusion chromatography. We used several chromatography buffers based on 20 mM Tris-HCl, pH-7.5, 5% glycerol, 0.5-2 mM DTT. These were then supplemented to produce a high salt buffer containing 150 mM KCl, the same plus 10 mM magnesium acetate, a medium salt buffer containing 50 mM potassium acetate plus 10 mM magnesium acetate, and a low salt buffer containing 5 mM magnesium acetate. Under all these conditions, gp43 gel-filtered as a monomer (Fig. 4). Using gel-exclusion chromatography, we also separated T4 DNA polymerase after preincubation with the other seven T4 replication proteins in various combinations. The only complex we observed was between gp43 and the gp45 clamp, which co-eluted with an apparent molecular mass of 150 kDa (results not shown). Subsequent gel-electrophoretic analysis of this peak showed that gp45 and gp43 co-eluted at ratios that differed in different fractions, suggesting that the half-life of the complex is less than the time of chromatography. This result is consistent with the observed molecular weight of the complex, 150 kDa, compared with its calculated molecular weight, 185 kDa. Taken together, these results indicate that interactions among T4 replication proteins in solution are weak and that DNA is required for the efficient assembly of T4 replication complexes.

View larger version (11K): [in this window] [in a new window]   Fig. 4.   T4 DNA polymerase is a monomer in solution as judged by gel filtration. A Superdex 200 high resolution 10/30 column was equilibrated with 20 mM Tris acetate, pH 7.8, 150 mM KCl, 10 mM magnesium acetate, 5% glycerol, 0.5 mM DTT and was calibrated with thyroglobulin (669 kDa), ferritin (440 kDa), aldolase (158 kDa), bovine serum albumin (67 kDa), and ovalbumin (43 kDa), designated 1, 2, 3, 4, and 5, respectively. Kav = (Ve  V0)/(Vt V0), where Ve = elution volume for the protein, V0 = column void volume, and Vt = total bed volume. MW, molecular weight.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Alberts et al. (12) suggested that two T4 gp43 DNA polymerase molecules remain coupled once loaded onto template DNA and then catalyze the coordinated replication of the leading and lagging strands of an entire genome. Based on our experience, however, the conditions in the early T4 experiment were not sufficient to test the model. Diluting active replication complexes of both phage T7 and E. coli showed that coordinated DNA synthesis of leading and lagging strands in those systems is resistant to dilution (14-17). The importance of the dilution method for characterizing DNA replication was highlighted by the finding that Sequenase can replace E. coli DNA polymerase III holoenzyme for leading-strand and lagging-strand synthesis in undiluted reactions but not after dilution of pre-formed replication complexes (16).

There are no previous reports of high dilution experiments using active T4 replication complexes catalyzing the coordinated synthesis of leading and lagging strands. Here, we show that active T4 replication complexes are resistant to high dilution provided that the dilution buffer is supplemented with the gp44/62 clamp loader, the gp45 clamp, and the gp32 ssDNA-binding protein. When any of those proteins is omitted from the dilution buffer, DNA synthesis becomes uncoordinated. These results suggest that gp44/62, gp45, and gp32 are derived from solution for each round of Okazaki fragment synthesis in vitro, although the action in vivo of an as yet unidentified protein linking these proteins physically to the lagging-strand complex is not excluded. Omitting gp32 has less impact on lagging-strand synthesis than omitting the other accessory proteins, suggesting that the role of gp32 in lagging-strand synthesis is less crucial than that of gp44/62 and gp45. However, we cannot exclude the possibility that gp32 is sometimes recycled because of its ability to bind ssDNA cooperatively (28).

In E. coli, the analog of gp45 is the  clamp, which also functions distributively during the synthesis of Okazaki fragments, whereas the analog of gp44/62 is the  complex, which functions processively (16). This distributive behavior of the T4 clamp, clamp loader, and gp32 suffices to explain the increasing average size of Okazaki fragments with increasing time of incubation and/or substrate concentration in undiluted reactions. This increased Okazaki fragment synthesis then decreases the ratio of accessory proteins per Okazaki fragment initiation event, which in turn increases the time required to initiate new Okazaki fragments and, thus, increases the average size of Okazaki fragments. The same reasoning can explain the larger Okazaki fragments in reactions started with 9.4 nM gp43 and then diluted 64-fold in our standard dilution buffer than in reactions started with 2.35 nM gp43 and then diluted 128-fold in our standard dilution buffer or in reactions started with 9.4 nM gp43 and then diluted 64-fold in dilution buffer containing 150 mM potassium glutamate. In the latter cases, Okazaki fragment synthesis was reduced so that the ratio of accessory proteins per initiation event increased, thus decreasing the average fragment size.

The fraction of active replication complexes that survive 64- and 128-fold dilution remains unknown. Direct comparisons of the diluted and undiluted reactions show 3.8- and 4.7-fold less incorporation into leading plus lagging strands in 64- and 128-fold diluted reactions, respectively, than in the corresponding undiluted reactions. However, this does not mean that only 26 or 21% of preformed complexes survive dilution because in undiluted reactions, replication proteins continue to form additional replication complexes during the entire reaction. For example, at 2 min in undiluted reactions, 2-fold more complexes had formed than by the 45 s, when the dilutions were started (Fig. 2D). These new complexes considerably increase the difference between the undiluted and diluted reactions in incorporation into both leading and lagging strands. It also should be noted that measurements of the fraction of DNA used in replication (Fig. 2D) do not provide information about the fraction of complexes, if any, that were loaded on pre-extended DNA where replication had collapsed. Such complexes would also increase the difference in incorporation between the diluted and undiluted reactions.

A recent report (13) analyzed DNA replication catalyzed by D219A exonuclease-deficient gp43 plus the other seven T4 proteins using a 70-nucleotide minicircle substrate. This minicircle lacks dG residues. When ddCTP was used to inhibit lagging-strand synthesis, leading-strand synthesis was also strongly inhibited, suggesting strong coupling. In similar experiments using a larger minicircle substrate, we observed that when lagging-strand synthesis was inhibited, leading strand was also inhibited, but only moderately. Moreover, in the absence of lagging-strand synthesis (i.e. in reactions lacking CTP and UTP), leading-strand synthesis was also moderately inhibited. However, the latter effect was not observed using an M13 DNA substrate.³ Therefore, we do not yet understand the properties of minicircle systems sufficiently well to interpret the results they generate.

T7 DNA polymerase differs from most other DNA polymerases in that T7 does not encode its own processivity factor. Instead, T7 DNA polymerase binds strongly to host thioredoxin, which endows the complex with high processivity (29). Sequenase retains the bound thioredoxin and, thus, requires only a helicase to conduct strong leading-strand synthesis. Our results with Sequenase demonstrate that it can efficiently replace the T4 holoenzyme for leading-strand synthesis. However, because lagging-strand synthesis by Sequenase is poorly coordinated with leading-strand synthesis, protein-protein and/or protein-RNA primer interactions appear to be essential for coordination. At T7 replication forks, the ssDNA-binding protein encoded by T7 gene 2.5 is absolutely required for coordinating lagging-strand synthesis with leading-strand synthesis (30).

Among unrelated DNA polymerases, only Sequenase (but neither Klenow fragment of E. coli DNA polymerase I nor T7 DNA polymerase without thioredoxin, both nonprocessive enzymes) can benefit from the presence of gp41 helicase in leading-strand synthesis (20). This result suggests that interactions between the leading-strand T4 DNA polymerase and the gp41 helicase hexamer are important for forming the complex. Alternatively, for processive DNA synthesis by a polymerase-helicase mixture, merely trailing a processive polymerase behind a helicase may suffice to mimic a complex between a helicase and a processive polymerase.

Early experiments with affinity column chromatography showed that a gp43 column efficiently binds soluble gp43 (12). A more recent analysis using a two-hybrid system suggests that T4 gp43 can exist as a dimer, and deletion and point mutation analyses further suggest that positions 401-600 contain the residues that are required for dimerization (13). However, neither our gel-filtration studies nor very recent ultracentrifugation studies (20) were able to detect a stable gp43 dimer in solution in the absence of DNA. Furthermore, in cross-linking experiments using glutaraldehyde, although some cross-linked gp43 dimers accumulated, trimers and tetramers also accumulated, suggesting that the interactions are nonspecific.³

The only complex between T4 gp43 and another T4 replication protein that we could detect by gel filtration is between gp43 and the gp45 clamp, but this complex is rather unstable. This complex was detected previously using ssDNA-cellulose chromatography with the crowding agent polyethylene glycol (22); in the absence of polyethylene glycol, the complex was not detected. DNA is required for the proteins of the helicase-primase complex to form a stable primosome, and these proteins do not form a complex in solution (Ref. 7 and this study). Thus, protein-protein contacts within the T4 replication complex seem to form primarily upon loading onto template DNA.

In E. coli, DNA polymerase holoenzyme III forms a stable 14-subunit complex consisting of two polymerase cores held together by the  subunit (18). The  subunit also interacts with the DnaB replicative helicase (19). Thus,  connects both the helicase and polymerase components of the E. coli replication complex. This connection dramatically increases the rate of movement of the replication complex. No analogs of the  subunit have been described in phages T4 or T7. Because rates of DNA synthesis in the reconstituted T4 and T7 systems in vitro agree well with the corresponding rates in vivo, it seems unlikely that such an analog exists. There is also no evidence that the T7 DNA polymerase can dimerize. Accordingly, it was suggested that the leading-strand and lagging-strand polymerases in the T7 replication fork are associated with the T7 gene 4 helicase-primase and do not directly associate with each other (17). The same pattern was suggested for the T4 replication complex (26). Further studies should elucidate whether the polymerase dimer forms upon loading onto DNA or whether the two molecules are only indirectly connected through the helicase-primase complex.

	ACKNOWLEDGEMENTS

We thank Jim Karam, William Konigsberg, Nancy Nossal, and James Wang for providing strains and plasmids and Kirill Lobachev for assistance with the hybridization experiments. We are grateful to Matt Longley for help and advice throughout the study and to Nancy Nossal, Matt Longley, and Ben Van Houten for critical readings of the manuscript.

	FOOTNOTES

^* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

To whom correspondence should be addressed: Laboratory of Molecular Genetics E3-01, NIEHS, P. O. Box 12233, Research Triangle Park, NC 27709. Tel.: 919-541-3029; Fax: 919-541-7613; E-mail: kadyrov@niehs.nih.gov.

Published, JBC Papers in Press, June 4, 2001, DOI 10.1074/jbc.M101310200

³ F. A. Kadyrov and J. W. Drake, unpublished results.

² The procedure is also described at www.basic.nwu.edu/biotools/proteincalc.html.

	ABBREVIATIONS

The abbreviations used are: gp, growth protein; DTT, dithiothreitol; ssDNA, single-stranded DNA; kb, kilobase(s); FPLC, fast protein liquid chromatography.

	REFERENCES

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

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