©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
DnaX Complex of Escherichia coli DNA Polymerase III Holoenzyme
CENTRAL ROLE OF IN INITIATION COMPLEX ASSEMBLY AND IN DETERMINING THE FUNCTIONAL ASYMMETRY OF HOLOENZYME (*)

(Received for publication, June 2, 1995; and in revised form, August 23, 1995)

H. Garry Dallmann (§) Roberta L. Thimmig Charles S. McHenry (¶)

From the Department of Biochemistry, Biophysics and Genetics and Graduate Program in Molecular Biology, University of Colorado Health Sciences Center, Denver, Colorado 80262

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
REFERENCES

ABSTRACT

The alternative forms of the DnaX protein found in Escherichia coli DNA polymerase III holoenzyme, and , were purified from extracts of strains carrying overexpressing plasmids mutated in their frameshifting sequences such that they produced only one subunit or the other. The purified subunits were used to reconstitute the and complexes which were characterized by functional assays. The complex-reconstituted holoenzyme required a stoichiometric excess of DNA polymerase III core, beyond physiological levels, for activity. The subunit stimulated the complex 2-fold, but could not be used to reconstitute a holoenzyme with complex and stoichiometric quantities of core. In the presence of adenosine 5`-O-(3`-thiotriphospate) (ATPS), the DNA polymerase III holoenzyme behaves as an asymmetric dimer; it can form only initiation complexes with primed DNA in one-half of the enzyme (Johanson, K. O., and McHenry, C. S.(1984) J. Biol. Chem. 259, 4589-4595). An asymmetric distribution of two products of the dnaX gene, and , has been postulated to underlie the asymmetry of holoenzyme. To provide a direct test for this hypothesis, we reconstituted holoenzyme containing only the or DnaX proteins. We observed that, although could function in the presence of ATP and high concentrations of DNA polymerase III core, it was nearly inert in the presence of ATPS. In contrast, -containing holoenzyme behaved exactly like native holoenzyme in the presence of ATPS. These results implicate as a key component required to reconstitute holoenzyme with native behavior and show that plays a key role in initiation complex formation. These results also show that is not a necessary component, since all of the known properties of native holoenzyme can be reproduced with a 9-subunit -holoenzyme.


INTRODUCTION

The DNA polymerase III holoenzyme (^1)of Escherichia coli can serve as a prototype for replicative complexes in all cells. Like many other complex mechanisms of macromolecular synthesis, the fundamental mechanisms of DNA replication are conserved throughout biology. The chemistry and direction of synthesis, the requirement for RNA primers, the mechanisms of semi-discontinuous replication with Okazaki fragments on the lagging strand, and the need for well defined origins are shared. There are also striking similarities between the individual components of the complex machinery responsible for this process.

The replicative polymerase assumes its special role because of its ability to interact with other specialized proteins at the replication fork. In E. coli, the DNA polymerase III core (alpha--) interacts with a beta subunit sliding clamp to enable maximum processivity (LaDuca et al., 1986). The gene 45 protein of T4 and eukaryotic PCNA exhibit similar properties (Burgers and Yoder, 1993; Reddy et al., 1993). The sliding clamp in all replication systems is a bracelet-like multimeric protein clasped around DNA. It contacts the polymerase, providing a tether to increase processivity. The crystal structures of yeast PCNA and E. coli beta are nearly superimposable (Kong et al., 1992; Krishna et al., 1994). Functional and structural studies of the T4 gene 45 sliding clamp indicate that it forms similar structures and functions like beta and PCNA (Hacker and Alberts, 1994; Kaboord and Benkovic, 1993; Gogol et al., 1992; Venkatesan and Nossal, 1982).

In E. coli, the beta sliding clamp is set by either of two DnaX complexes that contain either of two products of the dnaX gene, or , in a complex with -` and - (McHenry et al., 1986; Maki and Kornberg, 1988; O'Donnell and Studwell, 1990). The activity of this complex requires the ATPase activity of the DnaX proteins, presumably to couple ATP hydrolysis to beta assembly on primed DNA (Lee and Walker, 1987; Hawker and McHenry, 1987; O'Donnell et al. 1993, Oberfelder and McHenry, 1987). Two additional proteins, and `, are required to assemble a functional clamp loader (Onrust and O'Donnell, 1993). The - proteins are also participants; they bind to DnaX and increase its affinity for -` so that they can cooperatively assemble a functional complex at physiological protein levels (Olson et al., 1995). In eukaryotes, a 5-protein complex (Activator 1, RFC) is responsible for transferring the sliding clamp onto a primer terminus in an ATP-dependent reaction (Lee et al., 1991; Bunz et al., 1993). The proteins exhibit sequence homology between eukaryotes and both Gram-negative and -positive prokaryotes, and would be expected to act by a similar mechanism (Carter et al., 1993; O'Donnell et al., 1993).

Asymmetric replicative complexes have been proposed in both eukaryotic and prokaryotic systems. In eukaryotes the polymerase has been suggested as the leading strand polymerase and as the lagging strand (Turchi and Bambara, 1993; Araki et al., 1992; Hübscher and Thömmes, 1992; Burgers, 1991; Nethanel and Kaufmann, 1990). In E. coli, the replicative complex apparently forms a dimer (McHenry, 1982; Studwell-Vaughan and O'Donnell, 1991) that behaves asymmetrically (Johanson and McHenry, 1984; McHenry and Johanson, 1984), and proposals have been made concerning the enzyme having a distinguishable leading and lagging strand, capable of performing the distinct specialized functions expected of each polymerase (Hawker and McHenry, 1987; McHenry, 1988; Maki et al., 1988). The initial observation that provided functional evidence for asymmetry and led to the formulation of the asymmetric dimer hypothesis came from the use of the ATP analog ATPS in initiation complex formation in native holoenzyme in a concerted reaction where the DnaX complex assembled the beta clamp in the presence of associated polymerase (Johanson and McHenry, 1984; McHenry and Johanson, 1984). All of the enzyme formed initiation complexes in the presence of ATP, but only one-half of the enzyme entered functioning initiation complexes in the presence of ATPS (Johanson and McHenry, 1984). This asymmetry in the presence of ATPS was not an equilibrium artifact, since the 50/50 distribution was independent of ATPS concentration once the enzyme was saturated, and the reactions did not progress beyond the 50% level with increasing time. It was proposed that this asymmetric behavior arose not from two distinct populations of enzymes in solution, but from a difference between two halves of an asymmetric dimeric DNA polymerase with distinguishable leading and lagging strand halves.

Efforts to determine the basis of this functional asymmetry have focused on the two products of the dnaX gene, and (Hawker and McHenry, 1987; McHenry, 1988; Maki et al., 1988; O'Donnell and Studwell, 1990). Translation of dnaX mRNA produces full-length (71,000 Da) and a product resulting from -1 translational frameshifting into a frame with a stop codon, (47,400 Da) (McHenry et al., 1989; Tsuchihashi and Kornberg, 1990; Blinkowa and Walker, 1990; Flower and McHenry, 1990). These two products were found within the same holoenzyme assembly (Hawker and McHenry, 1987) and it was suggested that an asymmetric holoenzyme may arise from an asymmetric assembly of these two DnaX proteins (Hawker and McHenry, 1987; McHenry, 1988; Maki et al., 1988). Consistent with this hypothesis, has been shown to influence synthesis on the two strands asymmetrically on natural coupled replication forks (Wu et al., 1992).

Despite the attractive hypothesis concerning asymmetric placement of and within holoenzyme, a definitive test has not been made. In this report, we describe our use of a DnaX overproducing strain engineered to produce only or . This enabled purification of , free of possible proteolytic breakdown products of , and provided a rich source of the subunit, free of possible contaminating . We used these purified subunits to explore the functional differences in their contribution to the holoenzyme reaction and to definitively test whether DnaX protein placement underlies the functional asymmetry of holoenzyme.


EXPERIMENTAL PROCEDURES

Plasmids and Strains and Manipulations

Plasmid pJF119EH was obtained from Peter Fürste (Fürste et al., 1986) and pDM1 was obtained from Mead et al.(1985). Plasmid pMWZ1101, containing the wild-type apt, dnaX genes, and a portion of orf12, was constructed by M. Welch in this laboratory. E. coli strain HB101 (F, Delta(gpt-proA)62, leuB6, supE44, ara-14, galK2, lacY1, Delta(mcrC-mrr) rpsL20(Str^r), xyl-5, mtl-1, recA13) was used for transformations and for protein expression. Oligonucleotide 1 (see Fig. 1A) was inserted into EcoRI- and PstI-digested plasmid pJF119EH. Oligonucleotide 1 was constructed such that it introduced an optimal Shine-Dalgarno sequence (Reznikoff and Gold, 1986), a start codon, frequently used codons in place of the rare codons in the initial dnaX sequence, and a NarI cloning site (used in the next cloning step). The remaining dnaX sequence was obtained from a NarI-PstI fragment of pMWZ1101. The resulting plasmid, pBBMD11, produced both and when induced with isopropyl-beta-D-thiogalactoside.


Figure 1: Construction of overexpression plasmids that produce only or . A, plasmid map of plasmid pRT610. Plasmid pRT610 was constructed as described under ``Experimental Procedures,'' and expresses both the and subunits upon induction with isopropyl-beta-D-thiogalactoside. B, site-directed mutagenesis of dnaX. Plasmid pRT610 was mutagenized using oligonucleotides 2 and 3 to produce plasmids that expressed only or , as described under ``Experimental Procedures'' and ``Results.''



To provide a plasmid suitable for production of single-stranded DNA for mutagenesis, we cloned the fl phage origin (obtained as a HincII fragment from pDM1) into the NaeI site of PBBMD11 resulting in plasmid pRT610. Plasmid pRT610 was transformed into strain XL-1 Blue (recA1, endA1, gyrA96, thi, hsdR17 (rk,mk+), supE44, relA1, , lac, {F`, proAB, lacI^qZDeltaM15, Tn10(tet^r)}, Stratagene, La Jolla, CA) to obtain single-stranded DNA. Oligonucleotide-directed in vitro mutagenesis of pRT610 was performed as described by Kunkel (1985) and Kunkel et al.(1987). The resulting plasmids were sequenced to verify the incorporation of the proper mutations.

Buffers

These were: buffer A (50 mM Tris-HCl (pH 7.5), 10% (w/v) sucrose, 5 mM dithiothreitol, 5 mM benzamidine, 0.5 mM phenylmethylsulfonyl fluoride); buffer B (50 mM Tris-HCl (pH 7.5), 10% (w/v) sucrose, 2 M NaCl, 0.3 M spermidine); buffer TBP (50 mM Tris-HCl (pH 7.5), 20% (w/v) glycerol, 5 mM dithiothreitol, 5 mM benzamidine, 0.5 mM phenylmethylsulfonyl fluoride); buffer SP (50 mM Tris-HCl (pH 7.5), 50 mM NaCl, 10% (w/v) glycerol, 5 mM dithiothreitol); and buffer N (20 mM sodium phosphate (pH 7.4), 10% glycerol, 50 mM NaCl and 5 mM MgCl(2)) and Buffer H (25 mM Hepes-KOH (pH 7.5), 25 mM NaCl, 5% glycerol, 0.1 mM EDTA).

Growth and Induction of E. coli Overproducing Strains

The and subunit overproducing strains, HB101/pRT610-A and HB101/pRT610-B, were grown in 200 liters of F medium (14 g/liter yeast extract, 8 g/liter tryptone, 12 g/liter K(2)HPO(4), 1.2 g/liter; KH(2)PO(4), pH 7.2) + 1% glucose, and 50 µg/ml ampicillin at 37 °C. Cells were induced (OD = 1.2) with isopropyl-beta-D-thio-galactoside (1 mM final concentration) for 3 h before harvesting (OD = 3.1). Cells were harvested in a Sharples continuous flow centrifuge and resuspended in buffer A at 50% (w/v). Cells were flash frozen in liquid nitrogen and stored at -80 °C.

Cell Lysate Preparation and Ammonium Sulfate Precipitation

All procedures were performed at 4 °C unless otherwise stated. For both the and purifications, 120 g of frozen cells (240 g of cell paste) were resuspended in 330 ml of buffer A and 30 ml of buffer B. Lysozyme (120 mg in 8 ml of buffer A) was added, and cells were lysed for 1 h on ice and for 4 min at 37 °C. The lysate was centrifuged at 22,000 times g for 1 h to remove debris. Supernatants were pooled to yield fraction I (Cull and McHenry, 1995).

Fraction I was precipitated with 107 g of ammonium sulfate (0.226 g for each ml of fraction I, 40% saturation) and centrifuged at 22,000 times g for 30 min. Pellets were backwashed by resuspension in a Dounce homogenizer with 100 ml of buffer TBP + 0.1 M NaCl containing 0.2 g/ml ammonium sulfate (35% saturation), and recentrifuged as before. A second backwash using 50 ml of 0.17 g/ml ammonium sulfate (30% saturation) in buffer TBP + 0.1 M NaCl was done as above. The final pellets were flash frozen in liquid nitrogen and stored frozen at -80 °C as fraction II.

Purification of the Subunit

SP-Sepharose Ion Exchange Chromatography

One-half of fraction II pellets were dissolved in buffer SP and diluted to a conductivity of 50 mM NaCl (226 ml). This material was loaded onto a 900-ml SP Sepharose (Pharmacia Biotech Inc., 7.5 times 20 cm) column equilibrated with buffer SP. After loading, the column was washed with 1-column volume of buffer SP and developed with a 9-column volume gradient of 50-350 mM NaCl in buffer SP at a flow rate of 160-ml/h; 160-ml fractions were collected. Contaminant proteins did not bind to the column and were eluted during the initial wash. (fractions 31-33, conductivity = 250 mM NaCl equivalents) eluted as the only peak during the gradient. Fractions were pooled at one-half peak height (by activity) as fraction III.

Sephacryl S-400 HR Gel Filtration Chromatography

Fraction III was precipitated with ammonium sulfate (55% saturation) and centrifuged at 28,000 times g for 30 min. The fraction III pellet was dissolved in 19 ml of buffer H and centrifuged (28,000 times g, 30 min) to clarify. Fraction III was then applied to a 700-ml S-400 HR (Pharmacia) column (44:1 height:diameter ratio) equilibrated with buffer H. The column was developed in the same buffer at a flow rate of 22 ml/h, and 20-ml fractions were collected. (fractions 28 and 29) eluted as the only peak. These fractions were pooled, distributed in aliquots, which were flash frozen in liquid nitrogen and stored at -80 °C as fraction IV.

Purification of the Subunit

Q-Sepharose Chromatography

One-half of fraction II pellets were dissolved in buffer SP and diluted to a conductivity of 50 mM NaCl (244 ml). This material was loaded onto a 900-ml Q Sepharose (Pharmacia, 7.5 times 20 cm) column equilibrated in buffer SP. After loading, the column was washed with 1-column volume of buffer SP, then developed with a 9-column volume gradient of 50-400 mM NaCl in buffer SP at a flow rate of 160 ml/h, and 160-ml fractions were collected. (fractions 38-40, conductivity = 330 mM NaCl equivalents) was completely separated from a minor contaminant protein (40 kDa) which eluted five fractions earlier. Fractions were pooled at one-half peak height (by activity, fractions 38-40) as fraction III.

Sephacryl S-300 HR Gel Filtration Chromatography

Fraction III was precipitated with ammonium sulfate (55% saturation) and centrifuged at 28,000 times g for 30 min. The fraction III pellet was dissolved in 11 ml of buffer H and centrifuged (28,000 times g, 30 min) to clarify. Fraction III was applied to a 700-ml Sephacryl S-300 HR (Pharmacia) column (44:1 height:diameter ratio) equilibrated with buffer H. The column was developed in the same buffer at a flow rate of 17 ml/h, and 21-ml fractions were collected. (fractions 21-28) eluted as a single broad peak. These fractions were pooled and stored at -80 °C as fraction IV in small aliquots.

and Subunit Activity Assays

Assays of the and subunits contained 4 pmol of core (unless otherwise stated), 400 fmol of beta, 540 pmol of M13G (as nucleotide), 500 fmol each of , `, and , 60 units of DnaG primase, and 1.6 µg of E. coli single-stranded DNA binding protein (SSB). Reactions were carried out in a volume of 25 µl in a buffer containing 50 mM Hepes-KOH (pH 7.5), 10% (v/v) glycerol, 0.1 M potassium glutamate, 10 mM dithiothreitol, 10 mM magnesium acetate, 200 µg/ml bovine serum albumin, 0.02% (v/v) Tween 20, 10 µM ATP, 48 µM dATP, dCTP, and dGTP, and 18 µM [^3H]TTP (specific activity 520 cpm/pmol dTTP). Assays were incubated at 30 °C for 5 min and quenched by trichloroacetic acid precipitation. The unit definition for and activity is 1 pmol of total nucleotide incorporated into acid insoluble DNA in a 5-min reaction at 30 °C.

Reconstitution of and Complexes

The and complexes used here were prepared and assayed as described in Dallmann and McHenry(1995).

Initiation Complex Formation Assays

Initiation complexes were assayed as described by Johanson and McHenry(1984). Holoenzyme was reconstituted with core (typically 240 fmol unless otherwise stated), beta (400 fmol), and the or complex (100 fmol) and compared to native holoenzyme. The above proteins were mixed with DnaG primase-primed, SSB-coated, M13Gori in the presence of nucleotide (ATP or ATPS), and incubated for 5 min at 30 °C. Anti-beta-IgG (8 µg) was added as a trap to prevent further initiation complex formation (Johanson and McHenry, 1984). Initiation complexes were quantitated by measuring DNA synthesis after the addition of dATP, dCTP, and dGTP (to 48 µM) and [^3H]TTP (to 18 µM; specific activity, 520 cpm/pmol dTTP).

Determination of Molar Extinction Coefficients

The extinction coefficients of the and subunits were determined in 20 mM sodium phosphate (pH 7.4), 50 mM NaCl, and 5% glycerol, in the presence and absence of 6 M guanidine hydrochloride. Spectra of and were measured on a Hewlett-Packard 8450A diode array spectrophotometer between 200 and 320 nm. The (max) for both proteins was 280 nm. Extinction coefficients for and in 6 M guanidine hydrochloride were calculated from the number of tryptophan and tyrosine residues in each protein as described by Edelhoch(1967). The ratio of the absorbance of the native proteins to the absorbance of the proteins in 6 M guanidine hydrochloride was used to correct the calculated extinction coefficients to obtain the coefficient for the native protein. The extinction coefficient for (monomer) was determined to be = 42,577 liters mol cm, and that for (monomer), = 25,216 liters mol cm.

Other Procedures

Chromatographic supports, proteins, holoenzyme subunits, and nucleic acids were obtained and/or prepared as described in Olson et al.(1995) and Johanson and McHenry(1984). SDS-polyacrylamide gel electrophoresis and protein determinations were performed as de-scribed in Olson et al.(1995). ATPS was purified to remove residual ATP as described in Johanson and McHenry(1984).


RESULTS

Construction of Expression Plasmids That Overproduce and and Only or

To provide large quantities of and , we constructed a vector that contained the dnaX gene, modified to achieve maximal expression. This was accomplished by replacement of the region in front of the gene with oligonucleotide 1 (Fig. 1A) containing an optimal Shine-Dalgarno site and by replacement of rare codons in the first codons of the gene with common ones for the same amino acids (Reznikoff and Gold, 1986). This enabled expression of the two products of the dnaX gene to approximately 20% of the total E. coli protein (data not shown). Although this overexpression permitted purification of large quantities of and , this strategy suffered from two shortcomings: 1) the quantity of obtained was considerably lower than that of , presumably because of increased levels of frameshifting at high expression levels, and 2) the possibility existed that some of the obtained was contaminated by proteolyzed . (^2)To avoid these problems, we introduced specific mutations in dnaX at the site of frameshifting to construct plasmids that expressed only or .

The plasmid producing only , pRT610A, was created using oligonucleotide 2 (Fig. 1B), which eliminated the frameshift region by altering two codons in the sequence, AAA AAG, to AAG AAA, without altering the corresponding Lys-Lys in the protein. The G A mutation stabilized an A-U interaction between the mutant mRNA and the anticodon, perhaps discouraging slippage, but more importantly, the A G mutation eliminated frameshifting by placing a G residue at a nonwobble position in the -1 frame where G-U base pairs are not possible (Flower and McHenry, 1990). This construct allowed the expression of only. The plasmid producing only , pRT610B, was created using oligonucleotide 3 for mutagenesis. This included the same mutations as in the -overproducing plasmid, eliminating frameshifting, but also included an in-frame codon for glutamate, followed by a stop codon, allowing expression of a protein equivalent to authentic frameshifted .

Purification of and

The and subunits of DNA polymerase III holoenzyme were overexpressed and purified for use in functional analyses of and in holoenzyme and to provide sufficient material for structural studies (see Dallmann and McHenry (1995)). Both proteins were purified from E. coli strains containing expression plasmids that produced either or , not both subunits simultaneously (Fig. 1). and were purified to homogeneity ( Fig. 2and Fig. 3) and retained high specific activities throughout their purifications ( Table 1and Table 2). Both proteins are stable at -80 °C for over 1 year and for 1 week on ice.


Figure 2: Purification of . Samples of fractions I to IV were denatured in SDS sample buffer and subjected to electrophoresis on a 12% SDS-polyacrylamide gel. The gel was stained in Coomassie Brilliant Blue G-250. Lane 1, fraction I (cell lysate); lane 2, fraction II (ammonium sulfate); lane 3, fraction III (SP-Sepharose peak); lane 4, fraction IV (S-400 HR peak). All lanes contain 30,000 units of (equivalent to 5 µg of fraction IV).




Figure 3: Purification of . Samples of Fractions I to IV were boiled in SDS sample buffer and subjected to electrophoresis on a 12% polyacrylamide gel. The gel was stained with Coomassie Brilliant Blue G-250. Lane 1, fraction I (cell lysate); lane 2, fraction II (ammonium sulfate); lane 3, fraction III (Q-Sepharose peak); lane 4, fraction IV (Sephacryl S-300 HR peak). All lanes contain 80,000 units of (equivalent to 5 µg of fraction IV).







Preliminary experiments in which and were purified from a strain containing the expression plasmid pBBMD11, which contained the wild-type dnaX sequence and co-expressed both and , indicated that mixed / oligomers do not form to a significant extent in the absence of other overproduced subunits in vivo. Fig. 4A shows the protein and DNA synthesis activity profile of fraction II prepared from a strain containing pBBMD11 chromatographed on an S-Sepharose ion exchange column. Gel electrophoresis of the two major peaks in the profile revealed only in the first peak, while the second peak contained only (Fig. 4B). A mixed / oligomer would be expected to elute at a position between pure and ; however, no such peak was seen. This observation implies that and do not form mixed oligomers in vivo when co-expressed, consistent with the results of Tsuchihashi and Kornberg(1989), who also purified and from a co-expressing plasmid and observed no mixed oligomers of and .


Figure 4: Co-expression of and subunits does not yield mixed complexes. and were overexpressed from plasmid pBBMD11. Fractions I and II were prepared as described under ``Experimental Procedures.'' A, S-Sepharose chromatography. Fraction II was chromatographed over a 100-ml S-Sepharose (Pharmacia) column equilibrated in buffer TBP. and were eluted with a 10-column volume gradient of 0-300 mM NaCl in buffer TBP. Protein concentration (box), conductivity (bullet), and activity () are shown. B, gel electrophoresis of S-Sepharose column fractions. Samples of fractions I and II and selected column fractions were boiled in SDS sample buffer and subjected to electrophoresis on a 12% polyacrylamide gel. The gel was stained with Coomassie Brilliant Blue G-250. Lane 1, molecular weight markers; lane 2, fraction I (cell lysate), 10 µg; lane 3, fraction II (ammonium sulfate), 5 µg; lanes 3-20, column fractions 50, 53, 56, 59, 62, 65, 68, 71, 74, 77, 80, 83, 86, 89, 92, 95, and 97 (volume equivalent to 0.5 µl of each column fraction).



Initiation Complex Formation with and Complexes, Effect of ATPS

Functional asymmetry in DNA polymerase III holoenzyme was originally demonstrated by the observation that the ATP analogue, ATPS, promoted only half the amount of initiation complex formation than ATP (Johanson and McHenry, 1984). Although the basis of this asymmetry was unknown, it was suggested that an asymmetric placement of the and subunits in holoenzyme was responsible and reflected properties of the leading and lagging strand polymerases (Hawker and McHenry, 1987). Both subunits have been isolated in subcomplexes of holoenzyme. The subunit has been isolated in the complex (`) (McHenry et al., 1986; Maki and Kornberg, 1988), while has been isolated bound to core (alpha) in DNA polymerase III` (McHenry, 1982). Both and share the same 446 N-terminal residues and can interact with , `, and to form ``DnaX complexes'' (Onrust and O'Donnell, 1993; Xiao et al., 1993). Olson et al.(1995) have shown that the assembly of DnaX complexes is cooperative and that increases the affinity of -` for and . The availability of overproduced, purified subunits allowed reconstitution of DnaX complexes (Dallmann and McHenry, 1995) and reexamination of the effect of ATPS on holoenzyme reconstituted with either or to directly test whether one of these subunits uses ATPS uniquely.

Initiation complexes were formed using ATP or ATPS with complex, core, and beta (Fig. 5A), with complex, high concentrations of core, and beta (Fig. 5B) and with native holoenzyme purified from E. coli (Fig. 5C). Using native holoenzyme, ATPS resulted in approximately one-half the amount of initiation complexes formed compared to ATP, in agreement with the results of Johanson and McHenry (1984). As expected, ATP also supported initiation complex formation with both and complexes, whereas ATPS did not support initiation complex formation with complex and only supported half the amount of initiation complex formation with complex, compared to ATP. The result with complex-reconstituted holoenzyme was identical to that observed with native holoenzyme (Fig. 5C) (Johanson and McHenry, 1984). Thus, the subunit/complex must play a major role in loading beta subunit clamps onto primed DNA in native holoenzyme. A striking result from this experiment was that complex-reconstituted holoenzyme was indistinguishable from native holoenzyme, whereas complex-reconstituted holoenzyme was very different. Clearly the asymmetric behavior of holoenzyme, as revealed by the differential effects of ATP and ATPS, was not due to asymmetric placement of and within holoenzyme, but instead reflects properties of the subunit alone.


Figure 5: Differential effects of ATPS on and complexes. A, effect of ATPS on initiation complex formation by complex. complex (100 fmol), core (240 fmol), and beta (400 fmol) were assayed for the ability to form initiation complexes on primed, SSB-coated M13G (540 pmol as nucleotide) as a function of ATP () or ATPS (bullet) concentration as described under ``Experimental Procedures.'' B, effect of ATPS on initiation complex formation by complex. complex (100 fmol), core (4 pmol), and beta (400 fmol) were assayed for the ability to form initiation complexes on primed, SSB-coated M13G (540 pmol as nucleotide) as a function of ATP () or ATPS (bullet) concentration as described under ``Experimental Procedures.'' Note that a high level of polymerase III core was used to fully enable the catalytic potential of . Lower activity was observed with the low polymerase III core levels used in A. C, effect of ATPS on initiation complex formation by native DNA polymerase III holoenzyme. 70 units of DNA polymerase III holoenzyme were assayed for the ability to form initiation complexes on primed, SSB-coated M13G (540 pmol as nucleotide) as a function of ATP () or ATPS (bullet) concentration as described under ``Experimental Procedures.''



To exclude the possibility that the above result reflected partial proteolysis of the subunit to a -like protein (Blinkova et al. 1993), an experiment was carried out in which initiation complexes prepared using complex with ATP and ATPS were purified by gel filtration over Bio-Gel A5m (Bio-Rad) columns (Johanson and McHenry, 1984), and fractions were analyzed by SDS-polyacrylamide gel electrophoresis and Western blotting using an anti-/ polyclonal antibody. No cleavage of to a -like protein was observed. Furthermore, activity assays of the column fractions showed that only half the amount of initiation complex was obtained with ATPS compared to ATP (data not shown), consistent with the above results.

Titration of Core in and Complex Assays

Early in the development of assays for the and complexes, it was noted that, under our standard assay conditions containing 100 mM potassium glutamate, complex required large amounts of core before any significant DNA synthesis activity was observed. In experiments in which core was titrated into and complex assays (Fig. 6). complex showed maximal activity at the lowest amount of core used (240 fmol); however, complex required 4 pmol of core to achieve an activity comparable to that of complex. Evidently the interaction of the subunit with core (McHenry, 1982; Studwell-Vaughan and O'Donnell, 1991) produced a more active polymerase than enzyme reconstituted with complex and core, which are not known to interact directly.


Figure 6: Large excess of core is required for complex activity. The complex (100 fmol) (bullet) and complex (100 fmol) () were mixed with the indicated amounts of core, beta (400 fmol), SSB-coated M13G (540 pmol as nucleotide), and ATP (10 µM). A control experiment () containing only core and beta was also performed. All remaining assay components were added and incubated at 30 °C for 5 min as described under ``Experimental Procedures.''



Titration of into and Complexes

Because both and are present in native holoenzyme as currently isolated, we asked whether titration of the or subunits into the or complexes affected activity. Titration of the subunit into the or complex assays produced no effect (data not shown), whereas titration of the subunit into the or complex assays stimulated the reaction (Fig. 7). Titration of (0-1600 fmol) into complex (100 fmol/assay, 4 nM) caused a 2-fold inhibition of DNA synthesis, presumably due to competition between and complex in binding core. Titration of the subunit (0-1600 fmol) into complex (100 fmol/assay, 4 nM) consistently gave a 2-fold stimulation of DNA synthesis regardless of the amount of core used in the assays. As described above, complex required a large excess of core (40-fold) to achieve activity levels comparable to those with complex performed with stoichiometric amounts of core ( Fig. 6and, in Fig. 7, compare DNA synthesis levels at 0 fmol of in complex assays). The titrations of into complex assays in Fig. 7were performed at decreasing levels of core ( complex:core ratios, 1:10, 1:4, and 1:2) to determine whether could recruit the complex into a reconstituted holoenzyme. If acts as an organizing center for the assembly of a holoenzyme that includes the subunit, one might expect that the addition of to complex assays performed at stoichiometric levels of core would yield DNA synthesis activities at least comparable to that of complex alone. Alternatively, the effect of added in complex assays may only be related to the dimerization of core or due to conformational changes induced by the -alpha contact, and not due to a recruitment of complex into holoenzyme. At decreasing levels of core, only caused a 2-fold stimulation of complex activity and did not appear to promote more efficient recruitment of complex into holoenzyme (Fig. 7).


Figure 7: Titration of the subunit into and complex assays. The complex (100 fmol) was assayed with 200 fmol of core () and the indicated amounts of . The complex (100 fmol) was assayed with 200 fmol (), 400 fmol (circle), and 1 pmol () of core and the indicated amounts of . Each assay contained 400 fmol of beta, 540 pmol of SSB-coated M13G (as nucleotide), and 10 µM ATP. All remaining assay components were mixed and incubated at 30 °C for 5 min as described under ``Experimental Procedures.''



The 2-fold stimulation of complex by might also be explained by a reequilibration of the complex subunits during assays and the formation of significant amounts of complex in situ. However, when the subunit was preincubated with the complex at 15 °C for various times before assaying, no time-dependent increase in activity occurred (Fig. 8). A control assay in which no was added gave 39 pmol of DNA synthesis. All other assays showed a 2-fold stimulation independent of preincubation time. This result suggests that any reequilibration of complex into complex in these assays is not responsible for the observed stimulation.


Figure 8: Stimulation of complex by is independent of preincubation. The complex (100 fmol) was preincubated with (200 fmol) for the indicated times at 15 °C. Each assay contained core (280 fmol), beta (400 fmol), SSB-coated M13G (540 pmol as nucleotide), and ATP (10 µM). All remaining assay components were added and incubated at 30 °C for 5 min as described under ``Experimental Procedures.'' A control assay in which no subunit was added yielded 39 pmol of DNA synthesis, approximately 2-fold lower than the data shown, consistent with the results shown in Fig. 7.




DISCUSSION

The and subunits of DNA polymerase III holoenzyme were purified from overexpressing plasmids which produce only the or subunits (Fig. 1Fig. 2Fig. 3; Table 1and Table 2) in order to investigate the functions and physical properties of each subunit alone, without interference from the other. Experiments with an overproducing plasmid that co-expressed both and from the wild-type dnaX gene showed that the and subunits, overexpressed by themselves, do not form detectable quantities of mixed oligomers in vivo (Fig. 4), consistent with the observation of Tsuchihashi and Kornberg (1989). It has been suggested (Stukenberg et al., 1994; O'Donnell, 1994) that a mixed assembly can be formed in vitro by incubation of with in the absence of other holoenzme subunits. However, our observation that and do not form detectable mixed assemblies in vivo shows that this assembly is not favored, at least not in the absence of other overproduced proteins.

The purified and subunits were used to reconstitute the complex (`) and complex (`) (Dallmann and McHenry, 1995). These complexes were then mixed with polymerase III core and the beta subunit in order to test the hypothesis that an asymmetric distribution of and in holoenzyme (Hawker and McHenry, 1987) was responsible for the asymmetric activity of holoenzyme when assayed with ATPS (Fig. 5C) (Johanson and McHenry, 1984). In these experiments, ATPS supported only half the amount of initiation complex formation as ATP. A possible interpretation is that only one of the DnaX components, either or , can hydrolyze ATPS to form initiation complexes, reflecting their asymmetric placement in holoenzyme. The prediction for the reconstitution experiment was that one of the - or -reconstituted holoenzymes would be inactive with ATPS and the other would display the same activity with ATPS as with ATP. Consistent with this prediction, ATPS did not support initiation complex formation with complex-reconstituted holoenzyme; however, with complex-reconstituted holoenzyme, ATPS supported only half the amount of initiation complex formation, compared to ATP, i.e. the complex reconstituted holoenzyme behaved exactly as native holoenzyme. This clearly indicated that the asymmetric behavior of holoenzyme, as revealed by the effect of ATPS, was not due to an asymmetric placement of and within holoenzyme, but reflected properties of the subunit alone. This result also demonstrated that the subunit/complex plays a role in loading beta subunit clamps onto primed DNA in native holoenzyme, a point that was not previously clear.

The above results raised a number of questions about the role of in holoenzyme. The subunit also functions to dimerize core (McHenry, 1982; Studwell-Vaughan and O'Donnell, 1991) and might serve as an organizing center for holoenzyme, recruiting the complex into holoenzyme (Stukenberg et al., 1994; O'Donnell, 1994). The data in Fig. 6appeared to support this suggestion, since, in the absence of , complex required large amounts of core to yield activities comparable to that of complex with stoichiometric amounts of core. To address this possibility, we titrated into complex assays at various ratios of complex to core to determine whether can efficiently recruit complex into a holoenzyme that could function at stoichiometric levels of core (Fig. 7). Although caused a 2-fold stimulation of complex activity, this level of stimulation was seen at all concentrations of core used. This observation indicated that did not promote more efficient recruitment of complex into holoenzyme over the 5-min time course of the assay, a time sufficient for the synthesis of 300 Okazaki fragments. If it had, the stimulation by would have been expected to be more dramatic at lower enzyme concentrations, due to the greater effective concentration achieved because of proximity effects as complex was recruited into a holoenzyme complex. The possibility that dissociation of , `, and from the complex, and assembly of significant amounts of complex accounted for the results in Fig. 7was ruled out in an experiment in which the subunit and the complex were preincubated (Fig. 8). The lack of any increase over the 2-fold stimulation seen previously indicated the absence of any stripping of , `, and from complex by the subunit within the time frame of the experiment. Additionally, the half-life for the dissociation of DnaX complexes was 1.4 h at 20 °C (Olson et al., 1995), making the possibility of reequilibration of subunits unlikely in a 5-min assay.

Holoenzyme reconstituted with the complex as the sole beta clamp loader is functionally impaired when compared to native holoenzyme or to holoenzyme reconstitued with complex. In 100 mM potassium glutamate, a salt concentration that approximates the lower end of physiological ionic strength, the complex requires core at concentrations of 160 nM in order to achieve DNA synthesis levels approaching those of complex reconstituted with 10 nM core. If an E. coli cell contains 10-20 copies of holoenzyme (McHenry and Kornberg, 1977), its intracellular concentration would be 14-28 nM based on a cell volume of 1 fl (Ingraham et al., 1983). Clearly complex only appears to function at nonphysiological concentrations of core, whereas complex exhibits maximal activity at physiological concentrations. This study has provided a functional explanation for the observations of Blinkova et al.(1993) who found that mutant E. coli strains which do not express are viable, but that a temperature-sensitive mutation which interferes with function is lethal. Their additional observation of rapid proteolysis of to a -like protein, plus the functional aspects of activity presented here, raise the possibility that the subunit seen in the physical holoenzyme complex could result from cleavage of . Of course, the complex may have another ancillary role in DNA replication or repair. For example, mismatch repair requires a product of the dnaX gene (Lahue, et al. 1989) and would not be expected to require assembly of a dimeric replication fork. It is also possible that complex could serve a role for unloading beta sliding clamps left behind during holoenzyme cycling (Stukenberg et al., 1994).

After this manuscript was under review, a paper by O'Donnell and colleagues (Onrust et al., 1995) appeared reporting a - mixed complex requiring an excess of ; significant complex formation did not occur at a 1:1 ratio between and , even at high, nonphysiological protein concentrations. We interpret this as further evidence that - and - interactions are favored over mixed DnaX protein assemblies in the absence of other holoenzyme subunits. In the absence of other interactions that pull equilibria favoring mixed DnaX assemblies, these would not be expected to form at significant levels in vivo. However, our finding does not exclude more complex mechanisms involving other holoenzyme or cellular proteins not present at adequate levels in the DnaX-overproducing cells to redirect the assembly pathway.

In another report in the same series (Xiao et al., 1995), holoenzyme assembled with ATPase mutants of exhibited diminished replication activity, while enzyme assembled with ATPase mutants of appeared fully active. Our results with ATPS show that the full activity of holoenzyme can be reconstituted with only and that by itself is ineffective. The most direct interpretation of these results would be that serves as an ATPase to reproduce ATPS-dependent initiation complex formation with holoenzyme. Alternatively, could alter the conformation of within holoenzyme, enabling it to use ATPS as an effective substrate. We infer that can also participate in the ATPase-coupled initiation complex formation of native holoenzyme.


FOOTNOTES

*
This work was supported, in part, by National Institutes of Health Research Grant GM35695 and facilities support from the Lucille P. Markey Charitable Trust. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
Supported by a Postdoctoral Fellowship from the Natural Sciences and Engineering Research Council of Canada.

To whom correspondence should be addressed.

(^1)
The abbreviations used are: holoenzyme, E. coli DNA polymerase III holoenzyme; core, E. coli DNA polymerase III core (alpha--); -complex, --`--; complex, --`--; DnaX complex, a complex containing either product of the dnaX gene ( or ) with associated , `, and ; ATPS, adenosine 5`-O-(3`-thiotriphosphate); SSB, E. coli single-stranded DNA binding protein; PCNA, proliferating cell nuclear antigen.

(^2)
is particularly sensitive to E. coli proteases under specific conditions, yielding a protein close to in molecular weight (Blinkova et al. 1993).


REFERENCES

  1. Araki, H., Ropp, P. A., Johnson, A. L., Johnston, L. H., Morrison, A., and Sugino, A. (1992) EMBO J. 11, 733-740 [Abstract]
  2. Blinkova, A., Hervas, C., Stukenbarg, P. T., Onrust, R., O'Donnell, M. E. and Walker, J. R. (1993) J. Bacteriol. 175, 6018-6027 [Abstract]
  3. Blinkova, A. L., and Walker, J. R. (1990) Nucleic Acids Res. 18, 1725-1729 [Abstract]
  4. Bunz, F., Kobayashi, R., and Stillman, B. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 11014-11018 [Abstract]
  5. Burgers, P. M J. (1991) J. Biol. Chem. 266, 22698-22706 [Abstract/Free Full Text]
  6. Burgers, P. M J., and Yoder, B. L. (1993) J. Biol. Chem. 268, 19923-19926 [Abstract/Free Full Text]
  7. Carter, J. R., Franden, M. A., Aebersold, R., and McHenry, C. S. (1993) J. Bacteriol. 175, 3812-3822 [Abstract]
  8. Cull, M. A., and McHenry, C. S. (1995) Methods Enzymol. 262, 22-35 [Medline] [Order article via Infotrieve]
  9. Dallmann, H. G., and McHenry, C. S. (1995) J. Biol. Chem. 270, 29563-29569 [Abstract/Free Full Text]
  10. Edelhoch, H. (1967) Biochemistry 7, 1948-1954
  11. Flower, A. M., and McHenry, C. S. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 3713-3717 [Abstract]
  12. Fürste, J. P., Pansegrau, W., Frank, R., Blocker, H., Scholz, P., Bagdasarian, M., and Lanka, E. (1986) Gene 48, 119-131 [CrossRef][Medline] [Order article via Infotrieve]
  13. Gogol, E. P., Young, M. C., Kubasek, W. L., Jarvis, T. C., and von Hippel, P. H. (1992) J. Mol. Biol. 224, 395-412 [Medline] [Order article via Infotrieve]
  14. Hacker, K. J., and Alberts, B. (1994) J. Biol. Chem. 269, 24209-24220 [Abstract/Free Full Text]
  15. Hawker, J. R., Jr., and McHenry, C. S. (1987) J. Biol. Chem. 262, 12722-12727 [Abstract/Free Full Text]
  16. Hübscher, U., and Thömmes, P. (1992) Trends Biochem. Sci. 17, 55-58 [CrossRef][Medline] [Order article via Infotrieve]
  17. Ingraham, J, Maaløe, O., and Neidhardt, F. (1983) Growth of the Bacterial Cell , p. 269, Sinauer Assoc., Sunderland MA
  18. Johanson, K. O., and McHenry, C. S. (1984) J. Biol. Chem. 259, 4589-4595 [Abstract/Free Full Text]
  19. Kaboord, B. F., and Benkovic, S. J. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 10881-10885 [Abstract]
  20. Kong, X. P., Onrust, R., O'Donnell, M., and Kuriyan, J. (1992) Cell 69, 425-437 [Medline] [Order article via Infotrieve]
  21. Krishna, T. S-R., Kong, X. P., Gary, S., Burgers, P. M., and Kuriyan, J. (1994) Cell 79, 1233-1243 [Medline] [Order article via Infotrieve]
  22. Kunkel, T. A. (1985) Proc. Natl. Acad. Sci. U. S. A. 82, 488 [Abstract]
  23. Kunkel, T. A., Roberts, J. D., and Zakour, R. A. (1987) Methods Enzymol. 154, 367 [Medline] [Order article via Infotrieve]
  24. LaDuca, R. J., Crute, J. J., McHenry, C. S., and Bambara, R. A. (1986) J. Biol. Chem. 261, 7550-7557 [Abstract/Free Full Text]
  25. Lahue, R. S., Au, K. G., and Modrich, P. (1989) Science, 245, 160-164 [Medline] [Order article via Infotrieve]
  26. Lee, S. H., and Walker, J. R. (1987) Proc. Natl. Acad. Sci. U. S. A. 84, 2713-2717 [Abstract]
  27. Lee, S. H., Kwong, A. D., Pan, Z. Q., and Hurwitz, J. (1991) J. Biol. Chem. 266, 594-602 [Abstract/Free Full Text]
  28. Maki, H., Maki, S., and Kornberg, A. (1988) J. Biol. Chem. 263, 6570-6578 [Abstract/Free Full Text]
  29. Maki, S., and Kornberg, A. (1988) J. Biol. Chem. 263, 6555-6560 [Abstract/Free Full Text]
  30. McHenry, C. S. (1982) J. Biol. Chem. 257, 2657-2663 [Abstract/Free Full Text]
  31. McHenry, C. S. (1988) Biochim. Biophys. Acta 951, 240-248 [Medline] [Order article via Infotrieve]
  32. McHenry, C. S., and Johanson, K. O. (1984) in Proteins Involved in DNA Replication (Hubscher, U., and Spadari, S., eds) pp. 315-319, Plenum Press, New York
  33. McHenry, C. S., and Kornberg, A. (1977) J. Biol. Chem. 252, 6478-6484 [Abstract]
  34. McHenry, C., Oberfelder, R., Johanson, K., Tomasiewicz, H., and Franden, M. (1986) in Mechanisms of DNA Replication and Recombination (Kelly, T., and McMacken, R., eds) pp 47-61, Alan R. Liss, New York
  35. McHenry, C., Griep, M. Tomasiewicz, H., and Bradley, M. (1989) in Molecular Mechanisms in DNA Replication and Recombination (Richardson, C., and Lehman, I. R., eds) pp. 115-126, Alan R. Liss, New York
  36. Mead, D. A., Skorupa, E. S., and Kemper, B. (1985) Nucleic Acids Res. 13, 1103-1118 [Abstract]
  37. Nethanel, T., and Kaufmann, G. (1990) J. Virol. 64, 5912-5918 [Medline] [Order article via Infotrieve]
  38. O'Donnell, M. E. (1994) Annals N. Y. Acad. Sci. 726, 144-155 [Medline] [Order article via Infotrieve]
  39. O'Donnell, M., and Studwell, P. S. (1990) J. Biol. Chem. 265, 1179-1187 [Abstract/Free Full Text]
  40. O'Donnell, M., Onrust, R., Dean, F. B., Chen, M., and Hurwitz, J. (1993) Nucleic Acids Res. 21, 1-3 [Medline] [Order article via Infotrieve]
  41. Oberfelder, R., and McHenry, C. S. (1987) J. Biol. Chem. 262, 4190-4194 [Abstract/Free Full Text]
  42. Olson, M., Dallmann, H. G., and McHenry, C. S. (1995) J. Biol. Chem. 270, 29570-29577 [Abstract/Free Full Text]
  43. Onrust, R., and O'Donnell, M (1993) J. Biol. Chem. 268, 11766-11772 [Abstract/Free Full Text]
  44. Onrust, R., Finkelstein, J., Turner, J., Naktinis, V., and O'Donnell, M. (1995) J. Biol. Chem. 270, 13366-13377 [Abstract/Free Full Text]
  45. Reddy, M. K., Weitzel, S. E., and von Hippel, P. H. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 3211-3215 [Abstract]
  46. Reznikoff, W., and Gold, L. (1986) Maximizing Gene Expression , Butterworths, Stoneham, MA
  47. Stukenberg, P. T., Turner, J., and O'Donnell, M. E. (1994) Cell 78, 877-887 [Medline] [Order article via Infotrieve]
  48. Studwell-Vaughan, P. S., and O'Donnell, M. (1991) J. Biol. Chem. 266, 19833-19841 [Abstract/Free Full Text]
  49. Tsuchihashi, Z., and Kornberg, A. (1989) J. Biol. Chem. 264, 17790-17795 [Abstract/Free Full Text]
  50. Turchi, J. J., and Bambara, R. A. (1993) J. Biol. Chem. 268, 15136-15141 [Abstract/Free Full Text]
  51. Venkatesan, M., and Nossal, N. G. (1982) J. Biol. Chem. 257, 12435-12443 [Free Full Text]
  52. Wu, C. A., Zechner, E. L., Hughes, A., Franden, M. A., McHenry, C. S., and Marians, K. J. (1992) J. Biol. Chem. 267, 4064-4073 [Abstract/Free Full Text]
  53. Xiao, H., Dong, Z., and O'Donnell, M. (1993) J. Biol. Chem. 268, 11779-11784 [Abstract/Free Full Text]
  54. Xiao, H., Naktinis, V., and O'Donnell, M. (1995) J. Biol. Chem. 270, 13378-13383 [Abstract/Free Full Text]

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.