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Journal of Virology, August 2006, p. 7491-7499, Vol. 80, No. 15
0022-538X/06/$08.00+0     doi:10.1128/JVI.00435-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

Surface Mutagenesis of the Bovine Papillomavirus E1 DNA Binding Domain Reveals Residues Required for Multiple Functions Related to DNA Replication

Stephen Schuck and Arne Stenlund*

Cold Spring Harbor Laboratory, Cold Spring Harbor, New York 11724

Received 3 March 2006/ Accepted 9 May 2006


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The E1 protein from papillomaviruses is a multifunctional protein with complex functions required for the initiation of viral DNA replication. We have performed a surface mutagenesis of the well-characterized E1 DNA binding domain (DBD). We demonstrate that substitutions of multiple residues on the surface of the E1 DBD are defective for DNA replication without affecting the DNA binding activity of the protein. The defects of individual substitutions include failure to form the double trimer that melts the ori and failure to form the double hexamer that unwinds the ori. These results demonstrate that the DBD plays an essential role in multiple DNA replication-related processes apart from DNA binding.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Viral initiator proteins, such as the E1 proteins from the papillomaviruses and the large T antigens of the polyomaviruses, are multifunctional proteins that, in a single polypeptide, combine multiple activities that are required for the initiation of DNA replication (for a review, see references 2 and 33). These proteins therefore provide an avenue for dissecting the initiation process in systems with a minimal number of components. Because tasks such as the recognition of a site for initiation, melting of the template, and unwinding of the double-stranded DNA are common to all organisms that use double-stranded DNA as their genetic material, the mechanisms used by the viral initiators are likely to be informative for this process in general.

The manner by which initiator proteins such as E1 provide these diverse activities is not well understood, but recent data indicate that different oligomeric forms of the initiator protein provide the different functions. E1 is an ~70-kDa polypeptide which is capable of sequence-specific DNA binding, has DNA helicase activity, and can melt viral ori DNA (12, 17, 28, 31, 36, 41). It is believed that recognition of the ori is carried out by E1 in combination with E2 and that a highly sequence-specific complex consisting of a dimer of E1 and a dimer of E2 forms on the ori (24, 27). Recently it was also demonstrated that E1 forms a double trimer, which can melt DNA, and serves as a precursor for the E1 double hexamer, which in turn is the active helicase and unwinds the DNA (26). A plausible model, for which some support exists, is that the initiator protein, by binding to the DNA, modifies the DNA structure (4, 11). The modified DNA structure in turn becomes a substrate for a different initiator complex, which again modifies the DNA structure to provide the substrate for a third kind of initiator complex, etc. In this manner, an ordered progressive change of the template structure can occur.

The involvement of the initiator DNA binding domain (DBD) in these processes has, with few exceptions (32, 39), been viewed as confined to the initial sequence-dependent DNA binding step. The DBD of the papillomavirus E1 proteins is a well-conserved region of ~150 amino acids which structurally is highly homologous to the DBD of the T antigens from the polyomaviruses (3, 10, 16, 19). The DBD of the E1 protein and its DNA binding properties have been studied in great detail at both the biochemical and structural levels and a good understanding of how DNA binding and dimerization come about for these proteins now exists (3-5, 10, 11, 15).

The E1 DBD is known to take part in a complex interplay with the viral transcription factor E2, where a direct interaction between the DBDs of the two proteins results in structural changes in the template, which in turn trigger a second interaction between the helicase domain of E1 and the trans-activation domain of E2 (1, 13, 22). A dimer interaction between the two surfaces in the DBD is required for sequence-specific DNA binding and appears to occur only when the DBD is bound to DNA (35).

Our excellent understanding of how E1 DBD binds DNA has allowed us to address whether the E1 DBD is required for processes other than DNA binding. By performing a comprehensive mutational analysis of the surface of the DBD, while specifically avoiding residues involved in DNA binding, we have identified multiple residues on the surface of the DBD that affect DNA replication without affecting DNA binding. These results demonstrate that the E1 DBD clearly takes an active part in processes such as template melting and unwinding, and it may also be involved in processes such as cell cycle control of DNA replication and interactions with cellular replication factors.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
DNA helicase assays. DNA helicase assays were performed essentially as described previously (28, 30). A 50-mer oligonucleotide with 28 bp of complementarity to M13 was end labeled and annealed to M13 DNA, generating a substrate with a 22-nucleotide 3' tail. Three quantities of wild-type (wt) and mutant E1 (8, 15, and 30 fmol) were incubated with the substrate at 37°C for 30 min, and the reaction was terminated by the addition of sodium dodecyl sulfate (SDS) to 0.1% and 50 µg/ml proteinase K. After digestion for 15 min, the samples were run on a 5% acrylamide gel and the fraction of displaced oligonucleotide was determined.

Transient replication assays. Transient replication assays were performed as described previously (38). Briefly, Chinese hamster ovary (CHO) cells of the rodent cell line were electroporated with expression vectors for E1 and E2, together with the two ori plasmids 11/X/X and X/12/X containing a distal and a proximal E2 binding site (BS), respectively. These two plasmids each contain one E2 BS and are identical except that, in 11/X/X, the proximal E2 BS 12 has been mutated, and in X/12/X, the distal E2 BS 11 has been mutated (25). In addition, in 11/X/X, an EcoO109I site in the pUC19 backbone has been destroyed. Two and 3 days after transfection, low-molecular-weight DNA was isolated and digested with DpnI, HindIII, and EcoO109I, which distinguishes between the two ori plasmids. The digests were separated on agarose gels, and after transfer to nitrocellulose, replicated plasmid DNA was detected by hybridization with a 32P-labeled ori probe.

In vitro DNA replication assays. Assays for DNA replication in vitro were essentially carried out as described previously (27, 40). Briefly, standard in vitro DNA replication assays were carried out in 25-µl reaction mixtures containing 30 mM HEPES-KOH, pH 7.5; 7 mM MgCl2; 1 mM dithiothreitol; 4 mM ATP; 0.2 mM each of GTP, UTP, and CTP; 0.1 mM each of dATP, dGTP, and dTTP; 10 µM [32P]-dCTP; 40 mM creatine phosphate; 1 µg creatine kinase and 50 ng of template DNA; 10 µl of S100; and 0.5 µl nuclear extract from H293 cells. Reaction mixtures were incubated at 37°C for 60 min, treated with proteinase K (50 µg/ml) in the presence of 0.5% SDS, extracted with phenol-chloroform, precipitated, and analyzed by agarose gel electrophoresis.

E1 protein expression and purification. E1 mutants and wt E1 were expressed in Escherichia coli as N-terminal glutathione S-transferase-fusions, purified by affinity chromatography, cleaved, and purified by ion exchange chromatography as described previously (29). E1 purified in this manner is monomeric, as determined by glycerol gradient sedimentation and gel filtration (28).

EMSA. Four percent acrylamide gels (39:1 acrylamide/bis) containing 0.5x Tris-borate-EDTA lacking EDTA were used for all electrophoretic mobility shift assay (EMSA) experiments. E1 was added to the probe (~2 fmol) in 10 µl binding buffer (20 mM HEPES, pH 7.5, 100 mM NaCl, 0.7 mg/ml bovine serum albumin, 0.1% NP-40, 5% glycerol, 5 mM dithiothreitol, 5 mM MgCl2, and 2 mM ATP or ADP). After incubation at room temperature for 1 h, the samples were loaded and run for 2 h at 9 V/cm. The ability to generate discrete complexes, especially the double trimer and the double hexamer, was critically dependent on high-purity acrylamide, freshly prepared ammonium persulfate solution, overnight polymerization of the gels, and a precise prerunning time (9 V/cm for 4 h).

Combined EMSA and unwinding assays. Unwinding assays were performed by incubating 2 fmol of probe with E1 under EMSA conditions (binding buffer in the presence of 2 mM ATP), but at 32°C and in the presence of 10 ng/µl E. coli single-stranded DNA (ssDNA)-binding protein (SSB) (USB). The ssDNA is detected as an SSB/ssDNA complex under EMSA conditions.

Probes. All probes were generated by PCR using primers end labeled with [{gamma}-32P]ATP and T4 polynucleotide kinase. Probes were purified by polyacrylamide gel electrophoresis (PAGE), eluted by diffusion, and precipitated.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Surface mutagenesis of the E1 DBD. We mutated E1 DBD surface residues based on the X-ray crystal structure of a dimer of E1 DBD bound to its DNA binding site (11). We changed all exposed surface residues to alanine, with the exception of the residues predicted to be involved in DNA binding and dimerization, based on the X-ray cocrystal structure (Fig. 1). These 63 alanine substitutions were generated in the context of the mammalian E1 expression vector pCG E1 (37) by site-directed mutagenesis.


Figure 1
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  		FIG. 1. Space-filling representation of the E1 DBD dimer as determined by X-ray crystallography. The colored residues in panel A represent the 63 residues on the surface of the E1 DBD that were changed to alanines. In panel B, the 12 residues that had a significant deleterious effect on DNA replication when changed to alanine are shown.

 
We wanted to use this set of surface mutants for two purposes. Our primary objective was to identify surface residues on the E1 DBD that affect the ability of E1 to support DNA replication. As a secondary objective, we were interested in identifying residues on the surface of the E1 DBD that affect the interaction with the DBD of E2. The interaction between these two proteins is complex. A physical interaction between the E1 helicase domain and the E2 trans-activation domain is required for DNA replication in vivo. In the context of the bovine papillomavirus minimal ori, where the E1 and E2 BSs are in close proximity to each other, the interaction between the helicase domain and the activation domain does not occur in the absence of a second physical interaction between the E1 DBD and the E2 DBD, which bends the DNA between the two binding sites sharply (13). If the E1 and E2 binding sites are separated by two turns of the helix (21 bp), the interaction between the two DBDs is no longer necessary. Therefore, E2 protein, with point mutations on the surface of the E2 DBD that affect the interaction with E1 DBD, is incapable of supporting DNA replication of an ori where the E1 and E2 BSs are proximal but shows wt levels of activity for DNA replication of an ori where the E1 and E2 BSs are distal (6, 13). By using these two oris, we expected to simultaneously identify E1 DBD mutants that were defective for replication in general (which would fail to replicate either plasmid) as well as E1 DBD mutants that were defective for replication due to a failure to interact with E2 DBD (which would fail to replicate only the plasmid with the proximal E2 BS).

After sequence verification, the E1 expression vectors were cotransfected, together with the E2 expression vector pCG E2 and the two ori constructs, into CHO cells (which have been used extensively to characterize the bovine papillomavirus replicon) and assayed for transient DNA replication 2 and 3 days after transfection. The mutant E1 proteins were placed in one of four categories depending on their overall ability to support viral replication: ++ (50 to 100% of wt activity), + (20 to 50% of wt activity), –/+ (<20% of wt activity), and – (no detectable replication) (Table 1).


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  		TABLE 1. Replication activity of E1 DBD surface mutantsa

 
The substitutions that showed no (–) or low (+/–) DNA replication activity (16 substitutions indicated in Table 1) were then tested for expression of the mutant E1 proteins by transfection of the expression vectors into COS cells. Two days after transfection, whole-cell lysates were prepared and analyzed by SDS-PAGE and Western analysis using a monoclonal antibody directed against E1 (Fig. 2A). In repeated experiments, two of the substitutions failed to generate wt levels of the full-length protein (E294A and E304A) (Fig. 2A, lanes 5 and 8), while two mutants gave rise to significantly smaller products (R297A and D177A) (lanes 6 and 7). The remaining 12 substitutions were expressed at levels similar to that of the wt protein.


Figure 2
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  		FIG. 2. (A) In vivo expression of mutant E1 proteins. The E1 expression vectors that were unable to support DNA replication in vivo were transfected into COS cells. Forty-eight hours after transfection, the cells were lysed by the addition of Laemmli-loading dye and the resulting lysates were analyzed by SDS-PAGE, transferred to nitrocellulose, and probed with a monoclonal E1 antibody. (B) In vivo replication assays. Replication assays were carried out by transfection of the E1 and E2 expression vectors pCGE1 and pCGE2, together with the ori plasmids X/12/X (proximal E2 BS) and 11/X/X (distal E2 BS) (25). In lanes 1 and 2, wt E1 and wt E2 were used. In lanes 3 and 4, wt E1 and the mutant E2 (388/390) (13) were used. In lanes 5 through 28, the respective E1 mutants were cotransfected with wt E2. Low-molecular-weight DNA was harvested 48 and 60 h after transfection, digested with DpnI, EcoO109I, and HindIII, run on agarose gels, transferred to nitrocellulose, and probed with an ori probe. After digestion with EcoO109I and HindIII, the ori plasmid with the proximal E2 BS gives rise to a 2.5-kb band and the plasmid with the distal E2 BS gives rise to a 2.9-kb band.

 
Results of transient DNA replication assays for these 12 mutants are shown in Fig. 2B. To be able to distinguish between the two ori plasmids, we had destroyed an Eco109I restriction site in the plasmid backbone in the plasmid containing the distal E2 BS. Consequently, upon digestion with Eco109I and HindIII, the plasmid with the distal E2 BS gives rise to a 2.9-kb fragment, while the plasmid with the proximal E2 BS gives rise to a smaller (2.5-kb) fragment. Wild-type E1, in combination with wt E2, supported replication of the two oris with the proximal and distal E2 BS equally well (Fig. 2B, lanes 1 and 2). E2 with the 388/390 double mutation in the E2 DBD showed a strong preference for replication of the plasmid with the distal E2 BS, as observed previously (13) (Fig. 2B, lanes 3 and 4). Of the 12 mutants that were selected as having severe defects for DNA replication, two mutants, 252 and 256, showed preferences for replication of the plasmid with the distal E2 BS (lanes 9 through 12). We also found the same pattern for the mutants K212A and T239A (data not shown). However, due to the position of these substitutions, they are unlikely to be directly involved in interaction with E2 DBD, as will be discussed below.

We next generated the 12 substitutions in the E. coli E1 expression vector pET E1 and expressed and purified the mutant proteins to apparent homogeneity using previously established procedures (Fig. 3A). The behavior during purification was similar for all mutants with the exception of K279A, which was expressed at significantly higher levels than was the wt protein.


Figure 3
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  		FIG. 3. (A) Purification of mutant E1 proteins from E. coli. The mutant E1 proteins were expressed in E. coli and purified using established procedures. wt E1 (lane 1) and the mutant proteins were analyzed by SDS-PAGE and stained with Coomassie brilliant blue. (B) DNA binding activity of mutant E1 proteins. The 12 mutant E1 proteins were tested for sequence-specific ori binding by EMSA in the presence of ADP using a 39-bp probe centered on the E1 BS. Two quantities, 30 and 60 fmol, of the wt and mutant proteins were used in lanes 1 through 19. For the mutants W295A, I296A, K168A, and G290A, which had reduced DNA binding activity, 120 and 240 fmols of E1 were used (lanes 20 through 27).

 
We first tested these mutants in a DNA binding assay to verify that their DNA binding was not compromised (Fig. 3B). We used a recently developed EMSA for E1 (26). Using a short (39-bp) probe containing the E1 BS, E1 BS-dependent DNA binding can be measured. In the presence of ADP, E1 forms two kinds of complexes on this probe, an E1 BS-specific dimer and nonsequence-specific trimer. The majority of the mutants had binding activities that differed less than twofold compared to that of the wt (Fig. 3B, lanes 1 through 21). However, some of the mutants showed qualitative or quantitative differences. The mutants 295 (Fig. 3B, lanes 20 and 21) and 290 (lanes 26 and 27) formed exclusively trimer and no dimer. Our previous data have demonstrated that the formation of trimer and dimer competes (26). Consequently, the formation of exclusively trimer indicates a severe defect for dimer formation. Several mutants also showed reduced overall binding activity. For the mutants 295, 296, 168, and 290 (Fig. 3B, lanes 20 through 27), four-times-higher levels of protein were used, demonstrating that DNA binding for these mutants is reduced approximately two- to fourfold. Some of the other mutants, such as N252A (Fig. 3B, lanes 14 and 15) and R268A (lanes 6 and 7), may also have subtle defects for DNA binding.

We next measured the DNA replication activity of the mutant E1 proteins in an in vitro DNA replication assay (Fig. 4A). Such an assay measures the ability of the E1 protein to recognize the ori, melt and unwind the DNA, and cooperate with cellular factors for DNA synthesis (34). The results from Fig. 4A were quantified and are shown in Fig. 4B. We chose an E1 concentration where the level of E1 was limiting for the level of DNA synthesis (Fig. 4A, lane 2), as indicated by the approximately twofold increase in incorporation by the addition of a twofold-higher level of E1 (lane 3). All the mutant proteins, with the exception of T188A and N252A (Fig. 4A, lanes 4 and 5), showed defects in cell-free replication to various extents. Mutants 168, 256, 268, 269, 279, 290, 295, and 296 failed to give rise to any significant DNA synthesis (Fig. 4A, lanes 8 through 15). Mutants S280A and S281A also showed clear defects but retained 5 to 10% of wt activity.


Figure 4
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  		FIG. 4. (A) In vitro DNA replication using mutant E1 proteins. The 12 mutant E1 proteins were tested for their abilities to support DNA replication in a cell-free replication system, followed by analysis of the replication products by agarose gel electrophoresis. In lane 1, no E1 was added. In lane 2, 200 ng of wt E1 was added, and in lane 3, 400 ng of wt E1 was added. In lanes 4 through 15, 200 ng of the respective mutant E1 was added. (B) The level of incorporation of [{alpha}-32P]dCTP for wt E1 and for each mutant was quantitated and is shown in a bar graph.

 
We next measured DNA helicase activity of the mutant proteins in an oligonucleotide displacement assay (Fig. 5) (14). We measured the level of oligonucleotide displacement at three E1 concentrations for each mutant, which was then plotted as a function of protein concentration. The mutants fell roughly into three groups. First, two mutants, L268A and R269A, showed significantly increased DNA helicase activities compared to that of the wt E1 protein. The second group contains seven mutants (K168A, T188A, S280A, N252A, V256A, S281A, and G290A) that had helicase activities similar (less than twofold reduced) to that of the wt E1. Three mutants, K279A, W295A, and I296A, had significant (two- to threefold) reductions in DNA helicase activity. As discussed below, the mutants W295A and I296A likely have structural defects, which may explain their reduced helicase activities.


Figure 5
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  		FIG. 5. DNA helicase activity of mutant E1 proteins. Three quantities of wt and mutant E1 proteins (8, 15, and 30 fmol) were tested for the ability to displace a labeled oligonucleotide annealed to an M13 template at 37°C. The displaced labeled oligonucleotide was quantitated, and the fraction of input was plotted as a function of E1 levels.

 
We recently defined conditions under which various oligomeric E1 complexes bound to the ori can be identified by EMSA (26). Using a very short (32-bp) probe centered on the E1 BS, a double hexamer can form in the presence of hydrolysable ATP. The formation of this complex requires intact E1 BS but does not involve melting of the template. We tested the wt E1 and the DBD surface mutations in this assay (Fig. 6). All mutants were capable of forming the E1 dimer. The most striking effect was observed with the mutant K279A, which failed to form a double hexamer (Fig. 6, lanes 15 to 16). In addition, the mutants G290A (Fig. 6, lanes 21 and 22), W295A (lanes 23 and 24), and to some extent, I296A (lanes 25 and 26) formed a trimer (E13), indicating that these mutants are in some way defective for dimer formation, which is consistent with the results shown in Fig. 3. In addition, the mutant S281A showed a peculiar complex migrating significantly faster than the E1 dimer (Fig. 6, lanes 19 and 20). This difference in complex mobility is not due to the presence of E1 fragments or E1 degradation since no detectable lower-molecular-weight forms of E1 are present and may result from a conformational change in the protein as a result of this particular mutation. We have observed that some mutations in other domains of E1 can generate a complex with similar mobility, indicating that this altered mobility might be related to the conformation of the E1 protein.


Figure 6
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  		FIG. 6. Double hexamer formation by E1 mutants. EMSA was performed using the 32-bp ori probe. Sixty and 120 fmol of wt and mutant E1, respectively, were used in the presence of ATP.

 
We recently demonstrated that on a full-length (84-bp) ori probe, E1 forms double trimers and double hexamers in the presence of ADP and ATP, respectively (26). This is a stringent test for the ability to assemble into functional complexes on ori probes, and we therefore tested the mutants (with the exception of the mutants G290A, W295A, and I296A, which were defective for DNA binding) in this assay (Fig. 7). All mutants, with one exception, K279A (Fig. 7, middle panel, lanes 20 to 23), formed the double trimer in the presence of ADP, although the level of double trimer varied. All mutants, with two exceptions, R269A and K279A (Fig. 7, middle panel, lanes 17 to 19 and 23 to 25), formed the double hexamer, although again the level of complex formation varied. As expected, the mutant defective for double-trimer formation (K279A) was also defective for double-hexamer formation, since the double trimer is a required precursor for the double hexamer (26). This identifies two E1 DBD surface residues that clearly are required for the formation of the larger functional E1 complexes.


Figure 7
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  		FIG. 7. Formation of double trimers and double hexamers by E1 mutants. EMSA was performed using the 84-bp ori probe. Thirty, 60, or 120 fmol of wt and mutant E1 was used in the presence of ATP or ADP as indicated in the figure. Lane 1 in all three panels contained probe alone.

 
An ori fragment-unwinding assay combines multiple activities such as DNA binding, melting, and DNA helicase activities. This assay is essentially an EMSA in the presence of E. coli SSB and performed at 32°C and in the presence of ATP. We compared the abilities of all the mutants to unwind ori DNA and to generate an SSB/ssDNA complex (Fig. 8). The results were quantitated and are shown in Fig. 8B. Six mutants showed greater than twofold reductions in unwinding activity. For four of these mutants, K168A (Fig. 8A, upper panel, lanes 8 to 10), G290A (lower panel, lanes 13 to 15), W295A (lower panel, lanes 16 to 18) and I296A (lower panel, lanes 19 to 21), this defect is most likely caused by the defect in DNA binding observed for these mutants. In addition, however, the mutants R269A (Fig. 8A, upper panel, lanes 23 to 25) and K279A (lower panel, lanes 4 to 6), which have little or no defect for DNA binding, showed a reduction of more than twofold in the ability to unwind the ori, indicating that these residues may affect unwinding directly. Both of these two mutants have defects for double-hexamer formation.


Figure 8
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  		FIG. 8. (A) Ori fragment unwinding by E1 mutants. Ori fragment unwinding was carried out using the 84-bp ori probe. In lane 1, top panel, probe alone was added. In lanes 2 and 3, top panel, E. coli SSB was added to double-stranded probe (lane 2) or denatured probe (lane 3). In lane 4, top panel, wt E1 was added in the absence of SSB. In lanes 5 through 25, top panel, and lanes 1 through 21, bottom panel, 40, 80, and 160 fmol of wt E1 and the respective mutants were used. (B) The unwinding reactions in panel A were quantitated, and the fraction of single-stranded DNA generated was plotted as a function of E1 concentration for each mutant.

 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Many (27/63) of the alanine substitutions on the surface of the E1 DBD showed some defect for DNA replication in vivo. In many cases, these defects were modest and therefore difficult to pursue. We consequently focused on the group of mutants (16/63) which showed a minimum of fivefold reduction for DNA replication in vivo. For 4 of the 16 mutants, we could not detect wt levels of full-length protein. These mutants, D177A, E294A, R297A, and E304A, are likely to affect the stability of E1 either because folding is affected directly or because signals or modifications that control stability are affected.

The remaining 12 mutants were tested for activities related to DNA replication in a series of in vitro assays to identify which specific function was affected by a particular mutation. Although the mutations were chosen such as to not affect DNA binding or dimerization by the E1 DBD, we found four mutants (K168A, G290A, W295A, and I296A) that showed significant defects for DNA binding. Inspection indicates that of these, at least three of the mutations involved are likely to have structural effects on the DBD. Residue 290 is a Gly-to-Ala mutation, and both W295 and L296 are hydrophobic residues that are partially buried. Thus, structural effects on the DBD likely cause the DNA binding defect of these mutants, maybe by altering the position of the different DNA binding elements relative to each other. It is unclear why K168A showed a defect for DNA binding.

The results from the cell-free DNA replication assays were particularly informative (Fig. 4A and B). All mutants, with four exceptions, showed a complete lack of activity for DNA synthesis in vitro. Two of the exceptions (T188A and N252A) had wt activities for DNA replication in vitro. Consequently, these mutations are unlikely to affect the basic enzymatic machinery of E1, in spite of showing a severe defect for DNA replication in vivo. As expected, these mutants had wt or close to wt activity in all assays (Table 2). Two other mutants, S280A and S281A, had residual (5 to 10%) activity for in vitro replication.


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  		TABLE 2. Summary of biochemical activities of E1 DBD mutantsa

 
There are several possible causes for the in vivo replication defect of these four mutants, which we currently cannot distinguish between. A simple reason could be that the mutations involved affect intracellular localization of E1. Another possibility is that these mutations affect some aspect of replication control, which is required for DNA replication in vivo but not in vitro. For example, it is likely that replication in vivo is controlled by modifications of E1, such as cell cycle-dependent phosphorylation (9, 20). These mutations may affect a target or signal for such modification, although no such cell cycle-dependent phosphorylation has been mapped to the E1 DBD. Indeed, T188, S280, and S281 are all potential phosphate acceptors. Although the phenotype with a defect for replication in vivo but not in vitro is unusual, there are examples of similar phenomena for SV40 T antigen where a deletion N terminal to the DBD has similar effects (21). It is also possible, however, that the defects of S280A and S281A reflect qualitative effects on DNA binding. As shown in an EMSA, S281A, in addition to the standard dimer complex, forms an anomalous fast-moving complex (Fig. 6). This complex is not caused by the presence of E1 fragments and may be reflective of a conformational difference in the mutant protein. In support of such a conclusion, we have observed that some mutations in the helicase domain result in a similar anomalous fast-moving complex (S. Schuck and A. Stenlund, unpublished data). The remaining eight mutants (K168A, V256A, L268A, R269A, K279A, G290A, W295A, and I296A) lack detectable activity for DNA replication in vitro. These latter mutants clearly have defects affecting the biochemical activity of E1.

None of the mutants lacked helicase activity; indeed the largest effect on helicase activity was a reduction of approximately threefold for the mutants W295A and I296A, which are likely to have structural defects and a twofold reduction for the mutant K279A, which, as shown in Fig. 6 and 7, failed to form a double hexamer. The increased helicase activity (approximately twofold) by the mutants L268A and R269A was completely unexpected. Although we have no clear explanation for this increased activity, it is important to recognize that in contrast to most other helicases, the viral initiator proteins, due to their many activities, are not dedicated helicases and that the accommodation of these multiple different functions may require compromises that result in suboptimal activity for a given function. This would allow room for improvement.

We also tested the mutant E1 proteins in the recently developed assays for double-trimer and double-hexamer formation by EMSA as well as for unwinding using an EMSA with E. coli SSB that detects single-stranded DNA generated by E1. The double-trimer and double-hexamer results were very clear; only one single mutant, K279A, was defective for the formation of the ADP-dependent double trimer. This mutant was also defective for the formation of a double hexamer, as expected, since the double trimer is a precursor for the double hexamer on the ori-sized probe. The failure to form the double hexamer is also consistent with the failure of this mutant to form a double hexamer on the short probe (Fig. 6). The mutant R269A also failed to form the double hexamer on the ori-sized probe. However, because this mutant still formed a double hexamer on the short probe and also formed a double trimer, R269A likely affects a step in the transition from the double trimer to double hexamer, while the K279A also affects a step prior to double-trimer formation. With the exception of the DNA binding-defective mutants (K168A, G290A, W295A, and I296A), these two mutants also had the most severe defect for unwinding. The function of the E1 DBD in double trimer, double hexamer, or unwinding is currently not understood; however, these results demonstrate that the role of the E1 DBD in fact is critical.

In summary, the scan of the surface of the E1 DBD has resulted in multiple mutants that affect DNA replication. Based on the behavior of these mutants, we believe that we can identify at least four categories of mutants that affect the ability of E1 to support DNA replication in vivo (Table 2). The first category includes the substitutions that affect the DNA binding activity of E1 (K168A, G290A, W295A, and I296A), likely for structural reasons. Although these four mutants may also affect other activities, such effects would be obscured by the defect for DNA binding. Indeed, since the effects on DNA binding are modest, it is not likely that the DNA binding defects account entirely for the severe replication defects of these mutants.

The second category, which includes the mutants T188A, N252A, S280A, and S281A, is more interesting. Two of these mutants (T188 and N252) have no defect for DNA replication in vitro. They also lack obvious biochemical defects and therefore clearly must affect some step that is important for only replication in vivo, such as some aspect of replication control. The most obvious candidate for such a step is cell cycle signals, which play a role for DNA replication in vivo but likely not for replication in vitro. The other two mutants in this group (S280A and S281A) behave similarly, except that these mutants show substantial reductions for in vitro replication.

A third category is defined by the mutants L268A and V256A which have no obvious biochemical defects but still are defective for replication in vitro and in vivo. It seems likely that these mutations affect a function that is required specifically for DNA replication, possibly an interaction with cellular DNA replication factors. It is well established that E1 interacts with a number of cellular factors such as replication protein A, topoisomerase I, and DNA polymerase {alpha} (7, 8, 18, 23). At least in one case, this interaction has been mapped to the E1 DBD (7).

The fourth category includes mutants R269A and K279A. These mutants have very specific defects in formation of the E1 double hexamer on the ori probe and, as a consequence, show defects for unwinding. These are the only mutants (in addition to the mutants with DNA binding defects) where the defects observed in biochemical assays can clearly account for the lack of DNA replication activity in vitro and in vivo.

It is apparent that there are many ways to disable the DNA replication activity of the E1 protein, which is a testimony to the multitude of diverse functions that this class of protein performs. Furthermore, all of these activities appear to depend to some extent on the E1 DBD, including the DNA helicase activity. This provides good evidence that the DBD takes part in many biochemical activities that have been considered to reside in other domains.

Our screen has provided us with a number of candidate residues for several different functions, and further study will obviously be required to verify these tentative assignments. The mutants L268A and N256A would be good starting points to explore physical interactions between the E1 DBD and members of the cellular replication machinery. The mutants T188A and V252A and maybe also S280A and S281A could be used to determine whether these residues are acceptor sites for cell cycle-dependent phosphorylation. Clearly, the mutations R269A and K279A define residues that are involved in double-trimer and double-hexamer formation and will be important for the understanding of how these complexes melt and unwind double-stranded DNA.

Finally, our screen to identify mutations in the E1 DBD that interact with the E2 DBD failed, although we identified several mutants (N252A, V256A, T239A, and K212A) that had the desired properties, i.e., these mutants supported the replication of an ori with a distal E2 BS but not with a proximal E2 BS (Fig. 2). Due to the position of these residues, they are unlikely to interact directly with the E2 DBD. Because we have structural information for both E1 and E2 DBDs bound to DNA, and the E1 and E2 BS are located only a few base pairs apart, there are very severe steric constraints on which residues in E1 DBD could interact with E2 DBD. In addition, we know which residues on the surface of E2 DBD are required for the interaction with E1 DBD. Based on this information, none of these four residues is likely to interact with E2 DBD. A possible explanation for these "false positives" in our screen is that these mutants affect the DNA binding activity of E1 and these defects in ori recognition can be more readily rescued by E2 bound to the distal E2 BS than by E2 bound to the proximal E2 BS.


   ACKNOWLEDGMENTS
 
This work was supported by NIH CA 13106 to A.S.


   FOOTNOTES
 
* Corresponding author. Mailing address: Cold Spring Harbor Laboratory, 1 Bungtown Road, P.O. Box 100, Cold Spring Harbor, NY 11724. Phone: (516) 367-8407. Fax: (516) 367-8454. E-mail: Stenlund{at}cshl.edu. Back


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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Journal of Virology, August 2006, p. 7491-7499, Vol. 80, No. 15
0022-538X/06/$08.00+0     doi:10.1128/JVI.00435-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.



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