Search tips
Search criteria 


Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
J Biol Chem. Author manuscript; available in PMC 2011 February 3.
Published in final edited form as:
PMCID: PMC3033201



DNA replication, as with all macromolecular synthesis steps, is controlled in part at the level of initiation. Although the Origin Recognition Complex (ORC) binds to origins of DNA replication, it does not solely determine their location. To initiate DNA replication ORC requires Cdc6 to target initiation to specific DNA sequences in chromosomes and with Cdt1, loads the ring-shaped MCM2-7 DNA helicase component onto DNA. ORC and Cdc6 combine to form a ring-shaped complex that contains six AAA+ subunits. ORC and Cdc6 ATPase mutants are defective in MCM loading and ORC ATPase mutants have reduced activity in ORC-Cdc6-DNA complex formation. Here we analyzed the role of the Cdc6 ATPase on ORC-Cdc6 complex stability in the presence or absence of specific DNA sequences. Cdc6 ATPase is activated by ORC, regulates ORC-Cdc6 complex stability and is suppressed by origin DNA. Mutations in the conserved origin A element, and to a lesser extent mutations in the B1 and B2 elements, induce Cdc6 ATPase activity and prevent stable ORC-Cdc6 formation. By analyzing ORC-Cdc6 complex stability on various DNAs, we demonstrated that specific DNA sequences control the rate of Cdc6 ATPase, which in turn controls the rate of Cdc6 dissociation from the ORC-Cdc6-DNA complex. We propose a mechanism explaining how Cdc6 ATPase activity promotes origin DNA sequence specificity: on DNA that lacks origin activity, Cdc6 ATPase promotes dissociation of Cdc6, whereas origin DNA down regulates Cdc6 ATPase resulting in a stable ORC-Cdc6-DNA complex, which can then promote MCM loading. This model has relevance for origin specificity in higher eukaryotes.

Brenner and Jacob described the replicon model in 1963 (1), which outlined in general terms how DNA replication might start at a specific location within chromosomes. This model formed the basis for discovery of initiator proteins that recognize origins of DNA replication. The bacterial initiator, DnaA, (2) recognizes the bacterial origin of DNA replication, oriC, by binding to multiple 9mer DnaA boxes within the origin (3). However DnaA also binds frequently to DnaA boxes outside of DNA replication origins and these sites do not promote initiation of DNA replication (4), therefore the origin is defined by a combination of specific protein-DNA interactions, including DnaA, HU and IHF proteins and an arrangement of DNA sequences that form multiple binding sites in a specific conformation (5). It is only this particular conformation that allows the initiator to promote origin unwinding, to recruit other DNA replication proteins such as the DnaB helicase and helicase loading protein DnaC to the DNA and consequently to initiate DNA synthesis (5).

The mechanism that specifies the location of origins of DNA replication in eukaryotic chromosomes is less clear, particularly in mammalian cells, but it involves a combination of sequence specific protein-DNA interactions, epigenetic factors and cell cycle timing (6,7). In S. cerevisiae, specific DNA sequences are known to contribute the location of origins of DNA replication in chromosomes (810). Knowledge of such sequences lead to the discovery of the Origin Recognition Complex (ORC) (11), a six subunit complex. Homologues of ORC exist in all eukaryotic species. Yeast origins contain several conserved genetic elements (12): the A element, which serves as a binding site for ORC (11); the B1 element, which is partially involved in ORC binding (13,14) and the B2 element as a potential binding site for loading the MCM proteins (15,16). ORC in higher eukaryotes does not bind specifically to DNA, but has a preference for AT-rich (17,18) or supercoiled DNA (19).

Our understanding of how DNA sequences or locations within the chromosome are selected as sites of initiation of DNA replication is still poor since it is unclear what defines a start site for initiation of DNA replication. S. cerevisiae ORC binds to origins in a DNA sequence specific fashion, however this feature is not conserved in higher eukaryotes. But since the protein components required for initiation of DNA replication are conserved, it is expected that studies of how this process occurs in yeast will shed light on the process in all eukaryotes. In budding yeast it has been shown that the Orc1 subunit in ORC has to be in the ATP bound form for ORC to bind DNA (11,20). Orc1-dependent ATPase, stimulated by a arginine residue in an adjacent Orc4 subunit, is also regulated in a DNA sequence specific manner (20), but this ATPase activity has no influence on the selection of DNA sequences as origins of DNA replication (21). Although ATP-bound ORC can bind to origins of DNA replication specifically, its affinity cannot explain why some DNA sequences are selected as origins and others are not e.g. ORC binding sites have been identified by ChIP analysis which do not act as replication sites (10).

Another protein involved in the sequential assembly of pre-replicative complexes (pre-RCs) at origins of DNA replication is the Cdc6 protein (cdc18 in S. pombe) (2227). Cdc6 is a member of the AAA+ family of proteins and has striking amino acid sequence similarity with Orc1, the largest subunit of ORC (28,29). Together with ORC and Cdt1, Cdc6 is required for loading the mini-chromosome maintenance (MCM) proteins onto origin DNA (3036). Cdc6 binds to and changes the structure of ORC and together they bind cooperatively to origin DNA with higher DNA sequence specificity than ORC binding alone (37,38). Specific nucleotides within the origin DNA are required for ORC-Cdc6-DNA complex formation and Orc1 ATPase mutants are less efficient in forming the ORC-Cdc6-DNA complex (38). We were interested in understanding how Cdc6 contributes to the selection of certain DNA sequences in chromosomes of yeast to function as origins of DNA replication, a process with potential relevance to origin selection in higher eukaryotes. Since Cdc6 is also an AAA+ protein, like Orc1-5 (38,39), and many initiation proteins use their ATPase activity in a regulatory fashion (40), we analyzed the contribution of Cdc6 ATPase activity to ORC-Cdc6 and ORC-Cdc6-DNA complex assembly and its impact on the selection of DNA sequences as origins of DNA replication.

Experimental Procedures


ORC expressed in insect cells from recombinant baculoviruses and Cdc6 expressed in E. coli were purified to apparent homogeneity as described (11,20,38).

ATPase assays

ORC, Cdc6 ATP hydrolysis was carried out as described (20), with minor modifications. 2.5 pmol of ORC and 2.5 pmol of Cdc6 were incubated for 30 min on ice in 12 µl of ATPase buffer (25 mM HEPES, pH 7.6, 100 mM potassium glutamate, 5 mM magnesium acetate, 1 mM DTT, 1mM EDTA, 1mM EGTA, 0.1% (v/v) Triton X-100, 10% Glycerol) containing 2.5 pmol of DNA (when indicated) and 100 µM ATP (unless otherwise indicated). After the incubation 5 µCi of α-32P ATP (3000 Ci/mmol) was added and the reaction was started by moving the tubes from ice into a RT water bath. At 15, 30, 45 and 60 min 2µl aliquots were removed and stopped with 0.5 µl 2% SDS stop solution. 1 µl of the samples were consequently spotted on TLC plates and developed.

DNA probes

DNA probes and competitor DNA (290 bp GC-rich DNA) used in the gel shift and/or footprinting experiments were prepared as described (12,38). For the ATPase experiments a 91 bp fragment (residue 790–880) was amplified from plasmid pARS1WT or from the various mutant pARS1 plasmids (in the case of the A, B1, B2 fragment, an oligo (B1) primer and a A, B2- template DNA, was used). For ARS607 and ARS 607A, a 91 bp fragment was amplified from pARS607 with primer ARS 607 FWD 91bp (5’-CTTACGCTGGGTATTTTTTTTTTGG-3’) and primer ARS 607 REV 91bp (5’-GAGCTTTGTCTTGTTTATATTTAGTTACG-3’). For ARS607A a modified primer was used to create a linker scanning mutation as in pARS1 A using primer ARS 607 A- REV 91bp (5’-GAGCTTTGTCTTGTCCTCGAGGAGTTACGTTGGG -3’).

Glycerol gradients, footprints and gel shifts

Experiments were performed as described (38).

Determination of dissociation constants

Dissociation constants were determined using a gel shift assay. Gel shift reactions were performed as described previously (38) in the absence of competitor DNA. After challenging the complex with 100× fold cold competitor DNA (ARS1 291 bp) aliquots were loaded onto a continuously running gel in the cold room at the indicated time points. koff, R values and t1/2 rates were determined using KaleidaGraph 4.0 (Synergy software). All experiments were repeated at least twice with similar results.


Cdc6 ATPase is induced by ORC-Cdc6 complex formation

ORC and Cdc6 have been shown to form a stable complex in the presence of a slowly hydrolysable ATP analogue - ATPγS, but not in the presence of ATP (38). This finding suggests that ATP-hydrolysis regulates complex stability. To address this question in more detail we analyzed ATPase activities and complex formation of ORC and Cdc6 (Fig. 1). Cdc6 on its own had no ATPase activity as reported (31,41) (Fig. 1A). The ORC ATPase activity was similar to what has been reported (20). In contrast to a recent study (31), which used different salt conditions, we found very robust ATPase activity when Cdc6 was added to ORC, a 2.5 fold induction over the ATPase activity of ORC alone. This result suggests that ORC and Cdc6 form at least a transient complex in the absence of DNA, which induces either ORC or Cdc6 ATPase activity. To identify the active ATPases in the ORC-Cdc6 complex we analyzed two Cdc6 ATPase mutants (Cdc6 E224G and Cdc6 N263A) and an ORC ATP hydrolysis mutant (ORC4R/5A) in ATPase assays. Cdc6 E224G has a mutation in the highly conserved Walker B motif (31,35) and is dominant lethal when over-expressed in yeast. Cdc6 N263A (42,43) has a mutation in the sensor 1 motif, is temperature sensitive and lethal when over-expressed. Cdc6 belongs to the class of AAA+ proteins and mutations in Walker B or sensor 1 motifs frequently interfere with the ability of AAA+ proteins to hydrolyze ATP (40). The two Cdc6 mutants individually displayed no ATPase activity, similar to wild type Cdc6 (Fig. 1A). The addition of Cdc6 N263A to ORC did not change the ATPase activity, in contrast to wild type Cdc6. Cdc6 E224G addition to ORC lead to a reduction of overall ATPase activity. These results suggest that Cdc6 is a major ATPase within the ORC-Cdc6 complex. To analyze the contribution of ORC toward ATPase activity of the ORC-Cdc6 complex, we analyzed an ORC4R/5A double mutant (Fig. 1B). Mutations in arginine finger of ORC4 (ORC4R) have been shown to abolish Orc1-dependent ATPase (21). ORC5A carries a mutation in the Walker A motif of Orc5 which blocks ATP binding to ORC5, however this mutation has no significant phenotype in vitro and in vivo (20,44,45). We measured ATPase of the combined ORC4R/5A mutant in the absence and presence of Cdc6 (Fig. 1B). ORC4R/5A showed a very weak ATPase, which was dependent on high but physiological relevant ATP concentrations (up to 1mM). Addition of Cdc6 to the ORC4R/5A mutant lead to a significant increase in ATPase activity and this was dependent on high ATP concentrations as well. High ATP concentrations, however, had no effect on the ORC-Cdc6 induced ATPase (data not shown). These results show that ORC4R/5A can stimulate Cdc6 ATPase. Taken together, the ORC and Cdc6 ATPase data suggest that both ORC and Cdc6 are active ATPases in the ORC-Cdc6 complex, since ATPase mutants in both ORC and Cdc6 did not abolish ATPase activity completely.

Figure 1
Cdc6 and ORC interaction is regulated by ATPase activity. (A) ATP hydrolysis by ORC, Cdc6, Cdc6 E224G, Cdc6 N263A, ORC-Cdc6, ORC-Cdc6 E224G and ORC-Cdc6 N263A. ATP hydrolysis rates were determined in the absence of DNA. (B). ATPase activity in the presence ...

Cdc6 ATPase regulates ORC-Cdc6 stability

Using glycerol gradient sedimentation, we have shown that in the presence of ATP, ORC and Cdc6 did not form a stable complex, whereas in the presence of the non-hydrolysable ATP analogue, ATPγS, a stable ORC-Cdc6 complex was observed (38). This suggests that ATPase activity controls the stability of the ORC-Cdc6 complex in the absence of DNA. The ORC mutants ORC-4R, an Orc1-dependent ATP hydrolysis mutant (21), and ORC-1A, an Orc1 ATP binding mutant (20), did not form a stable ORC-Cdc6 complex in the presence of ATP (38). These results indicate that ORC ATPase in the ORC-Cdc6 complex does not regulate ORC-Cdc6 complex stability, suggesting that it is the Cdc6 ATPase that is responsible. This possibility was examined by analyzing Cdc6 ATPase mutants for their ability to form a stable complex with ORC (Fig. 1C and D). Cdc6 E224G and Cdc6 N263A were fractionated either individually or together with ORC in the presence of ADP, ATP or ATPγS by glycerol gradient sedimentation (Fig. 1C). Cdc6 E224G and Cdc6 N263A individually fractionated as monomers. The Cdc6 ATPase mutants co-fractionated with ORC in the presence of ATP and ATPγS, as seen by disappearance of monomeric Cdc6 and concomitant appearance of Cdc6 in the peak fraction containing ORC, which was verified by western blot with anti-Cdc6 antibodies (Fig 1D). In the absence of ORC, Cdc6 E224G and N263A were not found in the fraction corresponding to the ORC-Cdc6 (Fig 1D). Previously we have shown that in the presence of ADP ORC and Cdc6 do not form a complex (38). This is also true for Cdc6 E224G; however Cdc6 N263A formed a weak complex with ORC in the presence of ADP probably due to inefficient exchange of ATP for ADP.

The results so far show that Cdc6E224G and Cdc6N263A were capable of forming a stable complex with ORC in the presence of ATP, which was not true for wild type Cdc6 protein. The results suggest that, in the absence of DNA, the Cdc6 ATPase negatively regulates ORC-Cdc6 complex stability. Since Orc1 and Orc5 ATP binding or ATPase activity have no influence on ORC-Cdc6 complex stability (38), it is the Cdc6 ATPase within the ORC-Cdc6 complex that regulates complex stability.

DNA sequence dependent ATPase induction in the ORC-Cdc6-DNA complex

Since the ORC-Cdc6 complex is assembled on DNA, we were interested in determining how DNA modulates the ATPase activity in the complex. In performing these experiments, wild type and mutant versions of origin DNAs were used, as well as non-specific natural DNAs that do not function as an origin in yeast. Thus the relationship between origin binding and ATPase activity could be assessed. The ATPase of ORC, ORC4R/5A, Cdc6 and the two Cdc6 ATP hydrolysis mutants (Cdc6 E224G and Cdc6 N263A) were analyzed in the absence and presence of GC-rich non origin DNA, ARS1 origin DNA (containing the A, B1 and B2 elements), and mutant ARS1 A or AB1B2 DNA (Fig. 2). Note that both the A and B2 elements are ORC binding sites; B2 is only used as an ORC binding site in the absence of a functional A element (11,38).

Figure 2
ATPase of the ORC-Cdc6 complex is regulated by DNA sequence and is dependent on Cdc6 ATPase. ATP hydrolysis rates were determined in the absence of DNA (-DNA), or in the presence of GC-rich DNA (GC), ARS1, ARS1 with a mutant A element (ARS1 A ...

Cdc6 had no ATPase in the absence or presence of DNA (Fig. 2A, lanes 1, 4, 7, 10, 13). In the absence of DNA, ORC is an ATPase (lane 2) and addition of Cdc6 to ORC led to a 2.5 fold increase in ATPase compared to ORC alone (Fig. 2A, lane 3). GC-rich DNA and ARS1 AB1B2, which do not support initiation of DNA replication, ORC-DNA and ORC-Cdc6-DNA complex formation (38), did not suppress ORC ATPase activity (Fig. 2A, compare lane 2 with lanes 5, 14). Rather they induced ORC-Cdc6 ATPase activity 5-fold stronger than the ATPase in the presence of ORC alone (Fig. 2A, compare lane 2 with lanes 6, 15). This result suggests that transient binding of ORC and Cdc6 to GC-rich DNA activates the ATPase in either ORC or Cdc6. ARS1 origin DNA suppressed ORC ATPase by 8–10 fold (Fig. 2A, lanes 2, 8) as reported (20). ARS1 DNA also promoted stable complex formation of the ORC-Cdc6-ARS1 complex (38). The addition of ARS1 DNA to reactions containing ORC and Cdc6 suppressed the ATPase activity of ORC-Cdc6 about 5 fold, compared to the ATPase activity in the absence of DNA (Fig. 2A, compare lane 3, 9). An ARS1 A mutant, that supports weak binding of ORC, but does not support ORC-Cdc6 complex formation on DNA (38), suppressed ORC ATPase activity likely due to the weaker, non-perfect ORC binding site within the B2 element (Fig. 2A, compare lane 2 with lane 11). Interestingly, the ARS1 A mutant did not suppress ORC-Cdc6 ATPase but resulted in similar ATPase as the ORC-Cdc6 complex in the absence of DNA (Fig. 2A, compare lane 3 with lane 12). This result suggests that ORC can bind to the ARS1 A via the weak ORC binding site within the B2 element, but that addition of Cdc6 triggers ATPase activity that disrupts the ORC-Cdc6 binding to the ARS1 A DNA. These data indicate that ORC-Cdc6 complex formation on DNA is regulated by ATPase activity and that the ORC and the ORC-Cdc6 complexes differ in the way they respond to non-origin DNA sequences, as exemplified by the striking differences between ARS1 and ARS1 A DNA.

Cdc6 is the active ATPase in the ORC-Cdc6-DNA complex

Next we were interested in identifying the DNA regulated ATPase within the ORC-Cdc6-DNA complex. We therefore analyzed mutant versions of these proteins, including ORC4R/5A, Cdc6 E224G and N263A for their ability to regulate ORC-Cdc6 ATPase activity. ORC4R/5A, a very weak ATPase was reproducibly suppressed by ARS1 DNA, but not by GC-rich DNA or ARS1 AB1B2 (Fig. 2B, lanes 2, 5, 8, 11, 14). Addition of Cdc6 to ORC4A/5A resulted in stronger ATPase in the presence and absence of DNA, similar to the case of ORC and Cdc6, indicating that ORC ATPase is not the major ATPase within the ORC-Cdc6-DNA complex (Fig. 2B, lanes 3, 6, 9, 12, 15). Interestingly, based on the Cdc6-activated ATPase, this mutant complex was still able to distinguish between DNAs that acted as an origin and DNAs that did not.

The two Cdc6 mutants, Cdc6 E224G (31) and N263A did not increase the ATPase when compared with ORC alone, in the absence or presence of any DNA (Fig. 2C and D). Interestingly, the dominant negative Cdc6 E224G mutant resulted in reducing the ATPase of ORC in solution or in complex with non-origin GC-rich or ARS1 A and AB1B2 DNA (Figure 2D). Combined, these results suggest that Cdc6 is the major ATPase within the ORC-Cdc6 complex, independent of whether DNA is present or not. Furthermore, origin DNA, but not non-origin DNAs, suppressed the ORC-Cdc6 ATPase activity, suggesting that the ORC-Cdc6 complex, and the Cdc6 ATPase within it, could discriminate between functional and non-functional origin sequences. For example, even though the non-origin ARS1 A DNA has an ORC binding site within it and this sequence is capable of suppressing ORC ATPase activity (Fig. 2A, lanes 2, 10), this DNA was incapable of suppressing the ORC-Cdc6 ATPase activity.

B1 and B2 origin mutations induce ATPase within the ORC-Cdc6-DNA complex

The results presented in Figure 2, combined with those in a previous report (38), suggest that multiple DNA sequences within an origin regulate the ATPase activity of the ORC-Cdc6 complex. We were therefore interested in quantifying the contribution of the genetically conserved B1 and B2 origin elements to the ATPase activity (Fig. 3A). A linker scanning mutant in B1, a point mutant in B1 (A838G) and a B2 linker scanning mutant were analyzed. The linker mutation of the B1 element has been shown to reduce the affinity of ORC, however a point mutant in the B1 element, ARS1 A838G, and a linker scanning mutation of the B2 element have no influence on ORC’s affinity for DNA (13). The B1 linker scanning mutation abolished and the point mutant reduced ORC-Cdc6 complex formation on DNA (38). The mutation of the B2 element had only a very weak influence on ORC-Cdc6 complex formation on DNA (Speck and Stillman, unpublished data). In ATPase assays with ARS1 B1, B1 A838G and B2 mutants, the addition of Cdc6 to ORC resulted in a 3x, 2.3x and 1.8x fold activation of ATPase respectively, compared to the activity of ORC alone (Fig. 3A). ORC ATPase activity on its own was at the same time also slightly increased in these mutants compared with the activity on ARS1 DNA (Fig. 3A). In contrast the A mutant resulted in 8.5x fold activation of ORC-Cdc6 ATPase activity, indicating that mutations in the B elements have a weaker but significant impact on activation of ATPase. Thus Cdc6 ATPase activity in the ORC-Cdc6 complex was most influenced by ORC binding sites that contribute most to origin activity, but additional sequences that do not affect ORC binding also alter the regulation of Cdc6ATPase activity.

Figure 3
ATPase of the ORC-Cdc6 complex is regulated by origin DNA. (A) ATP hydrolysis rates were determined in the absence of DNA (-DNA), or in the presence of GC-rich DNA (GC), ARS1, ARS1 with a mutant A element (ARS1 A), ARS1 with mutant B1 element ...

Origins of replication suppress ORC-Cdc6 ATPase

We tested whether mutations in ARS A elements at other origins have a similar impact. The ARS607 and an ARS607 A mutant DNAs were analyzed. ARS607 suppressed ORC ATPase efficiently and the addition of Cdc6 resulted in increased ATPase activity, similar to ARS1 (Fig. 3B). Interestingly a mutation in the A element resulted in significant ORC ATPase, comparable to ORC ATPase in the absence of DNA, and the addition of Cdc6 induced very strong ATPase. This situation is similar to an ARS1 AB1B2 mutant, suggesting that ARS607 has no secondary ORC binding site within its B2 element. The ARS607 data suggest that Cdc6 induced ATPase is down-regulated at origins of replication due to ARS A elements.

Cdc6 ATPase mutants promote binding of the ORC-Cdc6 complex to nonspecific DNA

The above-described experiments established that Cdc6 ATPase in the ORC-Cdc6 complex was regulated in an ORC and DNA sequence specific manner. Furthermore, we have shown that Cdc6 influences the DNA binding specificity of the ORC-Cdc6 complex (37,38). We were therefore interested in determining if Cdc6 ATPase regulates the DNA sequence specificity of ORC-Cdc6 interactions with DNA.

Either wild type Cdc6 or the two Cdc6 ATPase mutants were tested for DNA binding in the absence or presence of ORC using a gel-shift assay (Fig. 4 A, B) and a DNaseI footprint assay (Fig. 4C). Cdc6 and its mutants had very weak and non-specific binding to DNA (Fig. 4A and B, lanes 2, 11 and 21). In the absence of competitor DNA the addition of Cdc6 E224G and N263A formed an ORC-Cdc6-ARS1 complex like wild type Cdc6 (Fig. 4A, compare lanes 4–9, 13–18 and 22–27). However in the presence of competitor DNA, binding of the two mutant Cdc6 proteins were significantly impaired (Fig. 4B, compare lanes 4–9, 13–18 and 22–27; note that in the presence of competitor DNA five fold more ORC was used to detect ORC binding). This result indicates that Cdc6 ATPase mutants reduced DNA sequence specific complex formation. The DNaseI footprint assay showed that Cdc6 ATPase mutants did not influence ORC binding at the A and B1 elements of the ARS1 origin, however Cdc6 induced protection adjacent to the B1 element was lost (Fig. 4C, dotted line). More significant, the two Cdc6 mutants, but not the wild type Cdc6, induced significant non-specific DNA binding in multiple regions outside the genetically defined origin (Fig 4C, arrows). Since the Cdc6 mutants in the absence of ORC had no DNaseI footprint, we suggest that the ORC-Cdc6 ATPase mutant stabilizes the ORC-mutant Cdc6 complex in regions outside the origin and therefore has reduced sequence specificity. Similar results have been seen when the ORC-Cdc6 complex was analyzed in the presence of ATPγS (38).

Figure 4
ORC-Cdc6 binds nonspecifically to DNA in the absence of Cdc6 ATPase,. (A) Gelshift assay of 32P-labeled ARS1 DNA with ORC, Cdc6, Cdc6 E224G and Cdc6 N263A in the absence of competitor DNA. Protein concentrations are indicated. (B) Gelshift assay of 32 ...

The mechanism of Cdc6 induced sequence specificity

The DNA binding experiments established that the Cdc6 ATPase activity contributes to sequence specific DNA binding to origin DNA, however the mechanism of how this activity contributes to origin selection remained unclear. Theoretically Cdc6 ATPase can regulate DNA sequence specific binding in several ways: 1) stabilizing ORC binding to the origin; 2) destabilizing ORC on non-origin DNA (37) or by regulating Cdc6 interaction with ORC that is not bound to origin DNA; 3) Binding preferentially to ORC that is origin bound; 4) Release of Cdc6 from non-origin DNA. Since many biological regulatory interactions are controlled at the level of dissociation, and the footprint experiments suggested a dissociation defect of the Cdc6 ATPase mutants when in a complex with ORC, we measured the dissociation rates of ORC, Cdc6 and Cdc6 N263A in the presence of 32P-labeled ARS1 or ARS1 A DNAs. ORC, Cdc6 and origin DNA were incubated in the absence of competitor DNA and then challenged by addition of 100x wild type ARS1 DNA. The reactions were loaded at time intervals onto a running native polyacrylamide gel and the dissociation rates were determined from the autoradiogram. The t1/2 of ORC on origin DNA was determined to be 64.9 min., indicating a very slow turnover (Fig. 5A, Table 1). The binding of ORC to ARS1 A displayed increased turnover, with a 17.5x decreased t1/2 of 3.7 min (Fig. 5B, Table 1). Wild type Cdc6 showed a biphasic dissociation rate from origin DNA, one fraction of Cdc6 coming off DNA immediately upon addition of competitor and another fraction had a t1/2 of 62 min., similar to ORC (Fig. 5C, Table 1). Interestingly, 15 seconds after competitor DNA addition, all the Cdc6 was dissociated from an ORC-ARS1 A DNA complex; the t1/2 was consequently below our detection limit (Fig. 5D, Table 1). The Cdc6 N263A mutant, incubated with ARS1 and ORC, showed in contrast to wild type Cdc6, a dissociation rate with a t1/2 of 29.7 min (Fig. 5E, Table 1). This lower t1/2 for ORC-Cdc6 N263A compared to the wild type ORC-cdc6 probably reflects the measurement of the combined dissociation of two separate complexes; 1) Cdc6 N263A that is non-specifically bound to the DNA and 2) the Cdc6 N263A that is within the ORC-Cdc6 complex. In contrast, for wild type Cdc6, the non-specifically bound Cdc6 immediately dissociated from the origin DNA upon addition of competitor DNA, leaving only the ORC-Cdc6 complex to dissociate.

Figure 5
Cdc6 ATPase is required for rapid dissociation of the ORC-Cdc6 complex from mutant origin DNA. Equilibrated binding reactions (time point 0 min) were challenged with a 100-fold molar excess of unlabeled ARS1 competitor DNA and then loaded onto a running ...
Table 1
Dissociation rates and t1/2 of ORC-DNA and ORC-Cdc6-DNA complexes. Based on the data exemplified in figure 5, dissociation rates (koff), R values and t1/2 were calculated using KaleidaGraph 4.0 (Synergy software). The rates were calculated for ORC or ...

In the case of ARS1 A the dissociation rate of Cdc6 N263A was measurable at t1/2 of 9.9 min (Fig. 5F, Table 1), which is in stark contrast to Cdc6 that had a dissociation rate 100x faster. Interestingly Cdc6 N263A led to a 2.7 fold stabilization of ORC on ARS1 A (compare Fig 5D and F), indicating that the ORC-Cdc6 N263A-ARS1 A complex is overall more stable than the ORC-ARS1 A complex, indicating that Cdc6 promotes moderately higher ORC-Cdc6-DNA complex stability in the absence of Cdc6 ATPase. This result also fits with the increased nonspecific binding of the ORC-Cdc6 N263A-DNA complex seen in the DNaseI footprint assay in regions outside the primary ORC binding site (Fig. 4C). Overall these results indicate that Cdc6 ATPase regulates the disassembly of the ORC-Cdc6 complex on non-origin DNA. The major mechanism is the release of Cdc6 from ORC bound to non-origin DNA due to Cdc6 ATPase.


In previous studies we have analyzed ORC-Cdc6 complex formation in the presence and absence of DNA and showed that Cdc6 confers increased sequence specificity to the ORC-Cdc6 complex (37,38). We also showed that ORC and Cdc6 form a ring-shaped complex with a surface of similar dimension and shape to the ring-shaped surface of the MCM-hexamer, which likely functions as a MCM loading machine prior to initiation of DNA replication (38). In this study we concentrated on analyzing the mechanism by which Cdc6 increases DNA sequence specificity or origin recognition and stability of the ORC-Cdc6-DNA complex, both of which are relevant to understanding why origins of DNA replication locate to specific regions within chromosomes for pre-RC formation and MCM loading (31).

Six out of the seven proteins within the ORC-Cdc6 complex are predicted to belong to the class of AAA+ proteins (38,39). ATP binding and hydrolysis frequently regulate the function of AAA+ proteins (40). In our previous study we focused on the role of ORC ATP binding and ORC ATPase in regulating ORC-Cdc6 complex (38). In this study, the contribution of Cdc6 ATPase on ORC-Cdc6 interaction and sequence specificity was investigated. Mutants interfering with Cdc6 ATPase have been characterized in vivo, Cdc6 E224G has a mutation in the highly conserved Walker B motif (31,35) and is dominant lethal when over-expressed in yeast. Cdc6 N263A (42,43) has a mutation in sensor 1, is temperature sensitive and lethal when over-expressed. In the absence of DNA we found that Cdc6 addition to ORC resulted in rapid Cdc6-induced ATP hydrolysis and dissociation from ORC. Cdc6-ADP, which is produced during ATP hydrolysis, is not capable of binding to ORC (38). Cdc6 mutants defective for ATPase did not show increased ATPase in the presence of ORC and resulted in stable complex formation. The Cdc6 E224G mutant actually inhibited ORC ATPase activity, perhaps explaining why this mutant is a dominant lethal. The biological function of ORC induced Cdc6 ATPase in the absence of DNA is likely the forced disassembly of the complex to ensure that the ORC-Cdc6 complex only forms on origin DNA (38).

In the presence of origin DNA a stable ORC-Cdc6 complex was formed (31,38) and Cdc6 ATPase activity was induced, but this low level ATPase could originate from partial binding of the ORC-CDC6 complex to regions outside the origin, which are unable to suppress Cdc6 ATPase. In the presence of non-origin DNA, however, the Cdc6 ATPase activity was much higher and the complex was not stable on DNA. Mutations in conserved genetic elements of ARS resulted in activation of ATPase activity, which was dependent on Cdc6 ATPase. The A element represents the primary binding site for ORC (11), the sequence of A elements are most conserved within different ARS and point mutants between the A-element have drastic phenotypes on plasmid stability (11,12). Mutations in the A element have also been shown to reduce ORC binding (11). We suggest that Cdc6 activated ATPase and consequently disassembly of the ORC-Cdc6-DNA complex is a reason for the low plasmid stability of ARS A box mutants, in addition to the reduced affinity of ORC for DNA. Similarly, mutations in the B elements are important for origin function, however in this case, only simultaneous deletion of at least two of the B elements result in nonfunctional origins (12). Mutations in the B1 element reduce slightly the affinity of ORC for origin DNA (13,38) and mutations in the B2 element can be compensated by Cdc6 over-expression (15). We found that mutations in the B elements resulted in increased Cdc6 ATPase activity, which promotes disassembly of the complex. Cdc6 over-expression may be sufficient to force ORC-Cdc6 complex formation on the defective origin and therefore could compensate for B2 mutations.

The ARS1 A838G mutation is of particular interest, since this mutant binds to ORC well, but is partially defective in the Cdc6-dependent extended footprint on the ARS1 origin DNA (38) and has reduced ability to suppress the Cdc6-induced ATPase activity of the ORC-Cdc6 complex. We therefore suggest that specific nucleotide sequences modulate origin utilization by targeting the ORC-Cdc6 complex rather than ORC alone, again suggesting that Cdc6 contributes to origin selection.

It is known that origins with weak ORC binding sites can still function efficiently (46). We suggest that these DNA sequences bind the ORC-Cdc6 complex more efficiently to promote pre-RC formation. Furthermore, when Cdc6 is over-expressed in certain circumstances, re-replication of only a subset of origins of DNA replication occurs (47,48). Since Cdc6 contributes to origin selection and utilization, this may be one reason why not all origins are re-replicated when Cdc6 is present in excess.

Cdc6 ATPase within the ORC-Cdc6-DNA complex, first described by Randell et al. (31), regulates the stability of the ORC-Cdc6 complex. The function of the ORC-Cdc6 complex is to load in cooperation with Cdt1 the potential MCM-helicase onto DNA to form a pre-replication complex (pre-RC) (49). Based on our new findings that Cdc6 ATPase controls the stability of ORC-Cdc6 complex on DNA, we suggest that it is the stability of the ORC-Cdc6 complex on origin DNA, but not on non-origin DNA that regulates directly the location in chromosomes and amount of MCM loading. Cdc6 ATPase contributes to the selection of DNA sequences that can promote MCM loading and hence formation of a pre-RC. The ORC-Cdc6 complex stability depends on several factors, many of which are hardwired into the DNA sequence at origins, namely the A and B elements, but we envision that chromatin structure might also affect complex stability.

It has been shown that Cdc6 availability is further regulated by association with cyclin (50) and Cdc6 destruction is regulated by ubiquitin mediated proteolysis (5153). Cyclin-Cdc6 interaction and proteolysis of Cdc6 regulate the overall availability of Cdc6 during the cell division cycle, but when Cdc6 is available to bind to ORC prior to pre-RC assembly, DNA sequence information is a significant determinant in modulating Cdc6 activity and hence where on the DNA pre-RCs are formed. Since subtle changes in the DNA sequence influence Cdc6 ATPase activity, Cdc6-ORC complex stability can also be represented as having a specific probability at any given DNA sequence and therefore contributes to selection of functional origins. This fits well with the recent data in fission yeast in which origins of DNA replication are selected in a stochastic manner dependent on DNA sequence and do not function every cell cycle (5456). We suggest that Cdc6 (cdc18 in S. pombe) ATPase activity influences those DNA sequences that are selected as origins of DNA replication and even the stochastic firing of origins in each S phase. Such a scenario may exist for the selection of sites of DNA replication in the chromosomes of plant and animal cells where ORC binding is not known to be sequence specific. In fact Cdc6 is a rate limiting component for the initiation of DNA replication in vertebrate cells (36,57) and we suggest that it is the interaction between ORC and Cdc6 and the subsequent ATPase activities of these proteins that determines when and where in chromosomes initiation of DNA replication takes place.

Cdc6 ATPase activity within the ORC-Cdc6 complex is suppressed by origin DNA when compared to the activity in the absence of DNA and activation of Cdc6 ATPase, due to non-origin DNA, leads to disassembly of Cdc6 from DNA. It is possible that ORC bound to non-origin DNA has an altered conformation, which stimulates Cdc6 ATPase upon complex formation. This results in ADP-Cdc6 which cannot interact with ORC and is released from the ORC-DNA complex. Once ADP-Cdc6 is in solution it has to be recharged with ATP. This could be a passive reaction or actively modulated by a protein. On the other hand a specific protein present during pre-RC assembly might modulate Cdc6 ATPase to promote several rounds of MCM loading. Alternatively, a protein may stimulate yeast Cdc6 ATPase activity on origin DNA to promote dissociation of Cdc6 from the origin before its destruction at the G1/S phase transition and hence hinder re-replication in a single cell cycle.


We wish to thank Arne Stenlund for reading the manuscript, S.P. Bell for ORC mutants and Patty Wendel for technical help in performing some of the experiments. This research was supported by a research grant from the National Institutes of Health (GM45436).

The abbreviations used are

Origin Recognition Complex
Mini-chromosome Maintenance Protein
Autonomously Replicating Sequence
sodium dodecyl sulfate
deoxyribonuclease 1
adenosine triphosphatase.


1. Jacob F, Brenner S. C R Hebd Seances Acad Sci. 1963;256:298–300. [PubMed]
2. Fuller RS, Kornberg A. Proc Natl Acad Sci U S A. 1983;80(19):5817–5821. [PubMed]
3. Fuller RS, Funnell BE, Kornberg A. Cell. 1984;38(3):889–900. [PubMed]
4. Roth A, Messer W. Mol Microbiol. 1998;28(2):395–401. [PubMed]
5. Messer W. FEMS Microbiol Rev. 2002;26(4):355–374. [PubMed]
6. Aladjem MI, Fanning E. EMBO Rep. 2004;5(7):686–691. [PubMed]
7. Bell SP, Dutta A. Annu Rev Biochem. 2002;71:333–374. [PubMed]
8. Diffley JF. Genes Dev. 1996;10(22):2819–2830. [PubMed]
9. MacAlpine DM, Bell SP. Chromosome Res. 2005;13(3):309–326. [PubMed]
10. Xu W, Aparicio JG, Aparicio OM, Tavare S. BMC Genomics. 2006;7:276. [PMC free article] [PubMed]
11. Bell SP, Stillman B. Nature. 1992;357(6374):128–134. [PubMed]
12. Marahrens Y, Stillman B. Science. 1992;255(5046):817–823. [PubMed]
13. Rao H, Stillman B. Proc Natl Acad Sci U S A. 1995;92(6):2224–2228. [PubMed]
14. Rowley A, Cocker JH, Harwood J, Diffley JF. Embo J. 1995;14(11):2631–2641. [PubMed]
15. Wilmes GM, Bell SP. Proc Natl Acad Sci U S A. 2002;99(1):101–106. [PubMed]
16. Zou L, Stillman B. Mol Cell Biol. 2000;20(9):3086–3096. [PMC free article] [PubMed]
17. Giordano-Coltart J, Ying CY, Gautier J, Hurwitz J. Proc Natl Acad Sci U S A. 2005;102(1):69–74. [PubMed]
18. Vashee S, Cvetic C, Lu W, Simancek P, Kelly TJ, Walter JC. Genes Dev. 2003;17(15):1894–1908. [PubMed]
19. Remus D, Beall EL, Botchan MR. Embo J. 2004;23(4):897–907. [PubMed]
20. Klemm RD, Austin RJ, Bell SP. Cell. 1997;88(4):493–502. [PubMed]
21. Bowers JL, Randell JC, Chen S, Bell SP. Mol Cell. 2004;16(6):967–978. [PubMed]
22. Cocker JH, Piatti S, Santocanale C, Nasmyth K, Diffley JF. Nature. 1996;379(6561):180–182. [PubMed]
23. Kelly TJ, Martin GS, Forsburg SL, Stephen RJ, Russo A, Nurse P. Cell. 1993;74(2):371–382. [PubMed]
24. Liang C, Weinreich M, Stillman B. Cell. 1995;81(5):667–676. [PubMed]
25. Piatti S, Lengauer C, Nasmyth K. Embo J. 1995;14(15):3788–3799. [PubMed]
26. Nishitani H, Nurse P. Cell. 1995;83(3):397–405. [PubMed]
27. Muzi Falconi M, Brown GW, Kelly TJ. Proc Natl Acad Sci U S A. 1996;93(4):1566–1570. [PubMed]
28. Bell SP, Mitchell J, Leber J, Kobayashi R, Stillman B. Cell. 1995;83(4):563–568. [PubMed]
29. Muzi-Falconi M, Kelly TJ. Proc Natl Acad Sci U S A. 1995;92(26):12475–12479. [PubMed]
30. Donovan S, Harwood J, Drury LS, Diffley JF. Proc Natl Acad Sci U S A. 1997;94(11):5611–5616. [PubMed]
31. Randell JC, Bowers JL, Rodriguez HK, Bell SP. Mol Cell. 2006;21(1):29–39. [PubMed]
32. Seki T, Diffley JF. Proc Natl Acad Sci U S A. 2000;97(26):14115–14120. [PubMed]
33. Weinreich M, Liang C, Stillman B. Proc Natl Acad Sci U S A. 1999;96(2):441–446. [PubMed]
34. Oehlmann M, Score AJ, Blow JJ. J Cell Biol. 2004;165(2):181–190. [PMC free article] [PubMed]
35. Perkins G, Diffley JF. Mol Cell. 1998;2(1):23–32. [PubMed]
36. Stoeber K, Mills AD, Kubota Y, Krude T, Romanowski P, Marheineke K, Laskey RA, Williams GH. Embo J. 1998;17(24):7219–7229. [PubMed]
37. Mizushima T, Takahashi N, Stillman B. Genes Dev. 2000;14(13):1631–1641. [PubMed]
38. Speck C, Chen Z, Li H, Stillman B. Nat Struct Mol Biol. 2005;12(11):965–971. [PMC free article] [PubMed]
39. Clarey MG, Erzberger JP, Grob P, Leschziner AE, Berger JM, Nogales E, Botchan M. Nat Struct Mol Biol. 2006;13(8):684–690. [PubMed]
40. Erzberger JP, Berger JM. Annu Rev Biophys Biomol Struct. 2006;35:93–114. [PubMed]
41. Elsasser S, Lou F, Wang B, Campbell JL, Jong A. Mol Biol Cell. 1996;7(11):1723–1735. [PMC free article] [PubMed]
42. Schepers A, Diffley JF. J Mol Biol. 2001;308(4):597–608. [PubMed]
43. Takahashi N, Tsutsumi S, Tsuchiya T, Stillman B, Mizushima T. J Biol Chem. 2002;277(18):16033–16040. [PubMed]
44. Takahashi N, Yamaguchi Y, Yamairi F, Makise M, Takenaka H, Tsuchiya T, Mizushima T. J Biol Chem. 2004;279(9):8469–8477. [PubMed]
45. Loo S, Fox CA, Rine J, Kobayashi R, Stillman B, Bell S. Mol Biol Cell. 1995;6(6):741–756. [PMC free article] [PubMed]
46. Palacios DeBeer MA, Muller U, Fox CA. Genes Dev. 2003;17(15):1817–1822. [PubMed]
47. Green BM, Morreale RJ, Ozaydin B, Derisi JL, Li JJ. Mol Biol Cell. 2006;17(5):2401–2414. [PMC free article] [PubMed]
48. Tanny RE, MacAlpine DM, Blitzblau HG, Bell SP. Mol Biol Cell. 2006;17(5):2415–2423. [PMC free article] [PubMed]
49. Mendez J, Stillman B. Bioessays. 2003;25(12):1158–1167. [PubMed]
50. Mimura S, Seki T, Tanaka S, Diffley JF. Nature. 2004;431(7012):1118–1123. [PubMed]
51. Drury LS, Perkins G, Diffley JF. Embo J. 1997;16(19):5966–5976. [PubMed]
52. Drury LS, Perkins G, Diffley JF. Curr Biol. 2000;10(5):231–240. [PubMed]
53. Perkins G, Drury LS, Diffley JF. Embo J. 2001;20(17):4836–4845. [PubMed]
54. Dai J, Chuang RY, Kelly TJ. Proc Natl Acad Sci U S A. 2005;102(2):337–342. [PubMed]
55. Heichinger C, Penkett CJ, Bahler J, Nurse P. Embo J. 2006;25(21):5171–5179. [PubMed]
56. Patel PK, Arcangioli B, Baker SP, Bensimon A, Rhind N. Mol Biol Cell. 2006;17(1):308–316. [PMC free article] [PubMed]
57. Lemaitre JM, Bocquet S, Mechali M. Nature. 2002;419(6908):718–722. [PubMed]