Control of DNA Rereplication via Cdc2 Phosphorylation Sites in the Origin Recognition Complex

Yeast methods, strains, and plasmids.

S. pombe methods were essentially as described by Moreno et al. (53): YES is rich medium, and EMM2 is defined medium and was supplemented with leucine (L), uracil (U), adenine (A), and histidine (H) as indicated. YSO is rich medium low in adenine (36). The vitamin B₁ thiamine (B1) was added to 2.7 mg/liter as indicated. Genotypes are listed in Table 1.

Potential phosphorylation sites [(S/T)P] encoded by orp2 were mutated by site-directed mutagenesis using PCR. To generate orp2-T4A, plasmid pJL207 (orp2⁺ cloned into pREP3 [40]) was used as a template. Primers P1 and P2 amplified the 5′ coding region of orp2⁺; P2 has nucleotide changes that altered Thr to Ala. Separately, primers P3 and P4 were used to amplify the region of orp2⁺ 3′ of the sequence amplified by P1-P2. The P3 primer has nucleotide changes to alter two Thr codons to Ala codons and introduces an XhoI site. The P1-P2 and P3-P4 products were gel isolated, phosphorylated, and ligated together, and the ligation product was amplified using P1 and P4. This product was cloned to replace the corresponding region of the orp2⁺, and the clone was sequenced. Resulting amino acid changes are shown in Fig. 1. The mutated region was subcloned into appropriate orp2⁺ vectors as follows: pJL211 (pRep1-GST-orp2 [40]) was changed to make pJL329 (pRep1-GST-orp2-T4A). pJL320 (genomic orp2⁺, leu1⁺ integrating vector [see below]) was changed to make pJL322 (genomic orp2-T4A, leu1⁺ integrating vector). Primer sequences were P1 (CTGAAGATACGTTTTCAACCATTTTTAAAG), P2 (CGCCTAAAAAAGGAGTTCTTGAAGAT), P3 (CAACAACTTCTGGTGCTAATGTAGAAGG), and P4 (CTCCTCGAGCACCAGGACATCGAA).

To express alleles of orp2 from the orp2 promoter in the genome, an orp2⁺ leu1⁺ vector (pJL320) was constructed. pJL320 contains the orp2 promoter and coding sequence followed by the nmt1 terminator cloned between the PstI-SacI sites of integrating vector pJK148 (leu1⁺) (35). Steps in construction of pJL320 were as follows: The 1.7-kb PstI-NdeI fragment from pER17 (40) was cloned into PstI/NdeI-digested pJL207 (pREP3-orp2⁺) (40), thus replacing the nmt1 promoter with the orp2 promoter. From this, the PstI-SacI fragment containing the orp2 promoter and coding sequence and the nmt1 terminator was moved into PstI-SacI-digested pJK148 to create pJL320. pJL320 was cut with PflMI (in the orp2 5′ noncoding region) to target integration at orp2Δ::ura4⁺ to create orp2Δ::ura4⁺::orp2⁺ (leu1⁺).

Construction of JLP230 orp2⁺ and JLP238 orp2-T4A strains was as follows: plasmids pJL320 (orp2⁺ leu1⁺) and pJL322 (orp2-T4A leu1⁺) were integrated at the orp2Δ::ura4⁺ locus in diploid JLP46. Transformants were sporulated, and strains JLP230 and JLP238 were isolated and checked by Southern blot analysis. JLP306 and 308 are Ura⁻ derivatives of JLP230 and JLP238 made by one-step gene replacement of the ura4⁺ within the orp2Δ::ura4⁺ by transformation with HindIII-digested pJL378 (pJL378 is an unmarked deletion of orp2 coding sequences cloned into the HindIII site of pBluescript). Ura⁻ transformants were obtained using YES–0.1% 5-fluoroorotic acid to create orp2Δ::ura4⁺::orp2⁺ (leu1⁺).

Construction of nmt1GST-Cdc18* strains was as follows: pAL27-GST-cdc18-T4A has Thr codons at positions 26, 98, 104 and 134 mutated to Ala (44). pAL27-GST-cdc18-T4A was linearized with StuI and integrated at ura4-294 in strain FWP162 h⁺ leu1-32 ura4-294. GST-Cdc18-T4A is referred to as Cdc18*. The transformant JLP515 h⁺ leu1-32 ura4-294::GST-cdc18-T4A, ura4⁺was selected on plates lacking Ura, and stable integrants were identified as isolates that failed to grow on 0.1% 5-fluoroorotic acid–YES plates. JLP515 was mated to JLP306 (orp2Δ::orp2-T4A leu1⁺) to obtain strains JLP518, JLP519, and additional congenic isolates. JLP515 was mated to JLP308 (orp2Δ::orp2⁺ leu1⁺) to obtain JLP523, JLP524, and additional congenic isolates. All isolates were obtained by tetrad dissection.

Cdc18 induction and flow cytometry.

For induction of Cdc18* from the nmt1 promoter (49), cells were grown overnight in YES-thiamine. This preculture was used to inoculate EMM2-LAUH-B1 to an optical density at 600 nm (OD₆₀₀) of 0.1 and grown for at least 10 h at 32°C to an OD₆₀₀ of not more than 1.0. Cells were harvested and washed twice in water to remove thiamine. Cells were reinoculated into EMM2-LAUH (without thiamine) to a calculated OD₆₀₀ of 0.01. These cultures were grown at 32°C and harvested for flow cytometry, Western blotting, and microscopy at the times indicated. Cells were processed for flow cytometry as described (14) except that cells were stained with 1 μM Sytox Green (Molecular Probes) instead of propidium iodide. Fluorescence was analyzed using a Becton Dickinson FACScan and CellQuest software.

Competitive transformations.

JLP306 and JLP308 were grown separately overnight at 30°C in 5 ml of YES and then were mixed 1:1, spread on a YES plate, and grown overnight at 30°C. For each transformation, approximately 5 μl of cells from the plate was mixed with 150 μl of 0.1 M lithium acetate (pH 4.7) and incubated at 25°C for 1 h; 0.5 to 1 μg of plasmid DNA was added; 350 μl of 50% polyethylene glycol was added; and cells were incubated overnight at 25°C, harvested, washed gently with EMM2, and resuspended in 1 ml of EMM2. A 5- to 10-μl aliquot of this suspension was diluted and spread for single colonies on EMM2-LAUH (total viable), and the remainder was spread on four to six plates of EMM2-LAH (transformants). Colonies were picked from EMM2-LAH (transformants) or EMM2-LAUH (total) onto YSO. Red or pink colonies on YSO were counted after 2 days.

Antibodies and immunoblotting.

GST-Orp2 made in Escherichia coli was used to raise rabbit antibodies. The 1.3-kb XhoI fragment from pACT-166 (which contains orp2 cDNA cloned into the two-hybrid vector pAS1) (40) was subcloned into the SalI site of pGEX-KG (Novagen). The resulting glutathione S-transferase (GST)–Orp2 fusion protein was produced in E. coli (BL21), partially purified by binding to GSH-Sepharose (Pharmacia), and used to raise rabbit polyclonal antisera (26). Crude antisera readily detected Orp2 overexpressed in fission yeast. To detect wild-type levels of Orp2, antibody was affinity purified by binding to purified GST-Orp2 protein immobilized on a nylon membrane. Bound antibodies were eluted with 100 mM glycine (pH 2.0), neutralized with Tris (pH 8.5) and stored in 50% glycerol at −20°C, and used at a 1:100 dilution.

To distinguish which immunoreactive bands were due to Orp2 protein, extract from cells lacking the orp2⁺ gene were compared to extracts from wild-type cells and to extracts from cells overexpressing orp2⁺ (Fig. 2B and data not shown). Because orp2⁺ is essential, selective spore germination was used to obtain the population of cells lacking the gene (orp2Δ::ura4⁺), as described (40). Other experiments with epitope-tagged Orp2 have since confirmed the assignment of Orp2 specific bands. Strong cross-reacting bands were not detected when the primary antibody incubation included sarcosyl and sodium dodecyl sulfate (SDS) (Fig. 3A); however, antibody activity was rapidly lost in this solution. Standard Tris-buffered saline–0.3% Tween 20–5% powdered milk (26) was used in all other experiments. For analysis of GST-Cdc18*, whole-cell extracts were made by vortexing with glass beads as described (40), total protein was quantitated using Bradford's reagent (Bio-Rad), and 10 μg of protein was loaded per lane on an SDS–10% polyacrylamide gel. GST-Cdc18* was detected using rabbit anti-GST (gift of L. Hengst) at a 1:500 dilution.

Kinase assays and analysis.

GST, GST-Orp2, and GST-Orp2-T4A were expressed in S. pombe from plasmids pJL205, pJL211 (40), and pJL329 (this study). GST proteins were isolated from yeast lysates by binding GSH-Sepharose (40), except that lysis and binding were in NETN.5 (50 mM Tris [pH 8], 500 mM NaCl, 5 mM EDTA, 10% glycerol, 1% NP-40) with protease and phosphatase inhibitors as previously described. Bound complexes were washed four times with NETN.5, and GST proteins were eluted with 40 mM glutathione (pH 8.0) and dialyzed against a solution containing 50 mM Tris (pH 8), 150 mM NaCl, 50% glycerol, and 1% NP-40.

Cdc2 kinase was obtained from S. pombe nuc2 mutant cells arrested at 36.5°C for 4 h. Anti-Cdc13 (gift of P. Russell) bound to protein A-Sepharose or p13^suc1 beads (R. Aligue and P. Russell) were used to precipitate Cdc2 kinase. Assays were as described (52).

Phosphorylation of GST fusion proteins was analyzed by SDS-polyacrylamide gel electrophoresis (PAGE), and the ³²P-labeled GST-Orp2 and GST-Orp2-T4A bands were excised after autoradiography. Tryptic peptide mapping was performed as described (7) using pH 1.9 buffer for the first dimension of electrophoresis and phosphochromatography buffer for the second dimension of ascending chromatography. Equal amounts of radioactivity were analyzed for each map, so it was necessary to use 1/10 as much of the Cdc2-GST-Orp2 sample as for the other samples. Phosphoamino acid analysis was performed as described (7), and results were quantitated using a phosphoimager (Molecular Dynamics).

Cdc2 phosphorylation of Orc2.

Consensus Cdc2 phosphorylation sites [(S/T)PX(K/R)] (29) are present in the Orc2 and Orp2 homolog proteins in fission yeast, budding yeast, Drosophila melanogaster, Xenopus, and mammals (Fig. 1). These sites occur exclusively within the N-terminal regions of the proteins. This region is poorly if at all conserved. Multiple sequence alignments and PSI-BLAST (1) were used to determine an N-terminal boundary for sequence similarity. The first Orc2 sequence motif conserved from yeast to humans, (F/V)(D/E)EYF, begins at amino acid 193 of S. pombe Orp2. This conserved motif is the boundary between the variable and conserved regions shown in Fig. 1. Even among mammals, there is a sharp change in sequence conservation between the N-terminal region (64% identical between human and mouse) and the C-terminal region (89% identical between human and mouse). The consensus Cdc2 phosphorylation sites occur exclusively within the variable N-terminal region.

The fact that consensus (and usually additional nonconsensus SP or TP) Cdc2 phosphorylation sites are always present in the N-terminal region of Orc2 homologs, coupled with the fact that Cdc2 kinase is important for regulating DNA synthesis, suggests that these Cdc2 sites could be targets of Cdc2 kinase. The first test of this idea is to find out whether Orc2 is a substrate of Cdc2 in vitro. Therefore, a GST-Orp2 fusion protein was mixed with active Cdc2 kinase from fission yeast, and it was found that the GST-Orp2 became phosphorylated (Fig. 2A). GST-Orp2 was a better substrate than was histone H1, while GST did not become phosphorylated at all (data not shown). Identical results were obtained using Cdc2 isolated from fission yeast lysates by binding to p13^suc1 beads or by immunoprecipitation of the cyclin Cdc13. These results show that Cdc2 and Cdc13 complexes can phosphorylate Orp2 in vitro.

Phosphorylation of Orp2 in vitro depends on the consensus Cdc2 phosphorylation sites.

To see if the phosphorylation of Orp2 in vitro depends on the Cdc2 phosphorylation sites, various (S/T)P sequences in Orp2 were mutated to nonphosphorylatable AP sequences by site-directed mutagenesis. Changing the four TP sequences between amino acids 43 and 66 (Fig. 1) yielded the mutant orp2-T4A, which is the mutant characterized in this study. GST-Orp2 and the mutant GST-Orp2-T4A were purified from fission yeast and compared as Cdc2 substrates in vitro. GST-Orp2 was readily phosphorylated, whereas GST-Orp2-T4A was not (Fig. 2A). Quantitation showed that 10-fold more radioactive phosphate was incorporated into GST-Orp2 than into GST-Orp2-T4A. That equal amounts of substrate proteins were added to each reaction was determined by Coomassie staining of the total proteins in the reactions after separation by SDS-PAGE (Fig. 2A). Hereafter, we will refer to Orp2-T4A as the “nonphosphorylatable” Orp2, even though it might be a substrate of other kinases and could even have weak residual or alternative sites for Cdc2 phosphorylation.

Phosphopeptide mapping showed that the major Cdc2 phosphorylation sites in Orp2 were eliminated by the mutation of four threonines in Orp2-T4A (Fig. 2B). Consistent with the idea that threonines are the phosphorylation targets of Cdc2 in Orp2, phosphothreonine was the main (>90%) phosphoamino acid detected in GST-Orp2 phosphorylated by Cdc2 (datum not shown). Some phosphoserine was also detected in this reaction. However, most of this phosphoserine was independent of the addition of Cdc2 and came from a low level of kinase associated with the GST-Orp2 (datum not shown). This other kinase is not Cdc2, because the phosphopeptides differ from those generated by Cdc2 (Fig. 2B), because no Cdc2 was detected by Western blotting in preparations of GST-Orp2 (data not shown), and because this other kinase phosphorylates GST-Orp2 and GST-Orp2-T4A equally well (Fig. 2A).

Phosphorylation of Orp2 in vivo requires the same sites that are phosphorylated by Cdc2 in vitro

Phosphorylation of Orp2 in vivo causes a shift to a slower-migrating species in SDS-PAGE analysis (45). If the slowly migrating Orp2 results from phosphorylation by Cdc2, then the same Cdc2 sites required for Orp2 phosphorylation in vitro should also be required for phosphorylation and mobility shift of Orp2 in vivo. To test this, isogenic strains were constructed having either wild-type orp2⁺ or orp2-T4A. The orp2-T4A strains were viable with no obvious phenotype (see below). Immunoblot analysis showed that asynchronous wild-type cells had several mobility forms of Orp2, whereas the mutant orp2-T4A had a single, rapidly migrating band (Fig. 3). This single form of Orp2-T4A comigrated with the single, rapidly migrating form of wild-type Orp2 observed in G₁ cells (Fig. 3). Mutation of the single consensus Cdc2 phosphorylation site was not sufficient to abolish the cell cycle-regulated phosphorylation (data not shown). That the same Cdc2 sites required for Orp2 phosphorylation in vitro are also required for generation of slowly migrating forms of Orp2 in vivo suggests that Orp2 is an in vivo substrate of Cdc2.

Cell cycle regulation of Orp2 phosphorylation.

To find out when in the cell cycle Orp2 is phosphorylated, we synchronized cells in G₂ using block and release of a cdc25-22 mutant and followed Orp2 protein through two cell cycles (Fig. 4A). Orp2 shifted from slowly migrating forms at the G₂ block to faster migrating forms at the end of mitosis. The fastest migrating (unphosphorylated) form of Orp2 appeared simultaneously with the peak in septation. Typically, fission yeast cells are in the S phase of the next cell cycle by the end of septation; thus, the peak in septation and in abundance of fast-migrating Orp2 occurs near the G₁/S boundary.

As a second method for examining cell cycle regulation of phosphorylation of Orp2, cells were blocked in various stages of the cell cycle using cell cycle mutants or the replication inhibitor hydroxyurea (HU). Orp2 isolated from the arrested cells was analyzed by Western blotting (Fig. 4B). Orp2 was in the fastest migrating form at the cdc10 arrest (G₁ phase). More slowly migrating (phosphorylated) forms first appeared in the S phase (cdc22 and HU arrests) and when the replication factor Cdc18 was inactivated (cdc18). In G₂ and early M (cdc25 and nuc2), Orp2 accumulated in the slowest-migrating forms. It is notable that Orp2 is fully modified at the cdc25 mutant arrest (G₂). Cdc25 is the tyrosine phosphatase required for activating Cdc2 to the high levels required for mitosis. If Cdc2 is the kinase modifying Orp2, then the pool of Cdc2 that phosphorylates Orp2 may not require activation by Cdc25.

The phosphorylation state of Orp2 was assayed in cdc2 mutant cells. Phosphorylation of Orp2 was reduced but not abolished in cdc2-L7 and cdc2-33 mutants at restrictive temperature (Fig. 4B). Because Cdc2 is essential both for G₁-to-S and G₂-to-M transitions, these cdc2 mutants arrest as a mixture of 10 to 20% G₁ cells and 80 to 90% G₂ cells. These data show that inactivation of Cdc2 that is sufficient to arrest cell cycle progression does not result in fully dephosphorylated Orp2.

The phosphorylation state of Orp2 was also assayed in cells lacking cdc13. Cdc13 is a B-type cyclin essential for mitosis and for once-per-cell-cycle control of DNA replication. Selective spore germination was used to obtain a culture of cells lacking Cdc13. These cells cannot proceed past G₂ into mitosis, and they rereplicate DNA. The Orp2 in these cells was in the fast-migrating (unphosphorylated) form (Fig. 4C). Since cells arrested in G₂ by other mutations (Fig. 4B) accumulate phosphorylated forms of Orp2, the lack of Orp2 phosphorylation in the cdc13 mutant suggests that the Cdc13-Cdc2 complex may be directly required for phosphorylation of Orp2. Furthermore, this observation suggests that phosphorylation of Orp2 may not be needed for replication, since cdc13 deletion mutants replicate and rereplicate DNA (27).

Selective spore germination was also used to examine the role of another kinase, Hsk1, in generating phosphorylated Orp2. Hsk1 is essential for DNA replication and is the fission yeast homolog of Cdc7 (47). Cdc7 is bound to replication origins via the ORC and is required for replication initiation at individual origins throughout the S phase (6, 18, 20, 25). Orp2 is fully shifted to the phosphorylated forms even in the hsk1 mutant (Fig. 4C). Furthermore, Hsk1 failed to phosphorylate Orp2 in vitro (data not shown). These results indicate that Hsk1 does not phosphorylate Orp2. This is consistent with the idea that the relevant Hsk1 substrate is Mcm2 and possibly additional MCMs (reviewed in reference 70).

Phosphorylation of Orp2 is not required for DNA replication.

Cdc2 is essential for initiation of replication, and so presumably it has some substrate that is important for initiation. We and others expected that Orp2 could be such a substrate (40, 45). Our data suggest otherwise. Cells carrying orp2-T4A as their only allele of orp2 were viable and healthy, with no obvious phenotype. Likewise, cells with orp2-T4A and also mutated at two additional (S/T)P sites (T159→A and S162→A) were also viable and healthy. Finally, cells expressing GST-orp2-Δ1-127 as the only allele of orp2 were viable and healthy. This GST-Orp2-Δ1-127 fusion protein lacks the first 127 amino acids of Orp2 (approximately two-thirds of the variable region), including all the T-P sequences mutated in orp2-T4A. Except for S139, which has not been tested, all (S/T)P sequences in the variable region of Orp2 can be mutated or deleted without significantly altering replication initiation.

The orp2-T4A allele was integrated into the chromosome at the orp2Δ::ura4⁺ locus under control of the orp2 promoter. Several experiments were done to see if this mutant had a defect in initiation. Flow cytometry showed that the S phase progressed similarly in orp2-T4A and wild-type cells (data not shown). There was no detectable change in cell size at division, which would have indicated a cell cycle delay. There was no genetic interaction between orp2-T4A and cdc18, cdc19, or cdc21 temperature-sensitive alleles defective for replication factors.

To perform a possibly more sensitive assay, we compared the efficiency with which orp2-T4A and wild-type strains could be transformed with an ars plasmid as has been done to evaluate ars sequences in S. pombe (10, 22). The assays described above may be insensitive detectors of initiation defects, because each chromosome has many origins. Even if the efficiency of initiation declined to, say, 30% of wild-type values, this 30% may be adequate to give a wild-type phenotype by growth rate or flow cytometry assays, etc. Since plasmids have only a single origin, any defect in origin usage lowers transformation frequency. To control for other factors affecting transformation and cell viability, the mutant and wild-type strains were mixed and transformed in one tube (Materials and Methods). The orp2-T4A strain was marked with ade6-M210, and the orp2⁺ strain was marked with ade6-M216. Both alleles are Ade⁻, but on rich medium low in adenine (YSO), ade6-M216 yields pink colonies while ade6M-210 yields red colonies. The orp2⁺ ade6-M216 and orp2-T4A ade6-M210 strains were mixed 1:1, grown overnight, and then transformed with ura4⁺ tester plasmids containing wild-type and mutant derivatives of ars3002 (Table 2) (22). Transformants were selected on medium lacking uracil. In addition, cells were plated on medium containing uracil to determine the number of cells of each genotype surviving the transformation protocol. Colonies from both surviving cells and from transformed (i.e., Ura⁺) cells were picked to YSO to determine whether they were orp2⁺ ade6-M216 or orp2-T4A ade6-M210. The ratio of transformants of each genotype was divided by the ratio of surviving cells to normalize for survival and determine the relative transformation efficiency (Materials and Methods).

As shown in Table 2, the orp2-T4A mutant had transformation efficiencies up to threefold higher than the wild-type strain. The plasmids used in this assay contained wild-type and mutant derivatives of ars3002. This test was performed with several different ars constructs. Deletion mutants of ars3002 were chosen because they have been extensively characterized (22). In fact, the wild-type ars showed the greatest difference between orp2-T4A and orp2⁺. Perhaps the differences detected in this assay are obscured by the defective function of the ars mutants. In any case, the results give no indication of any initiation defect in the mutant, but rather suggest that the Cdc2 phosphorylation sites in Orp2 could have an inhibitory function.

Introducing mutations that might mimic phosphorylation of Orp2 had little effect on Orp2 function. Threonines in three of the putative Cdc2 phosphorylation sites were changed to aspartate (Fig. 1). The resulting mutant, orp2-T3D, has no obvious defect in growth or replication (data not shown). This orp2-T3D mutant has a transformation efficiency 0.7- to 1.1-fold that of the wild type in the transformation assay (Table 2); the variations in relative efficiency depended upon the plasmid ars tested. As with orp2-T4A, the greatest difference in transformation efficiency was observed with the wild-type ars, although in this case, the difference is a decrease in transformation efficiency in the mutant; however, all experiments suggest the orp2-T3D mutant is wild type or nearly wild type for initiation.

The Cdc2 phosphorylation sites in Orp2 help prevent rereplication.

Cdc2 is essential for limiting DNA replication to once per cell cycle. Inactivation of Cdc18 is one mechanism used by Cdc2 to prevent reinitiation, but there must be redundant mechanisms, since loss of the phosphorylation sites on Cdc18 does not in itself allow reinitiation. In S. pombe, overexpression of Cdc18 does allow rereplication, probably by supplying functional Cdc18 and also by interfering with other pathways for control of rereplication. Phosphorylation of Orp2 might be one of these additional mechanisms. If so, then we expect no obvious phenotype of mutating Cdc2 phosphorylation sites in Orp2 in wild-type cells, but the role of these sites in regulating rereplication might be revealed if Cdc18 is de-regulated. Therefore, we compared rereplication in orp2-T4A and wild-type strains when cdc18* was overexpressed in each of them. cdc18* lacks four Cdc2 sites and induces reinitiation and rereplication when overexpressed (as does cdc18⁺ itself) (44).

The orp2-T4A mutation enhanced rereplication induced by overexpression of cdc18*. A strain with a stable integrated copy of cdc18* controlled by the inducible nmt promoter was crossed with isogenic orp2-T4A and wild-type orp2⁺ strains that have wild-type levels of Orp2 expressed from its endogenous promoter. nmt-cdc18* orp2-T4A and nmt-cdc18* orp2⁺ strains were isolated by tetrad analysis. Cdc18* was induced, and rereplication was quantitated by flow cytometry (fluorescence-activated cell sorter analysis). The nmt promoter is repressed for at least 10 h after removal of thiamine and is not fully induced until 16 h after removal of thiamine (49). Western blot analysis showed that Cdc18* protein was detectable 13 h after induction in both genotypes, and expression was the same in both genotypes (Fig. 5C). Data shown in Fig. 5A and B are representative of quantitative analysis of four independent isolates of each genotype. An increase in rereplication in orp2-T4A compared with orp2⁺ was first observed when the inducible Cdc18 was wild type for Cdc2 phosphorylation sites (data not shown). However, the difference between orp2-T4A and orp2⁺ was magnified when Cdc2 sites in Cdc18 were mutated.

Overexpression of Cdc18 is still required to drive rereplication even when Cdc2 phosphorylation sites in both Cdc18 and Orp2 are mutated as in the nmt-cdc18* orp2-T4A strain. One explanation for this could be that nmt-cdc18* is not sufficient to supply replication initiation function when the nmt promoter is repressed; note that these cells also have a wild-type cdc18⁺ gene and so are not dependent on nmt-cdc18* for viability. We confirmed the published results that repressed levels of cdc18* can complement the essential replication initiation function of Cdc18 in the cdc18-K9 mutant at restrictive temperature. This suggests that that Orp2 phosphorylation and Cdc18 phosphorylation are redundant with yet more mechanisms for preventing DNA rereplication and that high level expression of Cdc18 is needed to override these mechanisms.