Cdc45 Protein-Single-stranded DNA Interaction Is Important for Stalling the Helicase during Replication Stress

Background: Polymerase stalling is coupled with helicase stalling at a eukaryotic replication fork by an unknown mechanism.

Results: When a Cdc45-ssDNA binding mutant is expressed in budding yeast cells exposed to hydroxyurea, there is uncoupling of the helicase from the polymerase.

Conclusion: Cdc45-ssDNA interaction is important during replication stress.

Significance: A new model explains how polymerase stalling is coupled with helicase stalling.

Abstract

Replicative polymerase stalling is coordinated with replicative helicase stalling in eukaryotes, but the mechanism underlying this coordination is not known. Cdc45 activates the Mcm2-7 helicase. We report here that Cdc45 from budding yeast binds tightly to long (≥ 40 nucleotides) genomic single-stranded DNA (ssDNA) and that 60mer ssDNA specifically disrupts the interaction between Cdc45 and Mcm2-7. We identified a mutant of Cdc45 that does not bind to ssDNA. When this mutant of cdc45 is expressed in budding yeast cells exposed to hydroxyurea, cell growth is severely inhibited, and excess RPA accumulates at or near an origin. Chromatin immunoprecipitation suggests that helicase movement is uncoupled from polymerase movement for mutant cells exposed to hydroxyurea. These data suggest that Cdc45-ssDNA interaction is important for stalling the helicase during replication stress.

Introduction

DNA replication in eukaryotes is a highly regulated event (1–5). Central to the activation of DNA replication is assembly of the replication fork helicase, which is composed of the Cdc45 protein, the heterohexamer Mcm2-7 ring, and the tetrameric GINS complex (6–8). This large helicase assembly is also known as the CMG² (Cdc45·Mcm2-7·GINS) complex. The CMG complex forms during S phase, and it requires the activity of two S phase cellular kinases, the Dbf4-dependent kinase and the cyclin-dependent kinase (9–13). The activity of Dbf4-dependent kinase is specifically required for incorporation of Cdc45 with Mcm2-7, whereas cyclin-dependent kinase is required for the recruitment of GINS to replication origins (9–14).

The CMG complex has markedly higher helicase activity and ATPase activity compared with Mcm2-7 helicase alone (15). The CMG complex is a highly processive and active helicase with high ATPase rates, whereas Mcm2-7 is a very poor ATPase, and particular reaction conditions are required to observe even weak activity in vitro (15–17). These data suggest that Cdc45 and GINS are required for full activation of the replication fork helicase (15, 16). Recent electron microscopy data suggest that the binding of Cdc45 and GINS to Mcm2-7 may favor a closed-ring conformation of Mcm2-7 and that this closed-ring conformation may be required for efficient ATPase and unwinding activity (18). Cdc45, Mcm2-7, and GINS travel with one another away from replication origins (7, 19), and recent data show that the CMG complex surrounds a single strand of DNA during helicase unwinding (20).

Cdc45 binds to Sld3 during much of the cell cycle (21). Furthermore, Cdc45 binds tightly to Sld3 in vitro (22). Sld3 may be important for delivering Cdc45 to the Mcm2-7 complex, and this event may require the prior activity of Dbf4-dependent kinase (21–24). However, Sld3 does not travel with the replication fork, unlike Cdc45 (25). In vitro, interaction of Cdc45 with Sld3 is disrupted by GINS, suggesting an in vivo mechanism to dissociate Cdc45 from Sld3 (22). Cdc45 also binds directly to GINS in vitro (22). The N terminus of Cdc45 is homologous the Escherichia coli nuclease RecJ (26). The functional importance of Cdc45 homology with RecJ has not yet been elucidated. Moreover, the function of the C-terminal region of Cdc45 has not yet been defined.

During episodes of DNA polymerase stalling, a coordinated response occurs that functions to inhibit progression through S phase (27–31). Single-stranded DNA (ssDNA) is often exposed during DNA polymerase stalling events, and RPA-coated ssDNA acts as a signal to trigger the DNA damage response (27–31). The cascade of events involved during the DNA damage response has been characterized previously (27–31). One key downstream effect of the DNA damage response is the inhibition of DNA replication initiation activity (27–32).

When the DNA polymerases are induced to stall by hydroxyurea or other agents, roughly 100 base pairs of DNA are unwound by the replication fork helicase ahead of the polymerase, even in the absence of a DNA damage response (27, 33, 34). Thus, polymerase stalling is coupled with helicase stalling in eukaryotic cells in a manner that does not depend upon the DNA damage response. However, thus far, little is understood about how DNA polymerase stalling is directly coordinated with DNA helicase inhibition.

We report here that Cdc45 binds tightly and directly to yeast genomic ssDNA sequences. We also report that ssDNA specifically disrupts the interaction between Cdc45 and Mcm2-7, suggesting a mechanism to inhibit replication fork helicase activity. The C terminus of Cdc45 is responsible for the ssDNA-binding activity, and we identified a mutant of Cdc45 (Cdc45-M268) that does not bind to ssDNA. The Cdc45-M268 mutant exhibits wild-type binding to Sld3, Mcm2-7, and GINS. When the cdc45-m268 mutant is expressed in budding yeast cells, cell growth is severely inhibited in the presence of hydroxyurea or methyl methanesulfonate. cdc45-m268 cells exposed to hydroxyurea also exhibit increased RPA accumulation near an origin. ChIP data suggest that upon exposure to hydroxyurea, the cdc45-m268 mutant cells have helicase movement uncoupled from polymerase movement. These data suggest that Cdc45-ssDNA interaction is important for stalling the helicase during replication stress.

EXPERIMENTAL PROCEDURES

Proteins

Proteins were purified as described (14, 22). GST-Cdc45, PKA-Cdc45, native Cdc45, GST-Sld3, PKA-Sld3, GST-Mcm2-7, PKA-Mcm2-7, and PKA-GINS were purified as described. GST-Cdc45 (fragments and mutants) and PKA-Cdc45 (fragments and mutants) were purified as the wild-type, full-length Cdc45. Protein kinase A was a generous gift from Susan Taylor. SDS/PAGE/Coomassie analysis of purified proteins is shown in supplemental Fig. S1. α-Factor peptide pheromone was obtained from Zymo Research (catalog no. Y1001).

Strains

The cdc45-td degron strain (YBH42, MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2–3,112 can1–100, Gal:UBR1(HIS3), cdc45-td (KanMX)) (35) was a generous gift of Karim Labib (Paterson Institute for Cancer Research, University of Manchester, Manchester, UK). The degron strain was transformed with the pRS415 plasmid (low copy) containing a wild-type or mutant copy of cdc45 under control of its native promoter. Allelic replacement was performed as described (36). The wild-type or mutant CDC45 gene and native promoter were PCR-amplified from linearized pRS415 containing wild-type or mutant CDC45 to include a Leu marker and 5′ and 3′ sequences to allow for homologous recombination into the CDC45 locus. This PCR fragment was then used to transform a haploid -Leu strain (BY4741). Transformants were selected on complete supplement mixture-Leu medium.

GST Pull-down

The GST pull-down reactions were performed as described (37). GST pull-down reactions were performed in a volume of 100 μl and contained GST-tagged protein (GST-Cdc45, GST-Sld3, or GST-Mcm2-7) in GST-binding buffer (40 mm Tris-HCl (pH 7.5), 100 mm NaCl, 0.1 mm EDTA, 10% glycerol, 0.1% Triton X-100, 1 mm DTT, 0.7 μg/ml pepstatin, 0.1 mm PMSF, and 0.1 mg/ml BSA) and varying amounts of radiolabeled DNA or protein as described in each figure. Reactions were incubated at room temperature for 1 h. Following incubation, reactions were added to 40 μl of prepared glutathione-Sepharose and mixed gently. Binding of GST-tagged protein to the beads was performed for 20 min with gentle mixing every few minutes. When the binding was complete, the beads were allowed to settle, the supernatant was removed, and the glutathione beads were washed twice with 0.5 ml GST-binding buffer. After the last wash, 30 μl of 5× SDS sample buffer was added to each reaction, and the samples were boiled for 10 min. Samples (20 μl) were then analyzed by SDS-PAGE.

Biotin Pull-down

The biotin pull-down reaction were performed as described (38). Biotinylated DNA conjugated to streptavidin-agarose magnetic beads (Dynal) were incubated for 5 min at 30 °C with various concentrations of radiolabeled protein in a solution containing 0.1 mm EDTA, 0.2 mm DTT, 10 mm magnesium acetate, 10% glycerol, 40 μg/ml BSA, 100 mm NaCl, and 20 mm Tris-HCl (pH 7.5) in a final volume of 25 μl. After the 5-min incubation, the beads were collected at room temperature using a magnet (Dynal). The supernatant was aspirated, and the beads were washed twice with a solution containing 0.1 mm EDTA, 0.2 mm DTT, 10 mm magnesium acetate, 10% glycerol, 40 μg/ml BSA, 100 mm NaCl, and 20 mm Tris-HCl (pH 7.5). The beads were collected with a magnet, the supernatant was aspirated, and the beads were heated at 95 °C for 10 min in a solution containing 2% SDS, 2 mm DTT, 4% glycerol, 4 mm Tris-HCl, and 0.01% bromphenol blue. The reactions were analyzed by SDS-PAGE. The gel was dried for 1 h at 80 °C and exposed to a phosphorimaging screen for 1 h.

Fluorescence Anisotropy

Fluorescence anisotropy was performed as described (38). Cdc45-binding to DNA was measured by the increase in anisotropy as varying concentrations of Cdc45 were incubated with 10 nm fluorescent DNA (5-Carboxyfluorescein) for 5 min in a solution containing 0.1 mm EDTA, 0.2 mm DTT, 10 mm magnesium acetate, 10% glycerol, 40 μg/ml BSA, and 20 mm Tris-HCl (pH 7.5) in a final volume of 40 μl. Polarized fluorescence intensities were measured at excitation and emission wavelengths of 495 and 538 nm, respectively. The data were plotted as change in anisotropy versus protein concentration, and dissociation constants (K_d) were derived by fitting the data with the equation y = m0/(m0 + m1), where m1 = K_d.

Kinase Labeling

Kinase labeling was performed as described (22). Proteins containing a protein kinase A tag at the N terminus (Cdc45, Sld3, Mcm2-7 (Mcm3), and GINS (Psf1)) were radiolabeled in a reaction volume of 100 μl that contained 20 μm of protein kinase A-tagged protein in kinase reaction buffer (5 mm Tris-HCl (pH 8.5), 10 mm MgCl₂, 1 mm DTT, 500 μm ATP, 500 μCi [γ-³²P]ATP) containing 5 μg of protein kinase A. Reactions were incubated for 1 h at 30 °C.

EMSA

1 nm radiolabeled DNA (single-stranded polymers of dT of various lengths) was mixed with various concentrations of protein (as detailed in Fig. 2) in 1× EMSA buffer (containing 0.1 mm EDTA, 0.2 mm DTT, 10% glycerol, 40 mg/ml BSA, and 20 mm Tris-HCl (pH 7.5)) for 5 min at 30 °C. 4% Glycerol and 0.01% bromphenol blue were then added to the solution, and the reaction was analyzed by a 10% native polyacrylamide gel in 0.5 × Tris-Borate-EDTA buffer. The gel was dried for 1 h at 80 °C and exposed to a phosphorimaging screen for 1 h.

FIGURE 2.

EMSA. 1 nm radiolabeled DNA (single-stranded polymers of dT of various lengths) was mixed with various concentrations of protein (as detailed in the figure) and analyzed by EMSA as described under “Experimental Procedures.”

Size Exclusion Chromatography

Size exclusion chromatography was performed as described (22). Unlabeled protein was mixed with radiolabeled protein as described in the Figs. 3–5 legends in a final volume of 200 μl and incubated at 30 °C for 1 h in column buffer (50 mm Tris (pH 7.5), 100 mm NaCl, 1 mm EDTA, 5% glycerol). The protein mixture was then subjected to 24 ml Superose 6 (GE Life Sciences) size exclusion chromatography in the same column buffer. The radiolabeled protein from each 250 μl fraction was then quantified and plotted.

FIGURE 3.

Charge reversal mutations in the C-terminal region of Cdc45 inhibit Cdc45 binding to ssDNA. A, 3 pmol GST-Cdc45 full-length or fragments were incubated with varying concentrations of radiolabeled 80-mer ssDNA positioned 1 kb upstream of ARS1-2 (for sequence, see supplemental Fig. S2A). B, 3 pmol biotinylated 80-mer ssDNA positioned 1 kb upstream of ARS1-2 was used to pull down radiolabeled Cdc45 full-length or fragments. C, charge reversal mutations in the C-terminal region of Cdc45 were studied for interaction with 80-mer ssDNA. D, various concentrations of Cdc45-wild-type, Cdc45-M2, Cdc45-M6, or Cdc45-M8 were incubated with 10 nm fluorescently labeled ARS1-2 and analyzed by fluorescence anisotropy as described under “Experimental Procedures.” The data were fit to a single-site binding equation. Cdc45 mutants M1, M3-M5, M7, or M9-M16 bound ssDNA similar to wild-type Cdc45 (data not shown). E, 100 pmol of ³²P-Cdc45-wild-type, ³²P-Cdc45-M2, ³²P-Cdc45-M6, or ³²P-Cdc45-M8 alone or with equimolar Sld3 was subjected to size-exclusion chromatography as described under “Experimental Procedures.” The radioactive counts in each fraction were used to calculate the pmols of Cdc45 in each fraction. Cdc45 (pmols) was then plotted versus the elution of molecular weight standards. F, 100 pmol of ³²P-Cdc45 (wild-type or M2, M6, or M8 mutants) alone or with equimolar Mcm2-7 in the presence of 1 mm ATP-γS was subjected to size-exclusion chromatography as described under “Experimental Procedures.” G, 100 pmol of ³²P-Cdc45 (wild-type or M2, M6, or M8 mutants) alone or with equimolar GINS was subjected to size-exclusion chromatography as described under “Experimental Procedures.”

FIGURE 4.

Cdc45-M268 (R487E, K488E, K520E, and R540E) does not bind ssDNA, but Cdc45-M268 binds Sld3, Mcm2-7, and GINS-like wild-type Cdc45. A, various concentrations of Cdc45-wild-type or Cdc45-M268 were incubated with different fluorescent 80-mer ssDNA sequences (sequences are shown in supplemental Fig. S2A) and analyzed by fluorescence anisotropy. The data were fit to a single-site binding equation. B, 100 pmol of ³²P-Cdc45-wild-type or ³²P-Cdc45-M268 alone or with equimolar Sld3 was subjected to size-exclusion chromatography as described under “Experimental Procedures.” The radioactive counts in each fraction were used to calculate the pmols of Cdc45 in each fraction. Cdc45 (pmols) was then plotted versus the elution of molecular weight standards. C, 100 pmol of ³²P-Cdc45 (wild-type or M268 mutant) alone or with equimolar Mcm2-7 in the presence of 1 mm ATP-γS was subjected to size-exclusion chromatography as described under “Experimental Procedures.” D, 100 pmol of ³²P-Cdc45 (wild-type or M268 mutant) alone or with equimolar GINS was subjected to size-exclusion chromatography as described under “Experimental Procedures.”

FIGURE 5.

60-mer ssDNA releases Cdc45 from Mcm2-7. Sequences of DNA are found in supplemental Fig. S2B. A, 100 pmol of ³²P-Cdc45-wild-type alone or with equimolar Sld3 and/or 2-fold molar excess of DNA was subjected to size-exclusion chromatography as described under “Experimental Procedures.” The radioactive counts in each fraction were used to calculate the pmols of Cdc45 in each fraction. Cdc45 (pmols) was then plotted versus the elution of molecular weight standards. B, 100 pmol of ³²P-Cdc45-wild-type alone or with equimolar GINS and/or 2-fold molar excess of DNA was subjected to size-exclusion chromatography as described under “Experimental Procedures.” C, 100 pmol of ³²P-Cdc45-wild-type alone or with equimolar Mcm2-7 and/or 2-fold molar excess of DNA in the presence of 1 mm ATP-γS was subjected to size-exclusion chromatography as described under “Experimental Procedures.” D, 100 pmol of ³²P-Cdc45-M268 alone or with equimolar Mcm2-7 and/or 2-fold molar excess of DNA in the presence of 1 mm ATP-γS was subjected to size-exclusion chromatography as described under “Experimental Procedures.”

Serial Dilution Analysis

Serial dilution was performed as described (37). For degron experiments, overnight culture (complete supplement mixture-leu containing raffinose, 24 °C) was transferred into YPGal medium and incubated for 1 h at 24 °C. Then, 10-fold serial dilutions were performed on the indicated media and incubated at the indicated temperatures as described previously. For allelic replacement, cells were grown overnight and plated onto the indicated media at 30 °C.

Fluorescence-activated Cell Sorting

FACS was performed as described (37). For cell cycle analysis, the strains were grown overnight at 30 °C. For G₁ arrest and release experiments, 6 × 10⁶ cells/ml were transferred into yeast-peptone-dextrose medium and treated with α-factor (Zymo Research) for 3 h at 30 °C following extensive washes and addition of 50 μg/ml Pronase (Calbiochem) to fresh YPD. Cells were then incubated further for the indicated time. Cell cycle progression was followed by flow cytometry (FACS) stained with propidium iodide using FACSAria.

In Situ Autophosphorylation Assay

An in situ autophosphorylation assay was performed as described (39). Cells were grown in YPD at 30 °C and arrested with α-factor. Cells were then released into 0.2 m HU for 90 min, and samples were collected and analyzed. Protein samples were prepared and transferred by immunoblotting onto PVDF. The membrane was then subjected to denaturation (50 mm Tris-HCl (pH 8.0), 2 mm EDTA, 50 mm DTT, 7 m guanidine HCl, 1 h at 24 °C), washed twice with TBS, and renatured (10 mm Tris HCl (pH 7.5), 140 mm NaCl, 1% BSA, 0.04% Tween 20, 2 mm EDTA, 2 mm DTT) at 4 °C with 5 changes of buffer over a 16-h period. The membrane was washed with 30 mm Tris HCl (pH 7.5) for 1 h and then equilibrated in kinase buffer (40 mm Hepes NaOH (pH 8.0), 1 mm DTT, 0.1 mm EGTA, 20 mm MnCl₂, 100 μm sodium orthovanadate) for 30 min at room temperature. Fresh kinase buffer supplemented with 10 μCi/ml [γ-³²P]ATP was added to the membrane and incubated for 1 h at room temperature. The blot was washed extensively (10 min each) twice with 30 mm Tris HCl (pH 7.5), once with 30 mm Tris HCl (pH 7.5) and 0.1% Nonidet P-40, once with 30 mm Tris HCl (pH 7.5), once with 1 m potassium hydroxide, once with H₂O, once with 10% TCA, and once with H₂O. The membrane was dried and analyzed using a phosphorimager.

Antibodies

Antibodies directed against RPA (Pierce, catalog no. MA1-25889), Rad53 (Santa Cruz Biotechnology, Inc., catalog no. SC-6749), and Pol2 (Santa Cruz Biotechnology, Inc., catalog no. SC-6752) were purchased. Open Biosystems produced polyclonal antibodies directed against Cdc45 or Psf2. We supplied the antigens. Crude serum was purified against immobilized antigen to remove nonspecific antibodies. The specificity of each antibody was analyzed by Western blot analysis of purified protein and wild-type yeast extract as shown in supplemental Fig. S4.

ChIP

For G₁ arrest and release experiments, 6 × 10⁶ cells/ml were transferred into YPD medium, treated with α-factor (Zymo Research) for 3 h at 30 °C following extensive washes and addition of 50 μg/ml Pronase (Calbiochem) to fresh YPD. Cells were further incubated for the indicated time. Chromatin immunoprecipitation was performed as described (40, 41), except we performed PCR with [³²P-α]dCTP as a component of the PCR reaction to quantify the amplified product (42). Formaldehyde cross-linked cells were lysed with glass beads in a bead beater. DNA was fragmented by sonication (Branson 450). Antibody and magnetic protein A beads (Dynabeads protein A, Invitrogen, catalog no. 100.02D) were added to the cleared lysate to immunoprecipitate the DNA. Immunoprecipitates were washed extensively to remove nonspecific DNA. Eluted DNA was then subjected to PCR analysis using primers directed against ARS305 or +4 kb upstream of ARS305 as described (8, 42). The radioactive bands in the agarose gel, each representing specific PCR-amplified DNA product, were quantified by phosphorimaging and normalized by a reference standard run in the same gel. The reference standard was a PCR reaction accomplished with a known quantity of template DNA replacing the immunoprecipitate.

RESULTS

Cdc45 Binds Single-stranded DNA with Little Sequence Specificity

Cdc45 is part of the replication fork helicase assembly in eukaryotes, and we postulated that Cdc45 might bind directly to ssDNA as part of its function. We purified Cdc45, Sld3, Mcm2-7, and GINS to homogeneity as described previously (Coomassie-stained gels are shown in supplemental Fig. S1) (22, 43, 44). Because Cdc45 initially binds at a replication origin and then travels away from the origin (8), we investigated budding yeast Cdc45 interaction with ssDNA and sequences positioned at the origin ARS1 and non-origin sequences positioned 0.5 kb and 1.0 kb upstream of ARS1 (supplemental Fig. S2A). 80-mer ssDNA was fluorescently labeled and tested for interaction with Cdc45 by fluorescence anisotropy (Fig. 1A). Using this technique, we found that Cdc45 bound to 80-mer ARS1 ssDNA and 80-mer ssDNA positioned 0.5 kb and 1.0 kb upstream (Fig. 1A). We fit the data to a single-site binding equation and found that the dissociation constant for Cdc45 binding to these 80-mer ssDNA sequences varied from 59 + 5 nm to 79 + 6 nm. However, Cdc45 bound very weakly to double-stranded DNA (dsDNA) at the ARS1 origin or at a region positioned 1 kb upstream, suggesting that Cdc45-DNA interaction is specific for ssDNA (Fig. 1A).

FIGURE 1.

Budding yeast Cdc45 binds to 80-mer single-stranded DNA at the replication origin ARS1 and at regions positioned 0.5 or 1.0 kb upstream. In all panels, the concentration of DNA is for oligomers, not nucleotides. A, 10 nm fluorescently labeled ssDNA or dsDNA positioned at ARS1 or 0.5 or 1 kb upstream of ARS1 (for sequences, see supplemental Fig. S2A) was incubated with varying concentrations of Cdc45. The results were analyzed by measuring the change in anisotropy as described under “Experimental Procedures.” The data were fit to a single-site binding equation to obtain a K_d, indicated at the right side of the graph. B, the ssDNA from ARS1 was deleted sequentially from the 5′ end to obtain oligonucleotides of the length of 80, 60, 40, and 20 bases. 80-mer and 60-mer ssDNAs bind to Cdc45 with a higher affinity than shorter ssDNA or dsDNA. C, experiments are similar to B, but the ARS1 ssDNA is sequentially deleted from the 3′ end. N.D., not determined.

To determine whetherCdc45 binding to ssDNA was specific for ARS1 and neighboring sequences, we also examined the interaction between Cdc45 and the early origin ARS305 and sequences positioned 0.5 kb and 1 kb downstream of ARS305 (Supplemental Fig. S3A). Cdc45 bound to these 80-mer ssDNA sequences with a dissociation constant that varied from 54 + 4 nm to 74 + 5 nm (supplemental Fig. S3B). Binding of Cdc45 to dsDNA encompassing the same regions was substantially weaker (supplemental Fig. S3B). We confirmed the interaction between Cdc45 and the 80-mer ssDNA sequences using GST-Cdc45 and radiolabeled ssDNA (supplemental Fig. S3C). In summary, the data suggest that Cdc45 binds to 80-mer ssDNA with little sequence specificity but that Cdc45 binds very weakly to 80-mer dsDNA.

Cdc45 Binds Tighter to 60-mer or 80-mer ssDNA Than to Shorter ssDNA

To determine the effect of ssDNA length on Cdc45 binding, we sequentially deleted nucleotides from the 80-mer ARS1 ssDNA from the 5′ end (Fig. 1B). The K_d for interaction of Cdc45 with 60-mer ssDNA was determined to be 92 + 6 nm. The K_d for 40-mer DNA was 424 + 24 nm, and a K_d could not be determined for the 20-mer ssDNA sequence. Similar results were found for sequential deletions from the 3′ end of the same sequence (Fig. 1C). These data suggest that Cdc45 exhibits higher affinity for 60-mer and 80-mer ssDNA compared with shorter ssDNA.

We also used the electrophoretic mobility shift assay (EMSA) to analyze Cdc45 binding to oligomers of dT, including lengths of 60 dT, 40 dT, 30 dT, 20 dT, and 10 dT (Fig. 2). The data show a slowly migrating band for the 60-mer dT incubated with 30 nm Cdc45 (Fig. 2A), consistent with a single protein-DNA complex formed between Cdc45 and 60-mer dT. The binding for the 60dT occurs at lower concentration of protein compared with the 40dT (Figs. 2A and 2B). In contrast, little binding is observed for 30dT, 20dT, and 10dT (Figs. 2C-2E). Protein-monomer-DNA-binding-sites are typically shorter than 40 nucleotides. Cdc45 may be sliding along the DNA in a manner that experimentally reveals a larger DNA binding site than its actual discrete interaction. It is also possible that a dimer of Cdc45 is binding to 40-mer or 60-mer single-stranded DNA.

Charge Reversal Mutations in the C-terminal Region of Cdc45 Inhibit Binding to 80-mer ssDNA

To study the effect of Cdc45-ssDNA binding in vivo, we identified separation of function mutations that disrupt Cdc45-ssDNA binding but do not disrupt Cdc45-Sld3 binding, Cdc45-Mcm2-7 binding, or Cdc45-GINS binding. To identify these separation-of-function mutations, we first determined the region of Cdc45 that binds to 80-mer ssDNA (Fig. 3, A and B). Using the GST pull-down and biotin pull-down assays, we found that amino acids 450–650 of Cdc45 bind to 80-mer ssDNA (1 kb-upstream of ARS1, sequence in Fig. 3A). Region 1–450 or region 1–207 of Cdc45 do not bind to 80-mer ssDNA (Fig. 3, A and B).

There are 23 positively charged residues in the 450–650 region of Cdc45. Anticipating that a charge reversal mutation of a positively charged residue in this region may inhibit interaction with negatively charged ssDNA, we systematically engineered 16 mutations, labeled M1 to M16, that contain one or more charge reversal mutations (Fig. 3C). Mutations M2, M6, or M8 partially inhibit the interaction between Cdc45 and 80-mer ssDNA, as measured by fluorescence anisotropy (Fig. 3D). The M1, M3-M5, M7, and M9-M16 mutations exhibited no defect in ssDNA binding (not shown). Mutations M2, M6, or M8 had no effect on the interaction between Cdc45 and Sld3 (Fig. 3E), between Cdc45 and Mcm2-7 (F), or between Cdc45 and GINS (G). These results suggest that we have identified mutations that specifically inhibit the interaction between Cdc45 and ssDNA.

Cdc45-M268 Is a Mutant That Specifically and Completely Disrupts the Interaction between Cdc45 and ssDNA

M2, M6, or M8 each partially inhibited the interaction between Cdc45 and ssDNA (Fig. 3D). To find a Cdc45 mutant that is completely deficient in ssDNA binding, we combined mutations M2, M6, and M8 to form the M268 mutation of Cdc45 (Cdc45-M268: R487E, K488E, K520E, and R540E). Cdc45-M268 is completely defective in binding four different 80-mer ssDNA sequences examined, as measured by fluorescence anisotropy (Fig. 4A). To determine whether the Cdc45-M268 mutation is specifically defective in ssDNA binding or whether it is defective in some other biochemical property, we examined the interaction between Cdc45-M268 with Sld3, Mcm2-7, or GINS and compared the results with wild-type Cdc45 (Fig. 4, B–D). Cdc45-M268 exhibited wild-type behavior with regards to Sld3 binding (Figs. 4B), Mcm2-7 binding (C), or GINS binding (D). These data suggest that Cdc45-M268 is completely and specifically defective in ssDNA binding.

Single-stranded DNA Specifically Disrupts the Interaction between Cdc45 and Mcm2-7

We next examined the biochemical consequences of Cdc45-binding to ssDNA. We specifically studied how Cdc45-binding to ssDNA affects the interaction between Cdc45 and Sld3, GINS, or Mcm2-7 (Fig. 5 and supplemental Fig. S2B). Using size-exclusion chromatography, we found that ssDNA had no effect on the interaction between Cdc45 and Sld3 (Fig. 5A). ssDNA also had no effect on the interaction between Cdc45 and GINS (Fig. 5B). However, 60-mer ssDNA disrupted the interaction between Cdc45 and Mcm2-7 (Fig. 5C). To determine whether the disruption of interaction between Cdc45 and Mcm2-7 was specific for 60-mer ssDNA, we also examined the effect of 60-mer dsDNA or 20-mer ssDNA on the interaction between Cdc45 and Mcm2-7 (Fig. 5C). 60-mer dsDNA or 20-mer ssDNA did not disrupt the interaction between Cdc45 and Mcm2-7, suggesting that DNA disruption of Cdc45-Mcm2-7 interaction is specific for long, single-stranded DNA.

To determine whether the disruption observed is due to ssDNA-Cdc45 interaction or ssDNA-Mcm2-7 interaction, we next studied whether ssDNA affects the interaction between Cdc45-M268 and Mcm2-7. 60-mer ssDNA has no effect on the interaction between Cdc45-M268 and Mcm2-7 (Fig. 5D). These data suggest that 60-mer ssDNA binding to Cdc45 specifically disrupts the interaction between Cdc45 and Mcm2-7.

cdc45-m268 Cells Are Sensitive to Hydroxyurea and Methyl Methanesulfonate Exposure

We acquired a temperature-sensitive degron yeast strain of CDC45 (cdc45-td) from the laboratory of Karim Labib (35). When cdc45-td cells are shifted from 24 °C to 37 °C in the presence of galactose, wild-type Cdc45 is degraded. We transformed cdc45-td cells with a plasmid expressing CDC45 or cdc45-m268 under control of its native promoter. When these cells are studied for cell growth by 10-fold serial dilutions and plating onto normal growth media, the results show no growth defect for the cdc45-m268 cells at the permissive temperature (24 °C) and no growth defect at the restrictive temperature (37 °C) (Fig. 6, A and B).

FIGURE 6.

Cells expressing cdc45-m268 (R487E, K488E, K520E, and R540E) exhibit a severe growth defect in the presence of HU or MMS. A and B, 10-fold serial dilutions of cdc45-td cells expressing CDC45 or cdc45-m268 at the permissive (24 °C) (A) or restrictive temperature (37 °C) (B–D); gal, galactose. C and D, same as B, except cells are exposed to 0.2 m HU (C) or 0.01% MMS (D) upon plating. E, the endogenous locus for CDC45 was replaced with cdc45-m268. These cells are analyzed in E–J and in the text. Shown are 10-fold serial dilutions of cells expressing CDC45 or cdc45-m268 at 30 °C. F, same as E, except cells were exposed to 0.2 m hydroxyurea. G, same as E, except cells were exposed to 0.01% MMS. H, Western blot with antibody directed against Cdc45 from whole cell extracts from wild-type CDC45 or cdc45-m268 cells. I, Rad53 autophosphorylation from whole cell extracts from cells expressing CDC45 or cdc45-m268 in the absence or presence of 0.2 m hydroxyurea. Cells were presynchronized with α-factor prior to exposure to hydroxyurea.

We next subjected the cells to reagents that induce polymerase stalling. Hydroxyurea depletes the cells of nucleotide pools, thereby inhibiting DNA polymerases. Methyl methanesulfonate alkylates DNA, thereby inhibiting DNA polymerases. cdc45-td cells with a plasmid expressing CDC45 or cdc45-m268 under control of its native promoter cells were then plated on growth medium at the restrictive temperature (37 °C) containing 0.2 m HU or 0.01% Methyl methanesulfonate (MMS) (Fig. 6, C and D). Under these conditions, cell growth is substantially disrupted in the cdc45-m268 cells compared with wild-type CDC45 cells.

Cell growth is normal for cells expressing cdc45-m268 from a plasmid in the absence of HU or MMS (Fig. 6, A and B), and we therefore integrated cdc45-m268 in the genome at the endogenous locus for CDC45. This method ensures that Cdc45-M268 copy number is identical to that of wild-type cells. As expected, growth of CDC45-wt or cdc45-m268 (allelic replacement) cells was identical as measured by serial dilution on agar plates (Fig. 6E). However, when the cells were plated onto medium containing 0.2 m HU or 0.01% MMS, a severe growth defect was observed for cdc45-m268 cells (Figs. 6, F and G). Expression of Cdc45-M268 in cdc45-m268 cells is equal to that of Cdc45 expression in wild-type cells (Fig. 6H), suggesting that the defect in cell growth is not due to altered protein expression. These data suggest that Cdc45-ssDNA interaction is important for cell growth in the presence of agents that induce DNA polymerase stalling.

We then utilized the cells modified by allelic replacement for further analysis. We first determined if there is a defect in S phase progression in wild-type compared with mutant cells by FACS analysis (Fig. 7). Cells were synchronized in G₁ with α-factor and then released into medium lacking α-factor for 0–120 min (Figs. 7, A and B). The rate of progression of wild-type and mutant cells through S phase was similar for wild-type compared with mutant cells, suggesting that Cdc45-ssDNA interaction is not important for S phase progression under normal conditions.

FIGURE 7.

Cells expressing CDC45-wt or cdc45-m268 (allelic replacement) in the absence of hydroxyurea demonstrate similar rates of progression through S phase. Fluorescence-activated cell sorting was performed on cells expressing CDC45-wt (A) or cdc45-m268 (B). Cells were synchronized in G₁ with α-factor and then released into medium lacking α-factor for the indicated time points.

cdc45-m268 Cells Exhibit Hyperactive Rad53 Autophosphorylation and Excess RPA Accumulation at or Near an Origin upon Exposure to Hydroxyurea

We next examined activation of the DNA damage response in wild-type and mutant cells (modified by allelic replacement) in the absence or presence of hydroxyurea. The exposure of excess ssDNA coated by RPA, the eukaryotic single-stranded binding protein, induces the DNA damage response, and autophosphorylation of Rad53 is a signal for activation of the DNA damage response pathway (39). In the absence of hydroxyurea, wild-type and mutant cells exhibit similar levels of Rad53 autophosphorylation (Fig. 6I). However, in the presence of hydroxyurea, mutant cells exhibit modestly greater autophosphorylation of Rad53 (Fig. 6I). Phosphorylated Rad53 is the active form of the kinase. A slightly retarded mobility of Rad53 in the presence of HU is revealed by Western blot analysis, consistent with a phosphorylated species. These data suggest that Cdc45-ssDNA interaction is important to prevent hyperactivation of the DNA damage response.

Activation of Rad53 autophosphorylation is usually the result of excess RPA-coated ssDNA. Thus, we next determined whether excess RPA accumulates in mutant cells treated with hydroxyurea. To perform this analysis, we used chromatin immunoprecipitation to monitor the position of RPA relative to the early origin ARS305 or a site +4 kb upstream of ARS305 (Fig. 8, A and B). We synchronized the cells in G₁ with α-factor and then released the cells into HU for 15–45 min. The results indicate that for mutant cells released into hydroxyurea, there is a substantial increase in PCR signal compared with the wild type for DNA positioned at an early origin and at regions positioned +4 kb upstream, especially at the 30 and 45 min time points (Fig. 8, A and B). These data suggest that in mutant cells synchronized in G₁ and then released into hydroxyurea-containing media, there is an increase in RPA-bound DNA at or near an origin.

FIGURE 8.

Cells expressing cdc45-m268 in the presence of hydroxyurea demonstrate accumulation of RPA-coated DNA at or near an origin. A and B, chromatin immunoprecipitation of cells expressing CDC45-wt or cdc45-m268. Anti-RPA antibody was used. Cells generated by allelic replacement were presynchronized with α-factor prior to release into medium containing 0.2 m HU. PCR primers were used that target the early yeast replication origin ARS305 or a region positioned +4 kb upstream of ARS305. [³²P-α]dCTP was included in the PCR reaction for quantitation. Data from A were quantified and plotted in B. C and D, similar to A and B, except anti-Cdc45 antibody was used.

Helicase Movement May Be Uncoupled from Polymerase Movement in cdc45-m268 Cells Exposed to Hydroxyurea

We next used ChIP to monitor the position of the helicase or polymerase relative to the early origin ARS305 or a site +4 kb upstream of ARS305 (Fig. 9). We monitored the position of the replicative helicase by performing ChIP with antibodies direct against a subunit of GINS (anti Psf2 antibody, Fig. 9, A and B). Verification of new antibodies used in this study is shown in supplemental Fig. S4). Wild-type cells were synchronized with α-factor and then released into medium containing hydroxyurea for 0–45 min (Fig. 9, A and B). For wild-type cells, the PCR signal for ARS305 and +4 kb ARS305 peaks at 15 min, and the signal decreases very slightly at 30 and 45 min. These data are consistent with helicase stalling near an origin in wild-type cells exposed to hydroxyurea. It has been observed previously that when the polymerase stalls, the helicase also stalls in wild-type cells because the movement of the polymerase and helicase is coupled (33). Thus, these data are consistent with the published literature.

FIGURE 9.

Cells expressing cdc45-m268 in the presence of hydroxyurea exhibit uncoupled movement of the replicative helicase. Shown is chromatin immunoprecipitation of cells expressing CDC45-wt or cdc45-m268. Cells were presynchronized with α-factor prior to release into medium containing 0.2 m HU. PCR primers were used that target the early yeast replication origin ARS305 or a region positioned +4 kb upstream of ARS305. [³²P-α]dCTP was included in the PCR reaction for quantitation. A and B, anti-Psf2 (a GINS component) antibody was used. C and D, anti-Pol2 (the catalytic subunit of Pol ϵ) antibody was used.

We then conducted a similar ChIP experiment with cdc45-m268 mutant cells (Fig. 9, A and B). When cdc45-m268 cells are synchronized in G₁ and then released into hydroxyurea for 0–45 min, the PCR signal for 15 min is modest, suggesting that GINS is properly incorporated into the helicase in mutant cells. However, the signal at 30 and 45 min time points is very weak in mutant cells compared with the wild type. These data suggest that the helicase does not stall at or near an origin in the presence of hydroxyurea at the 30 or 45 min time points. These data suggest that HU does not stall the replicative helicase in the mutant cells and that Cdc45-ssDNA interaction may be important to stall the helicase during replication stress.

To determine whether helicase movement is uncoupled from polymerase movement in mutant cells in the presence of hydroxyurea, ChIP was performed with antibodies directed against the catalytic subunit of the leading strand polymerase (anti-Pol2 antibody, Fig. 9, C and D). The results indicate that in wild-type cells, the polymerase stalls near an early replication origin in the presence of hydroxyurea, as expected (Fig. 9, C and D). The results for the mutant cells are very similar for the wild-type cells (Fig. 9, C and D), suggesting that the polymerase stalls near an early replication origin in the presence of hydroxyurea. Taken together, the data from Fig. 9 suggest that for mutant cells exposed to hydroxyurea, the polymerase stalls whereas the helicase does not stall. Thus, helicase movement may be uncoupled from polymerase movement in mutant cells exposed to hydroxyurea.

We also performed ChIP analysis with antibodies directed against Cdc45 to determine the position of another components of the CMG complex (Figs. 8, C and D). At 15 min, the PCR signal for wild-type and mutant cells are similar at ARS305 or +4 kb upstream. However, at 30 and 45 min, the signal for the mutant cells is decreased substantially compared with wild-type cells. These data provide further support that the helicase does not stall at or near an origin in mutant cells released into hydroxyurea for 30 or 45 min.

DISCUSSION

We found that Cdc45 binds to eight of eight 80-mer ssDNA sequences examined by the fluorescence anisotropy, GST pull-down, or biotin pull-down assays. The ssDNA sequences examined were positioned at an origin or at a region 0.5 kb or 1 kb upstream or downstream from an origin. These data suggest that Cdc45 binds to 80-mer ssDNA with little sequence specificity. Cdc45 binds to 80-mer ssDNA with a K_d that varies from 54 + 4 nm to 79 + 6 nm for eight oligonucleotides examined. Binding to dsDNA was substantially weaker, suggesting that Cdc45-binding to 80-mer DNA is specific for single-stranded DNA. Binding to 60-mer ssDNA was similar to that for 80-mer ssDNA. 60-mer ssDNA disrupts the interaction between Cdc45 and Mcm2-7, but 60-mer ssDNA has no effect on the interaction between Cdc45 and Sld3 or Cdc45 and GINS.

Region 450–650 of Cdc45 binds to 80-mer ssDNA, and charge reversal of four of 23 positively charged amino acids in the 450–650 region resulted in a decrease in 80-mer ssDNA binding activity. When the four mutations that exhibited a DNA binding defect were combined into one mutant protein (Cdc45-M268, R487E, K488E, K520E, and R540E), the resulting mutant did not bind at all to 80-mer ssDNA. However, Cdc45-M268 bound to Sld3, Mcm2-7, and GINS like wild-type Cdc45. When cdc45-m268 was expressed in budding yeast cells, no growth defect was observed under normal growth conditions. However, in the presence of 0.2 m hydroxyurea or 0.01% methyl methanesulfonate, expression of cdc45-m268 resulted in a severe growth defect of budding yeast cells. Furthermore, expression of cdc45-m268 in the presence of hydroxyurea resulted in excess RPA accumulation near an origin with hyperactive autophosphorylation of Rad53.

ChIP data suggest that for wild-type or mutant cells exposed to hydroxyurea, Pol2 stalls near an early replication origin as expected (Fig. 9, C and D). Furthermore, in wild-type cells, Psf2 and Cdc45 stall upon exposure to hydroxyurea (Fig. 9, A and B and Fig. 8, C and D). However, in mutant cells, Psf2 and Cdc45 do not stall upon exposure to hydroxyurea. Furthermore, high levels of RPA accumulate in mutant cells in the presence of hydroxyurea (Fig. 8, A and B). These data suggest that for mutant cells exposed to hydroxyurea, the helicase unwinds DNA, whereas the polymerase is stalled, and that RPA ssDNA accumulates in the region between the helicase and polymerase (Fig. 10).

FIGURE 10.

Cdc45-ssDNA interaction is important for stalling the helicase during replication stress. A, CDC45-wt-coupled polymerase and helicase stalling. The CMG complex unwinds DNA by encircling the leading DNA strand. Cdc45 does not bind to DNA when the polymerase is active because the protein only binds to 60-mer or longer ssDNA. When the polymerases stall, the exposed ssDNA will bind to Cdc45. Cdc45 will then release from Mcm2-7, stopping the replication fork helicase and inhibiting excess DNA from unwinding. Cdc45 remains bound to GINS, and there are a number of additional proteins (not shown) that from stable interactions between helicase and polymerase components. B, cdc45-m268 uncoupled helicase unwinding. When the polymerases stall, Cdc45 remains bound to Mcm2-7. The helicase unwinds DNA without polymerase activity, and RPA-coated ssDNA accumulates between the helicase and polymerase. There are a number of additional proteins (not shown) that form stable interactions between helicase and polymerase components.

Cdc45 Exhibits Length Dependence but Not Sequence Dependence for ssDNA

Cdc45-binding to ssDNA exhibits high affinity when the ssDNA is 60 nucleotides or greater, but there is little sequence dependence for Cdc45-ssDNA binding. The observation that Cdc45 exhibits a long-length dependence but little sequence dependence suggests that it will bind any genomic ssDNA, provided that at least 60 base pairs are unwound. It is not clear whether this length of ssDNA would be present at a normal replication fork, but this type of DNA structure would likely exist under conditions of DNA polymerase stalling (27–31). DNA polymerase stalling may yield 60-mer ssDNA that may bind to Cdc45.

The C-terminal region of Cdc45 binds to ssDNA. This region of Cdc45 is not homologous to RecJ (26). There is a very recent report that human Cdc45 binds to ssDNA in vitro (45). It will be interesting to determine in the future whether the mechanism we revealed for Cdc45 dissociation from Mcm2-7 in budding yeast is also operative in mammalian cells. Boos et al. (46) recently speculated that Cdc45 may bind to the lagging strand during unwinding, given the homology between Cdc45 and RecJ. Our data suggest that if Cdc45 does indeed bind the lagging strand during unwinding, then this function is not essential under normal conditions because our Cdc45 DNA-binding mutant replicates DNA like wild-type cells in the absence of DNA damage.

It was reported very recently that Mcm10 is involved in activating the CMG complex at a replication origin (47). Mcm10 has also been reported to bind DNA (48), and this function is likely related to replication initiation, not helicase stalling during replication stress. Thus, Cdc45-DNA interaction is likely performing a different function than Mcm10-DNA interaction because we report here that Cdc45-DNA interaction is not required for helicase activation.

Cdc45-ssDNA Interaction Is Important for Stalling the Helicase during Replication Stress

When budding yeast cells expressing cdc45-m268 are exposed to hydroxyurea or methyl methanesulfonate, the cells exhibit a severe growth defect. Furthermore, cells expressing cdc45-m268 and exposed to hydroxyurea accumulate excess RPA near an origin and hyperactive autophosphorylation of Rad53. These data suggest that Cdc45 interaction with ssDNA may be important during DNA polymerase stalling. Exposure of budding yeast cells to hydroxyurea or methyl methanesulfonate results in the accumulation of ssDNA (27–31), and this exposed ssDNA may bind to Cdc45 under normal conditions (Fig. 10A). However, in the cdc45-m268 mutant cells, Cdc45-M268 does not bind to the exposed ssDNA during DNA damage, and the helicase unwinds DNA in an uncoupled manner (Fig. 10B).

We found that 60-mer ssDNA disrupts the interaction between Cdc45 and Mcm2-7. Cdc45, along with GINS, activates the Mcm2-7 helicase (15). Disruption of the interaction between Cdc45 and Mcm2-7 may cause the unwinding activity to stop (Fig. 10A). Helicase inhibition may be important for cell survival during polymerase stalling to prevent excess DNA unwinding (27–31). We speculate that for normal cells, once the DNA polymerase restarts and the DNA becomes double-stranded, Cdc45 will bind to Mcm2-7, and the helicase will once again be active. Taken together, our data suggest a cellular mechanism to couple the inhibition of replicative DNA polymerase activity with the inhibition of replicative DNA helicase activity. Namely, Cdc45 interaction with ssDNA dissociates Cdc45 from the Mcm2-7 complex to inhibit DNA unwinding during DNA replication stress (Fig. 10).

Our model (Fig. 10) predicts that the association of various components of the replication fork will remain associated with one another in mutant and wild-type cells in the presence of hydroxyurea because GINS can form a stable bridge between Cdc45 and Mcm2-7, and Mrc1 can form a bridge between Pol ϵ and Mcm2-7. Immunoprecipitation data accomplished at physiologic conditions support this model because we found no substantial difference in replisome protein associations in mutant cells compared with wild-type cells under physiologic conditions (data not shown).

A recent report describes homology between RecJ and Cdc45, and the report also demonstrates that human Cdc45 can bind ssDNA (45). We speculate here, and also in a report of a recent EM structure of the CMG complex (18), that Cdc45 may bind to the lagging strand that is sterically excluded from the replicative helicase. We also speculate that Cdc45-ssDNA binding may chaperone the lagging strand toward the polymerase (45). We find that our Cdc45 mutant defective in binding ssDNA is competent for DNA replication under normal conditions. Although we cannot definitively rule out the possibility that Cdc45 may bind to the lagging strand during normal cellular replication, we can assert that Cdc45 ssDNA activity is not essential for budding yeast DNA replication under normal conditions.

The CMG Complex May Be a Poor Helicase in Vitro because of the High Level of ssDNA Present in Biochemical Assays

Recent work by the Hurwitz group (17) demonstrates that the CMG helicase is processive only in the presence of a polymerase, RPA, or E. coli single-stranded binding protein. The data from this work suggest that the CMG complex may require E. coli SSB for processive helicase activity because single-stranded DNA present in these biochemical assays is toxic to the CMG. Single-stranded DNA in these assays may bind to Cdc45, thereby dissociating Cdc45 from the Mcm2-7 complex and inactivating the CMG helicase. E. coli SSB may bind the excess ssDNA and allow the CMG helicase to unwind DNA processively. Recent data from the Masukata laboratory (49) demonstrates that a mutation in polymerase ϵ stalls the replicative helicase in vivo. If the polymerase is inactive in vivo, single-stranded DNA will accumulate behind the helicase and stall the helicase. Thus, our model helps explain the recent published data from the Hurwitz and Masukata laboratories (17, 49).

Modular Architecture of Replication Fork Helicase Allows for Helicase Disassembly during Replication Stress

The helicase in bacteria is composed of the bacterial homohexamer DnaB. DnaB is highly processive and rapid in vitro without the addition of accessory factors (50, 51). In contrast, the eukaryotic replication fork helicase requires the heterohexameric Mcm2-7 complex functioning in coordination with GINS and Cdc45 for processive and rapid unwinding (15, 17). We show here that Cdc45 may also dissociate from the Mcm2-7 complex when the polymerase stalls to stop uncoupled helicase movement. Recent work demonstrates that GINS dissociates from Mcm2-7 upon replisome encounter with a DNA break (52). Thus, the modular architecture of the eukaryotic helicase may allow for complex disassembly and helicase stalling during episodes of replication stress.

It was shown recently that the viral hexameric replicative helicase Large T antigen is able to bypass protein cross-linked to the translocating DNA strand by transient opening of the helicase ring (53). It is not clear whether this mechanism of helicase opening to bypass a protein block is applicable to the cellular replicative helicase because the CMG complex forms a closed ring around single-stranded DNA (18). However, our observation that Cdc45 may disassociate from Mcm2-7 in response to long ssDNA suggests that a mechanism may in fact exist for the CMG complex to transiently open under certain circumstances. It still remains to be determined whether the CMG complex can bypass a protein block on the translocating strand during cellular replication.

Uncoupling of Helicase and Polymerase Activities May Lead to Genome Instability in Higher Eukaryotes

A hallmark of cancer cells is genome instability, and defective DNA replication or repair is associated with genome instability and cancer (54). An inappropriate response to DNA damage is also strongly associated with genome instability and cancer (55). The DNA damage response is a well characterized signaling cascade that is conserved from yeast to humans (56). Mutations in components of the DNA damage response can also lead to genome instability and cancer (56). Furthermore, one arm of the DNA damage response responds specifically to RPA-coated ssDNA, caused by uncoupling of the helicase with the polymerase (53). Because the DNA damage response is highly conserved and the accumulation of excess ssDNA at a replication fork can cause genome instability, it is logical that a system for coupling helicase and polymerase stalling will exist in most eukaryotes to preserve genomic integrity. Thus, it is very likely that the function that we uncovered in budding yeast that couples polymerase stalling with helicase stalling is likely active in higher eukaryotes as well and may be one of the cellular mechanisms that preserves genomic integrity and prevents cells from becoming cancerous.