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Mutations in the human ChlR1 gene are associated with a unique genetic disorder known as Warsaw breakage syndrome characterized by cellular defects in sister chromatid cohesion and hypersensitivity to agents that induce replication stress. A role of ChlR1 helicase in sister chromatid cohesion was first evidenced by studies of the yeast homolog Chl1p; however, its cellular functions in DNA metabolism are not well understood. We carefully examined the DNA substrate specificity of purified recombinant human ChlR1 protein and the biochemical effect of a patient-derived mutation, a deletion of a single lysine (K897del) in the extreme C terminus of ChlR1. The K897del clinical mutation abrogated ChlR1 helicase activity on forked duplex or D-loop DNA substrates by perturbing its DNA binding and DNA-dependent ATPase activity. Wild-type ChlR1 required a minimal 5′ single-stranded DNA tail of 15 nucleotides to efficiently unwind a simple duplex DNA substrate. The additional presence of a 3′ single-stranded DNA tail as short as five nucleotides dramatically increased ChlR1 helicase activity, demonstrating the preference of the enzyme for forked duplex structures. ChlR1 unwound G-quadruplex (G4) DNA with a strong preference for a two-stranded antiparallel G4 (G2′) substrate and was only marginally active on a four-stranded parallel G4 structure. The marked difference in ChlR1 helicase activity on the G4 substrates, reflected by increased binding to the G2′ substrate, distinguishes ChlR1 from the sequence-related FANCJ helicase mutated in Fanconi anemia. The biochemical results are discussed in light of the known cellular defects associated with ChlR1 deficiency.
Superfamily 2 DNA helicases that contain an iron-sulfur (Fe-S) motif within the conserved helicase core domain have attracted considerable interest because they are implicated in the maintenance of genomic stability and genetically linked to human diseases (1). Among this group of Fe-S domain proteins, the XPD helicase is a component of the transcription factor IIH complex and required for nucleotide excision repair and transcription. XPD mutations are genetically linked to the UV skin cancer disorder xeroderma pigmentosum as well as two other genetically distinct diseases known as trichothiodystrophy and Cockayne syndrome characterized by neurological and developmental impairments (2). Mutations in the FANCJ helicase gene result in the autosomal recessive disorder Fanconi anemia (3–5). FANCJ mutant cells, like other Fanconi anemia complementation groups, exhibit hypersensitivity to DNA interstrand cross-linking agents. Other members of the Fe-S domain helicase family include mouse RTEL and the nematode FANCJ homolog DOG-1, which have been implicated in telomere maintenance (6) and stability of sequences flanking G-tracts (7), respectively. Although a biological function of the Superfamily IB helicase PIF1 in mammalian cells has not yet been determined, yeast Pif1 is involved in the maintenance of nuclear and mitochondrial genome stability (8). Pif1 was found to prevent genomic instability of a G-quadruplex (G4)3 forming a human minisatellite sequence inserted into the Saccharomyces cerevisiae nuclear genome (9) and is involved in the DNA replication through G4 motifs (10) and coordination of checkpoint activation following telomere uncapping (11).
Interestingly, a newly discovered member of the Fe-S domain helicase family is the ChlR1 (also named DDX11) helicase genetically linked to Warsaw breakage syndrome (WABS) (12). The single patient with Warsaw breakage syndrome was observed to display severe microcephaly, pre- and postnatal growth retardation, and abnormal skin pigmentation. The WABS patient displayed a compound heterozygous K897del mutation in trans with a maternally inherited splice site mutation in intron 22 that leads to a premature stop codon in exon 23. This would lead to nonsense-mediated decay of the maternal allele and monoallelic expression of the K897del-containing allele. Thus, the only functional allele would be the K897del mutation that was inherited from the father and appears recessive (father was asymptomatic). Cells from the patient exhibit chromosomal instability characterized by sister cohesion defects, chromosomal breakage, and sensitivity to the DNA cross-linking agent mitomycin and topoisomerase inhibitor camptothecin (12). Based on the cellular phenotypes, it was suggested that WABS represented a unique disease with cellular features of both Fanconi anemia and the cohesinopathy Roberts syndrome.
A role of the ChlR1 helicase in sister chromatid cohesion was first evidenced by studies of the yeast homolog Chl1p. chl1 genetically interacts with two genes (CTF7/ECO1 and CTF18/CHL12) that function in sister chromatid cohesion (13). In addition to Chl1p, budding yeast require a specialized replication factor C (Ctf18/Dcc1/Ctf8) and DNA polymerase-associated Ctf4 to maintain sister chromatid cohesion in cells arrested for long periods during mitosis (14). chl1 was identified as a high copy suppressor of ctf18Δ spore lethality, and CHL1 was found to be essential for chromatid disjunction during meiosis II (14). chl1Δ is synthetically lethal with ctf18Δ (15). A dosage increase of chl1+ rescued the sensitivity of swi1Δ, a mutant of the “replication fork protection complex,” to agents that induce replicational stress, pointing toward a role of Chl1p in the maintenance of replication forks as a component of the S phase stress response. The genetic results and demonstration that Chl1p binds to components of the replication machinery (14, 16) implicate a role of yeast Chl1p to preserve genomic stability by promoting proper chromosome segregation and efficient sister chromatid cohesion during S phase.
Depletion of human (h) ChlR1 by RNA interference resulted in abnormal sister chromatid cohesion and a prometaphase delay leading to mitotic failure (17). Consistent with its role in sister chromatid cohesion, hChlR1 was found to reside in a complex with cohesion factors (17). Subsequent RNA interference studies with human cells showed that hChlR1 is required for proper chromosome cohesion at both the centromeres and along the chromosome arms and tight binding of cohesion complexes to chromatin (18). Other studies showed a role of hChlR1 in the maintenance of replicating papillomavirus viral episomes by loading the E2 protein onto mitotic chromosomes (19). To examine the potential importance of ChlR1 during development, a Ddx11 knock-out mouse was made (18). Loss of ChlR1 in the mouse resulted in embryonic lethality. The authors concluded that the aneuploidy apparent in Ddx11−/− embryos was a consequence of sister chromatid cohesion defects and placental malformation; however, the precise molecular pathways whereby ChlR1 maintains genomic stability are not well understood.
Compared with the wealth of genetic information, only limited biochemical studies of ChlR1 have been performed to date. Hirota and Lahti (20) purified recombinant hChlR1 from insect cells and showed that it is a DNA-dependent ATPase and that it unwinds partial duplex DNA substrates with low processivity and a preferred 5′ to 3′ directionality. More recently, Hurwitz and co-workers (21) purified a recombinant FLAG-tagged hChlR1 expressed in human 293 cells and confirmed its DNA-dependent ATPase activity and 5′ to 3′ helicase directionality. In the presence of replication protein A or the Ctf18-replication factor C complex, hChlR1 could unwind longer duplexes of 500 base pairs. hChlR1 was shown to interact with Ctf18-replication factor C, proliferating cell nuclear antigen, and FEN-1, and hChlR1 stimulated FEN-1 endonuclease activity on an equilibrating flap DNA structure, a model substrate intermediate that forms during lagging strand synthesis (21). It was proposed that the involvement of ChlR1 in lagging strand processing during cellular DNA replication may be important for sister chromatid cohesion. The Timeless-Tipin protein complex that is implicated in replication fork stabilization and sister chromatid cohesion also interacts with ChlR1 (22), supporting the notion that the DNA helicase collaborates with other factors to maintain a fork structure conducive to establishment of cohesion.
To better understand the molecular functions of hChlR1 that might be relevant to its roles in sister chromatid cohesion or resistance to agents that induce replication stress, we carefully examined its substrate specificity on a variety of DNA structures. These efforts revealed novel aspects of hChlR1 helicase function pertaining to its tail length requirements for unwinding simple duplex DNA and its ability to efficiently unwind a recombination intermediate D-loop structure as well as G4 DNA, the latter of which is proposed to affect normal cellular DNA metabolic processes such as replication. A WABS patient-derived ChlR1 mutation residing in the extreme C terminus of the protein negatively affected its biochemical functions including its ability to unwind simple duplex or G-quadruplex DNA, suggesting that the helicase function of ChlR1 is likely to be important in vivo. The biochemical findings will be discussed in light of the known cellular defects associated with ChlR1 deficiency.
Human ChlR1 cDNA in pcDNA3 (Invitrogen), kindly provided by Dr. Jill M. Lahti (Department of Genetics and Tumor Cell Biology, St. Jude Children's Research Hospital, Memphis, TN), was subjected to PCR amplification using primers ChlR1-F-HindIII and ChlR1-R-XhoI (supplemental Table S1). The PCR product, which includes an extra His6 sequence at the N terminus and 3× FLAG tag at the C terminus, was cloned into the HindIII and XhoI sites of pcDNA3. The K50R and K897del mutations were generated by a QuikChange site-directed mutagenesis kit (Stratagene) according to the manufacturer's instructions using the designed mutagenic primers shown in supplemental Table S1 and the plasmid pcDNA3–6XHis-ChlR1–3XFLAG. All plasmids were sequenced to verify that no undesired mutations were introduced during PCR and cloning.
Expression and purification of hChlR1 protein was based on the procedure of Hurwitz and co-workers (21) with modifications. Briefly, pcDNA3–6XHis-ChlR1–3XFLAG was transfected into 293T cells using Lipofectamine 2000 (Invitrogen) as recommended by the manufacturer. Six 10-cm plates of cells (~1 × 107 cells/plate) were transfected. Cells were harvested 48 h after transfection by trypsinization and centrifugation. Pelleted cells were washed with cold PBS and cold PBS with proteinase inhibitors (Roche Applied Science) sequentially, resuspended in 5 ml of hypotonic buffer (10 mm Tris·HCl, pH 7.4, 10 mm KCl, 1.5 mm MgCl2, 1 mm DTT, 0.5 mm phenylmethylsulfonyl fluoride (PMSF), proteinase inhibitors), and incubated on ice for 15 min. Cells were lysed by Dounce homogenization (20 strokes), and the mixture was centrifuged at 4 °C for 30 min at 2,400 × g. The supernatant (cytosol fraction) was collected and kept on ice; the nuclear pellet was resuspended in 5 ml of nuclear buffer (20 mm Tris·HCl, pH 7.4, 0.15 m NaCl, 10% glycerol, 1.5 mm MgCl2, 0.2 mm EDTA, 1 mm DTT, 0.5 mm PMSF, proteinase inhibitors) and incubated with rocking at 4 °C for 30 min. The cytosolic and nuclear fractions were combined together and centrifuged at 4 °C for 1 h at 43,500 × g. The supernatant (~10 ml) was incubated with 0.5 ml of FLAG M2-agarose resin (Sigma) in FLAG buffer (20 mm Tris·HCl, pH 7.4, 0.15 m NaCl, 10% glycerol, 0.05% Nonidet P-40, 1.5 mm MgCl2, 0.2 mm EDTA, 0.5 mm PMSF, proteinase inhibitors) at 4 °C for 2 h. The resin was then washed twice with high salt FLAG buffer (20 mm Tris·HCl, pH 7.4, 0.5 m NaCl, 10% glycerol, 0.05% Nonidet P-40, 1.5 mm MgCl2, 0.2 mm EDTA, 0.5 mm PMSF, proteinase inhibitors) followed by one washing with FLAG buffer. ChlR1 protein was eluted with 4 μg/ml 3× FLAG peptide (Sigma) in elution buffer (25 mm Tris·HCl, pH 7.5, 1 mm EDTA, 0.15 m NaCl, 1 mm DTT, 0.01% Nonidet P-40, proteinase inhibitors) for 1 h at 4 °C. The FLAG-tagged ChlR1 protein was then dialyzed at 4 °C for 2 h against elution buffer using a dialysis tube with a 50-kDa molecular mass cutoff (Tube-O-DIALYZERTM). Aliquots were frozen in liquid nitrogen and stored at −80 °C. Purified recombinant ChlR1 proteins were analyzed for purity by sodium dodecyl sulfate-polyacrylamide gel electrophoresis followed by Coomassie staining. Protein concentration was determined by the Bradford assay using BSA as a standard.
PAGE-purified oligonucleotides used for the preparation of DNA substrates were purchased from Lofstrand Labs and are listed in supplemental Table S2 except for oligonucleotides used for substrates with increasing 5′ or 3′ ssDNA tails that can be found in Ref. 23. The duplex substrates were 5′-32P-end-labeled and prepared as described previously (24). For the 5′ flap and 3′ flap substrates, oligonucleotides complementary to the 5′ ssDNA arm and 3′ DNA arms, respectively, were annealed to the 19-bp forked duplex (supplemental Table S2). Synthetic Holliday junction (HJ) substrate with a 12-nucleotide (nt) homologous core, designated HJ(X12), was made by annealing four 50-mer oligonucleotides (X12-1, X12-2, X12-3, and X12-4) as described previously (25). D-loop substrates were prepared as described previously (26). G-quadruplex DNA substrates (OX-1-G2′ and TP-G4) were prepared as described previously (27).
Helicase assay reaction mixtures (20 μl) contained 25 mm Hepes-NaOH, pH 7.5, 25 mm potassium acetate, 1 mm magnesium acetate, 1 mm DTT, 100 μg/ml bovine serum albumin, 1 mm ATP, 10 fmol of the specified duplex or G4 DNA substrate (0.5 nm DNA substrate concentration), and the indicated concentrations of the specified hChlR1 protein. Helicase reactions were initiated by the addition of hChlR1 and then incubated at 37 °C for 30 min unless otherwise indicated. Reactions were quenched with the addition of 20 μl of 2× stop buffer (17.5 mm EDTA, 0.3% SDS, 12.5% glycerol, 0.02% bromphenol blue, 0.02% xylene cyanol). For standard duplex DNA substrates, a 10-fold excess of unlabeled oligonucleotide with the same sequence as the labeled strand was included in the quench to prevent reannealing. Unlabeled oligonucleotide was omitted in the quench for helicase reactions with the HJ(X12) and D-loop substrates. The products of the helicase reactions for duplex DNA substrates and D-loop substrate were resolved on non-denaturing 12% (19:1 acrylamide:bisacrylamide) polyacrylamide gels. Products of G4 unwinding reactions were resolved on 8% (19:1 acrylamide:bisacrylamide) polyacrylamide gels with 10 mm KCl in the gel and the running buffer. Radiolabeled DNA species in polyacrylamide gels were visualized using a PhosphorImager and quantitated using ImageQuant software (GE Healthcare). The percentage of helicase substrate unwound was calculated by using the following formula: Percent unwinding = 100 × (P/(S + P)) where P is the product and S is the substrate. The values of P and S were corrected after subtracting background values in the no-enzyme and heat-denatured substrate controls, respectively.
Protein/DNA binding mixtures (20 μl) contained the indicated concentrations of ChlR1 and a 0.5 nm concentration of the specified 32P-end-labeled DNA substrate in the same reaction buffer as that used for helicase assays (see above) containing 1 mm ATPγS or no nucleotide. The binding mixtures were incubated on ice for 30 min after the addition of hChlR1. After incubation, 3 μl of loading dye (74% glycerol, 0.01% xylene cyanol, 0.01% bromphenol blue) was added to each mixture, and samples were resolved on native 5% (19:1 acrylamide:bisacrylamide) polyacrylamide gels at 200 V for 2 h at 4 °C. For G4 DNA binding experiments, 5% polyacrylamide gels were used, and running buffer contained 10 mm KCl. The radiolabeled species were visualized using a PhosphorImager and analyzed with ImageQuant software. The dissociation constant (Kd) was calculated by Scatchard analysis using Prism (GraphPad Software).
ATP hydrolysis was measured using [γ-32P]ATP (PerkinElmer Life Sciences) and analysis by thin-layer chromatography (TLC) on polyethyleneimine-cellulose plates (Mallinckrodt Baker). The standard reaction mixture (20-μl total volume) contained 25 mm Hepes-NaOH, pH 7.5, 25 mm potassium acetate, 1 mm magnesium acetate, 1 mm DTT, 100 μg/ml bovine serum albumin, 250 μm [γ-32P]ATP, and 60 nm hChlR1 protein and was incubated at 37 °C. Reactions were quenched with 50 mm EDTA (final concentration). The reaction mixture was spotted onto a polyethyleneimine-cellulose TLC plate and resolved by using 0.5 m LiCl, 1 m formic acid as the carrier solvent. The TLC plate was exposed to a phosphorimaging cassette for 1 h, visualized using a PhosphorImager, and analyzed with ImageQuant software.
For experiments to determine Km(ATP), M13mp18 ssDNA was 2.1 nm, the concentration of ATP ranged from 31 to 8000 μm, and the reaction was incubated for 30 min. For determination of kcat, the concentration of ATP was 8.5 mm. Five-microliter aliquots were removed and quenched with 5 μl of 0.1 m EDTA at 0, 7.5, 15, 30, and 45 min, respectively. For determination of Keff, the M13mp18 ssDNA concentration ranged from 0 to 10.6 nm, and the reaction was incubated for 30 min. The kinetic parameters were calculated by Enzyme Kinetics 1.3 (SigmaPlot, Systat Software Inc.) using the Michaelis-Menten equation. All experiments were repeated at least three times with standard deviation (S.D.) indicated by error bars.
The ATP binding assay (30 μl) was performed in the same reaction buffer as for the helicase or ATPase assay described above with 5μCi of [α-32P] ATP (3000 Ci/mmol; PerkinElmer Life Sciences). Assays were initiated by adding hChlR1 protein to a final concentration of 230 nm followed by incubating for 30 min at the indicated temperature. Reactions were then applied to Bio-Spin P30 Tris chromatography columns (Bio-Rad), which had been pre-equilibrated in a reaction buffer. One-drop (~45 μl) fractions were collected as flow-through under gravity from the column that was eluted with Tris-EDTA. The specific radioactivity of each fraction was determined by a liquid scintillation counter (MicroBeta TriLux, PerkinElmer Life Sciences). The first peak (three to four drops) was considered as protein-bound ATP, and the second peak was considered as unbound ATP.
Streptavidin displacement reaction mixtures (20 μl) contained 25 mm Hepes-NaOH, pH 7.5, 25 mm potassium acetate, 1 mm magnesium acetate, 1 mm DTT, 100 μg/ml bovine serum albumin, 1 mm ATP, 10 fmol of the specified DNA substrate (0.5 nm), and the indicated concentrations of hChlR1 protein. For streptavidin displacement reactions, 10 fmol of DNA substrate was preincubated with 100 nm streptavidin (Sigma) for 10 min at 37 °C. Streptavidin bound to the DNA substrate was detected by a slower migrating gel-shifted species on nondenaturing 12% polyacrylamide gel. Reactions were initiated by adding hChlR1 prior to the addition of biotin (1 μm) and incubated at 30 °C for 15 min (unless otherwise noted) followed by a quench with the addition of 10 μl of stop buffer (50 mm EDTA, 40% glycerol, 0.9% SDS, 0.05% bromphenol blue, 0.05% xylene cyanol). Products were resolved on nondenaturing 12% (19:1 acrylamide/bisacrylamide) polyacrylamide gels and quantitated. A PhosphorImager was used for detection, and ImageQuant software was used for quantitation of the reaction products. The percentage of displacement was calculated. The data represent the mean of at least three independent experiments with S.D. values shown by error bars.
A chlr1 mutation resulting in a single amino acid deletion of lysine 897 in the hChlR1 protein, designated K897del (Fig. 1A), was reported to be genetically linked to Warsaw breakage syndrome (12). To understand the molecular defects of the protein encoded by the mutant allele, we purified the recombinant K897del and wild-type hChlR1 proteins that were expressed in human 293T cells. We also purified an engineered Walker A box (motif I) K50R hChlR1 protein in which the invariant lysine residue was replaced with arginine (21). The recombinant wild-type and mutant hChlR1 proteins were purified to near homogeneity as judged by their appearance as single bands after electrophoresis on Coomassie-stained SDS-polyacrylamide gels (Fig. 1B). hChlR1-K897del recombinant protein that was expressed and purified from human cells consistently showed a reduced yield compared with hChlR1-WT that may be consistent with the low detection of endogenous K897del protein in cells from the affected patient (12).
We first examined DNA unwinding activity catalyzed by hChlR1-WT and hChlR1-K897del mutant. Using a forked 19-bp duplex DNA substrate (supplemental Table S2), hChlR1-WT unwound the substrate in the presence of ATP and failed to unwind the substrate in the absence of ATP (supplemental Fig. S1), suggesting that hChlR1 unwinding is dependent on hydrolysis of nucleoside triphosphate. Using the same duplex substrates, neither hChlR1-K897del nor hChlR1-K50R could unwind this type of substrate under conditions in which hChlR1-WT unwound the forked duplex DNA molecules to near completion at a concentration of 2.4 nm (Fig. 1C). The ability of hChlR1-WT, but not hChlR1-K50R, to unwind the forked duplex indicated that the helicase activity was intrinsic to hChlR1 and not due to a contaminant in the helicase preparation. We also increased the incubation time (60 min) or protein concentration of hChlR1-K897del up to 38.4 nm but failed to detect helicase activity on forked duplex (data not shown). hChlR1-WT unwound the forked duplex DNA substrate more efficiently than the sequence-related FANCJ helicase (supplemental Fig. S2).
The failure of hChlR1-K897del to unwind the DNA substrates might reflect an impairment of DNA binding activity. To address this issue, we performed EMSAs with the mutant and wild-type proteins in the absence of ATP using the same radiolabeled forked duplex DNA used for the helicase assays. Results demonstrated that hChlR1-WT bound the DNA molecule in a protein concentration-dependent manner; however, hChlR1-K897del was unable to bind the forked duplex (Fig. 1D). The presence of ATPγS or ADP did not enhance binding of either the wild-type or mutant protein to the forked duplex (data not shown). Thus, the WABS patient-derived hChlR1-K897del mutation interferes with the ability of ChlR1 to efficiently bind the forked duplex DNA substrate.
We next examined the DNA-dependent ATPase activity of hChlR1-K897del compared with hChlR1-WT (Table 1). As a control, we included purified recombinant hChlR1-K50R, which was reported previously to have significantly reduced ATPase activity (21). Using covalently closed M13 single strand DNA as the effector molecule, we determined a Km value of ATP hydrolysis for hChlR1-WT to be 0.59 mm (Table 1). Because of the very low ATPase activity of hChlR1-K897del and hChlR1-K50R, Km values could not be determined. Using an ATP concentration (8.5 mm) that was ~10-fold greater than the Km for hChlR1-WT, ATPase assays with hChlR1-WT and hChlR1-K897del yielded kcat values of 438 and 5.0 min−1, respectively For comparison, the kcat value for hChlR1-K50R was 4.7 min−1, a value very similar to that of hChlR1-K897del. The Keff value for ATP hydrolysis by hChlR1-WT using M13 ssDNA was 0.087 nm (Table 1); however, the Keff value for hChlR1-K897del could not be determined because its ATPase activity even in the presence of M13 ssDNA was very low and did not achieve a plateau. The poor ATPase activity of hChlR1-K897del observed even in the presence of high concentrations of M13 ssDNA is consistent with its poor ability to bind DNA (Fig. 1D). These results suggest that the K897del mutation seriously compromised the ability of hChlR1 to hydrolyze ATP in a DNA-dependent manner.
To compare ATP binding by hChlR1-K897del with hChlR1-WT, an equal amount of hChlR1-K897del or hChlR1-WT was incubated with [α-32P]ATP under identical conditions, and binding mixtures were analyzed by gel filtration chromatography. Scintillation counting of the eluted fractions demonstrated that hChlR1-K897del bound ATP modestly better than hChlR1-WT, whereas very little [α-32P]ATP was retained with BSA, which served as a nonspecific control (Fig. 2). To address the stability of hChlR1-K897del protein, we examined its thermostability for ATP binding and found that the mutant protein was more sensitive to temperature increase than ChlR1-WT (supplemental Fig. S3), suggesting that it may be less stable. Although hChlR1-K50R, like hChlR1-K897del, was defective in ATP hydrolysis, the protein was also able to bind ATP. From these results, we conclude that hChlR1-K897del bound ATP similarly to hChlR1-WT but did not efficiently hydrolyze the nucleotide in the presence of DNA effector.
Although hChlR1 was shown previously to preferentially unwind partial duplex DNA substrates with a 5′ to 3′ directionality (20, 21), the importance of single strand length for loading and unwinding duplex DNA was not examined. To determine the minimal loading dock requirements by hChlR1-WT to catalyze efficient DNA unwinding, we performed hChlR1 helicase assays on duplex (19-bp) DNA substrates flanked by a 5′ ssDNA tail of increasing lengths (Fig. 3A). Very little to no unwinding of the duplex by hChlR1-WT (2.4 nm) was detected on substrates lacking a 5′ tail altogether (blunt duplex) or one flanked by a 5′ tail of 5 nt. hChlR1-WT unwound a very small percentage (<5%) of the duplex substrate flanked by a 10-nt 5′ tail. When the 5′ ssDNA tail was increased to 15 nt, over 90% of the DNA substrate was unwound. Nearly all of the substrate was unwound when the 5′ tail length was increased to 26 or 40 nt. These results suggest that a minimal 5′ tail length of ~15 nt is required for hChlR1-WT to efficiently unwind the adjacent duplex.
We next addressed the importance of the length of the noncomplementary 3′ ssDNA tail of a forked DNA duplex that was flanked by a fixed 5′ ssDNA tail of optimal length, 15 nt (Fig. 3B). In these experiments, we used 480-fold less hChlR1-WT protein in the helicase reactions with forked duplex substrates compared with the series of substrates with only a 5′ ssDNA tail. Increasing the length of 3′ ssDNA tail from 0 to 5 nt significantly increased unwinding of the substrate. An optimum was reached for the forked duplex helicase substrate with a 10-nt 3′ tail. Approximately 70% of the duplex substrate flanked by a 5′ tail of 15 nt and 3′ tail of 10 nt was unwound by hChlR1-WT. A further increase of the 3′ tail to 15 or 40 nt resulted in reduced unwinding of the forked duplex. From these results, we conclude that for a simple 5′ tailed duplex substrate hChlR1-WT requires a 5′ tail of ~15 nt to catalyze appreciable unwinding and that the additional presence of a 3′ ssDNA tail of ~5 nt to create a forked duplex substrate dramatically improves ChlR1 unwinding efficiency.
Given the hypersensitivity of WABS cells to agents that interfere with DNA replication (mitomycin C and camptothecin) (12), ChlR1 helicase may function at the interface of replication-coupled DNA repair and sister chromatid cohesion. Because homologous recombination of replicated sister chromatids occurs during S phase, we wanted to determine whether ChlR1 could unwind key DNA intermediates of replication and homologous recombination. hChlR1-WT was tested for unwinding of forked DNA structures with a 5′ flap, 3′ flap, or duplex in both arms. hChlR1-WT was able to efficiently unwind the 5′ flap substrate (Fig. 4A) in a protein concentration-dependent manner that was similar to its activity on a forked duplex with single-stranded 3′ and 5′ arms (supplemental Fig. S2). The downstream 5′ flap oligonucleotide was displaced at the lowest protein concentrations, creating a 5′ tailed duplex. Higher concentrations of enzyme resulted in the displacement of the upstream oligonucleotide with the concomitant disappearance of the 5′ tailed duplex. Quantitative analysis of the products from helicase reactions with the forked DNA structures demonstrated that hChlR1-WT was poorly active upon the 3′ flap and synthetic replication fork structures compared with the 5′ flap substrate (Fig. 4B).
We next tested a four-stranded HJ structure with a branch-migrating 12-nt core (supplemental Table S2). hChlR1-WT poorly unwound the HJ substrate under conditions that the helicase efficiently unwound the forked duplex substrate consisting of two partially complementary arms of the HJ structure (Fig. 5A). No significant unwinding of the HJ was detected at the highest concentration of hChlR1-WT tested, 19.2 nm (supplemental Fig. S4A). A measurement of DNA binding by EMSA revealed that hChlR1-WT failed to bind the HJ (Fig. 5B) under conditions that the protein effectively bound forked duplex molecules (Fig. 1D). hChlR1-WT also bound poorly to a blunt duplex DNA substrate compared with ssDNA (supplemental Fig. S4B), suggesting that the helicase does not interact favorably with core or duplex arms of the synthetic HJ structure.
Because the hChlR1-WT helicase reactions contained equimolar concentrations of ATP and Mg2+ (1 mm), some free Mg2+ ion not chelated to ATP may have bound to the HJ substrate, creating a more stacked structure inaccessible for hChlR1-WT unwinding, as reported previously for Drosophila melanogaster RecQ5β (28), Escherichia coli RecG (29), and S. cerevisiae MER3 (30). To address this possibility, we tested hChlR1-WT for unwinding of the HJ substrate at lower Mg2+ concentrations but did not detect any helicase activity (supplemental Fig. S4C). RECQ1 helicase efficiently unwound this HJ substrate in the hChlR1-WT reaction conditions, indicating that the substrate could be unwound by a helicase (supplemental Fig. S4D). The inability of hChlR1-WT to unwind the HJ structure suggests that ChlR1 is unlikely to perform HJ branch migration, a function performed by a number of RecQ helicases (31).
We next examined the ability of hChlR1-WT to unwind a three-stranded D-loop DNA substrate (supplemental Table S2) that serves as a model for an early intermediate of homologous recombination formed by invasion of single-stranded DNA into a DNA duplex to anneal with its complementary strand. hChlR1-WT was able to efficiently act upon the D-loop substrate by unwinding the third invading strand in a protein concentration-dependent manner (Fig. 5C). However, hChlR1-K897del failed to release the invading strand from the D-loop structure (Fig. 5C) consistent with its lack of helicase activity on a forked duplex (Fig. 1C).
The interaction of ChlR1 with proteins of the replication fork machinery that are important for the establishment of sister chromatid cohesion raised the question whether ChlR1 might be able to use its motor ATPase function for displacing proteins bound to DNA. To address this issue, we tested whether hChlR1-WT could disrupt the high affinity interaction of streptavidin bound to a biotinylated ssDNA molecule. Increasing concentrations of hChlR1-WT were incubated in the presence of ATP with a radiolabeled oligonucleotide that had streptavidin bound to the biotin conjugated either 28 or 52 nt from the 5′-end of the 66-mer oligonucleotide (supplemental Table S2). As shown in Fig. 6A, hChlR1-WT displaced streptavidin bound to the DNA substrate with biotin positioned 52 nt away from the 5′-end in a protein concentration-dependent manner. Approximately 60% displacement was achieved by 20 nm hChlR1-WT during the 30-min reaction incubation (Fig. 6B). When hChlR1-WT was tested for streptavidin displacement from a similar substrate except that the biotin was positioned 28 nt from the 5′-end, significantly lesser percentages of displacement were observed at all helicase concentrations compared with the substrate with biotin located 52 nt from the 5′-end (Fig. 6, A and B). Quantitative analyses demonstrated that ~3-fold greater displacement was observed at a hChlR1-WT concentration of 9.6 nm protein. These results demonstrate that hChlR1-WT more efficiently displaces streptavidin bound to biotin when it is bound farther away (by 24 nt) from the 5′-end. Unlike hChlR1-WT, hChlR1-K897del failed to displace streptavidin from biotinylated oligonucleotide (supplemental Fig. S5A). Also hChlR1-K50R failed to strip streptavidin from biotinylated oligonucleotide (supplemental Fig. S5B), indicating the hChlR1-WT motor activity is ATP-dependent and intrinsic to the purified recombinant protein. Thus, ChlR1, like the sequence-related FANCJ helicase (32), can disrupt the high affinity interaction of streptavidin bound to a biotinylated oligonucleotide.
Although ChlR1 plays a role in the establishment of sister chromatid cohesion during replication, how the helicase functions in this capacity is not well understood. Alternate DNA structures such as G4 DNA that may impede the replication fork are proposed to form in G-rich regions that are quite abundant in the human genome (for review, see Ref. 33). It is plausible that ChlR1, like the sequence-related FANCJ (34, 35), may unwind G4 DNA to help preserve timely progression of the replication fork, which is likely to be important in the process of sister chromatid cohesion.
To determine whether hChlR1-WT was able to unwind G4 DNA, we began by testing the recombinant protein for its activity on a well characterized four-stranded parallel G4 substrate (TP-G4) derived from a mouse immunoglobulin sequence that bore 5′ single-stranded DNA tails of 21 nt (27) (supplemental Table S2). G4 DNA formation of the TP-G4 substrate was confirmed previously by methylation protection analysis (27). Using a range of hChlR1-WT protein concentrations up to 2.4 nm, there was very little unwinding (<2%) of the TP-G4 substrate (Fig. 7A). In contrast, hChlR1-WT unwound the forked duplex substrate to near completion at a protein concentration of 0.15 nm (Fig. 7A). Even at a concentration of 19.2 nm hChlR1-WT, only 5% of the TP-G4 substrate was unwound (Fig. 7, B and C). In contrast to the poor activity of hChlR1-WT on TP-G4, FANCJ (4.8 nm) effectively unwound TP-G4 to near completion (35), suggesting a difference in substrate preference between the two Fe-S domain helicases in their relative ability to unwind G4 DNA.
We next tested hChlR1-WT to unwind an alternative form of G-G-paired DNA, designated G2′, which can occur when hairpin dimers of two antiparallel strands form Hoogsteen hydrogen bonds between guanine residues. This particular G2′ structure formed from double T4G4 repeats derived from the Oxytricha telomeric sequence has been well characterized by both NMR studies (36–38) and x-ray crystallography (39, 40). As shown in Fig. 7B, hChlR1-WT efficiently unwound the G2′ quadruplex (OX-1-G2′), which bore a 5′ single-stranded DNA tail of 20 nt (supplemental Table S2), in a protein concentration-dependent manner as evidenced by the product of the helicase reaction that co-migrated with the unannealed control radiolabeled oligonucleotide. The percentage of OX-1-G2′ substrate unwound at a given hChlR1-WT concentration was significantly greater compared with the TP-G4 substrate (Fig. 7C). The difference in hChlR1-WT helicase activity on the two G4 substrates was striking. For example, at an hChlR1-WT concentration of 4.8 nm, the helicase unwound 75% of the OX-1-G2 substrate compared with less than 5% of the TP-G4 substrate.
To determine whether hChlR1-WT unwinding of the G4 substrates was ATP-dependent, hChlR1-WT was incubated with OX-1-G2′ or TP-G4 in the presence of ADP or the poorly hydrolyzable ATPγS (Fig. 7D). Neither nucleotide substituted for ATP in the G4 unwinding reaction. To further determine whether G4 unwinding was dependent on ATP hydrolysis intrinsic to hChlR1-WT, we tested the engineered Walker A box mutant hChlR1-K50R for unwinding the OX-1-G2′ substrate. The hChlR1-K50R mutant protein failed to unwind the G2′ substrate (Fig. 7E), confirming that G4 unwinding by hChlR1-WT was dependent on intrinsic ATP hydrolysis. Similarly, hChlR1-K897del failed to unwind the OX-1-G2′ substrate at all concentrations tested (Fig. 7E).
The preference of hChlR1-WT to unwind the OX-1-G2′ substrate raised the possibility that hChlR1-WT might bind this G4 DNA substrate more stably compared with the less efficiently unwound TP-G4. To address this, we performed EMSAs with hChlR1-WT and the corresponding radiolabeled DNA substrates used for the helicase assays. A shift of both the OX-1-G2′ and TP-G4 substrates to a more slowly migrating species was detected on a native 5% polyacrylamide gel in a hChlR1-WT concentration-dependent manner (Fig. 8, A and B). At higher hChlR1-WT protein concentrations, a second more slowly migrating species was detected, suggesting multiple ChlR1 protein molecules binding less specifically to the G-quadruplex substrates. Careful analysis of the DNA binding isotherms revealed that hChlR1-WT bound the OX-1-G2′ substrate better than TP-G4 throughout the protein titration (Fig. 8B). A maximal difference was observed at 2.4 nm hChlR1-WT in which 80% of the OX-1-G2′ substrate was bound compared with only 20% of the TP-G4 substrate. The EMSA binding data were analyzed according to Scatchard theory, and apparent dissociation constants (Kd) were determined. The Kd for TP-G4 was 7.50 ± 1.8 nm, and that for OX-1-G2′ was 2.13 ± 0.78 nm, suggesting that hChlR1-WT bound the OX-1-G2′ substrate with greater affinity compared with the TP-G4 substrate.
To independently assess the apparent substrate preference of hChlR1-WT for OX-1-G2′ over TP-G4, we performed sequestration assays in which the helicase was preincubated with radiolabeled TP-G4 or OX-1-G2′ in the absence of ATP prior to simultaneous addition of ATP and a dT45 oligonucleotide and further incubation at 37 °C (Fig. 9A). If hChlR1-WT remained preferentially bound to the G2′ DNA molecule, then a greater concentration of dT45 would be required to inhibit helicase activity on the G2′ DNA substrate compared with TP-G4. Indeed this was observed to be the case because a greater concentration of dT45 was required for inhibition of hChlR1 helicase activity on OX-1-G2′ versus TP-G4 (Fig. 9B). These experimental results are consistent with the results from DNA binding assays that suggest that ChlR1-WT was more stably bound to the two-stranded antiparallel G2′ structure compared with the four-stranded parallel TP-G4 DNA molecule.
Although there is evidence that hChlR1 and its homologs in yeast and mice are required for maintenance of genomic stability, the biochemical functions of ChlR1 have not been extensively studied; therefore, we evaluated its DNA substrate specificity and catalytic activities. From these studies, it was determined that hChlR1-WT requires a minimal 5′ ssDNA tail of 15 nt to efficiently unwind a simple 5′ tailed duplex substrate. The additional presence of a 3′ ssDNA tail as short as 5 nt significantly improved the ability of ChlR1 to unwind the DNA substrate, indicating that the helicase prefers fork structures for DNA unwinding. The requirement for a 5′ ssDNA tail for hChlR1-WT to unwind a standard duplex DNA substrate was reflected by its activity on a three-stranded D-loop substrate, a key intermediate that forms during an early step of homologous recombination. In contrast to its helicase activity on a 5′ tailed D-loop substrate, hChlR1-WT failed to unwind an HJ structure. Thus, it is unlikely that ChlR1 or the sequence-related FANCJ helicase (23) performs branch fork migration of HJ-like structures during homologous recombination or replication restart as proposed for the RecQ helicases (31). hChlR1-WT efficiently unwound the 5′ flap structure, a key intermediate of lagging strand processing, which may play a role in its functional interaction with FEN-1 (21).
Consistent with its improved ability to unwind DNA substrates flanked by a minimal length of ssDNA, gel shift analyses demonstrated that hChlR1-WT bound ssDNA molecules and forked duplex molecules quite well under conditions that it bound very poorly to blunt duplex DNA. The patient-derived hChlR1-K897del deletion of 3 base pairs results in a single amino deletion of a highly conserved lysine just 10 amino acids from the C terminus (12). The endogenous hChlR1-K897del protein was poorly detected by immunoblot analysis of lysates from fibroblasts or lymphoblasts of the affected individual (12). Biochemical analysis of the purified recombinant hChlR1-K897del protein revealed a dramatic inhibitory effect on DNA binding and DNA-dependent ATPase activity, suggesting an important role of the extreme C terminus of the protein for its stable interaction with nucleic acid. Interestingly, a single amino acid substitution in a conserved region of the helicase core domain of FANCJ protein impaired its ability to bind DNA and dimerize in solution; moreover, the dimeric form of FANCJ bound forked duplex DNA more tightly than the monomeric form and also had a higher specific activity for ATP hydrolysis and DNA unwinding.4 It will be of interest to determine the assembly state of hChlR1 and the potential importance of hChlR1 subunit interactions for its DNA substrate preference and catalytic activity.
The DNA substrate specificity of hChlR1-WT on many accounts closely mirrors that of the sequence-related FANCJ (23) and DinG (41) helicases. Spies and co-workers (42) demonstrated that the Fe-S cluster in an archaeal XPD protein has a structural role in the recognition of junctions between single-stranded and double-stranded DNA, which may provide insight into the DNA substrate preference of Fe-S domain helicases. Clinically relevant ChlR1 mutations may affect the integrity of the conserved Fe-S domain, its ability to interact with DNA junctions, or its propensity to form a proposed ssDNA channel with the Arch domain to perform ATPase-driven DNA unwinding.
In addition to its ability to unwind duplex DNA substrates harboring a 5′ ssDNA tail, we tested whether hChlR1-WT could use its ATPase motor activity to disrupt a protein-DNA complex modeled by the high affinity interaction of streptavidin bound to a biotinylated single-stranded DNA oligonucleotide. hChlR1-WT was able to displace streptavidin from DNA in a helicase protein concentration- and ATP-dependent manner. The efficiency of streptavidin displacement by hChlR1-WT was dependent on the length of the 5′ ssDNA tract adjacent to the bound streptavidin, suggesting that the longer 5′ ssDNA loading region (52 versus 28 nt) may provide more suitable loading of multiple hChlR1-WT molecules to increase force production for protein displacement and/or a more optimal assembly ChlR1 state. The fact that the streptavidin-biotin complex is a very tight interaction (Kd ~ 10−15 m), which cannot be disrupted by a number of DNA helicases (e.g. WRN, BLM, and RECQ1) (32), suggests that that the ability of ChlR1 to disrupt protein-DNA complexes is biochemically unusual and may be important for its cellular function to preserve fork structure necessary for sister chromatid cohesion and/or participate in DNA repair or other processes. For example, ChlR1 was shown to be required for papillomavirus E2 protein loading onto mitotic chromosomes during DNA replication for the viral genome to be maintained and segregated (19, 43). The functions of the ChlR1 motor ATPase in this process are not well understood.
From our investigation, hChlR1-WT was found to harness the energy from ATP hydrolysis to unwind a G4 DNA structure. G4 structures that form in G-rich regions of the genome, which are believed to be abundant, are proposed to influence a number of DNA transactions including replication fork progression, gene expression, telomere maintenance and capping, and genetic recombination in G-rich elements such as ribosomal DNA repeats and heavy chain immunoglobulin switch regions (33). We demonstrated that hChlR1-WT efficiently unwound a G4 substrate (OX-1) with an alternative form of G-G-paired DNA (designated G2′) that occurs when hairpin dimers of two antiparallel strands form Hoogsteen hydrogen bonds. Unwinding of the G2′ substrate by hChlR1-WT to create single strands was observed at equimolar concentrations of ChlR1 monomer and DNA substrate, dependent on the presence of ATP in the reaction and a consequence of the intrinsic ATP hydrolysis catalyzed by the recombinant hChlR1-WT protein. In contrast to the efficient unwinding of the G2′ substrate by hChlR1-WT, the helicase was poorly active on a well characterized four-stranded parallel G4 DNA substrate (TP-G4). hChlR1-WT displayed a stronger binding affinity for OX-1 compared with TP-G4, suggesting that the increased helicase activity of hChlR1-WT on the G2′ structure was at least partly attributable to its preferential binding to the G2′ DNA substrate. Both TP-G4 and OX-1 substrates have four single strand tails of similar length (20–21 nt), suggesting that hChlR1-WT recognizes an element of the two-stranded antiparallel G-quadruplex structure itself more favorably than the four-stranded parallel G4. The preferred interaction of hChlR1-WT with the G2′ structure over the four-stranded G4 molecule was borne out in the sequestration experiments as well. The preference of hChlR1-WT to unwind the G2′ substrate is distinct from that of FANCJ, which was observed to unwind OX-1-G2′ and TP-G4 similarly throughout the FANCJ protein titration (35). In the future, it will be important to study the biological DNA substrates upon which ChlR1 acts and whether at least some of these form in G-rich sequences of the genome that may be prone to forming a particular class of G4 structures.
Understanding the molecular substrates of ChlR1 helicase may provide insight into the basis for the spontaneous premature centromere division of metaphase chromosomes characteristic of ChlR1-deficient cells. The sister chromatid cohesion defects may reflect problems during replication of regions that are difficult to replicate. ChlR1 would act to preserve genomic stability by resolving alternate DNA structures such as G4 that destabilize or impede the progression of the replication fork. It is conceivable that the relevant G4 substrates upon which ChlR1 acts are those in the lagging strand template similar to those structures proposed to be acted upon by Caenorhabditis elegans DOG-1 helicase (7). It is plausible that the lagging strand gets stalled and causes cohesion problems as a secondary event. This idea would be attractive because genetic and biochemical data already suggest that ChlR1 is involved in lagging strand DNA processing events (see the Introduction and Ref. 21). ChlR1 preference for a subset of G4 substrates may be pertinent to its putative activity on such structures that preferentially form in single-stranded DNA tracts between Okazaki fragments. The ability of ChlR1 to stimulate FEN-1 incision (21) and unwind certain G4 structures (this study) may enable lagging strand processing of G-rich sequences to occur more expeditiously. In addition to Okazaki fragment processing, FEN-1 is required for telomere stability by facilitating replication fork reinitiation through the G-rich lagging strand telomere (44). Evidence that ChlR1 interacts with FEN-1 may be relevant to a possible role of ChlR1 in telomere metabolism. It is plausible that ChlR1 may help cells deal with certain forms of DNA damage in G4 DNA at telomeres or other G-rich regions of the genome. Further biochemical and cellular studies are required for mechanistic insights into the functions of ChlR1 that are relevant to the maintenance of genomic stability and prevention of human disease.
We thank Dr. Jill M. Lahti (Department of Genetics and Tumor Cell Biology, St. Jude Children's Research Hospital, Memphis, TN) for kindly providing human ChlR1 cDNA.
*This work was supported, in whole or in part, by the National Institutes of Health Intramural Research Program of the NIA (to R. M. B.). This work was also supported by the Fanconi Anemia Research Fund (to R. M. B.).
This article contains supplemental Figs. S1–S5 and Tables S1 and S2.
4Y. Wu, J. A. Sommers, J. Loiland, H. Kitao, J. Kuper, C. Kisker, and R. M. Brosh, Jr., manuscript in preparation.
3The abbreviations used are: