Logo of jbcAbout JBCASBMBSubmissionsSubscriptionsContactJBCThis Article
J Biol Chem. Jun 24, 2011; 286(25): 22372–22383.
Published online Apr 29, 2011. doi:  10.1074/jbc.M111.233908
PMCID: PMC3121385

Formation of a Stable RuvA Protein Double Tetramer Is Required for Efficient Branch Migration in Vitro and for Replication Fork Reversal in Vivo*An external file that holds a picture, illustration, etc.
Object name is sbox.jpg


In bacteria, RuvABC is required for the resolution of Holliday junctions (HJ) made during homologous recombination. The RuvAB complex catalyzes HJ branch migration and replication fork reversal (RFR). During RFR, a stalled fork is reversed to form a HJ adjacent to a DNA double strand end, a reaction that requires RuvAB in certain Escherichia coli replication mutants. The exact structure of active RuvAB complexes remains elusive as it is still unknown whether one or two tetramers of RuvA support RuvB during branch migration and during RFR. We designed an E. coli RuvA mutant, RuvA2KaP, specifically impaired for RuvA tetramer-tetramer interactions. As expected, the mutant protein is impaired for complex II (two tetramers) formation on HJs, although the binding efficiency of complex I (a single tetramer) is as wild type. We show that although RuvA complex II formation is required for efficient HJ branch migration in vitro, RuvA2KaP is fully active for homologous recombination in vivo. RuvA2KaP is also deficient at forming complex II on synthetic replication forks, and the binding affinity of RuvA2KaP for forks is decreased compared with wild type. Accordingly, RuvA2KaP is inefficient at processing forks in vitro and in vivo. These data indicate that RuvA2KaP is a separation-of-function mutant, capable of homologous recombination but impaired for RFR. RuvA2KaP is defective for stimulation of RuvB activity and stability of HJ·RuvA·RuvB tripartite complexes. This work demonstrates that the need for RuvA tetramer-tetramer interactions for full RuvAB activity in vitro causes specifically an RFR defect in vivo.

Keywords: Bacteria, DNA Damage, DNA Enzymes, DNA Recombination, DNA Repair, DNA Replication


The RuvAB complex is a highly sophisticated molecular machine, which carries out branch migration of Holliday junctions during homologous recombination. RuvA binds specifically to four-armed Holliday junctions (HJ)4 and guides the assembly of two RuvB hexameric rings onto diametrically opposite arms of the HJ. RuvB, an AAA+ ATPase (1), is the motor that drives branch migration of the crossover point (13). After branch migration, a dimer of RuvC resolves the HJ by making two sequence-specific symmetrical cuts, producing either patched or spliced linear products (46). Genetic studies showed that RuvC cannot function in vivo in the absence of RuvAB (7, 8), and it has been proposed that an RuvABC complex, known as the resolvasome, allows RuvC to scan for cleavable sequences (3, 911). RuvA binds to HJs in vitro as one tetramer (complex I) or two tetramers that sandwich the junction (complex II) in a concentration-dependent manner; however, it is not clear whether the RuvAB complex contains one or two tetramers of RuvA in vivo (1221). A RuvAB branch migration complex made of two RuvA tetramers would prevent access of RuvC to the Holliday junction. Whether to form the resolvasome the RuvC dimer displaces one of the two RuvA tetramers present in the RuvAB complex, or whether the RuvC dimer simply binds opposite a single RuvA tetramer present in the complex is a currently unanswered question.

In addition to processing Holliday junctions in homologous recombination, RuvAB plays an important role upon DNA replication inactivation. In certain E. coli replication mutants, stalled replication forks undergo a process known as replication fork reversal (RFR) (22). As the stalled fork is reversed, the newly synthesized strands are unwound from the daughter duplexes and base pair to form a Holliday junction, known as a reversed fork. Branch migration of the reversed fork by RuvAB drives extrusion of an arm with a duplex end, which allows entry of RecBCD. RecBCD can either reset the fork by degrading the duplex or load RecA, which carries out invasion and strand exchange with the homologous duplex at the replication fork, to reset a new fork. Both pathways result in PriA-dependent replication restart (23). Intriguingly, RuvAB are actually required for replication fork reversal to occur in dnaEts, holDQ10am and rep E. coli mutants (24). Yet in vitro, RuvAB preferentially unwinds synthetic replication forks in a direction that is opposite the direction for fork reversal (25). RuvAB could reverse model replication forks in vitro if RuvB was only allowed to form one hexameric ring on the parental duplex of the fork (25). It has been speculated that in vivo the asymmetric binding of a single RuvA tetramer onto a three-armed fork may result in asymmetric loading of a single RuvB hexamer onto the parental duplex (24). Alternatively, cellular factors may force RuvA to load RuvB in this manner.

In an RuvA octamer, the two tetramers do not only interact with DNA but also with each other through four contacts involving domain II of each monomer. Specifically, six ionic interactions form between the α-helix 6 in domain II of each opposite monomer resulting in four points of contact between tetramers (supplemental Fig. 1) (18). The role of RuvA octamers for efficient branch migration has been investigated using RuvA mutants designed to disrupt the tetramer-tetramer interface and prevent complex II formation. A triple Escherichia coli RuvA mutant, RuvA3m, was unable to form complex II at RuvA concentrations of up to 2 μm and was deficient in processing synthetic HJs in vitro and in vivo (26). Unexpectedly, RuvA3m helicase activity and branch migration of Y-junctions in vitro appeared unaffected, and it was proposed that complex I was able to support one RuvB hexameric ring, but complex II was needed to assemble two hexameric rings on the Holliday junction (26). A Thermus thermophilus “tetramer-only” RuvA mutant, RuvA(DK) (L125D and E126K) was also studied. Electron microscopy demonstrated that a single RuvA(DK) tetramer formed tripartite complexes containing two hexameric rings of RuvB on the HJ (27). RuvA(DK) displayed reduced ability to promote branch migration of Holliday junctions in vitro and, significantly, could not support branch migration with a single RuvB hexamer (28). The capacity for replication fork reversal of these tetramer-only mutants has not been tested.

Recently, two ruvA mutants called ruvAz3 and ruvAz87 (H29R/K129E/F140S and N79D/N100D, respectively) were isolated and characterized as separation-of-function mutants that can process Holliday junctions but cannot reverse replication forks (29). The RuvAz proteins contain several mutations in different domains of the proteins, making it difficult to ascertain the molecular cause for their phenotypes. One intriguing observation is the inability of RuvAz mutants to form complex II on Holliday junctions, which is common with the RuvA3m and RuvA(DK) mutants discussed above. However, the tetramer-only RuvAz mutants are fully capable of promoting homologous recombination in vivo, whereas the tetramer-only RuvA3m is not.

To understand how the ability of RuvA to form octamers (complex II) relates to branch migration of HJs and replication fork reversal in vitro and in vivo, we have designed a new tetramer-only RuvA mutant, RuvA2KaP. RuvA3m, which was used in previous work (26), is inactive in vivo and displays significant nonspecific DNA binding, which could explain some discrepancies compared with studies of RuvA(DK) (28). In contrast, RuvA2KaP was designed to only disrupt tetramer-tetramer interactions. In this study, we have investigated the ability of RuvA2KaP to bind and process HJs and synthetic forks in vitro and how these correlate with the ability of the mutant to process different substrates in vivo. We also compare certain biochemical activities of RuvA2KaP mutant with those of the separation-of function mutants RuvAz3 and RuvAz87.


Mutagenesis of RuvA2KaP

To test RuvA2KaP function in vivo, plasmids pGB-ruvA and pGB-ruvAB were mutagenized using mutagenic primers 259F and 260R (supplemental Table 2). Both of these primers incorporated a codon change from GAA to CGC at position 379 in the coding sequence. Site-directed mutagenesis (QuikChange® II site-directed mutagenesis kit) was used to produce pGB-ruvA1KaP and pGB-ruvA1KaPB, which contained the single amino acid substitution E127R. The thermal cycling procedure employed was as follows: 1 time at 95 °C for 30 s, 16 cycles of 95 °C for 30 s, 53 °C for 45 s, and 68 °C for 10 min. To produce pGB-ruvA2KaP and pGB-ruvA2KaPB, which contain two amino acid substitutions E127R and K119A, pGB-ruvA1KaP and pGB-ruvA1KaPB were used as templates, and the primers 401F and 402R were used to introduce the second amino acid substitution K119A. Both primers 401F and 402R incorporated a codon change from AAA to GCA at position 355 in the coding sequence. The mutagenesis was carried out as described to introduce the first substitution (E127R). Sequencing of the pGB-ruvA1KaP, pGB-ruvA2KaP, pGB-ruvA1KaPB, and pGB-ruvA2KaPB constructs was carried out by MWG-Biotech using the primers RkF, RkR, and internal primers RkiF and RkiR (supplemental Table 2).

Recombinant Protein Production

All protein fractions were diluted with loading buffer (0.125 m Tris-HCl, pH 6.8, 2% SDS, 10% glycerol, 0.01% bromophenol blue, 10% β-mercaptoethanol) and analyzed by SDS-PAGE analysis. RuvA or RuvA2KaP was cloned into pET21a, expressed in BL21-GOLD(DE3), and purified following a protocol described previously (26) with several modifications. For each protein, four frozen pellets (total volume 4 ml) were thawed on ice and resuspended in 21 ml of Lysis buffer (100 mm Tris-HCl, pH 8, 5% glycerol, 2 mm EDTA, 1 mm of DTT, and 1 mg·ml−1 of lysozyme) and incubated for 30 min at 4 °C. A final concentration of 1 m NaCl and 0.1% Triton X-100 was added to the solution and incubation continued for a further 10 min on ice. The solution was made up to 0.4% sodium deoxycholate and spun at 42,000 rpm for 60 min at 4 °C in a Type 70 Ti rotor in an OptimaTM L-100 XP ultracentrifuge (Beckman Coulter). The supernatant was dialyzed against 2 liters of TEGD buffer (20 mm Tris-HCl, pH 8, 1 mm EDTA, 0.5 mm DTT, 10% glycerol).

The crude lysate was loaded onto a DEAE column, and the column was washed with TEGD buffer, and a gradient of 0–500 mm KCl in TEGD buffer was employed to elute the protein. The eluted protein was dialyzed against 2 liters of H buffer (10 mm KPi, pH 6.8, 150 mm KCl, 0.5 mm DTT, 10% glycerol) at 4 °C. Dialyzed protein was loaded onto a hydroxylapatite column, and the column was washed with H buffer. A 100-ml 10–600 mm gradient of KPi in H buffer was used to develop the column. Eluted RuvA was dialyzed at 4 °C against 2 liters of H buffer supplemented with 5 mm β-mercaptoethanol, 0.1 mm PMSF, and 100 mm KCl with no DTT. Dialyzed RuvA was loaded onto a HiTrap heparin column that was washed with buffer, and a 10–600 mm KPi gradient was used to elute the protein that was dialyzed against 2 liters of TEGD buffer at 4 °C. RuvA was loaded onto a single strand DNA column, and the column was washed with TEGD buffer and developed with a 0 mm to 1 m KCl gradient. Eluted RuvA protein was dialyzed overnight at 4 °C against 2 liters of TEGD buffer. Dialyzed RuvA protein was loaded onto a 1-ml Mono Q column that was washed with TEGD buffer, and a gradient 0–1.5 m KCl in TEGD buffer was used to elute the RuvA protein. The protein concentrations of the purified RuvA protein were determined using a Bradford assay. RuvA was stored as either 10 or 50% glycerol stocks at −20 °C. RuvA2KaP protein was produced using an identical procedure except that after purification on a heparin HiTrap column the protein was loaded straight onto a Mono Q column; the single strand DNA column was not used.

A protocol for producing RuvBD113E was modified to produce wild type RuvB (26). RuvB was overexpressed from plasmid pET21a in BL21-GOLD(DE3) cells. Cultures of RuvB-pET21a were grown in LB (100 μg·ml−1 ampicillin) at 37 °C to an absorbance of 0.6 at 600 nm. The cultures were supplemented with 1 mm isopropyl 1-thio-β-d-galactopyranoside and incubated for 6 h at 37 °C at 250 rpm. The cultures were pelleted at 4000 relative centrifugal force for 10 min in an SLA 3000 rotor in a Sorvall RC6 Plus centrifuge and resuspended in 25 ml of LB. The suspension was further spun for 10 min at 4000 relative centrifugal force at 4 °C, and the resulting pellets were frozen at −20 °C. Purification was carried out as described previously (26). RuvC was expressed and purified using a protocol as described previously (5).

Size Exclusion Chromatography (SEC)

RuvA, RuvA mutants, and RuvB proteins were dialyzed against TEGD supplemented with 0.1 m NaCl overnight at 4 °C. A total volume of 200 μl of 250 μg of each protein was applied to the 25 ml of Superose 6TM 10/30 GL column. The proteins were eluted from the column in TEGD buffer supplemented with 0.1 m NaCl at a flow rate of 0.3 ml·min−1. Molecular weights of species were estimated by comparison with five molecular weight standards (Bio-Rad). Fractions were analyzed by SDS-PAGE analysis, and the UV absorbance profiles of the eluted proteins were recorded.

DNA Substrate Preparation

The DNA substrate X12 (HJ) was constructed (see supplemental Table 1) as described previously (26). Replication fork-like substrates F2 and HJY3m were also constructed. JBM5a and IT01 were synthesized with a fluoro tag IRD700 attached to the 5′ end.

The required combinations of oligonucleotides were included in annealing reactions, using 1 μg of each oligonucleotide in SSC buffer (150 mm sodium chloride and 15 mm trisodium citrate, pH 7.0) incubated at 95 °C for 2 min and slowly annealed by cooling the heat block to room temperature. Substrates were purified by electrophoresis on an 8% native gel in TBE buffer run at 15 V·cm−1 for 80 min at 4 °C. Gel bands were cut from the gel, and DNA was eluted from the gel pieces by electrophoresis in 0.5× TBE buffer for 1 h using a BioTrap multikit (Schleicher & Schuell). Each DNA substrate was then dialyzed against 2 liters of DNA storage buffer (10 mm Tris-HCl, pH 8, 1 mm EDTA, 50 mm NaCl) and stored at −20 °C.


EMSA reactions containing the indicated amount of protein and DNA substrate were incubated in DNA binding buffer (50 mm Tris-HCl, pH 8, 0.5 mm EDTA, 1 mm DTT, 100 μg·ml−1 BSA, 6% glycerol). Proteins were diluted using Dilution buffer (20 mm Tris-HCl, pH 8, 150 mm NaCl, 0.5 mm DTT, 10% glycerol). Reactions were incubated for 5 min on ice. For gel loading, 5 μl of 80% glycerol was added to each 20-μl reaction to allow the samples to sink into the wells; no dye was used as the dye interferes with detection of the signal. In some instances, protein·DNA complexes were resolved on native gels of different concentrations of polyacrylamide as follows: between 4 and 10% in TBE buffer or in low ionic TAE buffer (6.7 mm Tris-HCl, pH 8.1, 2 mm EDTA, 3.2 mm sodium acetate) at 6 V·cm−1 at 4 °C in a Bio-Rad miniprotean II gel system. The gels were run at 10 V·cm−1 for 4 h at 4 °C. Gels were scanned using the Odyssey® infrared imaging system (from LI-COR Biosciences) at 700 nm at an intensity of 10.

For EMSAs in Mg2+, reactions were carried out exactly as for DNA binding assays in EDTA, except EDTA was omitted from the DNA Binding buffer and replaced by 5 mm MgCl2. The samples were loaded after addition of 5 μl of 80% glycerol and resolved using 6% polyacrylamide native gels with the EDTA omitted from the gel and replaced with 5 mm MgCl2. The gels were run in 0.5× TBM buffer (45 mm Tris base, 45 mm boric acid, 200 μm MgCl2) at 6 V·cm−1 for 4 h with recircularization of the buffer on ice.

Branch Migration Assays

Reactions were carried out in branch migration buffer (20 mm Tris-HCl, pH 7.5, 15 mm MgCl2, 2 mm ATP, 2 mm DTT and 100 μg·ml−1 BSA). DNA and then protein were added to the reactions at the required concentrations. The reaction was incubated at 37 °C for 30 min, and 5 μl of 5× Stop buffer (100 mm Tris-HCl, pH 8, 200 mm EDTA, 2.5% SDS, and 10 mg·ml−1 proteinase K) was then added to each 20-μl reaction with incubation at room temperature for 10 min. 5 μl of 80% glycerol was added to each sample, and the samples were loaded onto 6% polyacrylamide native gels and run in 0.5× TBE buffer at 7 V·cm−1 for 2 h. DNA products were visualized on the Odyssey® infrared imaging system at 700 nm at intensity 10.

RuvA·B-DNA Complex Formation Assay

Reactions were incubated at 37 °C for 30 min in formation buffer (20 mm triethanolamine-HCl pH 7.5, 10 mm MgCl2, 0.25 mm ATPγS, 1 mm DTT, 50 μg·ml−1 BSA). The proteins and DNA were added as required and were fixed by adding 0.2% glutaraldehyde and incubating reactions for 20 min at 37 °C. 5 μl of 80% glycerol were added to each reaction, and samples were loaded onto a 6% polyacrylamide native gels and run in 1× TAE buffer. Gels were run at 10 V·cm−1 for 2 h at room temperature and scanned on the LI-COR fluoro imaging system at 700 nm intensity 10.

RuvC Cleavage Assay

Reactions were carried out in cleavage buffer (50 mm Tris-HCl, pH 8, 10 mm MgCl2, 50 mm KCl, 5 mm β-mercaptoethanol, 100 μg·ml−1 BSA and 5% glycerol). 2 μl of RuvA protein and 2 μl of RuvC protein were mixed and incubated with HJ in cleavage buffer at 37 °C for 1 h. 5 μl of 5× stop buffer was then added to the 20-μl reactions, which were incubated at 37 °C for a further 10 min. 5 μl of 80% glycerol was added to 25-μl reactions that were loaded onto 6% native polyacrylamide gels and run in TBE buffer at 10 V·cm−1 at room temperature. DNA products were visualized using the Odyssey LI-COR fluoro imaging system.

ATPase Assay

The indicated amounts of protein and DNA substrate were mixed, and reactions were performed in ATPase buffer (50 mm Tris-HCl, pH 7.5, 50 mm NaCl, 15 mm MgCl2, 0.1 mg·ml−1 BSA, 1 mm DTT, 15 mm MgCl2, and 0.5 mm ATP). A Malachite green kit was used to detect activity; the reactions were preincubated on ice, and a zero time point was taken with 20 μl of reaction added to 5 μl of 0.5 m EDTA. Reactions were then incubated at 37 °C with time points taken at 5, 15, and 30 min. 150 μl of ALS mix (Innova Biosciences) was added to each well, and the reactions were incubated for 30 min at room temperature. The 96-well plates were scanned in a Tecan Sunrise fluorometer and analyzed with software by Magellan. A calibration curve of 10 different KPi concentrations was used to determine the amount of phosphate released in each reaction.

Measurement of Recombinational DNA Repair

UV irradiation was performed as described previously (29). The survival ratio was a comparison of colonies that grew on a replica control plate compared with colonies that grew on a plate that was irradiated. For mitomycin C treatment, cells were grown at 37 °C in LB to an A600 = 0.5, and mitomycin C was added to the culture at a final concentration of 2 μg/ml, and incubation was continued at 37 °C for 90 min. An untreated culture was used as control. Appropriate dilutions were plated on LB plates and incubated overnight at 37 °C. Ratios of cfu of mitomycin C treated/cfu of untreated cells were calculated.

Measurement of Conjugational Recombination

Conjugations were performed as described using JJC145 as Hfr donor (29); donor and recipient cells were mixed for 25 min. Selective medium was M9 minimal medium supplemented with leucine, proline, threonine, and arginine (2% final concentration each) and 10 μg·ml−1 chloramphenicol.

Measure of Linear DNA by Pulse Field Gel Electrophoresis

Quantification of pulsed field gels was performed using in vivo [3H]thymidine-labeled chromosomes as described previously (22).


RuvA2KaP Binds Efficiently to Holliday Junctions as a Single Tetramer

A set of RuvA mutants was generated by introducing amino acid substitutions in α-helix-6 in domain II, which is the region involved in tetramer-tetramer interactions. Specifically, the RuvA2KaP used in this study carried E127R and K119A mutations in the tetramer-tetramer interface (supplemental Fig. 1).The overall electrostatic charge of this region changes only slightly compared with wild type, and the slightly more basic/positive charge does not alter RuvA2KaP interaction with DNA (Fig. 1A). The mutant RuvA3m used in a previous study formed aberrant complexes on HJs, probably caused by the significant increase in positive charge in α-helix-6 (26).

RuvA2KaP binding synthetic Holliday junction. A, representative EMSA is shown. Increasing concentrations of RuvA or RuvA2KaP were incubated with IRD700-labeled junction X12 in EDTA buffer and analyzed by native 4% PAGE. The expected positions of unbound ...

The ability of RuvA2KaP to form stable complexes was tested in vitro; RuvA2KaP was analyzed on SDS-polyacrylamide gels and formed tetramers comparable with wild type RuvA, which is stable enough not to dissociate in SDS (supplemental Fig. 2A). When boiled and loaded onto an SDS-polyacrylamide gel, RuvA2KaP dissociated into monomers, indicating that, like RuvA, the mutant forms stable tetramers that only dissociate upon boiling (supplemental Fig. 2A). RuvA2KaP was analyzed by SEC on a 25-ml Superose 6TM 10/30 GL column, and the elution profile of RuvA2KaP was comparable with RuvA (supplemental Fig. 2B). Additionally, RuvA2KaP mixed with RuvB was loaded onto a 25-ml Superose 6TM 10/30 GL column, and the formation of the RuvA2KaPB complex was equivalent to RuvAB complex formation (supplemental Fig. 2C). In conclusion, RuvA2KaP forms a stable tetramer and is able to form a complex with RuvB in solution with an efficiency comparable with that of wild type RuvA.

The binding of RuvA2KaP to a fluoro-tagged synthetic HJ (X12) was tested using EMSAs (Fig. 1A). In EDTA-containing buffer, RuvA formed both complex I and II at lower concentrations but exclusively formed complex II at protein concentrations of 150 nm and higher. Conversely, RuvA2KaP only formed complex I, even at protein concentrations as high as 2 μm. The amount of HJ bound by RuvA2KaP was similar to wild type RuvA (Fig. 1B), indicating that the mutations did not affect the affinity of RuvA2KaP for HJs. Holliday junction binding was further tested in the presence of Mg2+ as RuvB requires at least 5 mm of Mg2+ for efficient ATP hydrolysis and branch migration. In 5 mm MgCl2, wild type RuvA bound to X12 exclusively as complex II, even at low protein concentrations, leaving a significant amount of free junction (Fig. 1C). RuvA2KaP formed complex I at concentrations up to 75 nm, but at 250 nm of RuvA2KaP and above, complex II also formed (Fig. 1C). The overall RuvA2KaP binding of X12 in Mg2+ was comparable with RuvA (Fig. 1D). It was clear that RuvA2KaP complex II was stabilized by Mg2+, so we checked for a direct stabilizing effect of 5 mm MgCl2 on tetramer-tetramer interactions using SEC. As the SEC elution profile of RuvA remained unchanged (data not shown), the effect of Mg2+ was therefore dependent on the presence of DNA. Experiments in Mg2+ buffer revealed that a lack of tetramer-tetramer interaction does not fully prevent complex II formation but renders it dependent on a high concentration of RuvA protein.

Stability of RuvA2KaP Complex II in Solution

In vivo RuvC cannot function without RuvAB (7, 8, 29, 30); however, it has been shown that formation of RuvA complex II occludes the HJ and prevents RuvC-mediated cleavage in vitro (26, 31, 32). To assess the stability of the RuvA2KaP complex II in the presence of Mg2+, we compared the ability of both wild type and mutant proteins to protect a HJ from cleavage by RuvC (Fig. 2). Varying concentrations of RuvA or RuvA2KaP were incubated with X12, followed by addition of 100 nm RuvC. Significant inhibition of RuvC HJ cleavage was observed at wild type RuvA concentrations of 100 nm, and cleavage was completely abolished at 300 nm RuvA. The inhibition of RuvC cleavage correlated well with complex II formation in EDTA binding studies but only roughly with the concentration at which complex II forms in the Mg2+ binding studies. HJ cleavage by RuvC was also tested in the presence of up to 2 μm of RuvA2KaP. Cleavage was only slightly inhibited within the range of 100–500 nm RuvA2KaP and was still weakly observed in the presence of 2 μm of RuvA2KaP (Fig. 2A). This demonstrates that although RuvA2KaP forms complex II on Holliday junctions, the stability of RuvA2KaP complex II is reduced compared with RuvA complex II.

Stability of complex II of RuvA mutants on Holliday junctions. A, increasing concentrations of RuvA mutants were incubated with X12; 100 nm RuvC was added, and the reaction was incubated for 15 min at 37 °C. The cleavage products were analyzed ...

Because the RuvA2KaP protein confers to E. coli a separation-of-function phenotype (see below), we compared the previously isolated separation-of-function mutants RuvAz3 and RuvAz87 to RuvA2KaP using the same conditions. As with RuvA2KaP, both RuvAz mutants were unable to inhibit RuvC cleavage of the junction at concentrations up to 300 nm (Fig. 2B). The inability of RuvAz mutants to protect the HJ from RuvC cleavage confirms that they form unstable complex II (29). These results also suggest that a tetramer of RuvA bound to the HJ (complex I) does not inhibit HJ cleavage by RuvC, and it is possible that both RuvA and RuvC bind together to the junction. Alternatively, complex I might transiently dissociate from the junction allowing RuvC access, whereas complex II does not dissociate.

Stimulation of RuvB ATPase Activity by RuvA Mutants

To measure the functional interaction between RuvA2KaP and RuvB, we tested the ability of RuvA2KaP to stimulate the DNA-dependent ATPase activity of RuvB. Time courses of ATP hydrolysis using 100 nm of RuvA or RuvA2KaP, 500 nm RuvB, and 5 ng of X12 are shown in Fig. 3A. The DNA-dependent ATPase activity of RuvA2KaPB was about half of the activity of wild type RuvAB; RuvAB hydrolyzed 80 μmol of ATP in 30 min, whereas RuvA2KaPB hydrolyzed 40 μmol of ATP in 30 min (Fig. 3A). The ATPase activity was also measured with higher RuvA concentrations. ATP hydrolysis was 2–3-fold lower with RuvA2KaP compared with RuvA at all concentrations at which RuvA2KaP complex II formation was observed by EMSA (250 nm and above) (Fig. 3B). These data indicate that tetramer-tetramer interactions within complex II are necessary for optimal stimulation of the DNA-dependent ATPase activity of RuvB.

ATP hydrolysis of RuvA mutant·RuvB complexes on synthetic HJs. A, time course of ATP hydrolysis. 100 nm RuvA or RuvA2KaP was incubated with 500 nm RuvB and X12 at 37 °C. ATP hydrolysis is proportional to the release of inorganic phosphate ...

The ability of the RuvA2KaP mutant to stimulate the DNA-dependent ATPase activity of RuvB was compared with that of the separation of function mutants RuvAz3 and RuvAz87. Fig. 3C shows the amount of ATP hydrolyzed per mol of RuvB per min in the presence of [var phi]X174 virion DNA for different RuvA mutants. RuvB alone hydrolyzed about 8 mol of ATP per mol of RuvB·min−1. The presence of RuvA stimulated RuvB to hydrolyze 50 mol of ATP per mol of RuvB·min−1. RuvA2KaP, RuvAz3, and RuvAz87 stimulated RuvB to hydrolyze 18, 20, and 16 mol of ATP per mol of RuvB·min−1, respectively. In the absence of DNA, neither RuvA, RuvA2KaP, RuvAz3, nor RuvAz87 stimulated the hydrolysis of ATP by RuvB, as expected (data not shown). These data show that the three mutants RuvA2KaP, RuvAz3, and RuvAz87 are less efficient in stimulating RuvB ATPase activity than wild type RuvA.

RuvA2KaP Forms Unstable RuvA·RuvB·HJ Tripartite Complexes in Solution

The RuvA2KaP mutant was found to bind RuvB in solution by SEC analysis as wild type RuvA (supplemental Fig. 2, B and C). To analyze the interactions of RuvA2KaP with RuvB on Holliday junctions (X12), these large protein·DNA complexes were cross-linked with 0.25% glutaraldehyde and analyzed by native PAGE. When incubated with 5 ng of X12, 400 nm RuvA alone could be cross-linked on DNA as complex II but 400 nm RuvA2KaP could not (Fig. 4, lanes 10 and 11). The inefficient glutaraldehyde cross-linking of RuvA2KaP complex II on X12 reveals the reduced stability of the RuvA2KaP tetramer-tetramer interaction across the junction. Increasing concentrations of RuvA or RuvA2KaP were incubated with 300 nm RuvB and X12. RuvB alone could not be cross-linked to X12 under these conditions (Fig. 4, lane 12). RuvAB complexes on X12 are detected when 25 nm RuvA and above is used (Fig. 4, lanes 1 and 2). At 200, 300, and 400 nm RuvA, complex II is also observed in addition to RuvAB complexes on DNA (Fig. 4, lanes 4–6). The stoichiometry of RuvA:RuvB on the junction is predicted to be 8:12 monomers, respectively, and thus at equal concentrations of RuvA and RuvB, there is an excess of RuvA that forms complex II. We incubated 300 and 400 nm RuvA with 600 nm RuvB (Fig. 4, lanes 7 and 8). This altered the ratio of RuvA:RuvB to favor formation of RuvAB complexes on DNA rather than complex II (Fig. 4, lanes 7 and 8). Interestingly, only a fraction of X12 could be cross-linked with the mutant RuvA2KaPB compared with wild type RuvAB, and a significant amount of DNA remained protein-free (Fig. 4, lanes 13–18). Increasing the concentration of RuvB to 600 nm had no effect on tripartite RuvA2KaP·RuvB·HJ complex formation (Fig. 4, compare lanes 17 and 18 with 19 and 20). The cross-linking experiments demonstrate the reduced stability of RuvA2KaP complexes with RuvB on DNA. Thus the stability of the RuvA double tetramer is crucial for the formation of a stable tripartite RuvAB·HJ complex.

Interaction of RuvA2KaP with RuvB on HJ. Increasing amounts of RuvA or RuvA2KaP were mixed with X12 in the presence of 300 or 600 nm RuvB in triethanolamine buffer containing ATPγS. The protein·DNA complexes were cross-linked using 0.2% ...

RuvA2KaP-RuvB-mediated Branch Migration of Synthetic Holliday Junctions

The ability of RuvA2KaP and RuvB to promote branch migration of HJs was tested using fluoro-tagged X12, increasing concentrations of RuvA or RuvA2KaP and 250 nm RuvB. As shown in Fig. 5A, X12 was processed efficiently by wild type RuvAB to form two branch migration products. In contrast, RuvA2KaP was clearly deficient in branch migration with small amounts of products observed at RuvA2KaP concentrations of 75 nm and above. Around 75% of X12 was processed at 100 nm wild type RuvA, whereas less than 25% of X12 was dissociated at 100 nm RuvA2KaP (Fig. 5B). We conclude that branch migration was inefficient due to the weak stability of RuvA2KaP complex II.

Holliday junction processing of RuvA2KaP with RuvB. A, branch migration of X12. Increasing concentrations of RuvA and RuvA2KaP were incubated with 5 ng of IRD700-labeled X12 and 250 nm RuvB at 37 °C for 30 min. The reactions were stopped and analyzed ...

Holliday junction branch migration with a single RuvB hexamer was tested using the synthetic junction HJY3-hm (28). HJY3-hm is made of three short 14-bp arms and one 49-bp arm, so that only one RuvB hexamer can load onto the long arm of this junction. As with X12, the binding affinities of wild type RuvA and RuvA2KaP to HJY3-hm were comparable (data not shown). The assays with HJY3-hm showed highly efficient RuvAB-promoted branch migration of the junction through its short arms, but RuvA2KaPB complex was inefficient (Fig. 5C). At 25 nm RuvA, HJY3-hm dissociated completely, but in contrast, only 25% of the substrate was processed by 25 nm RuvA2KaP (Fig. 5C). At 200 nm RuvA2KaP, only 35% of the HJY3-hm was processed (Fig. 5C). These results show that a single RuvB hexamer assembled on DNA requires stable interactions between two RuvA tetramers for efficient processing of a four-way junction.

Binding and Processing of Synthetic Replication Forks by RuvA2KaP

We tested the ability of RuvA2KaP to bind and process a synthetic model replication fork (F2), in which the three branches of the forks are fully double-stranded. In EDTA buffer, RuvA bound to F2 exclusively as complex II at concentrations above 10 nm, but RuvA2KaP only formed complex I at all concentrations tested (Fig. 6A). Significantly, RuvA2KaP showed reduced affinity of binding to forks compared with RuvA (Fig. 6B) even though the binding affinity of RuvA2KaP to HJs was comparable with that of RuvA. Similar results were obtained in the presence of Mg2+ above 10 nm RuvA, and 100% F2 was in complex II, but even at 250 nm, RuvA2KaP only bound 40% of F2 as a mixture of complex I and II (data not shown). These results confirm that stable binding of RuvA to the fork requires interactions between the two tetramers. RuvA2KaPB complex was significantly defective in processing the fork (Fig. 6C). Over 50% of the synthetic fork F2 was dissociated in reactions containing 250 nm RuvA and 300 nm RuvB, whereas less than 10% of the substrate dissociated in parallel reactions using RuvA2KaPB (Fig. 6D). The majority of the F2 dissociation products represent processing in the direction opposite to fork reversal, as reported previously (25). These results show that the pronounced defect in the ability of RuvA2KaP to bind to the synthetic fork (F2) resulted in a significant defect in processing.

RuvA2KaP binding to a synthetic fork in EDTA. A, increasing concentrations of RuvA or RuvA2KaP were incubated with fluorescently labeled F2 for 5 min on ice. Complexes of DNA with RuvA or RuvA2KaP were analyzed by native 4% PAGE. A schematic representation ...

Processing of Holliday Junctions by RuvA2KaP in Vivo

As shown above, the mutant RuvA2KaP was defective in forming stable complex II on model Holliday junctions and replication forks. As a consequence, the mutant was inefficient in branch migration of Holliday junctions and replication forks in vitro. It is important to correlate the observed defects of the RuvA2KaP mutant with the currently known biological roles of RuvAB in vivo, namely processing of Holliday junctions and RuvAB-dependent reversal of stalled replication forks.

We first tested the ability of RuvA2KaP to resolve HJ formed in vivo by homologous recombination between intact DNA molecules. The E. coli strain JJC3207 (ΔruvA100::cat ΔrecG263::kan) (29) has a defect in homologous recombination that causes deficiency in Hfr conjugation. To test if RuvA2KaP can process HJs in vivo, the ability of RuvA2KaP to rescue the mutant phenotype of this strain was tested and compared with RuvA. The ruvA2KaP coding sequence was cloned into the low copy plasmid pGB2 or in combination with the ruvB coding sequence to produce pGB-ruvA2KaP and pGB-ruvA2KaPB. The genes were expressed under the control of the native ruvAB promoter. The plasmids pGB2, pGB-ruvA, pGB-ruvA2KaP, or pGB-ruvA2KaPB were transformed into the recipient JJC3207. As the pGB-ruvA2KaPB plasmid codes for both RuvA2KaP and RuvB, both proteins are expressed at the same levels when introduced into E. coli. This reduces any effects of altering the balance between RuvA2KaP and RuvB, when RuvA2KaP is overexpressed from the pGB2 plasmid, although RuvB is expressed at low levels from the chromosome. The cells were grown to log phase and then mixed with a His+ Hfr donor, plated on chloramphenicol minimal medium devoid of histidine, and incubated for 48 h. If successful homologous recombination had occurred, JJC3207 cells could grow on the plates lacking histidine as the His+ gene had been acquired from the Hfr donor via conjugation. Ratios of His+ versus total recipient cfu are shown (Fig. 7A). Strain JJC3207 carrying vector pGB2 showed very low conjugation levels, less than 10−5 conjugants per cfu. Transformation of JJC3207 with pGB-ruvA resulted in much higher levels of conjugation, ~10−3 conjugants per cfu. When the cells were transformed with pGBruvA2KaP or pGBruvA2KaPB, the conjugation level was restored to nearly 10−3 conjugants per cfu, which is comparable with the rescued phenotype demonstrated by pGB-ruvA. Thus RuvA2KaP is able to process HJs during conjugation as efficiently as wild type RuvA expressed from an exogenous plasmid. The complementation of the homologous recombination defect of JJC3207 by RuvA2KaP was the same when expressed alone or in combination with RuvB on the pGB2 plasmid. This indicates that expressing extra copies of RuvB was not required for RuvA2KaP-mediated rescue of the JJC3207 conjugational defect.

RuvA2KaP complex II binding stability and RuvA2KaPB HJ and fork processing in vivo. A, C, and E, error bars represent S.D. B and D, error bars indicate the minimum and maximum values. A, RuvA2KaP was tested for rescue of conjugation ability of the E. ...

The ability of RuvA2KaP to process HJs in vivo was additionally tested during the DNA single strand gap and double strand break repair, by assessing whether the mutant RuvA2KaP was able to suppress the UV or mitomycin C (MMC) sensitivity of the mutant strain JJC2971 (ΔruvA100::cat) (29). JJC2971 was transformed with pGB-ruvA, pGB-ruvAB, pGB-ruvA2KaP, and pGB-ruvA2KaPB, and the cells were exposed to increasing doses of UV light. JJC2971 cells transformed with pGB2 alone resulted in survival ratio of 10−3 cells at a UV dose of 40 J/m2 (Fig. 7B). In contrast, the survival of JJC2971 transformed with pGB-ruvA was 100% at 40 J/m2 UV dose, indicating that the cells were able to repair DNA lesions. JJC2971 cells transformed with either pGB-ruvA2KaP or pGB-ruvA2KaPB gave survival profiles comparable with pGB-ruvA transformation at all doses tested, indicating that RuvA2KaP is fully capable of resolving HJ made during the recombinational repair of UV lesions. The co-expression of RuvA2KaP with RuvB in JJC2971 cells very slightly improved survival, which suggests that there may be a minor defect in the RuvA2KaP interaction with RuvB. In an additional assay, the JJC2971 cells were treated with 2 μg·ml−1 MMC for 90 min to induce double strand breaks in the DNA. These cells were then plated on LB spectinomycin overnight, and the ratios of colony forming units in treated and untreated cultures were calculated (Fig. 7C). Introduction of pGB2 only resulted in a survival ratio of 10−5 cells. Introduction of pGB-ruvA, pGB-ruvA2KaP, or pGB-ruvA2KaPB all resulted in a survival ratio of 10−2 cells indicating that the RuvA mutants were able to restore the cells resistance to MMC to the levels observed after introducing pGB-ruvA. These data suggest that RuvA2KaP is able to process Holliday junctions formed during recombination-mediated repair of DNA lesions as efficiently as wild type RuvA.

Stability of RuvA2KaP on Holliday Junctions in Vivo

The defect of RuvA2KaP in maintaining stable binding to both sides of junction DNA as a double tetramer was demonstrated in vitro by its inability to inhibit cleavage of the junction by RuvC. We decided to examine this finding in vivo. In E. coli, the rusA gene encodes a Holliday junction resolvase carried on a cryptic prophage, but this gene in not normally expressed (8, 29). The E. coli strain JJC2761 (ΔruvABC rus-1) does not encode RuvABC, but the rus-1 mutation results in rusA expression, and the cells survive UV damage as RusA resolves Holliday junctions formed by recombination-mediated repair (24, 29). We used the JJC2761 strain as a tool to test the effect of RuvA2KaP on Holliday junction resolution by RusA in vivo. If a ruvA-carrying plasmid is added to the strain, RuvA binding occludes the Holliday junction and prevents the action of RusA. Holliday junctions cannot be resolved, resulting in UV sensitivity and cell lethality. JJC2761 cells were transformed with pGB2, pGB-ruvA, pGBruvA2KaP, or pGBruvA2KaPB, and the survival of plasmid-harboring cells subjected to different doses of UV-irradiation is shown in Fig. 7D. A similar level of protection against RusA-catalyzed HJ resolution, resulting in UV sensibility, was observed for pGB-ruvA, pGBruvA2KaP, or pGBruvA2KaPB at 40 J/m2. At higher doses, the survival ratio of JJC2761 cells with pGBruvA2KaP or pGBruvA2KaPB was intermediate between those observed with the vector pGB2 and with the control plasmid pGB-ruvA, ~15–20-fold more cells survived compared with cells carrying pGB-ruvA at 100 J/m2. These data demonstrate that RuvA2KaP alone or with RuvB could protect the Holliday junction from cleavage by RusA, but not as efficiently as wild type RuvA. The RuvAz3 and RuvAz87 mutants, which were deficient at inhibiting RuvC-mediated cleavage of HJs (Fig. 2B), were also defective in preventing RusA cleaving Holliday junctions in the JJC2761 strain (29).

RuvA2KaP Is Deficient for Replication Fork Reversal in Vivo

Finally, the ability of the mutant RuvA2KaP to reverse stalled replication forks in conjunction with RuvB was tested in vivo using the strain JJC3723 (dnaEts recB ruvA100). In cells carrying the temperature-sensitive replication mutant dnaEts, RuvA is required for RFR (24). In the strain JJC3723, RFR can be measured by re-introducing RuvA on a plasmid and measuring the linearized DNA formed in this recB mutant by RuvC cleavage of reversed forks (29).

JJC3723 cells transformed with pGB2, pGB-ruvA, pGB-ruvA2KaP, and pGB-ruvA2KaPB constructs were grown at 30 °C and then shifted to 42 °C for 3 h. The amount of linearized DNA in the cells was analyzed by pulse field gel electrophoresis and quantified (Fig. 7E). Cells carrying the wild type RuvA plasmid, pGB-ruvA, resulted in 55% linearization of DNA compared with 11% DNA linearization in control cells with empty vector pGB2. Cells transformed with pGB-ruvA2KaP resulted in about 11% DNA linearization, similar to the control cells transformed with pGB2 alone, and thus RuvA2KaP could not promote replication fork reversal. When the JJC3723 cells were transformed with RuvA2KaP and RuvB (pGB-ruvA2KaPB), the amount of double strand breaks increased from 11 to ~28%, but it was still significantly lower than the levels of 55% observed with pGB-ruvA and the ~70% breakage conferred by pGB-ruvAB in a similar strain (33). These data indicate that RuvA2KaP, which binds forks but is deficient in complex II formation in vitro, is unable to reverse forks in vivo. Thus, a single RuvA tetramer that cannot interact with a second RuvA tetramer on DNA is not sufficient to reverse forks in concert with RuvB. Because RuvA2KaP is proficient in processing Holliday junctions in vivo during homologous recombination but is unable to reverse stalled replication forks, RuvA2KaP is a separation-of-function mutant similar to RuvAz3 and RuvAz87.


The aim of this study is to understand the molecular mechanisms of RuvAB acting on two different branched substrates in vivo, Holliday junctions and stalled replication forks. To this end, we used a defined RuvA mutant, RuvA2KaP, designed as tetramer-only by replacing two amino acid residues (E127R and K119A) engaged in ionic pairs that stabilize interactions between two RuvA tetramers on the Holliday junction (supplemental Fig. 1). Several lines of evidence argue that the two substitutions in RuvA2KaP do not affect the quaternary structure of the protein. The size exclusion data indicated that RuvA2KaP forms the correct complexes in the correct amounts in solution (supplemental Fig. 2A). The protein is stable compared with wild type RuvA when analyzed on SDS-PAGE (no degradation products were detected). Furthermore, like RuvA, RuvA2KaP remains a tetramer when run in SDS, indicating that interactions within the tetramer are stable (supplemental Fig. 2A). Complex formation with RuvB analyzed by SEC (supplemental Fig. 2C) indicates no defect in RuvA2KaP-RuvB protein interaction, and finally RuvA2KaP HJ binding affinity is as wild type (Fig. 1B). Therefore, we consider it unlikely that the two substitutions in RuvA2KaP affect the protein structure beyond destabilizing the double tetramer. As expected, RuvA2KaP was found to bind predominantly to HJs as complex I.

Tetramer Only Mutants Are Impaired in Vitro for the Formation of an RuvA·RuvB·HJ Tripartite Complex for the Stimulation of RuvB ATPase and Branch Migration Activities; Nevertheless, They Promote Homologous Recombination in Vivo

We observed that in Mg2+ buffer the affinity of RuvA2KaP for DNA remains similar to that of the wild type protein, but binding of the second tetramer is affected. This result suggests that Mg2+ has a greater effect on tetramer-tetramer interactions compared with modulating the affinity of RuvA for DNA. Interestingly, this result also reveals that complex II devoid of tetramer-tetramer interactions might form if the affinity for DNA of each of the two tetramers is high enough, albeit less stably than wild type. It supports the idea of RuvA co-existing on a HJ with RuvC (34). In fact, proteins are able to bind the opposite faces of a HJ without protein-protein interactions, as simultaneous binding to HJ was observed using two unrelated proteins, RuvA and the yeast mitochondrial protein YDC2 (35). The formation of unstable complex II devoid of tetramer-tetramer interactions correlates with the reduced ability of RuvA2KaP to stimulate RuvB ATPase activity in vitro and a defect in tripartite complex formation. Even at concentrations that allowed RuvA2KaP complex II formation on HJs in Mg2+ buffer, RuvA2KaP could not stimulate RuvB ATPase activity to wild type levels. The loss of tetramer-tetramer interaction could lead to a deficiency in functional interactions with RuvB on DNA. SEC analysis confirmed that physical interactions between RuvA2KaP and RuvB in the absence of DNA were not compromised, suggesting that RuvA2KaP-RuvB interactions were not responsible for the reduced ATPase activity observed by RuvA2KaPB. However, formation of the tripartite complex was clearly affected. Additionally, RuvA2KaP was defective in forming a RuvB complex when cross-linked to DNA, suggesting that stable formation of complex II via tetramer-tetramer interactions is critical for stabilizing RuvB on DNA. The decreased ATPase activity of an RuvA2KaPB mutant complex could reflect a role of tetramer-tetramer interaction in RuvA-facilitated communication required for two RuvB hexamers to function in concert. However, this is unlikely because RuvA2KaP is deficient at supporting a single RuvB hexamer during branch migration of the four-way junction HJY3-hm in vitro, compared with a double tetramer of RuvA. This cannot be due to a requirement for communication between two RuvB hexamers as there is only a single RuvB hexamer, although it remains possible that communications between subunits of the hexamer are affected. It is most likely that the ATPase and branch migration deficiency of RuvB in the presence of RuvA2KaP is due to an inherent inability of an RuvA single tetramer, or of an RuvA double tetramer devoid of tetramer-tetramer interactions, to stably tether RuvB to DNA. Alternatively, the wild type double tetramer stabilized by tetramer-tetramer interactions may have an unknown mechanistic function required for stable RuvB branch migration, such as the separation of the strands by the central negative pin and channeling through the cruciform tunnels.

The in vitro defects of RuvA2KaP, namely inefficient complex II formation and poor branch migration in vitro, do not translate into any detectable recombination deficiency in vivo. In vivo studies indicated that RuvA2KaP supports RuvB-mediated processing of Holliday junctions during conjugation and recombinational repair of UV- and MMC-induced DNA damage. Therefore, tetramer-tetramer interactions within complex II are not required for the formation of recombinant molecules, although it is possible that branch migration tracks are shorter in the presence of RuvA2KaP compared with wild type RuvA. Similarly, a surprising lack of correlation between in vivo homologous recombination proficiency and in vitro defects was previously observed for other separation-of-function mutants (RuvAz3 and RuvAz87 (29)). This paradox is confirmed here, and although they are recombination-proficient in vivo, these mutant proteins show a defect in protection against RuvC-mediated resolution and in stimulation of RuvB-ATPase activity similar to that of RuvA2KaP. The recombination efficiency of mutant proteins that are less active than wild type in several in vitro assays is an interesting observation that is not yet understood. The efficient RuvA2KaPB Holliday junction processing in vivo may be due to cellular factors stabilizing the complex on DNA, or alternatively, cellular factors may exist that recruit RuvA to the Holliday junctions thus increasing RuvA2KaP-localized concentrations. Such a cellular factor could be RecA, which creates the HJ by strand invasion. Alternatively, because pGB2 is present as 10 copies per cell, it is possible that the extra RuvA2KaP molecules expressed from this plasmid are able to overcome the inherent defect of RuvA2KaP in forming complex II on HJs. Indeed, at high concentrations of RuvA2KaP in magnesium buffer, complex II was formed on HJs, allowing more efficient branch migration in vitro. A final possibility is that the RuvAB branch migration motor is overly efficient at processing HJs in vivo and that the ineffective branch migration of RuvA2KaPB is still above the threshold of activity needed for HJ processing during homologous recombination in vivo.

Tetramer-only Mutant Is a Separation-of-Function Mutant

Several ruvA and ruvB separation-of-function mutants were isolated based on in vivo genetic screening (29, 33) and two were characterized in vitro (ruvAz3 and ruvAz87) (see Ref. 29 and this work). These RuvAz mutants were previously shown to be defective for complex II formation in vitro and for replication fork reversal in vivo, suggesting a role for tetramer-tetramer interaction in replication fork reversal (29). However, these mutants were defective for several in vitro activities, complex II formation and DNA binding (particularly in the presence of divalent cations); and it was not possible to identify an RuvA function that was specifically required for RFR. RuvA2KaP allowed us to directly test whether one or two RuvA tetramers bind replication forks to support a single RuvB hexamer resulting in unidirectional fork processing during fork reversal. RuvA2KaP bound to synthetic replication forks with a lower affinity than wild type, contrary to the binding affinity of RuvA2KaP for HJs, which was as wild type. These data imply that RuvA2KaP binding to a synthetic fork is inherently unstable and that wild type RuvA tetramer-tetramer interactions are required for fork binding. When Mg2+ is included in the fork binding reaction, the affinity of RuvA2KaP is more defective compared with wild type than in the absence of Mg2+, and little RuvA2KaP complex I is observed. These data indicate that the formation of complex II on a synthetic replication fork has a stronger requirement for stable tetramer-tetramer interactions than formation of complex II on a HJ. This higher requirement of stable complex II formation on synthetic replication forks may be due to the nature of the substrate, as an RuvA tetramer binds DNA on its concave basic face that contains four grooves in a cross arrangement (14). Thus HJ bound to the grooves would make more DNA protein contacts than a three-armed substrate. However, RuvA2KaP and wild type RuvA bind Y-junctions with a similar affinity (data not shown). The Y-junction and fork used in this study are identical in DNA sequence with the only difference between them being a single nick between the sister duplex arms of the fork substrate. Thus, it is likely that the structural differences between the rigid Y-junction and more flexible fork substrate account for the different interactions between these substrates and RuvA2KaP. This is confirmed by in vitro evidence that RuvA2KaPB is defective at processing the fork substrate in comparison with RuvA and defective at fork reversal in a dnaEts mutant in vivo.

Like RuvAz3 and RuvAz87, RuvA2KaP is a separation-of-function mutant, i.e. able to branch migrate Holliday junctions but not to reverse forks. Furthermore, these data are contrary to the model of a single RuvA tetramer recruiting a single RuvB hexamer onto the parental duplex arms of a stalled fork and the subsequent processing in the direction required for fork reversal. Our data indicate that the formation of an RuvA complex II stabilized by tetramer-tetramer interactions is crucial for fork reversal in vivo. As discussed previously (29), this study supports the idea that RFR mediated by RuvAB is a process that requires the formation of an inherently unstable RuvAB complex (consisting of a single hexamer of RuvB) on a three-way junction that is less stable than an RuvAB complex (consisting of two RuvB hexamers) bound to a HJ. The requirement for stabilizing this complex on a three-way fork junction is a double tetramer of RuvA with wild type tetramer-tetramer interactions, whereas branch migration of a HJ can be carried out by an unstable double tetramer with two RuvB hexamers providing additional stabilization through their contacts with RuvA.

Separation-of-function phenotype might result from various RuvAB defects, because different separation-of-function mutants were isolated in the ruvA as in the ruvB gene (29, 33). Studies of ruvB mutants showed that mutations that presumably affect the ATPase activity of RuvB also confer a separation-of-function phenotype (33). This study identifies the tight binding of two RuvA tetramers via tetramer-tetramer interactions as a property crucial for replication fork reversal but not for homologous recombination.

Supplementary Material

Supplemental Data:


We thank C. V. Privezentzev for advice regarding the purification of RuvB, N. Coates for producing the RuvA2KaP-pET21a plasmid, and S. Duigou for helpful reading of the manuscript.

*This work was supported in part by the Biotechnology and Biological Sciences Research Council, the Department of Structural Molecular Biology, University College London, by Grants ANR-05-BLAN-0204-01 and ANR-BLAN08-3_309268, and by a “Prix Coup d'Élan” from the Bettencourt-Shueller Foundation (to the B. M. laboratory).

An external file that holds a picture, illustration, etc.
Object name is sbox.jpgThe on-line version of this article (available at http://www.jbc.org) contains supplemental Tables 1 and 2 and Figs. 1 and 2.

4The abbreviations used are:

Holliday junction
replication fork reversal
adenosine 5′-O-(thiotriphosphate)
size exclusion chromatography
mitomycin C.


1. Iwasaki H., Shiba T., Makino K., Nakata A., Shinagawa H. (1989) J. Bacteriol. 171, 5276–5280 [PMC free article] [PubMed]
2. Tsaneva I. R., Müller B., West S. C. (1993) Proc. Natl. Acad. Sci. U.S.A. 90, 1315–1319 [PMC free article] [PubMed]
3. Putnam C. D., Clancy S. B., Tsuruta H., Gonzalez S., Wetmur J. G., Tainer J. A. (2001) J. Mol. Biol. 311, 297–310 [PubMed]
4. Connolly B., Parsons C. A., Benson F. E., Dunderdale H. J., Sharples G. J., Lloyd R. G., West S. C. (1991) Proc. Natl. Acad. Sci. U.S.A. 88, 6063–6067 [PMC free article] [PubMed]
5. Dunderdale H. J., Benson F. E., Parsons C. A., Sharples G. J., Lloyd R. G., West S. C. (1991) Nature 354, 506–510 [PubMed]
6. Iwasaki H., Takahagi M., Shiba T., Nakata A., Shinagawa H. (1991) EMBO J. 10, 4381–4389 [PMC free article] [PubMed]
7. Mandal T. N., Mahdi A. A., Sharples G. J., Lloyd R. G. (1993) J. Bacteriol. 175, 4325–4334 [PMC free article] [PubMed]
8. Sharples G. J., Chan S. N., Mahdi A. A., Whitby M. C., Lloyd R. G. (1994) EMBO J. 13, 6133–6142 [PMC free article] [PubMed]
9. Davies A. A., West S. C. (1998) Curr. Biol. 8, 725–727 [PubMed]
10. van Gool A. J., Hajibagheri N. M., Stasiak A., West S. C. (1999) Genes Dev. 13, 1861–1870 [PMC free article] [PubMed]
11. van Gool A. J., Shah R., Mézard C., West S. C. (1998) EMBO J. 17, 1838–1845 [PMC free article] [PubMed]
12. Ariyoshi M., Nishino T., Iwasaki H., Shinagawa H., Morikawa K. (2000) Proc. Natl. Acad. Sci. U.S.A. 97, 8257–8262 [PMC free article] [PubMed]
13. Hargreaves D., Rice D. W., Sedelnikova S. E., Artymiuk P. J., Lloyd R. G., Rafferty J. B. (1998) Nat. Struct. Biol. 5, 441–446 [PubMed]
14. Rafferty J. B., Sedelnikova S. E., Hargreaves D., Artymiuk P. J., Baker P. J., Sharples G. J., Mahdi A. A., Lloyd R. G., Rice D. W. (1996) Science 274, 415–421 [PubMed]
15. Chamberlain D., Keeley A., Aslam M., Arenas-Licea J., Brown T., Tsaneva I. R., Perkins S. J. (1998) J. Mol. Biol. 284, 385–400 [PubMed]
16. Dickman M. J., Ingleston S. M., Sedelnikova S. E., Rafferty J. B., Lloyd R. G., Grasby J. A., Hornby D. P. (2002) Eur. J. Biochem. 269, 5492–5501 [PubMed]
17. Parsons C. A., Stasiak A., Bennett R. J., West S. C. (1995) Nature 374, 375–378 [PubMed]
18. Roe S. M., Barlow T., Brown T., Oram M., Keeley A., Tsaneva I. R., Pearl L. H. (1998) Mol. Cell 2, 361–372 [PubMed]
19. Tsaneva I. R., Illing G., Lloyd R. G., West S. C. (1992) Mol. Gen. Genet. 235, 1–10 [PubMed]
20. Yamada K., Miyata T., Tsuchiya D., Oyama T., Fujiwara Y., Ohnishi T., Iwasaki H., Shinagawa H., Ariyoshi M., Mayanagi K., Morikawa K. (2002) Mol. Cell 10, 671–681 [PubMed]
21. Yu X., West S. C., Egelman E. H. (1997) J. Mol. Biol. 266, 217–222 [PubMed]
22. Seigneur M., Bidnenko V., Ehrlich S. D., Michel B. (1998) Cell 95, 419–430 [PubMed]
23. Michel B., Boubakri H., Baharoglu Z., LeMasson M., Lestini R. (2007) DNA Repair 6, 967–980 [PubMed]
24. Baharoglu Z., Petranovic M., Flores M. J., Michel B. (2006) EMBO J. 25, 596–604 [PMC free article] [PubMed]
25. McGlynn P., Lloyd R. G. (2001) J. Biol. Chem. 276, 41938–41944 [PubMed]
26. Privezentzev C. V., Keeley A., Sigala B., Tsaneva I. R. (2005) J. Biol. Chem. 280, 3365–3375 [PubMed]
27. Mayanagi K., Fujiwara Y., Miyata T., Morikawa K. (2008) Biochem. Biophys. Res. Commun. 365, 273–278 [PubMed]
28. Fujiwara Y., Mayanagi K., Morikawa K. (2008) Biochem. Biophys. Res. Commun. 366, 426–431 [PubMed]
29. Baharoglu Z., Bradley A. S., Le Masson M., Tsaneva I., Michel B. (2008) PLoS Genet. 4, e1000012. [PMC free article] [PubMed]
30. Dunderdale H. J., Sharples G. J., Lloyd R. G., West S. C. (1994) J. Biol. Chem. 269, 5187–5194 [PubMed]
31. Mahdi A. A., Sharples G. J., Mandal T. N., Lloyd R. G. (1996) J. Mol. Biol. 257, 561–573 [PubMed]
32. Whitby M. C., Bolt E. L., Chan S. N., Lloyd R. G. (1996) J. Mol. Biol. 264, 878–890 [PubMed]
33. Le Masson M., Baharoglu Z., Michel B. (2008) Mol. Microbiol. 70, 537–548 [PMC free article] [PubMed]
34. Whitby M. C., Dixon J. (1998) J. Biol. Chem. 273, 35063–35073 [PubMed]
35. Oram M., Keeley A., Tsaneva I. (1998) Nucleic Acids Res. 26, 594–601 [PMC free article] [PubMed]

Articles from The Journal of Biological Chemistry are provided here courtesy of American Society for Biochemistry and Molecular Biology

Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...


Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...