DNA Repeat Rearrangements Mediated by DnaK-Dependent Replication Fork Repair

Molecular Cell 21, 595–604, March 3, 2006 ª2006 Elsevier Inc.

DOI 10.1016/j.molcel.2006.01.025

DNA Repeat Rearrangements Mediated by DnaK-Dependent Replication Fork Repair Stephen J. Goldfless,1,2 Aviv Segal Morag,1 Kurt A. Belisle,1 Vincent A. Sutera, Jr.,1 and Susan T. Lovett1,* 1 Department of Biology and Rosenstiel Basic Medical Sciences Research Center Brandeis University Waltham, Massachusetts 02454

Summary We propose that rearrangements between short tandem repeated sequences occur by errors made during a replication fork repair pathway involving a replication template switch. We provide evidence here that the DnaK chaperone of E. coli controls this template switch repair process. Mutants in dnaK are sensitive to replication fork damage and exhibit high expression of the SOS response, indicative of repair deficiency. Deletion and expansion of tandem repeats that occur by replication misalignment (‘‘slippage’’) are also DnaK dependent. Because mutations in dnaX encoding the g and t subunits of DNA polymerase III mimic dnaK phenotypes and are genetically epistatic, we propose that the DnaKJ chaperone remodels the replisome to facilitate repair. The fork remains largely intact because PriA or PriC restart proteins are not required. We also suggest that the poorly defined RAD6-RAD18-RAD5 mechanism of postreplication repair in eukaryotes occurs by an analogous mechanism to the DnaK template-switch pathway in prokaryotes. Introduction Studies in both prokaryotes and eukaryotes suggest that replication often arrests and requires factors to repair and restart the replication fork. Homologous recombination is one of the mechanisms enlisted to repair the fork. In E. coli, broken forks are processed by RecA and RecBCD; the PriA primosomal complex is required after repair to reassemble the fork helicase and DNA polymerase III replication complex to allow resumption of replication. The partial inviability of recA, recB, and priA mutants suggests that fork breakage occurs in a significant fraction of chromosomal replication events (Capaldo et al., 1974; Nurse et al., 1991). We considered whether other cellular mechanisms, in addition to homologous recombination, were required for maintenance of replication forks. To discover factors, we isolated mutants of E. coli that failed to survive modest inhibition of replication by growth in the presence of hydroxyurea (HU). HU is an inhibitor of ribonucleotide reductase, thereby reducing the cellular levels of deoxynucleotide precursors for DNA replication (Timson, 1975). Sensitivity to azidothymidine (AZT), a nucleotide analog that blocks replication elongation (Elwell et al., 1987), *Correspondence: [email protected] 2 Present address: Metabolix, Inc., 21 Erie Street, Cambridge, Massachusetts 02139.

and to UV irradiation, which produces replication-block pyrimidine dimers, was also tested. A mutant in dnaK was isolated in such a screen and is sensitive to all three treatments. We have proposed that errors during RecA-independent replication fork repair are responsible for the bulk of spontaneous tandem repeat rearrangements in E. coli (Feschenko and Lovett, 1998; Lovett et al., 1993; Morag et al., 1999). The connection between fork repair and rearrangements is apparent from several properties of tandem repeat rearrangements. First, deletion or expansion of tandem repeats occur at high frequencies in the population, much greater than spontaneous mutation, suggesting that rearrangements may be actively mediated, even when the cells lack homologous recombination factors (such as RecA). Second, the majority of tandem repeat deletions take place during or very shortly after replication, consistent with the mechanism of ‘‘replication slippage,’’ involving misalignment of nascent strands during replication (Lovett and Feschenko, 1996). Replication slippage has been proposed to account for rearrangements between repeated DNA sequences in a wide variety of organisms, from bacteria (Albertini et al., 1982) and yeast (Tran et al., 1995) to humans (Efstratiadis et al., 1980). In E. coli, these rearrangements are elevated by certain mutations of the DNA polymerase III holoenzyme, suggesting that they arise as a response to replication difficulties (Saveson and Lovett, 1997). Third, even in RecA2 strains, tandem repeat deletions and expansions are often accompanied by sister chromosome exchange (SCE), deduced by the appearance of circular replisome dimers (Lovett et al., 1993; Morag et al., 1999). This points to the existence of a RecA-independent recombination mechanism between sister chromosomes that could potentially serve as a repair pathway. A crossfork template-switching mechanism (Figure 1) can provide a means for cells to repair blocked replication forks and can give rise to observed rearrangements when repair occurs in the context of tandemly repeated sequences (Figure 2) (Feschenko and Lovett, 1998; Lovett et al., 1993; Morag et al., 1999). Such a mechanism provides an explanation for all the properties of tandem repeat rearrangements, including, among other features, its association with crossing over between sister chromosomes. This sister-strand exchange mechanism (Figure 1), like replication-fork regression (Michel et al., 2004), allows the nascent strands to act as for DNA synthesis for each other; however, fork regression does not lead to SCE as does crossfork switching. However, the importance of a crossfork template-switching mechanism to the cell has remained uncertain, because the genetic basis of this proposed mechanism has been undefined. The properties of the dnaK mutants we report here are consistent with DnaK being a required component of this template-switch mechanism, important for replication fork repair and sustaining cellular viability. DnaK mutants are defective in survival to replication fork inhibition, yield a reduced number of RecA-independent tandem repeat rearrangements, accumulate singlestrand DNA as evident by constitutive SOS induction,

Molecular Cell 596

Figure 1. Proposed Template-Switching Repair Mechanism (A) An arrested replication fork with loaded DnaB helicase (hexagon) and DNA polymerase III complex bound to processivity clamps (ovals). (B) DnaK may be required to dislodge the polymerase to allow the unwinding of nascent strands, while the DnaB fork helicase remains loaded. Unwinding may be aided by a DNA helicase-clamp interaction. (C) Nascent strands pair, forming a Holliday junction, concomitant with a template switch to allow replication of the gapped region. If the switch occurs in a region with directly repeated sequences, mispairing at this step can generate repeat deletion or expansion. (D) Processing of Holliday junction to yield a crossover between sister chromosomes. Alternative processing (data not shown) can yield noncrossover products.

and appear to be synthetically lethal in combination with deficiency in homologous recombination. We propose that DnaK chaperone function is required to remodel the DNA polymerase/helicase complex to permit repair, a hypothesis supported by the phenotypes of a dnaX2016 mutant that mimics dnaK mutant properties and is genetically epistatic. Results Requirement for Chaperone DnaK for Survival to Replication Inhibition and DNA Damage We isolated an insertion mutation in the dnaK gene in a screen for E. coli K-12 mutants that failed to survive modest inhibition of replication by HU. As is characteristic of mutants in dnaK, this insertion mutant grew poorly

Figure 2. Misalignment Mechanisms for Rearrangements and Genetic Assays for Rearrangements (A–D) Simple slippage of the nascent strand (gray lines) on its template (black lines) in the context of directly repeated sequences (gray or black boxes) can yield (A) deletion of direct repeats or (B) expansion of direct repeats. Crossfork misalignment of nascent strands (gray lines) causes a template-switch reaction that can give rise to (C) deletion or (D) expansion of direct repeats. Parental strands (black lines) may also pair, but for simplicity of presentation, this pairing is not shown in these schemes. Because of the potential to generate Holliday junctions (see Figure 1), crossfork slippage can be accompanied by sister chromosome crossing over; simple slippage cannot. (E) Genetic assays used to detect rearrangements between repeated sequences. Plasmid pSTL55 (and its chromosomal integrant in STL695 derivatives) detects deletion between 787 bp tandem repeats (shown as boxes) to yield an intact tetracycline-resistance tetA gene; plasmid pSTL57 similarly detects deletion between tandem 101 bp repeats. Plasmid pEXPBR detects expansion of two 787 bp repeats to a triplication, yielding an intact tetA gene. Plasmids pMB302 and pMB303 assay deletion between 101 bp tetA repeats interspersed by inverted repeats.

at 37ºC and was unable to plate at an elevated temperature of 42ºC. However, even at low growth temperatures (30ºC) permissive for growth, the dnaK mutant was sensitive to low doses of the replication inhibitors HU and AZT (Figure 3), as well as UV (Figure 3) and X irradiation (data not shown). Sensitivity to these agents was fully complemented by a plasmid expressing the dnaK dnaJ operon. We also tested other mutants in dnaK, including a complete deletion of the dnaK open reading frame and a partial loss-of-function mutation, dnaK306. Both

DnaK-Dependent Template Switch Repair 597

Figure 3. Survival Phenotypes of dnaK Mutant Derivatives MG1655 (wild-type), closed circles; STL7155 (dnaK::EZ-Tn5), closed squares; STL8517 (dnaKD), closed triangles; STL8709 (dnaK306), closed diamonds; STL3817 (recA::cat), open circles; STL7155 (dnaK::Tn5)/pBAD33, open squares; STL7155 (dnaK::Tn5)/pBAD33dnaKJ+, open triangles; and STL8747 (dnaK306 recA::cat), open diamonds. (A–C) Fractional survival to various doses of UV (A), HU (B), and AZT (C). Error bars represent standard deviations. (D) Colony morphology on LB medium after overnight growth at 30ºC of the MG1655 derivatives MG1655 (wild-type), STL7180 (recA::cat), STL8709 (dnaK306), and STL8747 (dnak306 recA::cat). (E) Genetic epistasis of dnaK and dnaX. UV survival of MG1655 (wild-type), closed circles; STL8709 (dnaK306), closed diamonds; STL8786 (dnaX2016), open inverted triangles; and STL9645 (dnaK306 dnaX2016), closed inverted triangles.

alleles promoted similar sensitivity to modest replication inhibition or DNA damage (Figures 3A–3C). The dnaK306 point mutation allele affects the various functions of DnaK differentially (Petit et al., 1994; Wild et al., 1992) and, unlike the null allele, has little effect on survival at high temperatures (Wild et al. [1992] and confirmed by us, data not shown). Because sensitivity to these DNA damaging agents was observed independent of growth temperature and affected almost equally by dnaK306, we presume the requirement for DnaK for tolerance to replication inhibition or fork damage is distinct from DnaK’s essential role for growth at high temperature. DnaK Has Synergistic Effects with RecA on Cell Viability and Survival of Replication Inhibition We were unable to stably combine a null allele of dnaK with a mutation in recA, required for genetic recombination and induction of the SOS response. Transductants could be obtained but lost viability and became suppressed after further cultivation. This suggested that

complete loss of both the DnaK and RecA was lethal to the cell. We were, however, able to combine the hypomorphic allele dnaK306 and a null allele of recA: as previously shown (Bredeche et al., 2001), this combination was slow growing and poorly viable (Figure 3D). This double mutant exhibits more extreme sensitivity to HU and AZT than either single mutant (Figures 3B and 3C). This genetic additivity suggests that DnaK controls a response to replication inhibition and DNA damage that functions independently of RecA-dependent genetic recombination and the SOS response. RecA-Independent Rearrangements Are Dependent on DnaK Because of the proposed connection between replication fork repair and tandem repeat rearrangements, we asked whether DnaK was required for recovery of tandem repeat rearrangements. Indeed, DnaK had strong effects on deletion or expansion rearrangements between repeated DNA sequences in our genetic assays

Molecular Cell 598

Table 1. Rates of Rearrangements of 101 bp and 787 bp tetA Tandem Repeats on Plasmids Rearrangement Rate 3 1025 (CI) Strain Genotype

Plasmid pSTL57 101 bp Repeat Deletion

Plasmid pSTL55 787 bp Repeat Deletion

Plasmid pEXPBR 787 bp Repeat Expansion

Wild-type (MG1655) dnaK::EZ-Tn5 dnaKD dnaK306 recA::cat dnaK306 recA::cat

4.4 (2.6–7.0) 0.50 (0.16–1.0) 0.20 (0.089–6.6) 0.34 (0.12–1.1) 4.1 (2.9–7.1) 0.16 (0.046–0.75)

17 (10–36) 15 (12–25) 21 (15–36) 24 (7.9–51) 7.2 (3.2–2) 0.76 (0.29–2.1)

4.4 (2.4–7.4) ND ND 9.7 (4.0–17) 0.58 (0.2–0.76) 0.028 (0.018–0.088)

Rearrangement rates determined with 12–30 independent cultures by method of the median (Lea and Coulson, 1949) with 95% confidence intervals (CIs) (Saveson and Lovett, 1997). Abbreviation: ND, not determined.

(Figure 2E), consistent with the hypothesis that dnaK is required specifically for the recA-independent replication misalignment pathway (Table 1). Mutations in dnaK reduced deletion between 101 repeats carried on low-copy plasmids about 10-fold: we have previously shown that deletion of these short repeats occurs by misalignment during replication, independent of RecA (Lovett and Feschenko, 1996). DnaK had little or no effect on deletion of 787 bp tandem repeats, which rearrange primarily via RecA-dependent recombination. However, when recA was mutated, deletion of the larger 787 bp repeats became DnaK dependent. A similar effect of dnaK on expansion of 787 bp repeats was seen: DnaK function was required for the RecA-independent, but not RecA-dependent, mechanism that produces expansion rearrangements (Table 1). Gel electrophoretic analysis of pSTL55 plasmid deletion products from the recA derivatives indicated that formation of both monomer and dimer products was equally dependent on dnaK. For the recA::cat strain, 11/16 deletion products were replicon dimers, indicative of sister chromosome crossing over, giving a rate of 5.0 3 1025 for deletion associated with SCE and 2.2 3 1025 for deletion without SCE; for the dnaK306 recA::cat strain, 11/16 products were dimeric, giving a rate of 5.2 3 1026 for deletion associated with SCE and 2.4 3 1026 for deletion without SCE. Because DnaK appears to be required for rearrangements with and without associated crossing over, DnaK may facilitate both ‘‘simple slippage’’ (Figures 2A and 2B) that can yield only monomer products in our assays and ‘‘crossfork slippage’’ (Figures 2C and 2D) that can yield either monomer or dimer products, depending on the resolution of the Holliday junction produced by this mechanism. (Figure 1 illustrates resolution to the crossover product). DnaK was also found to be required for deletion of 787 bp repeats on the E. coli chromosome. In contrast to the same repeats carried on plasmids, deletion of the chromosomal repeats is not detectably RecA dependent (Table 2 and Lovett et al. [1993] and Saveson and Lovett [1997]). Deletion of the chromosome repeats, however, was strongly dependent on dnaK, reduced over 20-fold by dnaK306 or dnaKD (Table 2). Why the DnaK pathway operates preferentially over the RecA pathway for deletion of chromosomal, but not for plasmid-borne repeats, is not clear but could reflect differences in replication, cellular localization, or chromosome structure. Deletion of the chromosomal repeats is similarly reduced by mutations in the DNA polymerase processivity clamp and

the clamp loader (Saveson and Lovett, 1997), implicating a role for the b clamp in DnaK-dependent templateswitch repair. In contrast, mutations in many other subunits of the replisome lead to markedly elevated rates of tandem repeat deletion (Saveson and Lovett, 1997). Deletion of chromosomal repeats did not require either of the primosomal proteins PriA or PriC (Table 2), suggesting that deletion occurs without the need for the known mechanisms to reload DnaB helicase (Heller and Marians, 2005). In fact, the priA recA mutant was hyperdeletionogenic, as previously observed for the recA+ priA derivative (Saveson and Lovett, 1997). We have proposed that uncoupling of leading and lagging strand synthesis in priA mutants may account for this increased genetic instability (Lovett, 2003). Secondary Structure Induced-Replication Slippage, but Not Single-Strand Annealing after Chromosome Breakage, Requires DnaK When an inverted repeat sequence, capable of secondary structure formation, is placed between direct repeats, deletion is highly stimulated by two genetically distinct mechanisms (Bzymek and Lovett, 2001a). The primary mechanism is dependent on the DNA secondary structure-specific endonuclease SbcCD and appears to occur via single-strand annealing, initiated by a doublestrand break delivered to cruciform structures formed by the inverted repeats. The secondary mechanism,

Table 2. Rates of Rearrangements of 787 bp tetA Tandem Repeats on the E. coli Chromosome at lac Deletion Rate 3 1026 (CI) Strain Genotype

Chromosomal 787 bp Repeat Deletion

Wild-type (STL695) recAD recA::cat dnaKD dnaK306 dnaX2016 dnaN169 dnaK306 recA::cat dnaK306 dnaX2016 priA recA::cat priC recA::cat

1.6 (1.3–5.3) 2.5 (1.5–6.8) 3.6 (2.5–4.4) 0.085 (0.022–0.10) 0.076 (0.017–0.19) 0.017 (0.011–0.062) 0.064 (0.060–0.11) 0.067 (0.054–0.074) 0.013 (0.0049–0.032) 200 (120–280) 1.8 (1.2–7.8)

Rearrangement rates determined with 9–16 independent cultures grown in LB at 30ºC by method of the median (Lea and Coulson, 1949) with 95% CIs (Saveson and Lovett, 1997). Data for the dnaN169 and dnaX2016 strains are from Saveson and Lovett (1997).

DnaK-Dependent Template Switch Repair 599

Table 3. Deletion Rates of 101 bp tetA Tandem Repeats, with and without Intervening Inverted Repeats that Form Secondary Structures Deletion Rate 3 1025 (CI) Strain Genotype

pSTL57 (no IR)

pMB302 (F14C IR)

pMB303 (F14S IR)

Wild-type (MG1655) sbcD::kan dnaK::EZ-Tn5 dnaK::EZ-Tn5 sbcD::kan

4.4 (2.6–7.0) 2.0 (1.1–9.8) 0.52 (0.26–1.1) 0.39 (0.15–0.60)

130 (80–200) 42 (24–56) 59 (50–84) 2.1 (0.27–7.8)

48 (28–75) 6.5 (4.5–17) 15 (5.0–18) 1.4 (0.7–1.9)

Deletion rates determined with 12–23 independent cultures by method of the median (Lea and Coulson, 1949) with 95% CIs (Saveson and Lovett, 1997).

genetically independent of SbcCD, appears to be replication slippage promoted by hairpin structures on the lagging strand template. Both of these processes occur independent of the RecA homologous recombination system. We tested whether DnaK was required for both RecAindependent mechanisms of deletion or specifically affected the replication misalignment mechanism. If DnaK were to be required only for replication slippage, dnaK would reduce the SbcCD-independent deletion promoted by inverted repeats, with little or no effect on SbcCD-dependent deletion by single-strand annealing. This was indeed the observed outcome. As shown previously (Bzymek and Lovett, 2001a), two different inverted repeats (w50 bp in length) stimulated deletion between flanking 101 bp repeats by 30-fold and 11-fold, respectively (Table 3). And, as before, most of the deletion events stimulated by these inverted repeats required SbcD function. Whereas dnaK has a strong effect on deletion between 101 bp tandem repeats without intervening palindromic sequences, dnaK has little or no effect on deletion between the same 101 bp repeat separated by an inverted repeat sequence. When the SbcCD-dependent annealing pathway was eliminated by mutation of sbcD, the remaining deletion events were more strongly influenced by dnaK: loss of dnaK in a sbcD mutant background reduced deletion from 11- to 30-fold relative to rates in the single sbcD mutant. This provides further evidence that dnaK specifically affects a replication misalignment pathway for genetic rearrangements. Homologous Recombination Is Independent of DnaK Homologous recombination in E. coli is mediated by two main genetic systems distinct in their preference for substrates and in the way they load RecA protein: one requires RecBCD and operates on double-strand ends, whereas the other requires RecFOR and is specialized for recombination at SSB-coated single-strand gaps (Kowalczykowski, 2000). We assayed recombination events mediated by these two genetic systems for their dependence on DnaK. In E. coli, P1 transduction occurs by a RecA/RecBCDdependent pathway (Lovett et al., 1988); we show here that this pathway is not strongly affected by DnaK. The dnaK::EZ-Tn5 insertion mutant was compared to wildtype E. coli MG1655 in assays for transductional inheritance after infection with P1 transducing lysates. In the dnaK mutant grown at 30ºC, inheritance of metE::Tn10, sfsB::Tn10, and metC::Tn10 was 0.30, 0.91, and 0.56, respectively, relative to wild-type MG1655.

In another assay for recombination, crossing over between two plasmids sharing a short amount of sequence homology was determined. We have previous showed that most of these events occur via a RecA/RecFOR-dependent mechanism (Lovett et al., 2002). The dnaK::EZTn5 mutant at 30ºC showed no defect in crossing over between plasmids pSTL330 and pSTL333 (which share 104 bp of homology), with a recombination rate of 1.6 3 1026 per cell generation (n = 19; 95% confidence interval = 1.4–2.5 3 1026) compared to the wild-type rate of 1.5 3 1026 per cell generation (n = 16; 95% confidence interval = 0.72–2.7 3 1026). No defect in this assay was seen even when the dnaK mutant was grown and assayed at 37ºC (data not shown). These assays, combined with the lack of effect of dnaK on RecA-dependent deletion of plasmid-borne 787 bp repeats, confirm that DnaK does not significantly influence RecA-dependent homologous recombination via either RecBCD or RecFOR pathways. Constitutive and Enhanced Induction of the SOS Response in dnaK Mutants If a DnaK-dependent pathway was required to repair blocked replication forks, we reasoned that ssDNA gaps might persist in dnaK mutants and require RecAdependent recombinational processes for their repair. Therefore, dnaK mutants may be constitutively induced for the SOS response, a coordinated set of processes induced by RecA binding to ssDNA, which is a signal of DNA damage (Van Dyk et al., 2001; Walker, 1995). We measured induction of the SOS response with luciferase gene fusions to two SOS-regulated promoters: recA and dinB (Van Dyk et al., 2001). In a dnaK306 mutant, both genes were indeed constitutively induced relative to wild-type strains and showed further enhanced induction after treatment with the replication inhibitor AZT (Figure 4). Control recA mutant strains showed lower constitutive and induced levels of these genes, verifying that AZT treatment does indeed induce a RecA-dependent SOS response. A dnaK Phenotypic Mimic in the DNA Polymerase III g/t Protein The dnaX2016 mutation, like those in dnaK, led to strongly reduced recovery of deletions between chromosomal tandem repeats (Table 2 and Saveson and Lovett [1997]). DnaX encodes via a programmed translation frameshift both g, a component of the clamp loading complex, and t, which coordinates interactions between the components of the replisome itself and with the DnaB replicative helicase (McHenry, 2003). We wondered

Molecular Cell 600

Figure 4. Expression of SOS-Induced Genes, recA, and dinB Luciferase activity was measured in the indicated strain with the use of luciferase gene fusions (Van Dyk et al., 2001), with (gray bars) and without (black bars) 45 min prior treatment with AZT at 1 mg/ml. Arbitrary expression values are derived from the scintillation counts per minute divided by total colony-forming units of the culture. (Note that different scales for luciferase units are used for the two assays because the recA promoter is stronger than that for dinB.) The data represent the mean and standard deviation of three to four experiments.

whether dnaX2016, causing the mutational change glycine 118 to aspartate in both encoded proteins, would manifest other similar phenotypes as dnaK mutants and, if so, whether the genetic effects were epistatic, indicating that they act in the same mechanism. At its permissive temperature for growth, we found that the dnaX2016 mutant was modestly sensitive to UV irradiation (Figure 3E) and to AZT treatment (data not shown). The double dnaX2016 dnaK306 mutant was no more sensitive than the dnaK306 single mutant strain, with a slightly stronger effect than dnaX2016 alone (Figure 3E). In the assay for deletion of 787 bp chromosomal repeats, both dnaX21016 and dnaK306 strongly reduced deletion rates; the double dnaX2016 dnaK306 mutant exhibited reduction similar to the stronger affecting allele dnaX2016 (Table 2). This is consistent with the idea that DnaK and DnaX (g and/or t) act in the same mechanism to mediate genetic rearrangements and to promote survival to DNA damage. Discussion DnaK is E. coli’s Hsp70, an ATP-dependent heat shock chaperone conserved among prokaryotes and eukaryotes (Ang et al., 1991). DnaJ (Hsp40) and GrpE stimulate the ATPase of DnaK, aiding substrate recognition and nucleotide release (Liberek et al., 1991). In the absence of dnaK, E. coli cells fail to survive a heat shock and cannot grow at elevated temperatures because the DnaKJ GrpE chaperone is required to disaggregate and refold thermally misfolded proteins within the cell. In addition to its role in the heat shock response, the DnaK chaperone is required even at low temperatures to remodel specific protein targets. Its name derives from an early observation that dnaK is required for DNA replication of bacteriophage lambda. DnaK is required to release the DnaB helicase held in tight association with the lambda P protein, which along with

lambda O protein is required to escort the helicase to the lambda phage template (Liberek et al., 1988). DnaK is also required for replication of plasmids such as F and P1 (Chattoraj, 1995): it is required to dissociate and/or refold the inactive dimer of their initiation protein factor (RepE and RepA, respectively) to its active monomeric form (Chattoraj et al., 1996; Matsunaga et al., 1997). DnaK, however, is not absolutely required for growth of E. coli at low temperatures and is therefore not essential for replication of E. coli initiated at its origin of replication, oriC. The results presented here provide evidence that the DnaK chaperone is required for survival to replication inhibition and DNA damage. Mutants in dnaK have been previously reported to be sensitive to UV irradiation (Zou et al., 1998). This result was interpreted as an inability to recycle the excision repair complex UvrABC. However, the broad sensitivity of dnaK mutants to agents such as replication inhibitors and g irradiation (not affected by defects in excision repair, data not shown) suggests that DnaK participates in a more global response. The requirement for DnaK for tolerance to replication fork arrest or DNA damage seems to be distinct from DnaK’s role in the heat shock response. We observed temperature-independent defects in survival of null mutants to DNA damage or replication inhibition; furthermore, the dnaK306 allele, with minimal effect on survival at high temperatures (Wild et al., 1992), showed similar properties. A role for DnaK in replication fork repair is consistent with earlier studies showing that replication is less robust or abortive in dnaK mutants (Bukau and Walker, 1989; Ohki and Smith, 1989). A failure to complete replication in a subpopulation of cells can also account for the aberrant chromosome segregation phenotypes of dnaK mutants, observed even at low temperatures (Bukau and Walker, 1989). A previous study by B. Michel and colleagues (Bredeche et al., 2001) is also congruent with our hypothesis that DnaK defines a RecA-independent replication fork repair mechanism. They observed that a mutant in the Rep helicase, which experiences difficulty in the progression of the replication forks (Lane and Denhardt, 1975; Uzest et al., 1995), requires either full function of RecA or DnaK for viability. They concluded that DnaK and RecA-dependent mechanisms acted additively to promote survival in the face of replication difficulties. In our strain background and using null alleles of dnaK, even normal growth appears to require either DnaK or RecA. DnaK was also required to recover RecA-independent genetic rearrangements that have been deduced to occur by misalignment during replication, suggesting that such rearrangements are actively catalyzed by a process requiring DnaK. DnaK function was necessary for a broad set of RecA-independent rearrangements, including those involving loss or gain in the number of tandem repeats, rearrangements with or without associated sister chromosome crossing over, as well as deletion induced by secondary structure elements. DnaK was not found to be essential for rearrangements believed to be elicited by chromosome breakage, such as deletion promoted by SbcCD cleavage of cruciform-forming sequences. DnaK did not affect rearrangements that occur by

DnaK-Dependent Template Switch Repair 601

homologous recombination nor did it affect other recombination events that occur by RecA-dependent pathways. Because RecA-independent rearrangements occur at high frequencies and are often accompanied by sister-chromosome exchange, we have proposed that slippage rearrangements arise during replication fork repair (Feschenko and Lovett, 1998; Lovett et al., 1993; Morag et al., 1999). The phenotypes of dnaK mutants, presented here and elsewhere (Bredeche et al., 2001; Bukau and Walker, 1989; Ohki and Smith, 1989), are entirely consistent with this hypothesis. The DnaK chaperone is therefore the first likely component of this RecAindependent fork repair mechanism and provides us with an important genetic and biochemical tool to investigate this process further. What is the molecular nature of the DnaK fork repair mechanism? Based on numerous genetic properties of tandem repeat rearrangements (reviewed in Bzymek and Lovett [2001b]), we have proposed that a crossfork template-switch reaction (Figure 1) may be employed as the first response to a blocked replication fork. Unwinding of the nascent strands from the blocked fork, with subsequent annealing allows replication to bypass a barrier, potentially without the need to dismantle the fork helicase. In the context of tandem repeats, misalignment can occur during this template switching, giving rise to deletions or expansions in repetitive sequence arrays (Figure 2). In addition, the crossed strands in the repair intermediate (Figure 1) can be processed to yield the crossing over observed between sister chromosomes. This model also explains why, although rearrangements are stimulated by replication difficulties, replication restart factors PriA and PriC are not required, because the strand switching can occur with DnaB helicase still bound to the fork. We also note that this sister-strand pairing mechanism can give rise to the hemicatenated structures observed in eukaryotic replication forks (Lopes et al., 2003; Lucas and Hyrien, 2000), if the 30 ends of the nascent strands return to their parental templates while being still partially interwound with each other. Currently, evidence for such structures in eubacterial replication forks is lacking. A likely explanation for the requirement for DnaK in a replication fork repair mechanism is that its chaperone activity is necessary to remodel the replication complex to allow repair or to promote replication restart after repair. The properties of the dnaX2016 mutation in the g/t proteins of the DNA polymerase III replisome, which mimic dnaK phenotypes, are consistent with DnaK acting on the replisome to promote repair. g is a component of the processivity clamp-loader complex, and t mediates numerous critical interactions between the a subunits of the paired leading- and lagging-strand polymerases, between a and the g complex, and between a and the replication fork helicase DnaB (McHenry, 2003). t is also required to recycle the polymerase, freeing it from the b processivity clamp upon completion of a replication gap, as might occur on the lagging strand (Leu et al., 2003). We do not know the biochemical nature of the dnaX2016 defect, but other temperature-sensitive alleles of dnaX (such as dnaX36) did not exhibit the same phenotype with respect to DNA damage survival (data not shown). The effect of dnaX2016 does not seem to derive simply from difficulties in replication, because

impairment of many other components of the replisome lead to markedly elevated levels of rearrangements rather than decreases in their recovery (Saveson and Lovett, 1997). DnaK has been found to physically associate with components of the replisome, including DNA polymerase III subunits a (dnaE), d (holA), c (holC), and q (holE) (Butland et al., 2005), although the meaning and functional relevance of such interactions is unclear because DnaK also interacts with many diverse cellular proteins. Because DnaK’s participation in replication of bacteriophage and plasmids is uniformly to dissociate stable complexes, we favor the hypothesis that DnaK may loosen some interaction to allow repair to occur. One appealing candidate interaction is of DNA polymerase III with the nascent strand and template, known to be modulated by t (Leu et al., 2003). Biochemical studies have shown that DNA polymerase holoenzyme binds tightly to the 30 nascent terminus as long as single-strand DNA is present within the replication template. Only upon complete replication of a gap, as in completion of an Okazaki fragment, does affinity of polymerase III weaken, allowing recycling of the lagging strand polymerase from the processivity clamp (Naktinis et al., 1996). If synthesis is blocked on the lagging strand, DnaK may be required to free the polymerase from the nascent terminus to allow its realignment with its sister strand. A role in dislodging DNA polymerase III from the b clamp can explain the fact that DnaK is required for polymerase V-dependent mutagenesis (Petit et al., 1994), because polymerase V also competes for association with the b clamp (Tang et al., 1998). Polymerase V is not required for tandem repeat rearrangements (our unpublished data), suggesting that DnaK controls two types of replication fork repair pathways: one ‘‘errorprone’’ involving translesion polymerase UmuCD0 and one relatively ‘‘error-free’’ but susceptible to producing tandem repeat rearrangements. The RAD6/RAD18 pathways of postreplication repair in yeast have strikingly similar features to the DnaK-dependent pathways we propose here. RAD6/18-dependent ubiquitination of the PCNA processivity clamp (Hoege et al., 2002) apparently permits two types of repair: one error-prone, involving recruitment of translesion DNA polymerases z and h, and an error-free pathway, requiring RAD5, MMS2, and UBC13. This latter branch has been proposed to occur by a templateswitching mechanism (Haracska et al., 2004) and is associated with gap-filling repair (Torres-Ramos et al., 2002). Like DnaK in E. coli, Rad5 in yeast is required to recover repeated sequence rearrangements (Johnson et al., 1992). We propose that, in analogy to the DnaK pathway, the Rad5-dependent template switch may involve the cross-strand pairing, as illustrated in Figure 1, rather than the fork regression reaction presently favored (Haracska et al., 2004). The association of SCE with Rad5-dependent processes is not known but is a prediction of the crossfork mechanism. Both prokaryotes and eukaryotes may have evolved distinct mechanisms to remodel the processivity clamp (involving chaperones, for prokaryotes, or protein modification, for eukaryotes) to permit common template-switching and translesion synthesis modes of replication fork repair.

Molecular Cell 602

Experimental Procedures Strains and Plasmids All strains are isogenic with wild-type MG1655 (F- rph-1), except those used for the chromosomal deletion assays employing an integrated tetA duplication, which were derived from STL695 ([Lovett et al., 1993]; F2 lacZ::(bla tetAdup787) hisG4 argE2 leuB6 D(gptproA)62 thr-1 thi-1 rpsL31 galK2 lacY1 ara-14 xyl-5 mtl-1 kdgK51 supE44 tsx-33 rfbD1 mtl-1 racD). Growth medium was Luria-Bertani medium (LB), with 1.5% agar for plates (Miller, 1992), except that 0.4% glucose and 5 mM CaCl2 were added to medium for preparation of P1 lysates or P1 transduction. Cultures were grown at 30ºC unless otherwise noted. Antibiotics were used at the following concentrations: ampicillin (Ap), 100 mg/ml; tetracycline (Tc), 8–15 mg/ml; kanamycin (Km), 30 mg/ml; and chloramphenicol (Cm), 20 mg/ml. Insertion mutants were isolated by electroporation of the EZ-Tn5 transpososome complex (Epicentre, Madison, WI) into MG1655, with subsequent selection for kanamycin resistance, according to methods provided by the vendor. The site of dnaK::Tn5 insertion in original isolate STL7155 was sequenced and determined to occur after nucleotide 1198 of the 1916 bp open reading frame. The dnaK open reading frame was completely deleted in MG1655 by standard methods (Datsenko and Wanner, 2000) and replaced with the kan gene flanked by FRT site-specific recombination sites, producing strain STL8517. All other isogenic strains were constructed by P1virA transduction and include MG1655 derivatives STL8709 (dnaK306 thrA3092::Tn10kan), STL7180 (recA::cat), STL8747 (dnaK306 thrA3092::Tn10kan recA::cat), STL7280 (sbcD::kan), and STL8358 (dnaK::EZ-Tn5 sbcD::kan), STL8786 (dnaX2016 zbb::Tn10), and STL9645 (dnaX2016 zbb::Tn10 dnaK306 thrA3092::Tn10kan). STL695 derivatives in which chromosomal deletion between 787 bp tandem repeats can be measured include STL9297 (dnaK306 thrA3092::Tn10kan), STL9983 (dnaKD::FRT kan), STL9307 (recA::cat), STL9308 (dnaK306 recA::cat), STL10046 (dnaK306 dnaX2016 zaj-2101::Tn10kan), STL4627 (priA2::kan recAD304) and STL46626 (priC303::kan recAD304), and previously published strains STL753 (recAD304), STL1902 (dnaX2016 zaj2101::Tn10kan), and STL1887 (dnaN159 zid-3162::Tn10-kan) (Saveson and Lovett, 1997). Sequence analysis of our original dnaK306 strain, JJC517 (Bredeche et al., 2001; Petit et al., 1994), revealed an A175T mutation, not the T154S mutation reported (Wild et al., 1992), although it displays the phenotypes previously described for this mutant. The dnaKJ operon, including its promoter, was PCR amplified from MG1655 chromosomal DNA with primers 50 -GCA CAAAAAATTTTTGCATCTCCC-30 and 50 -CTGACCAGTTATCGTGA GAGTAAT-30 and Pfu polymerase (Stratagene, Inc) and inserted into SmaI-cut vector pBAD33 (Guzman et al., 1995), producing plasmid pBAD33-DnaKJ+. Survival and Genetic Rearrangement Assays Exponential cultures prepared in LB at 30ºC were challenged with various doses of UV irradiation and/or plated on LB medium containing AZT or HU (Foti et al., 2005). Various rearrangement assay plasmids (Bzymek and Lovett, 2001a; Lovett et al., 1993, 1994; Morag et al., 1999) were introduced into isogenic strains by DNA transformation (Dower et al., 1988). Deletion and expansion assays were performed for 12–30 multiple independent cultures for each plasmidbearing strain or STL695 derivative at 30ºC, as previously described (Lovett et al., 1993). The number of progeny bearing rearrangements was determined by the number of Tc-resistant colonies among the Ap-resistant total population, and a rearrangement rate was derived by using the method of the median (Lea and Coulson, 1949) with 95% confidence intervals (Saveson and Lovett, 1997). Agarose gel electrophoresis of 16 independent plasmid deletion products from various recA mutant strain backgrounds was performed as previously described (Lovett et al., 1993) to determine those events associated with sister chromosome crossing over, leading to plasmid dimer formation. Recombination Assays Quantitative transduction were performed as in Miller (1992) with P1 virA lysates prepared on donor strains (Singer et al., 1989) CAG18491 (metE::Tn10), CAG12072 (sfsB::Tn10), and CAG18475 (metC::Tn10) and recipient strains MG1655 and STL7154 (dnaK::EZ-Tn5) with

a moi of w0.1. The number of transductants arising after 2–3 days on LB-Tc was counted and normalized to the number of viable cells in the recipient cultures, determined by microscopic examination of at least 400 individual cells after Live/Dead staining (Invitrogen, Inc.). Crossing over between plasmids pSTL330 and pSTL333 was measured as previously described (Lovett et al., 2002), selecting Tc resistance among the Ap Cm-resistant plasmid-bearing population. Recombination rates were determined by method of the median (Lea and Coulson, 1949) for 16–19 independent cultures and the 95% confidence interval determined as previously described (Saveson and Lovett, 1997). SOS Promoter Assays Plasmids carrying Photorhabdus luminescens luxCDABE fusions to the E. coli recA (pDEW238) or dinB promoters (pDEW236) (Van Dyk et al., 2001) were transformed into the appropriate strains, selecting Ap resistance. At least three exponentially growing cultures in LB, with and without 45 min prior treatment with 1 mg/ml AZT, were assayed for bioluminescence in a Wallac 1409 liquid scintillation counter. Arbitrary luciferase expression values were calculated by dividing the amount of bioluminescence (cpm) by the total number of viable cells determined from serially diluted cells plated on LB + Ap. Acknowledgments We thank M. Berlyn (E. coli Genetic Stock Center), B. Michel (INRA, France), S. Sandler (U. of Massachusetts, Worcester), and J. Walker (U. of Texas, Austin) for strains and J. Schienda and N. Thomas for early work on the dnaK mutant. This work was supported by National Institutes of Health grant RO1 GM51753 and a Howard Hughes Medical Institute undergraduate summer internship program to S.J.G. Received: December 13, 2005 Revised: January 11, 2006 Accepted: January 17, 2006 Published: March 2, 2006 References Albertini, A.M., Hofer, M., Calos, M.P., and Miller, J.H. (1982). On the formation of spontaneous deletions: the importance of short sequence homologies in the generation of large deletions. Cell 29, 319–328. Ang, D., Liberek, K., Skowyra, D., Zylicz, M., and Georgopoulos, C. (1991). Biological role and regulation of the universally conserved heat shock proteins. J. Biol. Chem. 266, 24233–24236. Bredeche, M.F., Ehrlich, S.D., and Michel, B. (2001). Viability of rep recA mutants depends on their capacity to cope with spontaneous oxidative damage and on the DnaK chaperone protein. J. Bacteriol. 183, 2165–2171. Bukau, B., and Walker, G.C. (1989). Delta dnaK52 mutants of Escherichia coli have defects in chromosome segregation and plasmid maintenance at normal growth temperatures. J. Bacteriol. 171, 6030–6038. Butland, G., Peregrin-Alvarez, J.M., Li, J., Yang, W., Yang, X., Canadien, V., Starostine, A., Richards, D., Beattie, B., Krogan, N., et al. (2005). Interaction network containing conserved and essential protein complexes in Escherichia coli. Nature 433, 531–537. Bzymek, M., and Lovett, S.T. (2001a). Evidence for two mechanisms of palindrome-stimulated deletion in Escherichia coli: single-strand annealing and replication slipped mispairing. Genetics 158, 527– 540. Bzymek, M., and Lovett, S.T. (2001b). Instability of repetitive DNA sequences: the role of replication in multiple mechanisms. Proc. Natl. Acad. Sci. USA 98, 8319–8325. Capaldo, F.N., Ramsey, G., and Barbour, S.D. (1974). Analysis of the growth of recombination-deficient strains of Escherichia coli K-12. J. Bacteriol. 118, 242–249. Chattoraj, D.K. (1995). Role of molecular chaperones in the initiation of plasmid DNA replication. Genet. Eng. (N.Y.) 17, 81–98. Chattoraj, D.K., Ghirlando, R., Park, K., Dibbens, J.A., and Lewis, M.S. (1996). Dissociation kinetics of RepA dimers: implications for

DnaK-Dependent Template Switch Repair 603

mechanisms of activation of DNA binding by chaperones. Genes Cells 1, 189–199. Datsenko, K.A., and Wanner, B.L. (2000). One-step inactivation of chromosomal genes in Escherichia coli K-12 using PCR products. Proc. Natl. Acad. Sci. USA 97, 6640–6645. Dower, W.J., Miller, J.F., and Ragsdale, C.W. (1988). High efficiency transformation of E. coli by high voltage electroporation. Nucleic Acids Res. 16, 6127–6145. Efstratiadis, A., Posakony, J.W., Maniatis, T., Lawn, R.M., O’Connell, C., Spritz, R.A., DeRiel, J.K., Forget, B.G., Weissman, S.M., Slightom, J.L., et al. (1980). The structure and evolution of the human beta-globin gene family. Cell 21, 653–668. Elwell, L.P., Ferone, R., Freeman, G.A., Fyfe, J.A., Hill, J.A., Ray, P.H., Richards, C.A., Singer, S.C., Knick, V.B., Rideout, J.L., et al. (1987). Antibacterial activity and mechanism of action of 30 -azido-30 -deoxythymidine (BW A509U). Antimicrob. Agents Chemother. 31, 274–280. Feschenko, V.V., and Lovett, S.T. (1998). Slipped misalignment mechanisms of deletion formation: analysis of deletion endpoints. J. Mol. Biol. 276, 559–569. Foti, J.J., Schienda, J., Sutera, V.A., Jr., and Lovett, S.T. (2005). A bacterial G protein-mediated response to replication arrest. Mol. Cell 17, 549–560. Guzman, L.M., Belin, D., Carson, M.J., and Beckwith, J. (1995). Tight regulation, modulation, and high-level expression by vectors containing the arabinose PBAD promoter. J. Bacteriol. 177, 4121–4130. Haracska, L., Torres-Ramos, C.A., Johnson, R.E., Prakash, S., and Prakash, L. (2004). Opposing effects of ubiquitin conjugation and SUMO modification of PCNA on replicational bypass of DNA lesions in Saccharomyces cerevisiae. Mol. Cell. Biol. 24, 4267–4274. Heller, R.C., and Marians, K.J. (2005). The disposition of nascent strands at stalled replication forks dictates the pathway of replisome loading during restart. Mol. Cell 17, 733–743. Hoege, C., Pfander, B., Moldovan, G.L., Pyrowolakis, G., and Jentsch, S. (2002). RAD6-dependent DNA repair is linked to modification of PCNA by ubiquitin and SUMO. Nature 419, 135–141. Johnson, R.E., Henderson, S.T., Petes, T.D., Prakash, S., Bankmann, M., and Prakash, L. (1992). Saccharomyces cerevisiae RAD5-encoded DNA repair protein contains DNA helicase and zinc-binding sequence motifs and affects the stability of simple repetitive sequences in the genome. Mol. Cell. Biol. 12, 3807–3818. Kowalczykowski, S.C. (2000). Initiation of genetic recombination and recombination-dependent replication. Trends Biochem. Sci. 25, 156–165. Lane, H.E., and Denhardt, D.T. (1975). The rep mutation. IV. Slower movement of replication forks in Escherichia coli rep strains. J. Mol. Biol. 97, 99–112. Lea, D.E., and Coulson, C.A. (1949). The distribution of the numbers of mutants in bacterial populations. J. Genet. 49, 264–285. Leu, F.P., Georgescu, R., and O’Donnell, M. (2003). Mechanism of the E. coli tau processivity switch during lagging-strand synthesis. Mol. Cell 11, 315–327. Liberek, K., Georgopoulos, C., and Zylicz, M. (1988). Role of the Escherichia coli DnaK and DnaJ heat shock proteins in the initiation of bacteriophage lambda DNA replication. Proc. Natl. Acad. Sci. USA 85, 6632–6636. Liberek, K., Marszalek, J., Ang, D., Georgopoulos, C., and Zylicz, M. (1991). Escherichia coli DnaJ and GrpE heat shock proteins jointly stimulate ATPase activity of DnaK. Proc. Natl. Acad. Sci. USA 88, 2874–2878. Lopes, M., Cotta-Ramusino, C., Liberi, G., and Foiani, M. (2003). Branch migrating sister chromatid junctions form at replication origins through Rad51/Rad52-independent mechanisms. Mol. Cell 12, 1499–1510. Lovett, S.T. (2003). Connecting replication and recombination. Mol. Cell 11, 554–556. Lovett, S.T., and Feschenko, V.V. (1996). Stabilization of diverged tandem repeats by mismatch repair: evidence for deletion formation via a misaligned replication intermediate. Proc. Natl. Acad. Sci. USA 93, 7120–7124.

Lovett, S.T., Luisi-DeLuca, C., and Kolodner, R.D. (1988). The genetic dependence of recombination in recD mutants of Escherichia coli. Genetics 120, 37–45. Lovett, S.T., Drapkin, P.T., Sutera, V.A., Jr., and Gluckman-Peskind, T.J. (1993). A sister-strand exchange mechanism for recA-independent deletion of repeated DNA sequences in Escherichia coli. Genetics 135, 631–642. Lovett, S.T., Gluckman, T.J., Simon, P.J., Sutera, V.A., Jr., and Drapkin, P.T. (1994). Recombination between repeats in Escherichia coli by a recA-independent, proximity-sensitive mechanism. Mol. Gen. Genet. 245, 294–300. Lovett, S.T., Hurley, R.L., Sutera, V.A., Jr., Aubuchon, R.H., and Lebedeva, M.A. (2002). Crossing over between regions of limited homology in Escherichia coli. RecA-dependent and RecA-independent pathways. Genetics 160, 851–859. Lucas, I., and Hyrien, O. (2000). Hemicatenanes form upon inhibition of DNA replication. Nucleic Acids Res. 28, 2187–2193. Matsunaga, F., Ishiai, M., Kobayashi, G., Uga, H., Yura, T., and Wada, C. (1997). The central region of RepE initiator protein of mini-F plasmid plays a crucial role in dimerization required for negative replication control. J. Mol. Biol. 274, 27–38. McHenry, C.S. (2003). Chromosomal replicases as asymmetric dimers: studies of subunit arrangement and functional consequences. Mol. Microbiol. 49, 1157–1165. Michel, B., Grompone, G., Flores, M.J., and Bidnenko, V. (2004). Multiple pathways process stalled replication forks. Proc. Natl. Acad. Sci. USA 101, 12783–12788. Miller, J.H. (1992). A Short Course in Bacterial Genetics (New York: Cold Spring Harbor Press). Morag, A.S., Saveson, C.J., and Lovett, S.T. (1999). Expansion of DNA repeats in Escherichia coli: effects of recombination and replication functions. J. Mol. Biol. 289, 21–27. Naktinis, V., Turner, J., and O’Donnell, M. (1996). A molecular switch in a replication machine defined by an internal competition for protein rings. Cell 84, 137–145. Nurse, P., Zavitz, K.H., and Marians, K.J. (1991). Inactivation of the Escherichia coli priA DNA replication protein induces the SOS response. J. Bacteriol. 173, 6686–6693. Ohki, M., and Smith, C.L. (1989). Tracking bacterial DNA replication forks in vivo by pulsed field gel electrophoresis. Nucleic Acids Res. 17, 3479–3490. Petit, M.A., Bedale, W., Osipiuk, J., Lu, C., Rajagopalan, M., McInerney, P., Goodman, M.F., and Echols, H. (1994). Sequential folding of UmuC by the Hsp70 and Hsp60 chaperone complexes of Escherichia coli. J. Biol. Chem. 269, 23824–23829. Saveson, C.J., and Lovett, S.T. (1997). Enhanced deletion formation by aberrant DNA replication in Escherichia coli. Genetics 146, 457– 470. Singer, M., Baker, T.A., Schnitzler, G., Deischel, S.M., Goel, M., Dove, W., Jaacks, K.J., Grossman, A.D., Erickson, J.W., and Gross, C.A. (1989). A collection of strains containing genetically linked alternating antibiotic resistance elements for genetic mapping of Escherichia coli. Microbiol. Rev. 53, 1–24. Tang, M., Bruck, I., Eritja, R., Turner, J., Frank, E.G., Woodgate, R., O’Donnell, M., and Goodman, M.F. (1998). Biochemical basis of SOS-induced mutagenesis in Escherichia coli: reconstitution of in vitro lesion bypass dependent on the UmuD’2C mutagenic complex and RecA protein. Proc. Natl. Acad. Sci. USA 95, 9755–9760. Timson, J. (1975). Hydroxyurea. Mutat. Res. 32, 115–132. Torres-Ramos, C.A., Prakash, S., and Prakash, L. (2002). Requirement of RAD5 and MMS2 for postreplication repair of UV-damaged DNA in Saccharomyces cerevisiae. Mol. Cell. Biol. 22, 2419–2426. Tran, H.T., Degtyareva, N.P., Koloteva, N.N., Sugino, A., Masumoto, H., Gordenin, D.A., and Resnick, M.A. (1995). Replication slippage between distant short repeats in Saccharomyces cerevisiae depends on the direction of replication and the RAD50 and RAD52 genes. Mol. Cell. Biol. 15, 5607–5617. Uzest, M., Ehrlich, S.D., and Michel, B. (1995). Lethality of rep recB and rep recC double mutants of Escherichia coli. Mol. Microbiol. 17, 1177–1188.

Molecular Cell 604

Van Dyk, T.K., Wei, Y., Hanafey, M.K., Dolan, M., Reeve, M.J., Rafalski, J.A., Rothman-Denes, L.B., and LaRossa, R.A. (2001). A genomic approach to gene fusion technology. Proc. Natl. Acad. Sci. USA 98, 2555–2560. Walker, G.C. (1995). SOS-regulated proteins in translesion DNA synthesis and mutagenesis. Trends Biochem. Sci. 20, 416–420. Wild, J., Kamath-Loeb, A., Ziegelhoffer, E., Lonetto, M., Kawasaki, Y., and Gross, C.A. (1992). Partial loss of function mutations in DnaK, the Escherichia coli homologue of the 70-kDa heat shock proteins, affect highly conserved amino acids implicated in ATP binding and hydrolysis. Proc. Natl. Acad. Sci. USA 89, 7139–7143. Zou, Y., Crowley, D.J., and Van Houten, B. (1998). Involvement of molecular chaperonins in nucleotide excision repair. Dnak leads to increased thermal stability of UvrA, catalytic UvrB loading, enhanced repair, and increased UV resistance. J. Biol. Chem. 273, 12887–12892.