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Chapter 16 : Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance

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Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, Page 1 of 2

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Abstract:

This chapter briefly reviews the functions and properties of key proteins involved in homologous recombination, with a particular emphasis on RecA, a versatile protein with additional roles in the induction of the SOS response and in translesion DNA synthesis and mutagenesis. Parallels can be drawn between these bacterial recombination proteins and the eukaryotic recombination proteins. Evidence supporting the existence of recombinational repair of DNA in bacteria is discussed, followed by the developing views of how recombination proteins can help cells deal with replication forks whose progress has been blocked by endogenous or exogenous DNA damage. Although many discussions of recombinational repair, replication fork repair, and other mechanisms of DNA damage tolerance consider only the formal set of simple DNA structures, it is important to keep in mind the numerous observations indicating that the DNA structures present in vivo after DNA damage may be considerably more complex. It is important to recognize that the various replication fork recovery strategies involving homologous recombination functions are not necessarily independent of translesion DNA polymerases and that such translesion DNA polymerases may participate in recombinogenic replication fork recovery strategies at times. Much remains to be learned about the relationship between homologous recombination functions, replication fork repair/recovery, and translesion synthesis.

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16

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Image of Figure 16–1
Figure 16–1

DNA damage can interfere with the progress of replication forks and lead to the generation of various structures. (A) DSB can be created directly by ionizing radiation and certain other agents. (B and C) A DSB can be generated by a replication fork encountering a nick in the leading-strand template (B) or by a replication fork encountering a nick in the lagging-strand template (C) (note that the DSB could have a 3’ single-strand overhang). (D and E) A single-strand gap can be generated by a replication fork encountering a lesion in the leading-strand (D) or lagging-strand (E) template. (F and G) Replication forks can regress on encountering a lesion blocking both strands (F) or just one strand (G) to form a particular fork of Holliday junction commonly referred to as a chicken foot structure.

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–2
Figure 16–2

Model of the replication fork. In the replicative helicase DnaB acts to disrupt parental DNA and expose the two individual strands. One of these strands acts as the template for synthesis of the leading strand by a DNA Pol III complex, which proceeds uninterrupted in the 5’ → 3’ direction (arrows represent the 3’ ends of DNA strands). The anchoring of the leading-strand polymerase on this template strand by the β sliding clamp allows synthesis of the leading strand to continue for many thousands of bases. By contrast, because DNA synthesis occurs in the 5 ‘ — 3 ‘ direction, synthesis of the second DNA strand proceeds in segments using RNA primers made by the primase DnaG. This allows DNA synthesis to repeatedly initiate on the lagging-strand template. Thus, the lagging-strand-DNA Pol III complex continually associates and dissociates with the lagging-strand template to extend each RNA primer and form so-called Okazaki fragments. These Okazaki fragments are 1,000 to 2,000 bp in bacteria but only 40 to 300 bp in eukaryotes. β sliding clamps that are associated with the lagging-strand polymerase are also reloaded continually onto the lagging-strand template by the γ-complex clamp loader. A single continuous strand is formed from these discontinuous lagging strands by degradation of the RNA primers and the subsequent filling in of the resultant gaps by DNA Pol I followed by ligation of the 5’ end of one Okazaki fragment with the 3’ end of the adjacent fragment. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–3
Figure 16–3

Model of Okazaki fragment synthesis by a replication fork encountering an AP lesion on the leading-strand template. Thin grey lines indicate the template strand DNA. Dark gold and light gold lines with arrowheads indicate leading- and lagging-strand DNA synthesis, respectively. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–4
Figure 16–4

Effect of UV irradiation on the rate of DNA synthesis (“induced replisome reactivation/replication restart”). An exponentially growing culture of wild-type strain AB1157 was UV irradiated (10 J/m); 0.5 ml of culture was removed at intervals and pulse-labeled with [H]thymidine for 2 min, and trichloroacetic acid-precipitable counts were determined. Gold line, [H]thymidine cpm; black line, optical density at 450 nm (A). (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–5
Figure 16–5

Alkaline sedimentation profiles of labeled DNA showing postreplication repair and excision repair occurring in wild-type following UV irradiation. Gold lines indicate DNA uniformly labeled with [C]thymidine; black lines indicate DNA pulse-labeled with [H]thymidine. (A) A thymidine-requiring repair-proficient strain of K-12 was uniformly labeled with [C]thymidine, washed, pulse-labeled with [H]thymidine for 10 min, and then lysed for sedimentation in an alkaline sucrose gradient. (B) Same as panel A except that the C-labeled bacteria were exposed to 254-nm UV light at 6 J/m and then pulse-labeled with [H]thymidine for 10 min. (C) Same as panel B except that the pulse-labeled cells were washed and incubated in unlabeled medium for 50 min. (D) Bacteria were uniformly labeled with [C]thymidine, washed, exposed to UV light at 6 J/m, incubated for 60 min, and then pulse-labeled with [H]thymidine for 10 min. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–6
Figure 16–6

Comparison of structural domains in the RecA protein with the T4 UvsX, yeast (S. Rad51 and Dmc1, and archaeal RadA proteins. Core domain conservation is depicted in light gold. N-terminal domain conservation between Rad51, Dmc1, and RadA is shown in dark gold. Regions with no sequence homology include the light grey regions of the N-terminal domains and all regions C-terminal to the core, including the medium gold C-terminal domains of the RecA and T4 UvsX proteins. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–7
Figure 16–7

Examples of recombination reactions promoted by RecA protein. (A) Renaturation of complementary single strands; (B) asymmetric (nonreciprocal) strand exchange following the pairing of single-stranded and double-stranded DNA; (C) symmetric (reciprocal) strand exchange following the pairing of duplex DNA and partially single-stranded DNA. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–8
Figure 16–8

Electron micrographs of frozen-hydrated (A and B) and negatively stained (C) RecA-DNA-ATP-γS filaments. The RecA filaments in panels A and C are on double-stranded DNA, while the RecA in panel B is on a circular single-stranded DNA molecule. The white arrows in panel A show the general direction of fluid flow that occurred on the grid during blotting, prior to rapid freezing, as judged by the preferred orientation of filaments in this direction on the grid. The pitch of the filament sections between the white arrows are 114 and 108 Å (11.4 and 10.8 nm), while the pitch of the filament section that runs perpendicular to the flow direction (black arrow) is 96 A (9.6 nm). All panels are at the same magnification, and the tobacco mosaic virus particles in panel C are about 200 A (20 nm) in diameter. (Reproduced from reference with permission.)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–9
Figure 16–9

Model for the RecFOR-facilitated formation of a RecA nucleoprotein filament on gapped DNA. (A) The RecQ helicase and RecJ exonuclease might process the gapped DNA prior to recognition by the RecFOR proteins. (B and C) The SSB-coated gap (B) is first recognized by the RecFR complex (or RecF) (C). (D) The RecOR complex (or RecO) interacts with the RecFR-gapped DNA complex. (E) The RecFOR proteins serve to nucleate RecA protein filament assembly, which then extends over the single-strand region by growth in the 5’ → 3’ direction, displacing SSB. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–10
Figure 16–10

RecA filament. Two views of the RecA protein filament (three subunits are shown) crystallized in the absence of DNA ( ). ADP binds to the central domain of RecA near the subunit interface. An N-terminal α-helix of each subunit packs against the neighboring subunit, and the C-terminal domain protrudes from the inner radius of the helical filament.

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–11
Figure 16–11

Model for homologous pairing within a RecA nucleoprotein filament. (Courtesy of S. C. West.)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–12
Figure 16–12

General model for homologous recombination that depicts the DSB repair model suggested by Szostak et al. ( ). The text in parentheses designates proteins from that function at the designated steps. Light grey lines indicate newly synthesized DNA. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–13
Figure 16–13

Early steps of homologous recombination in are coordinated by RecA protein, RecBCD enzyme, and χ. RecBCD enzyme binds to the end of a DSB. It unwinds the double-stranded DNA while preferentially degrading the strand that was 3’ terminal at the entry point. Interaction with a DNA sequence known as χ results in attenuation of the 3’ → 5’ nuclease activity, activation of a weaker 5’ → 3’ nuclease activity, and facilitated loading of RecA protein onto the χ-containing single-stranded DNA that was produced by continued DNA unwinding beyond x. The resulting RecA protein-single-stranded DNA filament invades homologous double-stranded DNA (dsDNA) to produce a D-loop structure. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–14
Figure 16–14

Resolving a chicken foot Holliday junction generates a DSB. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–15
Figure 16–15

RuvA and RuvB function as a branch migration motor that remodels Holliday junctions. (A) A tetramer of RuvA (two gold subunits and two white subunits) binds to one face of a four-way DNA junction (Holliday junction; colored black) in a square planar conformation ( ). A second RuvA tetramer (not shown) binds to the opposite face of the DNA junction in a clam shell-like arrangement. (B) Side view of the RuvA-DNA complex shown in panel A. (C) Domain 3 of RuvA (disordered in the crystal structure shown in panel A) interacts with the RuvB helicase. RuvB has an AAA ATPase protein fold ( ). ATP (gold) binds in a cleft between the N-and C-terminal domains of RuvB. (D) Model showing the locations of the RuvB hexamer and the RuvA octamer during branch migration of a four-way DNA junction. RuvA manages the exchange of DNA strands at the center of the junction. RuvB is the motor that translocates double-stranded DNA from two branches of the four-way junction to cause branch migration. Domain 3 (small sphere) of RuvA mediates the interaction with RuvB (shown in panel C).

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–16
Figure 16–16

RecG is a replication fork-remodeling enzyme. The crystal structure of RecG bound to a fork-shaped DNA suggests a mechanism for reversal of a stalled replication fork ( ). Domains 2 and 3 of RecG are typical of a superfamily 1 helicase, with ATP (black) binding in the interdomain cleft. Domain 1 manages the DNA substrate, interacting with two double-stranded DNA branches and inserting a “wedge” at the junction to promote strand separation during translocation of the DNA. In the proposed mechanism, the dsDNA marked would be pulled toward helicase domains 2 and 3 to cause remodeling of a stalled replication fork.

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–17
Figure 16–17

Model for daughter strand gap repair. The thick black lines represent irradiated parental DNA containing a cyclobutane dimer. The thin gold lines represent daughter DNA synthesized after irradiation. The Holliday junction can be resolved in one of two ways. The cleavages marked “a” result in an exchange event involving the parental strand without a dimer ( ); the dimer stays in the parental strand. The cleavages marked “b” lead to an exchange event involving the parental strand with the dimer; the dimer is exchanged into the daughter strand (Adapted from references , and .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–18
Figure 16–18

cells that are proficient in both excision repair (uvrA) and discontinuous DNA synthesis with gap filling (recA) are resistant to killing by UV radiation. Cells that are defective in both of these processes are extremely UV sensitive. The survival curve of the double mutant is reproduced on the left with an expanded UV dose scale. From this curve, it can be estimated that one pyrimidine dimer per genome equivalent is lethal to a strain. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–19
Figure 16–19

DNA synthesized immediately after UV irradiation of has a lower molecular weight than normal. cells were exposed to UV radiation and pulse-labeled briefly with [H]thymidine. The cells were analyzed by sedimentation in alkaline sucrose gradients either immediately after being labeled (no incubation; dotted gold line) or following incubation for 70 min (solid gold line). Immediately following the pulse-label, the newly synthesized DNA is of low molecular weight and sediments near the top of the gradient. However, over time the newly synthesized DNA approaches the size of the unirradiated control (no UV; black line). (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–20
Figure 16–20

(A) Relative yield of Lac colonies after an excision-defective Flac strain was mated to mutant recipients. When the recipient is defective in the gene, the yield of Lac colonies is significantly reduced relative to that observed in a wild-type recipient. However, there is no detectable difference in the yield of Lac colonies in recipients that are either excision repair proficient or excision repair deficient (B) FLac DNA transferred from a UV-irradiated donor to an unirradiated recipient shows enhanced survival if, following transfer, the cells are exposed to photoreactivating light. This result is consistent with (but does not prove) the presence of replicative gaps opposite pyrimidine dimers. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–21
Figure 16–21

(A) Design and interpretation of an experiment ( ) to detect exchanges between DNA strands during postreplicative gap filling. cells were grown for several generations in medium containing the heavy isotopes [C]thymine and [N]thymine as well as [C]thymine. The cells were then exposed to various doses of UV light and grown for less than one generation in a medium without density markers (light medium) containing [H]thymidine. The newly replicated DNA is of hybrid density and has the C label uniquely in the heavy strand and the H label uniquely in the light strand. (B) If the cells are not irradiated and their DNA strands are separated by heat denaturation and equilibrium CsCl density gradient centrifugation, the strands separate cleanly as C-labeled heavy strands and H-labeled light strands. (C) If the cells are irradiated, strands of intermediate density are observed after heat denaturation and equilibrium CsCl density gradient centrifugation. This observation has been interpreted as indicating that exchanges have occurred between sister duplexes in the UV-irradiated cells so that light (H-labeled) DNA becomes covalently attached to heavy (C-labeled) DNA. This DNA of intermediate density could be resolved into heavy and light components after shearing to a molecular weight of less than 5 × 10, suggesting that the exchanges involved segments of at least this size.

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–22
Figure 16–22

Model assigning possible proteins to steps in daughter strand gap repair. See the text for details.

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–23
Figure 16–23

Both discontinuous replication and stalled replication generate small nascent DNA molecules that subsequently are converted into high-molecular-weight ones. Thus, it can be very difficult, if not impossible, to differentiate these processes by using the sedimentation velocity of radiolabeled newly synthesized DNA. The thicker lines in the lower left diagram indicate regions of gap filling by recombinational exchange and by nonsemiconservative DNA synthesis (see Fig. 16–17 ).

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–24
Figure 16–24

Schematic depiction of possible relationships of DNA lesions to replication forks in Replication forks A1 and A2 were established during the initiation of an initial round of DNA replication. Replication forks B1, B2, B1 ‘, and B2’ were established before the completion of this initial round of DNA replication. Lesions 1 and 2 differ with respect to their positions relative to these replication forks. See the text for details. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–25
Figure 16–25

Model showing how replication of DNA by a factory combined with subcellular localization of the DNA after synthesis could limit the time during which the two daughter strands could undergo recombinational repair. After DNA synthesis, the DNA becomes localized to the incipient daughter cells, so that the two daughter DNAs would not be able to undergo recombinational interactions although the two daughter strands are still with the same cell. The heavy arrow indicates a possible spatial/temporal window during which the two DNA molecules might be able to undergo such recombinational interactions.

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–26
Figure 16–26

Outline of an in vivo assay system for gap filling by recombinational repair. The gap-lesion plasmid GP21 (Kan) was introduced into cells along with a homologous partner plasmid (FGP20/Tamp; Amp). Parallel experiments were carried out with a heterologous plasmid (pUC18). Only gap-filling repair allows GP21 to transform the cells and confer a Kan phenotype. This can be done by translesion DNA replication, which does not depend on the partner plasmid, or by a process involving recombination functions, which depends on a homologous partner plasmid. The homologous partner plasmid carries a T opposite the site corresponding to the lesion in GP21, so that GP21 filled in by recombination will have a T at that position. In contrast, GP21 filled in by DNA Pol V-dependent translesion synthesis will have primarily an A at that location ( ). (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–27
Figure 16–27

Fork regression offers an avenue for damage tolerance (and replication fork repair) that does not involve DSB or DNA strand invasion.

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–28
Figure 16–28

Possible scheme for PriA-dependent control of replication fork establishment. A broken arm of the replication fork is processed by RecBCD. RecA (white ovals) is loaded, and a D loop recombination intermediate is formed between the broken arm and the sister chromosome. If PriA does not bind, DNA synthesis from the invading 3’ end would merely extend the D loop without forming a processive replication fork. If PriA (grey hexagon) does bind to the invading 3’ end, DNA synthesis is inhibited while PriA (with PriBC DnaCT) loads DnaB helicase (grey oval) to the displaced strand of the D loop. DnaB recruits DnaG primase (gold circle), and this relieves PriA inhibition to allow both lagging- and leading-strand DNA synthesis to begin via DNA Pol III holoenzyme (gold triangles). In contrast, a broken chromosome may engage both broken arms into the D loop, thereby potentially blocking DnaB loading; repair may be accomplished with a small amount of repair synthesis without replication fork establishment. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–29
Figure 16–29

UV-induced DNA replication intermediates observed during the recovery of replication. (A) Diagram of the migration pattern of PvuII-digested pBR322 during two-dimensional analysis. Nonreplicating plasmids run as a linear 4.4-kb fragment. Normal replicating fragments form Y-shaped structures and migrate more slowly due to their larger size and nonlinear shape, forming an arc that extends out from the linear fragment. Double Y- or X-shaped molecules migrate in the cone region. (B) The replication intermediates persist until a time correlating with replication recovery and lesion removal. Replication recovery, lesion repair, and the relative amount of replicating fragments (gold) and cone region intermediates (black) are plotted. Replication recovery was assayed by [H]thymine incorporation for UV-irradiated (black line) or mock-irradiated (gold line) cultures. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–30
Figure 16–30

Three possible mechanisms for the regression of replication forks. (A) Fork regression mediated by RecA. (B) Fork regression mediated by RecG. (C) Fork regression mediated by positive supercoils accumulated ahead of the replication fork. Plus sign represents positive supercoiling of the DNA; dark grey circle represents a lesion.

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–31
Figure 16–31

Models for restarting regressed replication forks that have been arrested by a lesion affecting just one strand of the DNA template. See the text for details. (Adapted from references and .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–32
Figure 16–32

Formation of the cone region intermediates depends on the presence of UV-induced lesions, RecA, and active replication. Replication recovery was assayed by [H]thymine incorporation, as in Fig. 16–29 , and the replication intermediates observed during the normal recovery period were monitored by two-dimensional gel analysis. (A) mutants fail to recover replication after UV-induced DNA damage, and the cone region intermediates persist and accumulate. (UV-irradiated cultures, gold; mock-irradiated cultures, black). (B) mutants fail to recover replication after UV-induced DNA damage and the cone region intermediates do not accumulate (UV-irradiated cultures, gold; mock-irradiated cultures, black). (C and D) y-structure (C) and cone region (D) are plotted for UV-irradiated (gold) and (grey) strains. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–33
Figure 16–33

Models for restarting DNA replication by junction cleavage. Formation of a Holliday junction from a replication fork stalled at a lesion (grey circle) might allow processing by the RuvABC helicase-endonuclease complex in one of two ways. (A) RuvABC might cleave the Holliday junction at the stalled fork before recombination from the free double-stranded DNA end has occurred, while formation of the Holliday junction might facilitate unmasking and subsequent removal of the block. The released double-stranded DNA end would then be recombined back into the homologous duplex to form a D-loop. Cleavage of the Holliday junction formed at the D-loop by RuvABC and reassembly of the replication machinery at the fork would reconstitute an active replisome. (B) The double-stranded DNA end that is spooled out from the Holliday junction might be recombined with the homologous sequences in the reannealed parental strands to form a D-loop intermediate that is linked to the original Holliday junction at the fork. A second Holliday junction would also be formed at the D-loop. Assuming that the original block could be removed, cleavage of both Holliday junctions by RuvABC would generate a forked DNA structure onto which the replication machinery could be reloaded. The 3’ ends of DNA strands are shown by arrowheads, and cleavage of Holliday junctions by RuvABC is shown by black triangles. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Image of Figure 16–34
Figure 16–34

Recovery from inhibition of DNA synthesis after UV irradiation: synergism in double mutants combining and Black dashed line, black dotted line, gold dotted line, gold dashed line, The UV dose was 3 J/m. Cells growing exponentially at 37°C were pulse-labeled with [H]thymidine for 2 min at various times before and after UV irradiation. The optical density (OD) curve defines the range of values obtained for all strains. (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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Figure 16–35

Rates of post-UV DNA synthesis in strains carrying substitutions in and/or Experiments were performed and plotted as described previously ( ). (Adapted from reference .)

Citation: Errol C, Graham C, Wolfram S, Richard D, Roger A, Tom E. 2006. Recombinational Repair, Replication Fork Repair, and DNA Damage Tolerance, p 569-612. In DNA Repair and Mutagenesis, Second Edition. ASM Press, Washington, DC. doi: 10.1128/9781555816704.ch16
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References

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1. Anderson, D. G.,, J. J. Churchill, and, S. C. Kowalczykowski. 1999. A single mutation, RecB(D1080A), eliminates RecA protein loading but not Chi recognition by RecBCD enzyme. J. Biol. Chem. 274:2713927144.
2. Anderson, D. G.,and, S. C. Kowalczykowski. 1997. The recombination hot spot chi is a regulatory element that switches the polarity of DNA degradation by the RecBCD enzyme. Genes Dev. 11:571581.
3. Anderson, D. G.,and, S. C. Kowalczykowski. 1997. The translocating RecBCD enzyme stimulates recombination by directing RecA protein onto ssDNA in a chi-regulated manner. Cell 90:7786.
4. Arenson, T. A.,, O. V. Tsodikov, and, M. M. Cox. 1999. Quantitative analysis of the kinetics of end-dependent disassembly of RecA filaments from ssDNA. J. Mol. Biol. 288:391–401.
5. Ariyoshi, M.,, D. G. Vassylyev,, H. Iwasaki,, H. Nakamura,, H. Shinagawa, and, K. Morikawa. 1994. Atomic structure of the RuvC resolvase: a Holliday junction-specific endonuclease from E. coli. Cell 78:10631072.
6. Arnold, D. A.,and, S. C. Kowalczykowski. 2000. Facilitated loading of RecA protein is essential to recombination by RecBCD enzyme. J. Biol. Chem. 275:1226112265.
7. Asai, T.,, D. B. Bates, and, T. Kogoma. 1994. DNA replication triggered by double-stranded breaks in E. coli: dependence on homologous recombination functions. Cell 78:10511061.
8. Asai, T.,, S. Sommer,, A. Bailone, and, T. Kogoma. 1993. Homologous recombination-dependent initiation of DNA replication for DNA damage-inducible origins in Escherichia coli. EMBO J. 12:32783295.
9. Barre, F. X.,, M. Aroyo,, S. D. Colloms,, A. Helfrich,, F. Cornet, and, D. J. Sherratt. 2000. FtsK functions in the processing of a Holliday junction intermediate during bacterial chromosome segregation. Genes Dev. 14:29762988.
10. Barre, F. X.,, B. Soballe,, B. Michel,, M. Aroyo,, M. Robertson, and, D. Sherratt. 2001. Circles: the replication-recombination-chromosome segregation connection. Proc. Natl. Acad. Sci. USA 98:81898195.
11. Beam, C. E.,, C. J. Saveson, and, S. T. Lovett. 2002. Role for radA/sms in recombination intermediate processing in Escherichia coli. J. Bacteriol. 184:68366844.
12. Bennett, R. J.,, H. J. Dunderdale, and, S. C. West. 1993. Resolution of Holliday junctions by RuvC resolvase. Cell 74:10211031.
13. Benson, F. E.,, G. T. Illing,, G. J. Sharples, and, R. G. Lloyd. 1988. Nucleotide sequencing of the ruv region of Escherichia coli K-12 reveals a LexA-regulated operon encoding two genes. Nucleic Acids Res. 16:15411549.
14. Berdichevsky, A.,, L. Izhar, and, Z. Livneh. 2002. Error-free recombinational repair predominates over mutagenic translesion replication in E. coli. Mol. Cell 10:917924.
15. Bernstein, H.,, H. C. Byerly,, F. A. Hopf, and, R. E. Michod. 1985. Genetic damage, mutation and the evolution of sex. Science 229:12771281.
16. Bianco, P. R.,, L. R. Brewer,, M. Corzett,, R. Balhorn,, Y. Yeh,, S. C. Kowalczykowski, and, R. J. Baskin. 2001. Processive translocation and DNA unwinding by individual RecBCD enzyme molecules. Nature 409:374378.
17. Blakely, G.,, S. Colloms,, G. May,, M. Burke, and, D. Sherratt. 1991. Escherichia coli XerC recombinase is required for chromosomal segregation at cell division. New Biol. 3:789798.
18. Bockrath, R.,, L. Wolff,, A. Farr, and, R. J. Crouch. 1987. Amplified RNase H activity in Escherichia coli B/r increases sensitivity to ultraviolet radiation. Genetics 115:3340.
19. Bolt, E. L.,and, R. G. Lloyd. 2002. Substrate specificity of RusA resolvase reveals the DNA structures targeted by RuvAB and RecG in vivo. Mol. Cell 10:187198.
20. Bonura, T.,and, K. C. Smith. 1975. Enzymatic production of deoxyribonucleic acid double-strand breaks after ultraviolet light irradiation of Escherichia coli K-12. J. Bacteriol. 121:511517.
21. Bonura, T.,and, K. C. Smith. 1975. Quantitative evidence for en-zymatically-induced DNA double-strand breaks as lethal lesions in UV irradiated pol+ and polAl strains of E. coli K-12. Photochem. Photobiol. 22:243248.
22. Bosco, G.,and, J. E. Haber. 1998. Chromosome break-induced DNA replication leads to nonreciprocal translocations and telomere capture. Genetics 150:10371047.
23. Bouche, J. P.,, K. Zechel, and, A. Kornberg. 1975. dnaG gene product, a rifampicin-resistant RNA polymerase, initiates the conversion of a single-stranded coliphage DNA to its duplex replicative form. J. Biol. Chem. 250: 59956001.
24. Bridges, B. A., 1972. Evidence for a further dark repair process in bacteria. Nat. New Biol. 240:5253.
25. Casaregola, S.,, M. Khidhir, and, I.B. Holland. 1987. Effects of modulation of RNase H production on the recovery of DNA synthesis following UV-irradiation in Escherichia coli. Mol. Gen. Genet. 209:494498.
26. Cassuto, E.,, J. Musalim, and, P. Howard-Flanders. 1978. Homology-dependent cutting in trans in extracts of Escherichia coli: an approach to the enzymology of genetic recombination. Proc. Natl. Acad. Sci. USA 75: 620624.
27. Chan, S. N.,, L. Harris,, E. L. Bolt,, M. C. Whitby, and, R. G. Lloyd. 1997. Sequence specificity and biochemical characterization of the RusA Holliday junction resolvase of Escherichia coli. J. Biol. Chem. 272:1487314882.
28. Chase, J. W.,and, C. C. Richardson. 1974. Exonuclease VII of Esch-erichia coli. Mechanism of action. J. Biol. Chem. 249:45534561.
29. Chen, Y. J.,, X. Yu, and, E. H. Egelman. 2002. The hexameric ring structure of the Escherichia coli RuvB branch migration protein. J. Mol. Biol. 319:587591.
30. Churchill, J. J.,, D. G. Anderson, and, S. C. Kowalczykowski. 1999. The RecBC enzyme loads RecA protein onto ssDNA asymmetrically and independently of chi, resulting in constitutive recombination activation. Genes Dev. 13:901911.
31. Clark, A. J., 1973. Recombination deficient mutants of E. coli and other bacteria. Annu. Rev. Genet. 7:6786.
32. Clark, A. J.,and, S. J. Sandler. 1994. Homologous genetic recombination: the pieces begin to fall into place. Crit. Rev. Microbiol. 20:125142.
33. Connelly, B.,, C. A. Parsons,, F. E. Benson,, H. J. Dunderdale,, G. J. Sharples,, R. G. Lloyd, and, S. C. West. 1991. Resolution of Holliday junctions in vitro requires the Escherichia coli ruvC gene product. Proc. Natl. Acad. Sci. USA 88:60636067.
34. Cooper, P., 1982. Characterization of long patch excision repair of DNA in ultraviolet-irradiated Escherichia coli: an inducible function under rec-lex control. Mol. Gen. Genet. 185:189197.
35. Cooper, P. K.,and, P. C. Hanawalt. 1972. Heterogeneity of patch size in repair replicated DNA in Escherichia coli. J. Mol. Biol. 67:110.
36. Courcelle, J.,, C. Carswell-Crumpton, and, P. C. Hanawalt. 1997. recF and recR are required for the resumption of replication at DNA replication forks in Escherichia coli. Proc. Natl. Acad. Sci. USA 94:37143719.
37. Courcelle, J.,, J. R. Donaldson,, K. H. Chow, and, C. T. Courcelle. 2003. DNA damage-induced replication fork regression and processing in Escherichia coli. Science 299:10641067.
38. Courcelle, J.,, A. K. Ganesan, and, P. C. Hanawalt. 2001. Therefore, what are recombination proteins there for? Bioessays 23:463470.
39. Courcelle, J.,and, P. C. Hanawalt. 1999. RecQ and RecJ process blocked replication forks prior to the resumption of replication in UV-irradiated Escherichia coli. Mol. Gen. Genet. 262:543551.
40. Courcelle, J.,and, P. C. Hanawalt. 2001. Participation of recombination proteins in rescue of arrested replication forks in UV-irradiated Escherichia coli need not involve recombination. Proc. Natl. Acad. Sci. USA 98: 81968202.
41. Courcelle, J.,and, P. C. Hanawalt. 2003. RecA-dependent recovery of arrested DNA replication forks. Annu. Rev. Genet. 37:611646.
42. Cox, M. M., 1993. Relating biochemistry to biology. How the recombinational repair function of RecA protein is manifested in its molecular properties. Bioessays 15: 617673.
43. Cox, M. M., 1994. Why does RecA protein hydrolyze ATP? Trends Biochem. Sci. 19:217222.
44. Cox, M. M., 1999. Recombinational DNA repair in bacteria and the RecA protein. Prog. Nucleic Acid Res. Mol. Biol. 63:311366.
45. Cox, M. M., 2001. Historical overview: searching for replication help in all of the rec places. Proc. Natl. Acad. Sci. USA 98:81738180.
46. Cox, M. M., 2001. Recombinational DNA repair of damaged replication forks in Escherichia coli: questions. Annu. Rev. Genet. 35:5382.
47. Cox, M. M., 2002. The nonmutagenic repair of broken replication forks via recombination. Mutat. Res. 510:107120.
48. Cox, M. M., 2003. The bacterial RecA protein as a motor protein. Annu. Rev. Microbiol. 57:551577.
49. Cox, M. M.,, M. F. Goodman,, K. N. Kreuzer,, D. J. Sherratt,, S. J. Sandler, and, K. J. Marians. 2000. The importance of repairing stalled replication forks. Nature 404:3741.
50. Cox, M. M.,and, I. R. Lehman. 1982. RecA protein-promoted DNA strand exchange. Stable complexes of RecA protein and single-stranded DNA formed in the presence of ATP and single-stranded DNA binding protein. J. Biol. Chem. 257:85238532.
51. Cox, M. M.,and, I. R. Lehman. 1987. Enzymes of general recombination. Annu. Rev. Biochem. 56:229262.
52. Cromie, G. A.,and, D. R. Leach. 2000. Control of crossing over. Mol. Cell 6:815826.
53. Cromie, G. A.,and, D. R. Leach. 2001. Recombinational repair of chromosomal DNA double-strand breaks generated by a restriction endonuclease. Mol. Microbiol. 41:873883.
54. Datta, S.,, N. Ganesh,, N. R. Chandra,, K. Muniyappa, and, M. Vijayan. 2003. Structural studies on MtRecA-nucleotide complexes: insights into DNA and nucleotide binding and the structural signature of NTP recognition. Proteins 50:474485.
55. Datta, S.,, R. Krishna,, N. Ganesh,, N. R. Chandra,, K. Muniyappa, and, M. Vijayan. 2003. Crystal structures of Mycobacterium smegmatis RecA and its nucleotide complexes. J. Bacteriol. 185:42804284.
56. Datta, S.,, M. M. Prabu,, M. B. Vaze,, N. Ganesh,, N. R. Chandra,, K. Muniyappa, and, M. Vijayan. 2000. Crystal structures of Mycobacterium tuberculosis RecA and its complex with ADP-AlF(4): implications for decreased ATPase activity and molecular aggregation. Nucleic Acids Res. 28: 49644973.
57. Dillingham, M. S.,and, S. C. Kowalczykowski. 2001. A step backward in advancing DNA replication: rescue of stalled replication forks by RecG. Mol. Cell 8: 734736.
58. Dillingham, M. S.,, M. Spies, and, S. C. Kowalczykowski. 2003. RecBCD enzyme is a bipolar DNA helicase. Nature 423:893897.
59. Dixon, D. A.,and, S. C. Kowalczykowski. 1991. Homologous pairing in vitro stimulated by the recombination hotspot Chi. Cell 66:361371.
60. Dixon, D. A.,and, S. C. Kowalczykowski. 1993. The recombination hotspot chi is a regulatory sequence that acts by attenuating the nuclease activity of the E. coli RecBCD enzyme. Cell 73:8796.
61. Donaldson, J. R.,, C. T. Courcelle, and, J. Courcelle. 2004. RuvAB and RecG are not essential for the recovery of DNA synthesis following UV-induced DNA damage in Escherichia coli. Genetics 166:16311640.
62. Dunderdale, H. J.,, F. E. Benson,, C. A. Parsons,, G. J. Sharples,, R. G. Lloyd, and, S. C. West. 1991. Formation and resolution of recombination intermediates by E. coli RecA and RuvC proteins. Nature (London) 354:506510.
63. Echols, H.,and, M. F. Goodman. 1990. Mutation induced by DNA damage: a many protein affair. Mutat. Res. 236:301311.
64. Echols, H.,and, M. F. Goodman. 1991. Fidelity mechanisms in DNA replication. Annu. Rev. Biochem. 60:477511.
65. Egelman, E., 2000. A common structural core in proteins active in DNA recombination and replication. Trends Biochem. Sci. 25:179182.
66. Egelman, E., 2000. A ubiquitous structural core. Trends Biochem. Sci. 25:183184.
67. Egelman, E. H., 1993. What do X-ray crystallographic and electron microscopic structural studies of the RecA protein tell us about recombination? Curr. Opin. Struct. Biol. 3:189197.
68. Egelman, E. H., 2001. Structural biology. Pumping DNA. Nature 409:573575.
69. Egelman, E. H.,and, A. Stasiak. 1993. Electron microscopy of RecA-DNA complexes: two different states, their functional significance and relation to the solved crystal structure. Micron 24:309324.
70. Eggler, A. L.,, S. L. Lusetti, and, M. M. Cox. 2003. The C terminus of the Escherichia coli RecA protein modulates the DNA binding competition with single-stranded DNA-binding protein. J. Biol. Chem. 278:1638916396.
71. Eggleston, A. K.,, A. H. Mitchell, and, S. C. West. 1997. In vitro reconstitution of the late steps of genetic recombination in E. coli. Cell 89: 607617.
72. Eguchi, Y.,, T. Ogawa, and, H. Ogawa. 1988. Stimulation of RecA-mediated cleavage of phage Φ80 cI repressor by deoxydinucleotides. J. Mol. Biol. 204:6977.
73. Eichman, B. F.,, M. Ortiz-Lombardia,, J. Aymami,, M. Coll, and, P. S. Ho. 2002. The inherent properties of DNA four-way junctions: comparing the crystal structures of Holliday junctions. J. Mol. Biol. 320:10371051.
74. Eichman, B. F.,, J. M. Vargason,, B. H. Mooers, and, P. S. Ho. 2000. The Holliday junction in an inverted repeat DNA sequence: sequence effects on the structure of four-way junctions. Proc. Natl. Acad. Sci. USA 97:39713976.
75. Finch, P. W.,, P. Chambers, and, P. T. Emmerson. 1985. Identification of the E. coli recNgene product as a major SOS protein. J. Bacteriol. 164: 653658.
76. Flores, M. J.,, H. Bierne,, S. D. Ehrlich, and, B. Michel. 2001. Impairment of lagging strand synthesis triggers the formation of a RuvABC substrate at replication forks. EMBO J. 20:619629.
77. Flores, M. J.,, S. D. Ehrlich, and, B. Michel. 2002. Primosome assembly requirement for replication restart in the Escherichia coli holDG10 replication mutant. Mol. Microbiol. 44:783792.
78. Friedberg, E. C., 1985. DNA Repair. W. H. Freeman & Co., New York, N.Y.
79. Funnell, B. E.,, T. A. Baker, and, A. Kornberg. 1987. In vitro assembly of a prepriming complex at the origin of the Escherichia coli chromosome. J. Biol. Chem. 262:1032710334.
80. Galitski, T.,and, J. R. Roth. 1997. Pathways for homologous recombination between chromosomal direct repeats in Salmonella typhimurium. Genetics 146:751767.
81. Ganesan, A. K., 1974. Persistence of pyrimidine dimers during post-replication repair in ultraviolet light-irradiated Escherichia coli K-12. J. Mol. Biol. 87:103119.
82. Ganesan, A. K., 1975. Distribution of pyrimidine dimers during postreplication repair in UV-irradiated excision-deficient cells of Escherichia coli K12, p. 317320. In P. C. Hanawalt and, R. B. Setlow (ed.), Molecular Mechanisms for Repair of DNA. Plenum Press, New York, N.Y.
83. Ganesan, A. K.,and, P. C. Seawell. 1975. The effect of lexA and recF mutations on post-replication repair and DNA synthesis in Escherichia coli K-12. Mol. Gen. Genet. 141:189205.
84. George, J. W.,, B. A. Stohr,, D. J. Tomso, and, K. N. Kreuzer. 2001. The tight linkage between DNA replication and double-strand break repair in bacteriophage T4. Proc. Natl. Acad. Sci. USA 98:82908297.
85. Giraud-Panis, M. J.,and, D. M. Lilley. 1998. Structural recognition and distortion by the DNA junction-resolving enzyme RusA. J. Mol. Biol. 278:117133.
86. Goodman, M. F., 2000. Coping with replication ‘train wrecks’ in Escherichia coli using Pol V, Pol II and RecA proteins. Trends Biochem Sci. 25:189195.
87. Gregg, A. V.,, P. McGlynn,, R. P. Jaktaji, and, R. G. Lloyd. 2002. Direct rescue of stalled DNA replication forks via the combined action of PriA and RecG helicase activities. Mol. Cell 9:241251.
88. Grompone, G.,, S. D. Ehrlich, and, B. Michel. 2003. Replication restart in gyrB Escherichia coli mutants. Mol. Microbiol. 48:845854.
89. Grompone, G.,, M. Seigneur,, S. D. Ehrlich, and, B. Michel. 2002. Replication fork reversal in DNA polymerase III mutants of Escherichia coli: a role for the beta clamp. Mol. Microbiol. 44:13311339.
90. Guo, F.,, D. N. Gopaul, and, G. D. van Duyne. 1997. Structure of Cre recombinase complexed with DNA in a site-specific recombination synapse. Nature 389:4046.
91. Haber, J. E., 1999. DNA recombination: the replication connection. Trends Biochem. Sci. 24:271275.
92. Hall, J. D.,and, D. Mount. 1981. Mechanisms of DNA replication and mutagenesis in ultraviolet-irradiated bacteria and mammalian cells. Prog. Nucleic Acid Res. Mol. Biol. 25:53126.
93. Hanawalt, P. C., 1966. The UV sensitivity of bacteria: its relationship to the DNA replication cycle. Photochem. Photobiol. 5:112.
94. Hanawalt, P. C.,and, I. Brempelis. 1967. Selective degradation of newly-replicated DNA after inhibition of DNA synthesis in Escherichia coli, p., 650. In Proceedings of the 7th International Congress on Biochemistry. The Science Council of Japan, Ueno Park, Tokyo, Japan.
95. Hanawalt, P. C.,, P. K. Cooper,, A. Ganesan, and, C. A. Smith. 1979. DNA repair in bacteria and mammalian cells. Annu. Rev. Biochem. 48:783836.
96. Hargreaves, D.,, D. W. Rice,, S. E. Sedelnikova,, P. J. Artymiuk,, R. G. Lloyd, and, J. B. Rafferty. 1998. Crystal structure of E. coli RuvA with bound DNA Holliday junction at 6 Å resolution. Nat. Struct. Biol. 5:441446.
97. Hegde, S.,, S. J. Sandler,, A. J. Clark, and, M. V. Madiraju. 1995. recO and recR mutations delay induction of the SOS response in Escherichia coli. Mol. Gen. Genet. 246:254258.
98. Hegde, S. P.,, M. H. Qin,, X. H. Li,, M. A. Atkinson,, A. J. Clark,, M. Rajagopalan, and, M. V. Madiraju. 1996. Interactions of RecF protein with RecO, RecR, and single-stranded DNA binding proteins reveal roles for the RecF-RecO-RecR complex in DNA repair and recombination. Proc. Natl. Acad. Sci. USA 93:1446814473.
99. Higgins, N. P.,, K. Kato, and, B. Strauss. 1976. A model for replication repair in mammalian cells. J. Mol. Biol. 101:417425.
100. Higuchi, K.,, T. Katayama,, S. Iwai,, M. Hidaka,, T. Horiuchi, and, H. Maki. 2003. Fate of DNA replication fork encountering a single DNA lesion during oriC plasmid DNA replication in vitro. Genes Cells 8:437449.
101. Hirii, Z. I.,and, K. Suzuki. 1968. Degradation of the DNA of Escherichia coli rec- (JC1569b) after irradiation with ultraviolet light. Photochem. Photobiol. 8:93105.
102. Holliday, R., 1964. A mechanism for gene conversion in fungi. Genet. Res. 5:282304.
103. Hooper, I.,and, J. B. Egan. 1981. Coliphage 186 replication requires host initiation functions DnaA and DnaC. J. Virol. 40:599601.
104. Hooper, I.,, W. H. Woods, and, J. B. Egan. 1981. Coliphage 186 infection requires replication but is delayed when the host cell is irradiated before infection. J. Virol. 40:341349.
105. Howard-Flanders, P.,and, R. P. Boyce. 1966. DNA repair and genetic recombination: studies on mutants of Escherichia coli defective in these processes. Radiat. Res. Suppl. 6:156184.
106. Howard-Flanders, P.,and, W. D. Rupp. 1981. Measurement of postreplication repair in prokaryotes, p. 459470. In E. C. Friedberg and, P. C. Hanawalt (ed.), DNA Repair: a Laboratory Manual of Research Procedures. Marcel Dekker, Inc., New York, N.Y.
107. Howard-Flanders, P.,, W. D. Rupp,, B. M. Wilkins, and, R. S. Cole. 1968. DNA replication and recombination after UV-irradiation. Cold Spring Harbor Symp. Quant. Biol. 33:195205.
108. Howard-Flanders, P.,and, L. Theriot. 1966. Mutants of Escherichia coli K-12 defective in DNA repair and in genetic recombination. Genetics 53 (Suppl.): 11371150.
109. Howard-Flanders, P.,, L. Theriot, and, J. B. Stedeford. 1969. Some properties of excision-defective recombination-deficient mutants of Escherichia coli K-12. J. Bacteriol. 97:11341141.
110. Howard-Flanders, P.,, S. C. West, and, A. J. Stasiak. 1984. Role of RecA spiral filaments in genetic recombination. Nature 309:215220.
111. Hsieh, P.,, C. S. Camerini-Otero, and, R. D. Camerini-Otero. 1990. Pairing of homologous DNA sequences by proteins: evidence for three-stranded DNA. Genes Dev. 4:19511963.ss
112. Itaya, M., 1990. Isolation and characterization of a second RNase H (RNase H II) encoded by the rnhB gene. Proc. Natl. Acad. Sci. USA 87:85878591.
113. Iwasaki, H.,, T. Shiba,, K. Makino,, A. Nakata, and, H. Shingawa. 1989. Overproduction, purification, and ATPase activity of the Escherichia coli RuvB protein involved in DNA repair. J. Bacteriol. 171:52765280.
114. Iwasaki, H.,, M. Takahagi,, A. Nakata, and, H. Shinagawa. 1992. Escherichia coli RuvA and RuvB proteins specifically interact with Holliday junctions and promote branch migration. Genes Dev. 6:22142220.
115. Iwasaki, H.,, M. Takahagi,, T. Shiba,, A. Nakata, and, H. Shinagawa. 1991. Escherichia coli RuvC protein is an endonuclease that resolves the Holliday structure. EMBO J. 10:43814389.
116. Iyer, V. N.,and, W. D. Rupp. 1971. Usefulness of benzoylated naphthoylated DEAE-cellulose to distinguish and fractionate double-stranded DNA bearing different extents of single-stranded regions. Biochim. Biophys. Acta 228:117.
117. Jain, S. K.,, M. M. Cox, and, R. B. Inman. 1994. On the role of ATP hydrolysis in RecA protein-mediated DNA strand exchange. III. Unidirectional branch migration and extensive hybrid DNA formation. J. Biol. Chem. 269:2065320661.
118. Jaktaji, R. P.,and, R. G. Lloyd. 2003. PriA supports two distinct pathways for replication restart in UV-irradiated Escherichia coli cells. Mol. Microbiol. 47:10911100.
119. Jessberger, R., 2002. The many functions of SMC proteins in chromosome dynamics. Nat. Rev. Mol. Cell. Biol. 3:767778.
120. Johnson, R. C.,and, W. F. McNeill. 1978. Electron microscopy of UV-induced postreplication repair of daughter strand gaps, p., 9599. In P. C. Hanawalt,, E. C. Friedberg, and, C. F. Fox (ed.), DNA Repair Mechanisms. Academic Press, Inc., New York, N.Y.
121. Johnson, R. D.,and, L. S. Symington. 1993. Crossed-stranded DNA structures for investigating the molecular dynamics of the Holliday junction. J. Mol. Biol. 229:812820.
122. Jones, J. M.,and, H. Nakai. 2000. PriA and phage T4 gp59: factors that promote DNA replication on forked DNA substrates microreview. Mol. Microbiol. 36:519527.
123. Kadyrov, F. A.,and, J. W. Drake. 2003. Properties of bacteriophage T4 proteins deficient in replication repair. J. Biol. Chem. 278:2524725255.
124. Kadyrov, F. A.,and, J. W. Drake. 2004. UvsX recombinase and Dda helicase rescue stalled bacteriophage T4 DNA replication forks in vitro. J. Biol. Chem. 279:3573535740.
125. Kaguni, J. M.,and, A. Kornberg. 1984. Replication initiated at the origin (oriC) of the E. coli chromosome reconstituted with purified enzymes. Cell 38:183190.
126. Kantake, N.,, M. V. Madiraju,, T. Sugiyama, and, S. C. Kowal-czykowski. 2002. Escherichia coli RecO protein anneals ssDNA complexed with its cognate ssDNA-binding protein: a common step in genetic recombination. Proc. Natl. Acad. Sci. USA 99:1532715332.
127. Kato, T., 1977. Effects of chloramphenicol and caffeine on postreplication repair in uvrA- umuC- and uvrA- recF- strains of Escherichia coli K-12. Mol. Gen. Genet. 156:115120.
128. Kelman, Z.,and, M. O’Donnell. 1995. DNA polymerase III holoenzyme: structure and function of a chromosomal replicating machine. Annu. Rev. Biochem. 64:171200.
129. Khidhir, M. A.,, S. Casaregola, and, I. B. Holland. 1985. Mechanism of transient inhibition of DNA synthesis in ultraviolet-irradiated E. coli: inhibition is independent of recA whilst recovery requires RecA protein itself and an additional, inducible SOS function. Mol. Gen. Genet. 199:133140.
130. Kim, J. I.,, M. M. Cox, and, R. B. Inman. 1992. On the role of ATP hydrolysis in RecA protein-mediated DNA strand exchange. J. Biol. Chem. 267:1643816443.
131. Kim, J. L.,, K. A. Morgenstern,, J. P. Griffith,, M. D. Dwyer,, J. A. Thomson,, M. A. Murcko,, C. Lin, and, P. R. Caron. 1998. Hepatitis C virus NS3 RNA helicase domain with a bound oligonucleotide: the crystal structure provides insights into the mode of unwinding. Structure 6:89100.
132. Kobayashi, I.,and, H. Ikeda.. 1983. Double Holliday structure: a possible in vivo intermediate form of general recombination in Escherichia coli. Mol. Gen. Genet. 191:213220.
133. Kogoma, T.., 1996. Recombination by replication. Cell 85:625627.
134. Kogoma, T.., 1997. Stable DNA replication: interplay between DNA replication, homologous recombination, and transcription. Microbiol. Mol. Biol. Rev. 61:212238.
135. Kogoma, T.,, G. W. Cadwell,, K. G. Barnard, and, T. Asai.. 1996. The DNA replication priming protein, PriA, is required for homologous recombination and double-strand break repair. J. Bacteriol. 178:12581264.
136. Kojima, M.,, M. Suzuki,, T. Morita,, T. Ogawa,, H. Ogawa, and, M. Tada.. 1990. Interaction of RecA protein with pBR322 DNA modified by N-hydroxy-2-acetylaminofluorene and 4-hydroxyaminoquinoline 1-oxide. Nucleic Acids Res. 18:27072714.
137. Kolodner, R.,, R. A. Fishel, and, M. Howard.. 1985. Genetic recombination of bacterial plasmid DNA: effect of RecF pathway mutations on plasmid recombination in Escherichia coli. J. Bacteriol. 163:10601066.
138. Kowalczykowski, S. C.., 1991. Biochemistry of genetic recombination: energetics and mechanism of DNA strand exchange. Annu. Rev. Biophys. Biophys. Chem. 20:539575.
139. Kowalczykowski, S. C.., 2000. Initiation of genetic recombination and recombination-dependent replication. Trends Biochem. Sci. 25:156165.
140. Kowalczykowski, S. C.,, D. A. Dixon,, A. K. Eggleston,, S. D. Lauder, and, W. M. Rehrauer.. 1994. Biochemistry of homologous recombination in Escherichia coli. Microbiol. Rev. 58:401465.
141. Kowalczykowski, S. C.,and, R. A. Krupp.. 1987. Effects of Escherichia coli SSB protein on the single-stranded DNA-dependent ATPase activity of Escherichia coli RecA protein. Evidence that SSB protein facilitates the binding of RecA protein to regions of secondary structure within single-stranded DNA. J. Mol. Biol. 193:97113.
142. Kowalczykowski, S. C.,and, R. A. Krupp.. 1995. DNA-strand exchange promoted by RecA protein in the absence of ATP: implications for the mechanism of energy transduction in protein-promoted nucleic acid transactions. Proc. Natl. Acad. Sci. USA 92:34783482.
143. Krasin, F.,and, F. Hutchinson.. 1977. Repair of DNA double-strand breaks in Escherichia coli, which requires RecA function and the presence of a duplicate genome. J. Mol. Biol. 116:8198.
144. Krasin, F.,and, F. Hutchinson.. 1981. Repair of DNA double-strand breaks in Escherichia coli cells requires synthesis of proteins that can be induced by UV light. Proc. Natl. Acad. Sci. USA 78:34503453.
145. Kraus, E.,, W. Y. Leung, and, J. E. Haber.. 2001. Break-induced replication: a review and an example in budding yeast. Proc. Natl. Acad. Sci. USA 98:82558262.
146. Kreuzer, K. N.., 2000. Recombination-dependent DNA replication in phage T4. Trends Biochem. Sci. 25:165173.
147. Kuempel, P. L.,, J. M. Henson,, L. Dircks,, M. Tecklenburg, and, D. F. Lim.. 1991. dif, a recA-independent recombination site in the terminus region of the chromosome of Escherichia coli. New Biol. 3:799811.
148. Kuzminov, A.., 1996. Recombinational Repair of DNA Damage. R. G. Landes, Georgetown, Tex.
149. Kuzminov, A., 1999. Recombinational repair of DNA damage in Escherichia coli and bacteriophage lambda. Microbiol. Mol. Biol. Rev. 63:751813.
150. Kuzminov, A.. 2001. DNA replication meets genetic exchange: chromosomal damage and its repair by homologous recombination. Proc. Natl. Acad. Sci. USA 98:84618468.
151. Kuzminov, A.. 2001. Single-strand interruptions in replicating chromosomes cause double-strand breaks. Proc. Natl. Acad. Sci. USA 98:82418246.
152. Kuzminov, A., and, F. W. Stahl. 1999. Double-strand end repair via the RecBC pathway in Escherichia coli primes DNA replication. Genes Dev. 13:345356.
153. Lark, K. G.,and, C. A. Lark. 1979. recA-dependent DNA replication in the absence of protein synthesis: characteristics of a dominant lethal replication mutation dnaT, and requirement for recA+ function. Cold Spring Harbor Symp. Quant. Biol. 43:537549.
154. LeBowitz, J. H.,and, R. McMacken. 1986. The Escherichia coli dnaB replication protein is a DNA helicase. J. Biol. Chem. 261:47384748.
155. Lederberg, J. 1947. Gene recombination and linked segregations in Escherichia coli. Genetics 32:505525.
156. Lee, E. H.,and, A. Kornberg. 1991. Replication deficiencies in priA mutants of Escherichia coli lacking the primosomal replication n’ protein. Proc. Natl. Acad. Sci. USA 88:30293032.
157. Lee, J. W.,and, M. M. Cox.. 1990. Inhibition of RecA protein promoted ATP hydrolysis. 1. ATPγ-S and ADP are antagonistic inhibitors. Biochemistry 29:76667676.
158. Lee, J. W.,and, M. M. Cox. 1990. Inhibition of RecA protein promoted ATP hydrolysis. 2. Longitudinal assembly and disassembly of RecA protein filaments mediated by ATP and ADP. Biochemistry 29:76777683.
159. Ley, R. D. 1973. Postreplication repair in an excision-defective mutant of Escherichia coli: ultraviolet light-induced incorporation of bromo-deoxyuridine into parental DNA. Photochem. Photobiol. 18:8795.
160. Ley, R. D. 1975. Ultraviolet-light induced incorporation of bro-modeoxyuridine into parental DNA of an excision-defective mutant of Esch-erichia coli, p. 313316. In P. C. Hanawaltand, R. B. Setlow (ed.)., Molecular Mechanisms for Repair of DNA. Plenum Publishing Corp., New York, N.Y.
161. Lindsley, J. E.,and, M. M. Cox. 1989. Dissociation pathway for RecA nucleoprotein filaments formed on linear duplex DNA. J. Mol. Biol. 205:695711.
162. Liu, J.,and, K. J. Marians.. 1999. PriA-directed assembly of a primosome on D loop DNA. J. Biol. Chem. 274:2503325041.
163. Liu, J.,, L. Xu,, S. J. Sandler, and, K. J. Marians.. 1999. Replication fork assembly at recombination intermediates is required for bacterial growth. Proc. Natl. Acad. Sci. USA 96:35523555.
164. Livneh, Z.,, O. Cohen-Fix,, R. Skaliter, and, T. Elizur.. 1993. Replication of damaged DNA and the molecular mechanism of ultraviolet light mutagenesis. Crit. Rev. Biochem. Mol. Biol. 28:465513.
165. Livneh, Z.,and, I. R. Lehman.. 1982. Recombinational bypass of pyrimidine dimers promoted by the RecA protein of Escherichia coli. Proc. Natl. Acad. Sci. USA 79:31713175.
166. Lloyd, R. G.,, F. E. Benson, and, C. E. Shurvinton.. 1984. Effect of ruv mutations on recombination and DNA repair in Escherichia coli. Mol. Gen. Genet. 194:303309.
167. Lloyd, R. G.,and, C. Buckman.. 1991. Genetic analysis of the recG locus of Escherichia coli K-12 and of its role in recombination and DNA repair. J. Bacteriol. 173:10041011.
168. Lloyd, R. G.,, S. M. Picksley, and, C. Prescott. 1983. Inducible expression of a gene specific to the recF pathway for recombination in Escherichia coli K12. Mol. Gen. Genet. 190:162167.
169. Lloyd, R. G.,and, G. J. Sharples. 1991. Molecular organization and nucleotide sequence of the recG locus of Escherichia coli K-12. J. Bacteriol. 173:68376843.
170. Lloyd, R. G.,and, G. J. Sharples. 1993. Dissociation of synthetic Holliday junctions by E. coli RecG protein. EMBO J. 12:1722.
171. Lovett, S. T. 2003. Connecting replication and recombination. Mol. Cell 11:554556.
172. Lovett, S. T.,and, R. D. Kolodner. 1989. Identification and purification of a single-stranded-DNA-specific exonuclease encoded by the recJ gene of Escherichia coli. Proc. Natl. Acad. Sci. USA 86:26272631.
173. Luder, A.,and, G. Mosig.. 1982. Two alternative mechanisms for initiation of DNA replication forks in bacteriophage T4: priming by RNA polymerase and by recombination. Proc. Natl. Acad. Sci. USA 79:11011105.
174. Lusetti, S. L.,and, M. M. Cox. 2002. The bacterial RecA protein and the recombinational DNA repair of stalled replication forks. Annu. Rev. Biochem. 71:71100.
175. Madiraju, M. V.,, A. Templin, and, A. J. Clark. 1988. Properties of a mutant recA-encoded protein reveal a possible role for Escherichia coli recF-encoded protein in genetic recombination. Proc. Natl. Acad. Sci. USA 85:65926596.
176. Magee, T. R.,, T. Asai,, D. Malka, and, T. Kogoma. 1992. DNA damage-inducible origins of DNA replication in Escherichia coli. EMBO J. 11:42194225.
177. Magee, T. R.,and, T. Kogoma. 1990. The requirement for RecBC enzyme and an elevated level of activated RecA for induced stable DNA replication in Escherichia coli. J. Bacteriol. 172:18341839.
178. Maisnier-Patin, S.,, K. Nordstrom, and, S. Dasgupta. 2001. Replication arrests during a single round of replication of the Escherichia coli chromosome in the absence of DnaC activity. Mol. Microbiol. 42:13711382.
179. Malkova, A.,, E. L. Ivanov, and, J. E. Haber.. 1996. Double-strand break repair in the absence of RAD51 in yeast: a possible role for break-induced DNA replication. Proc. Natl. Acad. Sci. USA 93:71317136.
180. Mandal, T. N.,, A. A. Mahdi,, G. J. Sharples, and, R. G. Lloyd. 1993. Resolution of Holliday intermediates in recombination and DNA repair: indirect suppression of ruvA, ruvB, and ruvC mutations. J. Bacteriol. 175:43254334.
181. Marians, K. J. 1999. PriA: at the crossroads of DNA replication and recombination. Prog. Nucleic Acid Res. Mol. Biol. 63:3967.
182. Marians, K. J. 2000. PriA-directed replication fork restart in Escherichia coli. Trends Biochem. Sci. 25:185189.
183. Marians, K. J. 2000. Replication and recombination intersect. Curr. Opin. Genet. Dev. 10:151156.
184. Masai, H.,, T. Asai,, Y. Kubota,, K. Arai, and, T. Kogoma. 1994. Escherichia coli PriA protein is essential for inducible and constitutive stable DNA replication. EMBO J. 13:53385345.
185. Masai, H.,, M. W. Bond, and, K. Arai. 1986. Cloning of the Esch- erichia coli gene for primosomal protein i: the relationship to dnaT, essential for chromosomal DNA replication. Proc. Natl. Acad. Sci. USA 83:12561260.
186. McCall, J. O.,, E. M. Witkin,, T. Kogoma, and, V. Roenger-Maniscalco.. 1987. Constitutive expression of the SOS response in recA718 mutants of Escherichia coli requires amplification of RecA718 protein. J. Bacteriol. 169:728734.
187. McGlynn, P.,, A. A. Al-Deib,, J. Liu,, K. J. Marians, and, R. G. Lloyd. 1997. The DNA replication protein PriA and the recombination protein RecG bind D-loops. J. Mol. Biol. 270:212221.
188. McGlynn, P.,and, R. G. Lloyd.. 2000. Modulation of RNA polymerase by (p)ppGpp reveals a RecG-dependent mechanism for replication fork progression. Cell 101:3545.
189. McGlynn, P.,and, R. G. Lloyd. 2001. Action of RuvAB at replication fork structures. J. Biol. Chem. 276:4193841944.
190. McGlynn, P.,and, R. G. Lloyd.. 2001. Rescue of stalled replication forks by RecG: simultaneous translocation on the leading and lagging strand templates supports an active DNA unwinding model of fork reversal and Holliday junction formation. Proc. Natl. Acad. Sci. USA 98:82278234.
191. McGlynn, P.,and, R. G. Lloyd. 2002. Genome stability and the processing of damaged replication forks by RecG. Trends Genet. 18:413419.
192. McGlynn, P.,and, R. G. Lloyd. 2002. Recombinational repair and restart of damaged replication forks. Nat. Rev. Mol. Cell Biol. 3:859870.
193. McGlynn, P.,and, R. G. Lloyd.. 2002. Replicating past lesions in DNA. Mol. Cell 10:700701.
194. McGlynn, P.,, R. G. Lloyd, and, K. J. Marians. 2001. Formation of Holliday junctions by regression of nascent DNA in intermediates containing stalled replication forks: RecG stimulates regression even when the DNA is negatively supercoiled. Proc. Natl. Acad. Sci. USA 98:82358240.
195. McGrew, D. A.,and, K. L. Knight. 2003. Molecular design and functional organization of the RecA protein. Crit. Rev. Biochem. Mol. Biol. 38: 385432.
196. McHenry, C. S. 2003. Chromosomal replicases as asymmetric dimers: studies of subunit arrangement and functional consequences. Mol. Microbiol. 49:11571165.
197. McInerney, P.,and, M. O’Donnell. 2004. Functional uncoupling of twin polymerases: mechanism of polymerase dissociation from a lagging-strand block. J. Biol. Chem. 279:2154321551.
198. Mendonca, V. M.,, H. D. Klepin, and, S. W. Matson. 1995. DNA helicases in recombination and repair: construction of a delta uvrD delta helD delta recQ mutant deficient in recombination and repair. J. Bacteriol. 177:13261335.
199. Menetski, J. P.,, D. G. Bear, and, S. C. Kowalcyzkowski. 1990. Stable DNA heteroduplex formation catalyzed by the Escherichia coli RecA protein in the absence of ATP hydrolysis. Proc. Natl. Acad. Sci. USA 87: 2125.
200. Michel, B.,, S. D. Ehrlich, and, M. Uzest. 1997. DNA double- strand breaks caused by replication arrest. EMBO J. 16:430438.
201. Michel, B.,, M. J. Flores,, E. Viguera,, G. Grompone,, M. Seigneur, and, V. Bidnenko. 2001. Rescue of arrested replication forks by homologous recombination. Proc. Natl. Acad. Sci. USA 98:81818188.
202. Michel, B.,, G. Grompone,, M. J. Flores, and, V. Bidnenko. 2004. Multiple pathways process stalled replication forks. Proc. Natl. Acad. Sci. USA 101:1278312788.
203. Michel, B.,, G. D. Recchia,, M. Penel-Colin,, S. D. Ehrlich, and, D. J. Sherratt. 2000. Resolution of Holliday junctions by RuvABC prevents dimer formation in rep mutants and UV-irradiated cells. Mol. Microbiol. 37:180191.
204. Morimatsu, K.,and, S. C. Kowalczykowski. 2003. RecFOR proteins load RecA protein onto gapped DNA to accelerate DNA strand exchange. A universal step of recombinational repair. Mol. Cell 11:13371347.
205. Morimatsu, K.,, M. Takahashi, and, B. Norden. 2002. Arrangement of RecA protein in its active filament determined by polarized-light spectroscopy. Proc. Natl. Acad. Sci. USA 99:1168811693.
206. Morrow, D. M.,, C. Connelly, and, P. Hieter. 1997. “Break copy” duplication: a model for chromosome fragment formation in Saccharomyces cerevisiae. Genetics 147:371382.
207. Mosig, G. 1987. The essential role of recombination in phage T4 growth. Annu. Rev. Genet. 21:347371.
208. Mosig, G. 1998. Recombination and recombination-dependent DNA replication in bacteriophage T4. Annu. Rev. Genet. 32:37913.
209. Muller, B.,, I. Burdett, and, S. C. West. 1992. Unusual stability of recombination intermediates made by E. coli RecA protein. EMBO J. 11: 26852693.
210. Muller, B.,, I. R. Tsaneva, and, S.C. West. 1993. Branch migration of Holliday junctions promoted by the Escherichia coli RuvA and RuvB proteins: comparison of the RuvAB- and RuvB-mediated reactions. J. Biol. Chem. 268:1717917184.
211. Murialdo, H. 1988. Lethal effect of λ terminase in recombination-deficient Escherichia coli. Mol. Gen. Genet. 213:4249.
212. Myers, R. S.,and, F. W. Stahl.. 1994. Chi and the RecBC D enzyme of Escherichia coli. Annu. Rev. Genet. 28:4970.
213. Nassif, N.,, J. Penney,, S. Pal,, W. R. Engels, and, G. B. Gloor. 1994. Efficient copying of nonhomologous sequences from ectopic sites via P-element-induced gap repair. Mol. Cell. Biol. 14:16131625.
214. Neuwald, A. F.,, L. Aravind,, J. L. Spouge, and, E. V. Koonin. 1999. AAA+: a class of chaperone-like ATPases associated with the assembly, operation, and disassembly of protein complexes. Genome Res. 9: 2743.
215. Nishinaka, T.,, Y. Ito,, S. Yokoyama, and, T. Shibata. 1997. Conformational distortion of sugar moieties in the RecA-bound DNA structure determined by a simulated annealing method. Nucleic Acids Symp. Ser. 37: 269270.
216. Nurse, P.,, J. Liu, and, K. J. Marians.. 1999. Two modes of PriA binding to DNA. J. Biol. Chem. 274:2502625032.
217. Nurse, P.,, K. Zavitz, and, K. Marians. 1991. Inactivation of the Escherichia coli PriA DNA replication protein induces the SOS response. J. Bacteriol. 173:66866693.
218. Olavarrieta, L.,, M. L. Martinez-Robles,, J. M. Sogo,, A. Stasiak,, P. Hernandez,, D. B. Krimer, and, J. B. Schvartzman.. 2002. Supercoiling, knotting and replication fork reversal in partially replicated plasmids. Nucleic Acids Res. 30:656666.
219. Opperman, T.,, S. Murli,, B. T. Smith, and, G. C. Walker. 1999. A model for a umuDC-dependent prokaryotic DNA damage checkpoint. Proc. Natl. Acad. Sci. USA 96:92189223.
220. Opperman, T.,, S. Murli, and, G. C. Walker. 1996. The genetic requirements for UmuDC-mediated cold sensitivity are distinct from those for SOS mutagenesis. J. Bacteriol. 178:44004411.
221. Otsuji, N.,, H. Iyehara, and, Y. Hideshima.. 1974. Isolation and characterization of an Escherichia coli ruv mutant which forms nonseptate filaments after low doses of ultraviolet light irradiation. J. Bacteriol. 117:337344.
222. Pages, V.,and, R. P. Fuchs. 2003. Uncoupling of leading and lagging-strand DNA replication during lesion bypass in vivo. Science 300:13001303.
223. Panyutin, I. G.,and, P. Hsieh. 1993. Formation of single base mismatch impedes spontaneous DNA branch migration. J. Mol. Biol. 230:413424.
224. Parsons, C. A.,, I. Tsaneva,, R. G. Lloyd, and, S. C. West. 1992. Interaction of Escherichia coli RuvA and RuvB proteins with synthetic Holli- day junctions. Proc. Natl. Acad. Sci. USA 89:54525456.
225. Parsons, C. A.,and, S. C. West. 1993. Formation of a RuvAB-Holliday junction complex in vitro. J. Mol. Biol. 232:397405.
226. Picksley, S. M.,, P. V. Attfield, and, R. G. Lloyd.. 1984. Repair of double-strand breaks in Escherichia coli requires a functional recN gene product. Mol. Gen. Genet. 195:267274.
227. Pollard, E.,and, E. P. Randall.. 1973. Studies on the inducible inhibitor of radiation-induced DNA degradation of E. coli. Radiat. Res. 55:265279.
228. Pollard, E. C.,and, P. M. Achey. 1975. Induction of radioresistance in Escherichia coli. Biophys. J. 15:11411154.
229. Pollard, E. C.,, D. J. Fluke, and, D. Kazanis. 1981. Induced radioresistance: an aspect of induced repair. Mol. Gen. Genet. 184:421429.
230. Pollard, E. C.,, S. Person,, M. Rader, and, D. J. Fluke. 1977. Relationship of ultraviolet light mutagenesis to a radiation-damage inducible system in Escherichia coli. Radiat. Res. 72:519532.
231. Postow, L.,, C. Ullsperger,, R. W. Keller,, C. Bustamante,, A. V. Vologodskii, and, N. R. Cozzarelli.. 2001. Positive torsional strain causes the formation of a four-way junction at replication forks. J. Biol. Chem. 276: 27902796.
232. Pugh, B. F.,and, M. M. Cox.. 1987. Stable binding of recA protein to duplex DNA. Unraveling a paradox. J. Biol. Chem. 262:13261336.
233. Pugh, B. F.,and, M. M. Cox.. 1988. High salt activation of RecA protein ATPase in the absence of DNA. J. Biol. Chem. 263:7683.
234. Putnam, C. D.,, S. B. Clancy,, H. Tsuruta,, S. Gonzalez,, J. G. Wetmur, and, J. A. Tainer.. 2001. Structure and mechanism of the RuvB Holliday junction branch migration motor. J. Mol. Biol. 311:297310.
235. Quinones, A.,, C. Kucherer,, R. Piechocki, and, W. Messer. 1987. Reduced transcription of the rnh gene in Escherichia coli mutants expressing the SOS regulon constitutively. Mol. Gen. Genet. 206:95100.
236. Radding, C. M.., 1991. Helical interactions in homologous pairing and strand exchange driven by RecA protein.