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Microbiology and Molecular Biology Reviews, December 1999, p. 751-813, Vol. 63, No. 4
1092-2172/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.

Recombinational Repair of DNA Damage in Escherichia coli and Bacteriophage lambda

Andrei Kuzminov*

Institute of Molecular Biology, University of Oregon, Eugene, Oregon 97403

SUMMARY
TWO-STRAND DNA DAMAGE, RECOMBINATIONAL REPAIR, SOS RESPONSE, AND DNA REPLICATION
    Mechanisms of DNA Damage and Repair
        Damage reversal and one-strand repair.
        Two-strand repair.
        Homologous recombination versus recombinational repair.
        The two mechanisms of two-strand damage.
        The two recombinational repair pathways of E. coli.
        Frequency of two-strand lesions.
        Recombinational repair capacity of E. coli cells.
    SOS Response: Reaction of E. coli to DNA Damage
        Repair instead of DNA damage checkpoints: the prokaryotic strategy.
        Organization of the SOS regulon.
        Levels of SOS induction.
    Cellular Processes That Surround and Complicate Recombinational Repair
        Initiation of chromosomal DNA replication in E. coli.
        Elongation phase of DNA replication in E. coli.
        Initiation of plasmid DNA replication.
        Nucleoid segregation and the problem of accessibility.
    Summary
RECA: HOMOLOGOUS PAIRING ACTIVITY
    recA Gene and Mutants
        recA and peculiarities of recA null mutants.
        Cellular processes dependent on RecA.
    In Vitro Activities of RecA
        RecA without DNA.
        Filament formation by RecA around ssDNA.
        Two DNA-binding sites in RecA filament.
        Cleavage of LexA repressor by RecA filament.
        Detection by RecA filament of homology to ssDNA bound in the primary site.
        Strand exchange between DNA1 and DNA2 catalyzed by RecA filament.
        Assistance for RecA by SSB at all stages.
    Supervision of RecA Activity
        Inhibition by MutS and MutL of pairing between homeologous sequences.
        Possible disruption of pairing of insufficient length by helicase II.
    Summary
RESOLVING RECOMBINATION INTERMEDIATES
    The Three Ways To Remove a Pair of DNA Junctions
RUV LOCUS: PHENOTYPES OF MUTANTS AND GENETIC STRUCTURE
    Interaction of Ruv Proteins In Vitro with Holliday Junctions
    Pairwise Interactions of Ruv Proteins: RuvABC Resolvasome
    RuvAB Translocase
    RecG Helicase
    Three-Strand Junctions and the Hypothetical RecG Pathway
    Summary
REPAIR OF DAUGHTER STRAND GAPS
    Origin of Daughter Strand Gaps and Mechanism of Their Repair: Early Studies
    Presynaptic Phase of Daughter Strand Gap Repair: RecF, RecO, and RecR
        recF, recO, and recR: mutant phenotypes.
        recF, recO, and recR: possible replisome connection revealed by gene structure.
        Properties of RecF, RecO, and RecR and their influence on RecA-promoted reactions in vitro.
        Replisome reactivation and model for RecFOR catalysis of RecA polymerization at daughter strand gaps.
    DNA Topoisomerases and Synaptic Phase of Daughter Strand Gap Repair
        DNA gyrase.
        Topoisomerase I.
    Postsynaptic Phase of Daughter Strand Gap Repair
        One-strand repair: lesion removal and filling in of the gap.
        Removal of DNA junctions and associated RecA filaments.
    Backup Repair of Daughter Strand Gaps: Translesion DNA Synthesis
    Summary
DOUBLE-STRAND END REPAIR
    Origin and Repair of Double-Strand Ends
        Evidence for replication fork disintegration.
        Evidence for replication fork repair by recombination.
        DNA replication primed by double-strand end-promoted recombination.
        Overview of double-strand end repair.
    Preparation of Double-Strand Ends by RecBCD Nuclease for RecA Polymerization
        RecBCD: Genes and mutants.
        RecBCD: Biochemical activities.
        RecBCD: Mechanism of DNA hydrolysis before Chi.
        RecBCD: Mechanism of DNA hydrolysis after Chi.
        RecBCD: RecA filament assembly.
    Postsynaptic Phase of Double-Strand End Repair
        DNA-keeping enzymes.
        Replication fork restart.
        The two pathways for DNA junction removal in double-strand end repair.
    Role of ExoV in Chromosomal DNA Replication
        ExoV and stability of replication forks.
        Excessive DNA degradation affects survival after ionizing radiation more than after UV.
        DNA degradation as a possible backup strategy.
    Double-Strand End Repair in the Absence of RecBCD
        RecE pathway.
        RecF pathway.
        Unified mechanism of double-strand end repair.
    Summary
SITE-SPECIFIC MONOMERIZATION OF THE CHROMOSOME AFTER RECOMBINATIONAL REPAIR
    Genetics of the XerCD-dif System
    In Vivo Biochemistry of the XerCD-dif System
    A Supramolecular Chromosomal Structure around dif?
    Summary
GLOBAL REGULATION OF RECOMBINATIONAL REPAIR
    Regular DNA Replication
    SOS-Induced Conditions
    SOS Expression as a Compensation
    Summary
SINGLE-STRAND ANNEALING: THE PHAGE WAY TO LINK HOMOLOGOUS DOUBLE-STRAND ENDS
    Single-Strand Annealing in DNA Metabolism of Lambdoid Phages
        Overview of SSA repair.
        SSA enzymes of phage lambda .
        SSA enzymes of the Rac prophage.
        Mechanisms of double-strand end repair in lambda  infection: invasion versus annealing.
        Possible role of SSA repair in the life cycle of lambda .
    Single-Strand Annealing Recombination in Plasmids in the Absence of RecA
        Double-strand break repair in plasmids with direct repeats.
        Double-strand break repair in plasmids with inverted repeats.
    Summary
CONCLUSION AND FUTURE DIRECTIONS
ACKNOWLEDGMENTS
REFERENCES


SUMMARY
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Although homologous recombination and DNA repair phenomena in bacteria were initially extensively studied without regard to any relationship between the two, it is now appreciated that DNA repair and homologous recombination are related through DNA replication. In Escherichia coli, two-strand DNA damage, generated mostly during replication on a template DNA containing one-strand damage, is repaired by recombination with a homologous intact duplex, usually the sister chromosome. The two major types of two-strand DNA lesions are channeled into two distinct pathways of recombinational repair: daughter-strand gaps are closed by the RecF pathway, while disintegrated replication forks are reestablished by the RecBCD pathway. The phage lambda  recombination system is simpler in that its major reaction is to link two double-stranded DNA ends by using overlapping homologous sequences. The remarkable progress in understanding the mechanisms of recombinational repair in E. coli over the last decade is due to the in vitro characterization of the activities of individual recombination proteins. Putting our knowledge about recombinational repair in the broader context of DNA replication will guide future experimentation.


TWO-STRAND DNA DAMAGE, RECOMBINATIONAL REPAIR, SOS RESPONSE, AND DNA REPLICATION
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Homologous recombination was described in Escherichia coli in the mid-1940s (351), and for many years it was thought to be the result of a sexual process, analogous to that found in eukaryotes. When the sensitivity to DNA damage of the first recombination-deficient mutants was noticed, it was realized that recombination in this bacterium may serve the needs of DNA repair as well (105, 107, 266, 267). Subsequently, genetic studies delineated two recombinational pathways---the primary, RecBC pathway, serving the needs of "sexual" recombination, and the secondary, RecF pathway, kicking in when the primary pathway is inactive and moonlighting at "postreplication repair" of daughter strand gaps (102, 106, 108). Still later, biochemical characterization of recombinational activities suggested that their primary role is in DNA repair (131, 132). Finally, the realization that disintegrated replication forks are reassembled by recombination justified the "repair" purpose for the RecBC pathway (130, 333) and prompted a revision of our ideas about the relationships of DNA replication and recombination.

The goal of this review is to consolidate genetic data on homologous recombination, physical data on DNA damage and repair, and biochemical data on recombinational enzymes under a different idea in an attempt to highlight new areas for the future in vitro and in vivo experiments. The different idea is that the primary role of the homologous recombination system in E. coli is to repair lesions associated with DNA replication of damaged template DNA (130, 336). Therefore, this review differs from other recent reviews on homologous recombination in E. coli (108, 320, 377) in that its two main emphases are on (i) the evidence for recombinational repair in bacteria and (ii) the interactions of various recombinational repair proteins with each other and with the replication machinery. The recombinational repair machinery is conserved among eubacteria, and so the same two basic pathways are present in such dissimilar species as E. coli and Bacillus subtilis. Therefore, although concentrating on the E. coli recombinational repair paradigm, occasionally I use evidence from other eubacteria.

Mechanisms of DNA Damage and Repair

Damage reversal and one-strand repair. Bacterial genomic DNA, like any macromolecule, is subject to constant chemical and physical assault. Repair of the resulting lesions is essential if DNA is to serve as the template for transcription and its own reduplication. In the course of evolution, a complex enzymatic machinery has evolved to maintain this centrally important molecule in usable form (195). Repair of some DNA modifications simply reverses the damage, returning DNA directly to its original state. For instance, photolyase, using near UV-visible light, splits UV-induced pyrimidine dimers (reviewed in reference 545). Another example is the suicidal Ada protein of E. coli, which transfers a methyl group from the modified base O6-methylguanine to itself (reviewed in reference 580).

Repair of other types of lesions requires removal of a segment of the DNA strand around the lesion. The double-strandedness of DNA provides the means for repairing the resulting single-strand gaps: the removed bases can be resynthesized by using the intact complementary strand as a template. One example of such a strategy is the repair of modified bases that do not cause DNA distortion. The so-called base excision repair system acts with precision---an enzyme called DNA glycosylase removes a modified base to produce an abasic site, the phosphodiester bond at the 5' side of the site is broken, and the repair is completed by a single-base nick translation by DNA polymerase (151) and sealing of the nick by DNA ligase. Another repair system, nucleotide excision repair, deals with DNA-distorting lesions. An excinuclease removes a 12- to 13-nucleotide segment of a single strand centered around the lesion, and the resulting gap is filled in by repair synthesis (reviewed in reference 544). The third repair system, methyl-directed mismatch repair, can liberate up to 1,000 nucleotides from one strand in its efforts to correct a single mismatch arising during DNA replication (reviewed in reference 440). A lesion affecting a single DNA strand is referred to in this review as one-strand lesion, and repair of such DNA damage is referred to as one-strand repair.

Two-strand repair. Although the bulk of DNA damage affects one strand of a duplex DNA segment, occasionally both DNA strands are damaged opposite each other, resulting in two-strand damage, a term proposed by Howard-Flanders (266). To repair two-strand damage without the loss of sequence information, a cell needs a higher level of redundancy, an extra homologous sequence whose strands could be used to fix both DNA strands of the damaged sequence. The principle of such two-strand repair is depicted in Fig. 1. An affected duplex homologously pairs and exchanges strands with an intact homologous duplex (Fig. 1B). The resulting joint molecule is "resolved" by symmetric single-strand cuts in homologous strands, yielding two new DNA molecules, each containing a single one-strand lesion (Fig. 1C). Now the damaged strands can be mended by one-strand repair with the complementary strands as templates (Fig. 1D).


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FIG. 1.   The idea of two-strand repair. (A) A DNA molecule with a two-strand lesion (small open rectangles in the solid duplex) is shown side-by-side with an intact homolog (open duplex). (B) The two sequences have exchanged strands in a homologous region, converting the two-strand lesion into a pair of one-strand lesions. (C) Junction resolution (the strands to cut are shown in panel B) separates the chromosomes from each other. (D) Excision repair removes the one-strand lesions, completing the overall repair reaction. Note that if black and white "parental" DNAs are not identical, the resulting chromosomes may become "recombinant."

Thus, the strategy of the two-strand repair is to convert a two-strand lesion into a pair of one-strand lesions by strand exchange with an intact homologous DNA sequence. Three common phases of the two-strand repair are evident from this scheme. The central phase, during which a damaged DNA sequence trades strands with an intact homologous sequence to form a joint molecule, is called synapsis. In E. coli and other eubacteria, this phase is catalyzed by RecA protein. Accordingly, the preparatory phase preceding the synapsis is called presynapsis, while the resolution of joint molecules is referred to as postsynapsis (103). The four-strand junctions holding the joint molecules together are usually referred to as Holliday junctions, after Holliday, who recognized their importance in one of the early models of homologous recombination (256).

Homologous recombination versus recombinational repair. Since the machinery for the two-strand repair is complex and not copious and since the repair incidents are rather infrequent, this type of repair is more accessible to genetic than to biochemical study. The principal genetic assay for two-strand repair is to monitor the formation of new chromosomes resulting from alternative resolution of joint molecules. A joint molecule (Fig. 2A) can be redrawn to show that the DNA junctions are able to isomerize (Fig. 2B). This isomerization of the junctions creates two possible ways of resolving each junction (shown by numbers beside the arrows [Fig. 2B]). If the resolution is random, in 50% of the cases the participating chromosomes will exchange shoulders, forming two "recombinant" chromosomes (Fig. 2C). If the parental chromosomes were genetically marked, progeny carrying recombinant chromosomes would be detected genetically as having traits that initially resided on separate parental chromosomes.


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FIG. 2.   The four ways to resolve a joint molecule with a double junction. Lowercase letters w, x, y, and z designate unique sites which serve as markers on the homologous chromosomes. A junction is resolved by two symmetrical single-strand cuts (small black arrows in panel B) across each other. Each such diagonal pair of cuts is numbered either 1 or 2 for each junction. If junctions freely isomerize and are resolved independently of each other, four outcomes of the resolution are expected. In two of the outcomes, the chromosome arms will be exchanged, resulting in recombinant chromosomes. (A) A joint molecule with two junctions as shown in Fig. 1B. (B) The same joint molecule isomerized to show both junctions in the open planar configuration (498). (C) The four resolution outcomes, numbered according to the resolution options realized at the left and the right junctions.

Because of this association of the two-strand repair with homologous recombination, the former is better known as recombinational repair. The availability of the homologous recombination assay was a mixed blessing for the development of recombinational-repair concept. On the one hand, most recombinational-repair mutants of E. coli were isolated because of their deficiencies in homologous recombination. On the other hand, since genetic recombination has important evolutionary consequences (181, 418, 736), the recombinational-repair system of E. coli was for a long time viewed from the perspective of its long-term evolutionary value rather than its short-term repair value.

The typical "repair" features of the recombinational-repair system in E. coli are sometimes used as an argument that it could not have evolved due to its role in genetic exchange (131, 132). However, the recombination system of E. coli might have arisen purely for repair purposes but eventually have been integrated into the evolutionary tools of the long-term survival system. Therefore, the strongest argument in favor of the repair role of homologous recombination and against its evolutionary role should come from comparison of the selective values of repair and genetic exchange for the long-term survival of bacteria. Since, due to their decreased viability (see "Frequency of two-strand lesions" below) and high sensitivity to DNA-damaging agents, recombination-deficient mutants are unlikely to survive outside the laboratory, repair must have a high selective value. In contrast, the role of homologous recombination in the E. coli evolution is obscure, since the E. coli genome in nature evolves as a collection of clonal lineages with little recombinational cross talk among the clones and little proven selective value for such horizontal transfer (419, 436). Thus, for the short-term survival, the system of homologous recombination in E. coli has a higher evolutionary value as a DNA repair mechanism than as a mechanism for creating new allelic combinations. This does not mean that the "exchange" consequences of homologous recombination are unimportant for the long-term survival of E. coli; it only means that their selection coefficient is smaller.

Formation of recombinant chromosomes must be a direct consequence of DNA damage repair, because (i) DNA damage greatly stimulates homologous recombination (111, 360, 451, 529) and (ii) the genetic requirements of this damage-stimulated recombination are the same as those of the "spontaneous" recombination. Still, it is possible that this stimulation of homologous recombination in E. coli by DNA damage occurs because DNA damage makes cells "hyper-rec" towards all DNA molecules rather than only towards the damaged ones. However, damage on one DNA stimulates its recombination only with homologous DNA, arguing against the idea of nonspecific activation of recombination by DNA damage (530). Even when damage on one DNA molecule stimulates recombination between sequences absent from the damaged molecule which are situated on other, intact molecules, such "teleactivation" is observed only when the damaged molecule carries homology to the recombining molecules (213). This strict homology requirement for the recombination activation by DNA damage also suggests that repair of certain DNA lesions requires interactions with homologous chromosomes.

The two mechanisms of two-strand damage. Two-strand lesions appear in DNA in two distinct ways. DNA synthesis in a region increases recombination in this region (447), suggesting that one source of two-strand lesions is DNA replication. The fact that replication of DNA containing one-strand lesions stimulates recombination between this DNA and an intact homolog (360) suggests that DNA replication causes two-strand lesions when it runs into unrepaired one-strand lesions. There are at least two mechanisms of replication-dependent conversion of one-strand damage into two-strand damage. In vivo, a noncoding lesion (for example, an abasic site) is an absolute block to DNA replication in growing cells (347); similarly, in vitro, a noncoding lesion in template DNA blocks the progress of the major E. coli DNA polymerases (59). In the chromosome, replication is likely to reinitiate downstream of a noncoding lesion (see "Elongation phase of DNA replication in E. coli" below), leaving behind an unfillable single-strand gap (Fig. 3) (see "Origin of daughter strand gaps and mechanism of their repair: early studies" below). Such an unfillable gap is called a daughter strand gap, since it appears in one of the two daughter branches after the replication fork passage (538, 734). Another type of one-strand lesion, a single-stranded interruption in template DNA, is proposed to cause a disintegration (collapse) of a replication fork (see "Evidence for replication fork disintegration" below) (234, 597). As a result, a double-strand end is detached from the full-length duplex molecule (Fig. 3). Finally, inhibited replication forks are broken, similarly releasing one of the replicating branches as a free double-stranded end (263, 334).


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FIG. 3.   The two major types of replication-induced two-strand lesions. A replication fork moves from left to right along the template DNA with unrepaired one-strand lesions. The left template contains a noncoding lesion (T=T, thymine dimer), the right template has a single-strand interruption. Additional explanations are given in the figure and in the text.

The other principal source of two-strand DNA damage is direct induction. Ionizing radiation (X rays and gamma rays), when passing through a solution, generates free radicals, which damage and break molecules in their immediate vicinity. The energy deposition by gamma radiation allows the formation of clusters of several radicals, so that a big molecule near such clusters can suffer multiple instances of damage (717). Besides chemically modified bases and interruptions in one DNA strand, ionizing radiation also causes double-strand breaks (48, 219). On the average, for every 20 single-strand breaks induced by X rays in DNA, there is one double-strand break (reviewed in reference 324). Another direct two-strand lesion, a cross-link, is observed in DNA treated with psoralen plus UV-light or with mitomycin C (101, 671). In summary, two-strand damage is induced in DNA either directly or as a result of DNA replication on a template DNA containing one-strand damage.

The two recombinational repair pathways of E. coli. The two types of replication-induced two-strand lesions are repaired in E. coli by two separate pathways, both dependent on the recA gene but named after the critical genes that distinguish between them (Fig. 4). Daughter strand gaps are repaired by the RecF pathway (see "Repair of daughter strand gaps" below), while disintegrated replication forks are repaired by the RecBC pathway (see Double-strand and repair" below). The three common phases (see "Two-strand repair" above) of the two repair reactions are (Fig. 4) (i) presynapsis, during which the damaged DNA is prepared for homology search, followed closely by synapsis, during which homologous pairing and strand exchange with the intact sister duplex occur; (ii) DNA replication restart; and (iii) postsynapsis, during which the recombination intermediates are resolved.


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FIG. 4.   The two pathways of recombinational repair in E. coli. On the left, the RecF (daughter strand gap repair) pathway is shown; on the right, the RecBC (double-strand end repair) pathway is shown. Additional explanations are given in the text.

Direct two-strand lesions are repaired by the same two pathways. Double-strand breaks are fixed by the RecBC pathway (555); most probably, they are treated as pairs of independent double-strand ends (see "Overview of double-strand end repair" below). Cross-links are repaired by the combined effort of both the RecBC and the RecF pathways (595), since a fraction of them are apparently converted to double-strand breaks while the rest are converted to unfillable single-strand gaps.

The two recombinational repair pathways are equally important for the repair of DNA damage during normal growth of enteric bacteria, since (i) both the recBC and recF null mutants reduce the viability of E. coli to approximately the same degree (85, 86, 547), and (ii) in Salmonella typhimurium, recombinational repair in the chromosome, detected as a deletion formation between long repeats, is not blocked by single recB or recF mutations but is prevented in a recB recF double mutant (198); physically detected sister chromatid exchange in the E. coli chromosome depends on both the recB and recF genes (630).

Frequency of two-strand lesions. Under conditions of laboratory growth, two-strand lesions are too infrequent to be detectable in wild-type (WT) cells directly by physical techniques, although they are detectable in recombinational repair mutants (434). After massive DNA damage, daughter strand gaps are detected as single-stranded regions of several hundreds of nucleotides in the chromosomal DNA (278, 710) or as interruptions in the newly synthesized DNA (538, 714), double-strand breaks are detected as an immediate chromosome fragmentation (61, 323, 683), and disintegrated replication forks are detected as replication-induced chromosome fragmentation (60, 715).

A more sensitive although less precise indication of the frequency of two-strand lesions during normal growth is the viability of various recombinational repair mutants. Under laboratory conditions, mutants defective at the presynaptic and synaptic phases of recombinational repair (see "Two-strand repair" above) have 25 to 50% viability (85, 86, 547) while those blocked at the postsynaptic phase are 25% viable (370). These approximate values suggest that under laboratory conditions, E. coli experiences two-strand lesions in almost every generation. The importance of this seemingly rare occurrence is raised by the following considerations: (i) a single unrepaired two-strand lesion is a "kiss of death" for the chromosome (268, 595), and (ii) judging by the significant capacity of the E. coli cells to undergo recombinational repair, E. coli cells occasionally experience massive two-strand DNA damage in the wild (see "SOS response: reaction of E. coli to DNA damage" below).

Recombinational repair capacity of E. coli cells. WT E. coli cells grown in a nutritionally poor medium are able to survive 53 to 71 cross-links per chromosome (595). It can be calculated on the basis of the data with excision repair-deficient strains (714) that E. coli cells are still viable after repairing 100 to 200 daughter strand gaps per chromosome. E. coli cells should also be able to tolerate multiple disintegration of replication forks, because recombinational repair should reattach the resulting double-stranded ends to the circular domains of the chromosome. The only two-strand DNA lesion that has proved to be deadly for E. coli is a double-strand break. E. coli survives only two or three double-strand breaks in its chromosome (325, 683), which suggests that whenever a double-strand break occurs in an unreplicated portion of the chromosome, it cannot be repaired. Whether E. coli is an exception among bacteria in its inability to repair multiple double-strand breaks remains to be determined. There is a eubacterium, Deinococcus radiodurans, which can repair >100 double-strand breaks per chromosome (437), but this extreme resistance to DNA damage stands out in the bacterial world.

SOS Response: Reaction of E. coli to DNA Damage

When growing in the laboratory an average E. coli cell may experience two-strand damage once or twice (see "Frequency of two-strand lesions" above). However, its capacity to repair this damage is many times this value (see "Recombinational repair capacity of E. coli cells" above), suggesting that in nature, E. coli may suffer massive DNA damage.

The two main E. coli reservoirs in nature are (i) the animal gut, where the microbe is dividing and concentrated; and (ii) the natural water of lakes and ponds, where the microbe is starving and diluted (560). In the gut, that is, in the environment rich in nutrients and protected from the elements, E. coli is likely to replicate its DNA for many generations without much need to repair it. However, when E. coli finds itself in the water, where DNA replication stops and DNA repair is anemic while the possibilities for damage of DNA are significant, the E. coli genome must accumulate a tremendous amount of DNA damage. Unfortunately, the gut is a discontinuous niche, since the animal the gut belongs to will eventually die; therefore, to survive in the long run, E. coli has to exit the old gut and recolonize a young one. When the battered E. coli from the water eventually makes it to a new gut and starts replicating, it finds its DNA riddled with unrepaired lesions.

The sporadic occurrence of massive DNA damage separated by long periods of undisturbed growth calls for a modest standby repair system, capable of rapid induction in response to increased DNA repair needs. Such an arrangement is indeed found in E. coli; the rapid increase in its DNA repair capacity is called the SOS response (506).

Repair instead of DNA damage checkpoints: the prokaryotic strategy. The bulk of two-strand DNA lesions in enterobacteria are probably generated as a result of DNA replication on template DNA containing one-strand lesions. An easy way to prevent this aggravation would be to stop DNA synthesis altogether when one-strand lesions are sensed. This is exactly what eukaryotic cells do---they employ checkpoint mechanisms to delay chromosomal replication when their DNA is damaged (reviewed in references 295 and 401). Since prokaryotes would also benefit from such a strategy, it was argued that E. coli might have a system to delay DNA synthesis when its chromosome is damaged (73).

However, several observations contradict this attractive idea. The initial inhibition of the DNA synthesis rate in E. coli is dose dependent, so that even after irradiation with almost lethal UV doses, when one would expect a complete replication stop if a checkpoint mechanism operated, the rate of DNA replication is still 20 to 50% of the maximal rate (162, 300). Furthermore, no E. coli mutation has been isolated that would prevent the inhibition of chromosomal replication by DNA damage (as rad9 mutations in yeast or p53 mutations in mammalian cells do). Not surprisingly, preventing the initiation of DNA synthesis with chloramphenicol during irradiation and for a couple of hours thereafter significantly improves the survival of E. coli, especially of recombinational repair-deficient mutants (47, 207, 234, 496, 607). If cells restarted DNA replication only when a "safe" level of DNA damage was attained as a result of repair, there would have been no effect of this drug-mediated inhibition of DNA synthesis on cell survival. Finally, the idea that E. coli has a mechanism to inhibit replication of damaged DNA is incompatible with the observations that E. coli initiates extra rounds of DNA replication from the origin when its DNA is heavily damaged (46, 300, 501). All these phenomena seem to indicate that, in contrast to eukaryotes, E. coli lacks a mechanism to stop chromosomal replication when its DNA is damaged and instead relies on enhanced repair and damage tolerance in its attempt to faithfully replicate the damaged genome.

E. coli and other eubacteria may have evolved such a minimalistic strategy because DNA replication is often the limiting step in their cell cycle (68). Eukaryotes can easily afford a replication delay, since their S phase is only a fraction of their overall cell cycle. In contrast, rapidly dividing E. coli cells have to race against time, since their chromosomal replication may take 1.5 times as long as their cell cycle (see "Cellular processes that surround and complicate recombinational repair" below).

Organization of the SOS regulon. DNA lesions inhibit DNA replication. Inhibition of DNA replication in E. coli induces the SOS response: an increased expression of some 20 genes aimed at restoring the capacity of the chromosome to replicate (Table 1). The resulting enhancement of the ability of the cell to repair and tolerate DNA damage is achieved in several independent ways. The capacity of the cell for excision repair (see "Damage reversal and one-strand repair" above) is enhanced by overproduction of the UvrD helicase and the UvrA and UvrC subunits of the UvrABC excinuclease. Induced amounts of DNA polymerase II increase the capacity of the cell for DNA synthesis across abasic sites (58, 484, 660). Up to a 50-fold increase in the amount of RecA protein (292, 543) and a similar increase in the expression of RecN protein (494) enhance the recombinational repair. The SOS induction makes possible repeated disengagement of replisomes stalled at the lesions in template DNA to allow resumption of the synthesis downstream, a phenomenon known as replisome reactivation (see "Replisome reactivation and model for RecFOR catalysis of RecA polymerization at daughter strand gaps" below). When recombinational repair cannot fix certain DNA lesions, the UmuD'C complex catalyzes translesion DNA synthesis (see "Backup repair of daughter strand gaps: translesion DNA synthesis" below). Overproduction of SfiA protein inhibits cell division (41), providing extra time for completion of recombinational repair. If all these measures fail to restore DNA replication, the lingering SOS induction awakens colicinogenic plasmids and dormant prophages, whose expression lyses the cell. The lysis of doomed cells benefits the viable cells of the same clone when resources are limited, since inviable bacterial cells can multiply for several generations, wasting precious nutrients. The lysis by induction of a prophage or colicinogenic plasmid is therefore an example of "bacterial apoptosis," which could have evolved to increase the number of viable cells in a clone.

                              
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TABLE 1.   E. coli proteins with known functions induced during the SOS response

During the undisturbed growth, induced expression of the SOS genes is prevented by the LexA repressor. LexA dimer binds to a palindromic sequence, the SOS box, in the promoter regions of the SOS genes, precluding initiation of transcription. The SOS box has an inverted repeat consensus 5'-TACTGTATATATATACAGTA-3', where the positions in bold are absolutely conserved (356). The substantial uninduced levels of certain SOS gene products (Table 1) are maintained due to imperfect SOS boxes in their operator regions or due to alternative promoters. Tight regulation of the genes with low-affinity SOS boxes is achieved by the high intracellular concentration of the LexA repressor (more than 1,000 molecules per cell) (558) and by the presence of two SOS boxes in the operators of lexA, recN and colicin genes (563).

The increased expression of the SOS genes in response to DNA synthesis inhibition is a result of inactivation of LexA repressor. The inactivation of LexA repressor is by autocleavage catalyzed by a recombinationally active form of RecA (see "Cleavage of LexA repressor by RecA filament" below). Only LexA molecules that are free in solution can be inactivated (367), which accounts for the late induction of the SOS genes with high-affinity SOS boxes. The two major types of two-strand DNA lesions (Fig. 3) induce the SOS response along the corresponding repair pathways (see "The two recombinational repair pathways of E. coli" above) (426).

Levels of SOS induction. The SOS response is by no means a desperate attempt to stay alive, as its name inaccurately implies (506), but, rather, an orderly and measured reaction of the cell to DNA synthesis inhibition. General information on the E. coli genes induced during the SOS response is summarized in Table 1. The strength of SOS boxes in the operator regions of the SOS genes correlates well with the in vitro LexA repressor affinities for the corresponding promoters and is likely to determine the timing of expression of a given gene during the SOS induction. According to thus inferred order of expression during the SOS induction, the genes of the SOS regulon could be loosely grouped into three categories. The first genes to be induced are mostly those responsible for one-strand repair (uvrA, uvrB, and uvrD) or damage tolerance (polB) (Fig. 5). The LexA repressor itself is also induced immediately. The DinI gene product, which delays activation of translesion DNA synthesis (753), is likely to be synthesized at this stage, too. Increase in expression of the immediately induced genes is usually less than 10 times that of their constitutive expression. If the increased expression of the one-strand repair genes does not help to regain normal rates of DNA synthesis, the genes of recombinational repair, recA and recN, are induced (Fig. 5). The maximal induction of these genes is higher, 20- to 50-fold over their regular levels. When DNA damage is massive, so that even the enhanced recombinational repair cannot overcome the inhibition of DNA replication, the third group of genes, represented by sfiA and umuDC, is called into action. Since these genes are expressed at very low levels during regular DNA synthesis, their SOS induction could be more than 100-fold. Expression of the umuDC operon inhibits recombinational repair and makes possible translesion DNA synthesis (see "Backup repair of daughter strand gaps: translesion DNA synthesis" below), while SfiA protein delays cell division. As DNA replication rates return to normal, the three expression groups of the SOS genes are likely to become repressed in the reverse order (Fig. 5). Alternatively, if a cell cannot repair its DNA damage and is doomed to generate a dead lineage, prophages and colicin plasmids are induced to lyse it (Fig. 5).


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FIG. 5.   An idealized induction kinetics of the four groups of LexA-controlled genes (also, see reference 611). The graph illustrates the well-regulated nature of the SOS response. Both the x axis (time) and y axis (the level of the SOS induction) are in arbitrary units; therefore, the heights of the three curves are not to be compared.

Cellular Processes That Surround and Complicate Recombinational Repair

The poor capacity of E. coli to repair double-strand breaks (see "Recombinational repair capacity of E. coli cells" above) suggests that this type of two-strand DNA damage is unusual in this organism. If one excludes double-strand breaks and DNA cross-links, the remaining two-strand lesions (daughter strand gaps and disintegrated replication forks) are the result of DNA replication on a damaged template DNA. In other words, recombinational repair acts to carry DNA replication through the template DNA containing unrepaired one-strand lesions. From this perspective, recombinational repair is surrounded by DNA replication: it starts when DNA replication stalls, and when it is finished, DNA replication resumes. Therefore, no discussion of recombinational repair is complete without a discussion of the DNA replication mechanisms.

The entire 4.7-Mbp circular chromosome of E. coli is traversed by a single replication bubble emanating from the unique replication origin. Both replication forks of the replication bubble are active; they meet in a chromosome region called the terminus, which is situated across from the origin. The terminus is delineated by termination sites arranged so as to form a replication fork trap---replication forks can enter the terminus, but they cannot exit it (253). To replicate the whole chromosome within a less-than-1-h bacterial cell cycle, replication forks have to proceed at about 650 bp/s (68). However, even the higher speed of almost 800 bp/s is insufficient when the cell cycle of E. coli is squeezed into 24 min in a rich medium. To prevent underreplication, E. coli starts a new round of DNA replication well before the completion of the ongoing round. Thus, in cells growing in a rich medium, there are one to three replication bubbles (two to six replication forks) (244).

Conceptually, replication of the E. coli chromosome is subdivided into three major phases: initiation, elongation, and termination (reviewed in references 25 and 406). Termination is the least understood phase (253) and is not immediately relevant to the needs of recombinational repair, although inhibition of DNA replication associated with termination sometimes causes disintegration of replication forks with their subsequent recombinational repair (263, 573). Elongation is the phase at which the two-strand lesion formation occurs and the recombinational repair machinery meets the replication machinery. Initiation of chromosomal DNA replication is helpful in defining interactions of the key replication proteins. An initiation strategy of multicopy plasmids is relevant because it utilizes the host reinitiation mechanism after the completion of recombinational repair.

Initiation of chromosomal DNA replication in E. coli. For a replication fork to start, the DNA duplex must be open. At the origin of chromosomal DNA replication, this opening is effected by binding of the initiator protein, DnaA. DnaA recognizes and binds to a degenerate nonanucleotide (T/C)(T/C)(A/T/C)T(A/C)C(A/G)(A/C/T)(A/C) (562). At the origin of chromosomal DNA replication, four DnaA recognition sites are found in a cluster. Binding of 10 to 20 DnaA monomers to this cluster of DnaA binding sites leads to an opening of DNA duplex nearby.

In vivo, single-stranded DNA (ssDNA) is immediately complexed with ssDNA-binding protein (SSB), which precludes the binding of many other proteins to this ssDNA. To load DNA replication machinery onto SSB-complexed ssDNA, help from other proteins bound to neighboring duplex regions is needed. In the E. coli chromosomal origin, DnaA itself, still sitting on the adjacent duplex region, assists in this loading.

Since DNA polymerases cannot start DNA synthesis without primers, the 10-nucleotide riboprimers are laid by a special RNA polymerase called primase (DnaG protein). DnaG primase works in a complex with a DNA helicase (encoded by dnaB) that drives DNA unwinding at the replication fork. The complex of DnaG and DnaB proteins is called a mobile primosome; it propagates along the ssDNA in the 5'-to-3' direction, laying primers every 1.5 to 2.0 kb. The mechanics of primosome assembly at the origin of chromosomal DNA replication is as follows. In solution, DnaB protein is always complexed with its inhibitor, DnaC protein. DnaC delivers DnaB helicase to ssDNA if DnaA protein is bound nearby. When DnaB helicase is loaded onto ssDNA and is associated with dnaG primase, the replicative primosome is formed.

The final stage of the replication fork formation is the association of a multisubunit DNA polymerase III (pol III) with the nascent replication bubble. First, a DnaN protein dimer is clamped around a primed segment of DNA to form a ring that slides along the RNA-DNA hybrid or duplex DNA. The DnaN clamp is called the processivity subunit of DNA pol III, since it ensures that DNA polymerase stays bound to DNA during polymerization. DNA synthesis begins when DNA polymerase holoenzyme is loaded onto the DnaN clamp at the primer. There are up to 300 DnaN monomers per cell (79), some 10-fold excess of DnaN dimers over DNA pol III holoenzyme, which is present at 10 to 20 copies per cell (747).

Elongation phase of DNA replication in E. coli. In an established replication fork, DnaB helicase (maybe with the help of auxiliary helicases like Rep and UvrD) unwinds parental duplex DNA while the associated DnaG primase lays primers for both the leading and the lagging strands. DnaN clamps are formed around the primed DNA segments, while the single-stranded regions between primers are complexed with SSB. When the stretch of DNA between the two adjacent primers is duplicated (SSB is apparently displaced), DNA pol III is transferred from its current DnaN ring onto a new DnaN ring, awaiting on the next primer (this explains the requirement for the excess of DnaN subunit over the holoenzyme). The two adjacent newly synthesized DNA stretches, called Okazaki fragments, are separated by a single-strand interruption between the 3' side of one of the fragments and the RNA primer, attached to the 5' side of the other fragment. A one-subunit repair DNA polymerase (DNA pol I) starts DNA synthesis from the 3' side of the interruption, simultaneously degrading the downstream RNA primer with its unique 5'-to-3' exonuclease activity. After the complete removal of the RNA primer, the single-strand interruption is sealed by DNA ligase.

This description corresponds to the mechanism of the lagging-strand DNA synthesis elucidated in vitro. In the reconstituted in vitro systems of the E. coli DNA replication, the lagging-strand synthesis is discontinuous, requiring periodic reloading of DNA pol III, while the leading-strand synthesis is continuous, so that DNA pol III is loaded only once and then is able to replicate megabases of DNA before dissociation (
406). It is said that in vitro the processivity of the leading-strand DNA synthesis is greater than that of the lagging-strand DNA synthesis. If DNA synthesis on the leading and the lagging strands has different processivity in vivo as well, the distribution of the length of daughter strand gaps (see "The two mechanisms of two-strand damage" above), produced during replication of templates with noncoding lesions, would be bimodal, with the gaps in the leading strand being longer than those in the lagging strand. However, the length distribution of daughter strand gaps is unimodal, suggesting that the processivity of DNA synthesis in vivo is comparable for the two strands (710). Indeed, the initial products of DNA synthesis in vivo, detectable in DNA ligase or DNA pol I mutants, are small fragments of the same length (the Okazaki fragments), which argues that E. coli DNA replication in vivo is discontinuous on both strands (383, 471, 483). This conclusion was questioned when it was found that the continuous leading-strand DNA synthesis in vitro may appear discontinuous if the nascent DNA misincorporates uracils instead of thymines, which are then subject to excision repair (472). However, in vivo Okazaki fragments are still generated even when excision of uracils, nucleotide excision repair, base excision repair, and mismatch repair are all inactivated (681, 709, 711), confirming that DNA replication in E. coli cells is discontinuous on both strands. Experiments of a different kind are needed to resolve this discrepancy between the in vitro and in vivo results.

Initiation of plasmid DNA replication. Small multicopy plasmids of E. coli initiate their DNA replication in a different way---they use the priming mechanism employed in the host recombinational repair of disintegrated replication forks but substitute a transcription intermediate for the recombination intermediate. DNA at their replication origins is first transcribed, and a portion of the resulting several-hundred-nucleotide transcript forms a stable RNA-DNA hybrid with its template, displacing the complementary DNA strand into the so-called R loop. Then the RNA portion of the hybrid is cleaved at a specific site to provide a primer for a limited DNA synthesis carried out by DNA pol I.

The complementary DNA strand, displaced as a result of this DNA pol I-catalyzed synthesis, is complexed with SSB. For this reason, it would be inert for any further transaction if not for the second mechanism of primosome assembly available in E. coli. In contrast to the first mechanism, which starts with DnaA multimer binding to a cluster of its recognition sequences, the second mechanism begins with PriA binding to what looks like a replication fork framework. As determined in vitro (
368, 466, 467), the sequence of events in the PriA-dependent primosome formation is as follows: (i) PriA binds to ssDNA near the branching point with the duplex DNA; (ii) PriB binds to PriA and stabilizes PriA binding to ssDNA; (iii) DnaT binds to the PriA-PriB complex, which is then further stabilized by binding of PriC; (iv) DnaB (replicative helicase) is delivered from the complex with DnaC (replicative helicase inhibitor) onto this branched DNA-PriABC-DnaT complex to form a preprimosome; and (v) DnaG (primase) associates with the preprimosome to lay primers for DNA synthesis. The key event in the primosome assembly is the competition for DnaB, with PriABC plus DnaT at the DNA substrate versus DnaC in solution. In vitro, the PriA-dependent primosome has a composition different from the DnaA-dependent primosome in that PriA and PriB proteins seem to stay associated with DnaB during the ensuing DNA synthesis (6).

Nucleoid segregation and the problem of accessibility. Nucleoid segregation sets the "window of opportunity" for recombinational repair. In rapidly growing E. coli cells, nuclear bodies, or nucleoids, are seen in the electron microscope as dimers (688, 741), and even after being freed from their "cells," many purified nucleoids have a doublet appearance (240, 458, 490), indicating that the separation of nascent nucleoids is concomitant with DNA replication. It has been proposed that replicated daughter branches of the parental chromosome do not stay entangled but from the very beginning form separate bodies, growing out from the replication point in opposite directions (381, 740). Spatial separation of the origin-proximal markers after origin duplication was visualized in live cells (215).

This continuous separation of daughter nucleoids should create a problem for reactions that rely on interactions between sister chromosomes. Still, it could be argued that although sister chromosomes may appear separate, they interact at the molecular level as if residing in the same space. This question was addressed by comparing site-specific recombination between plasmid molecules in vitro with the same interplasmidic reaction in vivo (250). The efficiency of the in vitro reaction depends on the plasmid concentration; the plasmid DNA concentration in vivo can be estimated from the plasmid copy number. It was expected that the effective in vivo concentration would be higher due to a variety of cellular factors. Contrary to that expectation, it was found that the effective in vivo concentration is an order of magnitude lower, suggesting that when homologous DNAs are searching for each other in vivo, they face the problem of restricted accessibility (250). In other words, if recombinational repair is to mend two-strand damage in one of the daughter DNAs, it has a certain time frame to do it before the daughter sequences are segregated into separate nucleoids.

Summary

DNA damage can be classified as affecting either one or both strands in a particular sequence. Similarly, cellular DNA repair mechanisms are categorized as either one-strand or two-strand repair. Since the two-strand repair frequently spins off recombinant chromosomes, it is generally known as recombinational repair. The bulk of the two-strand damage is generated by DNA replication, when a replisome stumbles upon an unrepaired one-strand lesion. The two major replication-induced two-strand lesions are daughter strand gaps and disintegrated replication forks. In E. coli, daughter strand gaps are repaired by the RecF pathway whereas disintegrated replication forks are repaired by the RecBCD pathway. Two-strand DNA lesions occur infrequently during regular growth in the laboratory, but in real life E. coli must occasionally experience massive DNA damage---hence the inducible DNA repair capacity, called the SOS response.

Recombinational repair acts to carry the replication apparatus through the template DNA containing unrepaired one-strand lesions and, in this respect, must collaborate with the chromosomal replication and the nucleoid segregation machinery. This puts recombinational repair reactions in a specific context, with their own idiosyncrasies, unresolved problems, and gray areas. One such controversy, bearing on the damage formation mechanisms, is whether in vivo replication is discontinuous on both DNA strands. One of the major complications for the recombinational repair, which depends on the availability of an intact sister duplex, is the accessibility of this duplex, because the sister nucleoids are continuously segregated as the cell grows. This aspect of the in vivo chromosomal metabolism is almost unstudied.


RECA: HOMOLOGOUS PAIRING ACTIVITY
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For damaged DNA to be repaired with the help of an intact homologous sequence, the two DNAs need to (i) find each other among numerous unrelated sequences and (ii) trade strands to make possible one-strand repair of the damage in the affected sequence. In E. coli, these intricate and seemingly intelligent reactions are catalyzed by a single, relatively small enzyme called RecA. The 38-kDa RecA searches for homology both catalytically and stoichiometrically, since the active species is a polymer comprising hundreds of RecA monomers.

recA Gene and Mutants

recA and peculiarities of recA null mutants. recA happened to be the first E. coli recombinational repair gene to be discovered (107). recA is not a part of any operon, is surrounded by genes unrelated to DNA metabolism, and has its own promoter and terminator (260, 546). Normally, recA expression maintains 1,000 to 10,000 RecA monomers per cell (292, 445, 543, 558). RecA production is induced by DNA-damaging treatments such as UV irradiation or nalidixic acid, resulting in up to a 50-fold increase in the amount of the protein (see "Organization of the SOS regulon" above) (292, 543).

No extragenic suppressors that would cancel the phenotypes of null recA alleles have been found, suggesting that recA is the only gene of its kind in the E. coli genome. recA mutations are unusually pleiotropic (for an early yet informative review, see reference 102). recA cells are extremely sensitive to DNA damage (107, 267, 689); nevertheless, recA null mutants are viable, although they grow slower than the WT cells. The slower growth of recA cultures is due to the continuous generation of dead cells (229) rather than because of growth defects. The fraction of dead cells in laboratory cultures of recA mutants reaches 50% (85).

WT cells stop cell division in response to inhibition of DNA synthesis, but recA mutant cells continue to divide under these conditions, producing anucleate cells (272); RecA inhibits cell division via SOS induction when there are irregularities with DNA replication (see "Organization of the SOS regulon" above). Under regular growth conditions, about 10% of recA cells lack chromosomal DNA; up to 20% of the total DNA in recA mutant cultures is degraded at any given moment (84, 105). This DNA degradation must target particular nucleoids, hence the asynchrony phenotype displayed by recA mutant cultures: whereas most cells in the WT cultures grown in a rich medium have either four or eight nucleoids, recA mutant cells have all numbers of nucleoids from zero to eight (598, 599).

Cellular processes dependent on RecA. The induction of the SOS response, the reaction of the cell to massive DNA damage, is absolutely dependent on RecA. RecA is activated by damaged DNA, and the activated form of RecA catalyzes self-cleavage of LexA repressor (see "Cleavage of LexA repressor by RecA filament" below). The SOS response increases the capacity of the cell to repair and tolerate DNA damage and also delays cell division. The damage to bacterial DNA also causes prophage induction, i.e., the lytic development of latent bacteriophages. Similarly to LexA repressor, bacteriophage repressors cleave themselves in the presence of activated RecA, but the phage induction has nothing to do with repair of cellular DNA and is in fact lethal to the host cell.

RecA plays a pivotal role in recombinational repair of such two-strand DNA lesions as daughter strand gaps, double-strand breaks, and interstrand cross-links. For example, while WT E. coli cells survive 53 to 71 cross-links per chromosome, recA cells are killed by a single cross-link (
595). RecA-dependent mechanisms of recombinational repair are discussed below (see "Resolving recombination intermediates" and "Repair of daughter strand gaps").

A bacterial cell can acquire a linear piece of chromosome from another cell in a variety of ways (reviewed in reference 9). During conjugation, this piece is transferred from another live cell, which has a conjugative plasmid integrated into its chromosome. During transduction, this piece is delivered by a bacteriophage, whose capsid had mistakenly packaged a fragment of the host DNA instead of the phage chromosome. During transformation, a cell picks up a piece of DNA from the environment, from a dead decomposing cell. Such an exogenous piece of DNA can be inserted, in whole or in part, into the chromosome in a RecA-dependent process; recA mutants are profoundly defective in all types of chromosomal recombination (reviewed in reference 399).

Two-strand DNA damage induces RecA-dependent mutagenesis. In general, DNA modification is mutagenic in that it causes point mutations, and especially strong mutagens are those that make DNA bases change their coding interactions. For example, guanine recognizes cytosine in the opposite position, but oxidation of guanine could make it recognize adenine, causing a misincorporation and, ultimately, a point mutation (433). However, RecA-dependent mutagenesis has a completely different nature. Some DNA modifications generate the so-called noncoding lesions, i.e., bases that are missing or so distorted that they are no longer recognized by DNA polymerases. DNA replication can bypass such a noncoding lesion in a RecA-dependent reaction, during which a DNA polymerase sometimes has to incorporate a random nucleotide in the new DNA chain across the damaged position, which often results in mutations (see "Backup repair of daughter strand gaps: translesion DNA synthesis" below).

In Vitro Activities of RecA

The variety of phenotypes of recA mutant cells stems from a single deficiency, the inability to form an active RecA filament. The in vitro properties and activities of RecA filament still bewilder and fascinate those who study them. For in-depth treatment of the enchanting RecA biochemistry, see the excellent reviews by Kowalczykowski (319) and Roca and Cox (524).

RecA without DNA. In high-concentration solutions, RecA aggregates to form oligomers, filaments, and bundles (70, 71, 246, 737). One of the major species in these aggregates consists of rings of six to eight monomers (70, 71, 246). These rings are characterized by electron microscopy for RecA from Thermus aquaticus, due to their greater stability (758). Surprisingly, they resemble in gross details both the hexameric rings of helicases like DnaB or RuvB and the F1-ATPase (168, 760).

The crystal structure of RecA, solved at 2.3-Å resolution, shows a spiral filament with six RecA monomers per turn (632). There is enough space inside the filament to accommodate two interacting DNA molecules. Although the crystals were formed either in the presence of ADP or without nucleotide cofactor, and so represent RecA species inactive in recombinational reactions, they show the overall arrangement of the structural elements within the RecA monomer as well as the way in which the monomers are arranged into filaments.

Filament formation by RecA around ssDNA. In vitro, in the presence of physiological concentrations of Mg2+ and ATP, RecA assembles around ssDNA into a helical filament (Fig. 6), an entity proficient in all known RecA activities (in the absence of the nucleotide cofactor or in the presence of ADP, RecA forms similar filaments but with different parameters; since such filaments are inactive in RecA-promoted reactions, they are not discussed in this review). At physiological pH, RecA filament does not readily assemble on duplex DNA; however, a filament assembled on ssDNA extends into a contiguous double-stranded region (362, 572). Every RecA monomer within a filament binds a single ATP molecule (307). RecA filament assembled on ssDNA slowly hydrolyses ATP at a rate of about 30 ATP molecules per min per monomer (69); duplex DNA-bound filament has an even lower ATPase activity (362, 502).


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FIG. 6.   RecA filament in vitro: photographs and molecular model. (A) A relaxed circular duplex DNA completely covered by RecA filament. Naked duplex DNA of the same length, lying mostly inside the RecA filament, illustrates that DNA within RecA filament is stretched 1.5 times. Reprinted from reference 625 with permission of the publisher. (B) A RecA-covered circular ssDNA molecule; the arrow points to a segment complexed by SSB. Below, another ssDNA circle of the same length can be seen, but since it is entirely complexed with SSB, it appears very different, i.e., much smaller and kinky. Note that while RecA filament stretches ssDNA, SSB compacts it. Reprinted from reference 246 with permission of the publisher. (C) A crystal structure-based molecular model of an 18-monomer segment of RecA filament with three symmetry-related monomers in gray. Each of these monomers is enlarged on the right to show residues conserved among eubacterial RecA proteins (see reference 524 for details). In such a filament, the 5' end of DNA1 would be at the top. Reprinted from reference 524 with permission of the publisher.

The ATP hydrolysis is not needed for the assembly of RecA filament on DNA, since the same recombination-proficient filament is formed in the presence of a nonhydrolyzable ATP analog, ATPgamma S (169). One role for ATP hydrolysis may be to promote filament disassembly, since RecA filaments formed in the presence of ATPgamma S do not disassemble on their own (reference 523 and references therein). Most of the measurements of recombination-proficient RecA filaments were done in the presence of ATPgamma S because of the greater filament stability in the absence of ATP hydrolysis; however, parameters of ATP-containing filaments are very similar (627).

The width of ATP-containing RecA filament is about 10 nM (100 Å) (165, 169), which is five times the width of duplex DNA. The ATPgamma S-containing filament has about six RecA monomers per 95-Å turn, with each RecA monomer binding about 3 nucleotides of ssDNA (311). The stoichiometry of duplex DNA binding is the same: one RecA monomer binds 3 bp, or a single 95-Å helical turn of RecA filament holds about 18 bp (the axial spacing between adjacent base pairs is 5.1 Å) (153, 161). Since the axial spacing between adjacent base pairs in native DNA duplex is 3.4 Å, it is said that duplex DNA inside RecA filament is extended 1.5 times (165, 626) (Fig. 6A). This extension, which is surprisingly close to the maximally extended DNA state of 1.7 (113, 608), is thought to facilitate homology recognition between two DNA molecules, captured by RecA filament (see "Detection by RecA filament of homology to ssDNA bound in the primary site" below).

RecA filament grows at its ends. Growth in the 5'-to-3' direction relative to the bound ssDNA is several times more efficient than growth in the opposite direction (362, 514, 571, 572). The maximal rate of RecA filament assembly in vitro is 30 to 40 monomers per s (514). Assuming a DNA binding stoichiometry of one RecA monomer per 3 nucleotides, a growing RecA filament engulfs about 100 nucleotides of ssDNA per second. In vitro, at the same time as the 3' end of RecA filament is growing, the 5' end may begin slowly disassembling (362, 569). In effect, the growing RecA filament under these conditions treadmills along the DNA.

Two DNA-binding sites in RecA filament. Soon after the beginning of biochemical characterization of RecA protein, it was realized that, to promote homologous pairing, RecA must have at least two DNA-binding sites: the primary site accommodating DNA1, around which the filament was assembled, and the secondary site for DNA2, to be compared with DNA1 (269). Since then, the idea of at least two DNA-binding sites within RecA filament has been substantiated with a variety of evidence.

In vitro, RecA promotes both three-strand exchange (between an ssDNA1 and a duplex DNA2) and four-strand exchange (between a duplex DNA1 with a single-stranded tail and a fully duplex DNA2), implying the ability of RecA filament to handle up to four DNA strands. However, the only DNA strands in these reactions fully protected by RecA filaments against DNase degradation are the ssDNA1 or the outgoing identical strand (in these experiments, SSB was absent), suggesting that either the hybrid duplex or the alternative duplex is excluded from the filament (98, 99). Moreover, RecA cannot catalyze strand exchange restricted to fully duplex DNA regions (120, 363), indicating that it cannot handle four DNA strands at the same time (reviewed in reference 129).

RecA filament has the primary site that binds ssDNA during filament assembly but can also accommodate duplex DNA. In addition, RecA filament has the secondary binding site, which can transiently bind duplex DNA if the primary site is occupied by ssDNA. If the primary site is occupied by duplex DNA, the secondary binding site can transiently bind ssDNA. If the primary site is occupied by ssDNA, the secondary site can stably bind an unrelated ssDNA (326, 421). Finally, in the presence of ATPgamma S and high Mg2+ concentrations, RecA filament can stably bind two DNA duplexes, but it is unclear whether they have to be homologous (762). Therefore, it seems that RecA has a primary binding site deep within the filament, accommodating up to two DNA strands, and a secondary binding site at the periphery of the filament, again accommodating up to two DNA strands.

ssDNA1 is bound by RecA filament along its sugar-phosphate backbone, so that DNA bases face inward (154, 349) and are ordered perpendicularly to the filament axis (326). Duplex DNA1 is bound by RecA filament along its minor groove (154, 161, 329). In contrast, binding by a RecA-dsDNA1 filament of the second duplex does not involve its minor groove (762).

Cleavage of LexA repressor by RecA filament. RecA filament holding a single DNA strand promotes autocleavage of the SOS response repressor, LexA (259, 366), as well as autocleavage of phage repressors (172, 522, 559) and of the UmuD protein (77). It is said that in these reactions RecA plays a role of coprotease, because there are conditions under which LexA, phage lambda  repressor, and UmuD cleave themselves in the absence of RecA (77, 364). The LexA-binding site lies deep within the filament groove and overlaps with the secondary DNA-binding site (759), explaining why, when both DNA-binding sites are occupied by ssDNA, LexA cleavage is inhibited (152, 515, 642). RecA filament assembled on duplex DNA promotes LexA cleavage at 5 to 20% of the rate observed with a filament assembled on ssDNA (152, 642). This feature of the SOS repressor cleavage makes biological sense---if all the single-stranded regions associated with DNA lesions are made double stranded (supposedly by pairing with intact homologous sequences), there is no reason to boost the repair capacity of the cell any further.

The RecA-promoted LexA autocleavage is rapid and is independent of Mg2+ concentrations in the range of 1 to 10 mM (345). In contrast, RecA-promoted autocleavage of UmuD and phage repressors is slow and is observed only at Mg2+ concentrations of 10 mM or higher (77, 172, 522, 559). The supposed biological significance of these differences, in relation to the idea that the SOS response needs to be induced early while phage repressors need to be cleaved only in moribund cells, will become clear later (see "SOS-induced conditions" below).

Detection by RecA filament of homology to ssDNA bound in the primary site. Although RecA forms a filament around ssDNA1 in a sequence-independent manner, the filament itself is "a sequence-specific DNA-binding entity, with the specificity determined by the bound DNA" (524). The amount of nonhomologous DNA in vivo is overwhelming, since even an identical sequence, shifted a single nucleotide out of register, becomes perfectly heterologous to DNA1. Heterologous DNA is not neutral in homology searches: preincubation of presynaptic filaments with heterologous DNA inhibits subsequent homologous pairing (214). The problem of the complexity of natural DNA is compounded by the enormous intracellular DNA packing densities, measured in E. coli at 20 to 100 mg/ml (57), and the restricted accessibility due to the nucleoid segregation (see "Nucleoid segregation and the problem of accessibility" above). Under these conditions, RecA has to find homology to the damaged DNA within minutes, as illustrated by the extreme DNA damage sensitivity of a partially active recA allele, proficient in homologous recombination in vivo and capable of "slow" recombinational reactions in vitro (273). If not repaired quickly, the damaged DNA could be degraded or segregated from its intact sister or could bring about even greater damage if the upcoming replication fork runs into it. On the other hand, if recombinational repair mends DNA damage mostly at replication forks, the affected and the intact homologous DNA segments should initially be in close proximity with each other.

The mechanism of homology search by RecA filament is still an enigma. An algorithm of homology search is likely to require repeated juxtaposition of short segments of DNA1 with short segments of duplex DNA. If RecA filament is able to juxtapose two potentially nonhomologous sequences, how does it then let go a duplex which proved to be nonhomologous? One possibility was that a nonhomologous duplex is expelled from the filament with the help of ATP hydrolysis. However, in vitro RecA catalyzes homologous pairing in the presence of nonhydrolyzable ATP analogs (258, 322, 428). Moreover, a mutant RecA protein with a 100-fold in vitro defect in ATP hydrolysis, which is activated by ATP as if it were ATPgamma S, still catalyzes homologous pairing, both in vivo and in vitro (82), so the homology search does not require ATP hydrolysis.

Kinetic experiments show that the homology search in vitro is reversible, follows second-order kinetics (i.e., it depends on the concentrations of both interacting DNAs), and is rapid compared to the next stage of pairing (31, 752). Under these conditions, a short segment of RecA-DNA1 complex is estimated to try 102 to 103 various duplex DNA segments per s, with the high iteration frequency demanding that the search be based on soft interactions only (752). However, these interactions are strong enough to cause partial unwinding of nonhomologous DNA within RecA filaments (137, 536). This unwinding is most probably caused by DNA2 extension inside the filament, as RecA puts it in register with DNA1. The duplex DNA2 is approached along its minor groove by the RecA-complexed ssDNA1 (27, 329, 495), and in the synaptic complex, all three DNA strands are underwound to the same extent of 19 nucleotides per turn (301). This underwinding may allow ssDNA1 to be accommodated in the otherwise too narrow minor groove of dsDNA2 (40).

The configuration of the three DNA strands in the synaptic complex has been a matter for debate and experimentation (129). An interesting idea was that the three strands form a DNA triplex, in which the homology recognition occurs. However, since the minor groove of the duplex DNA does not have enough determinants for homology recognition, the idea of a triplex requires ssDNA1 to interact with dsDNA2 via the major groove of the latter, which contradicts the available experimental evidence (see above). Also, no triplexes are detected in the synaptic complexes; instead, ssDNA1 is seen already paired with the complementary strand of dsDNA2 whereas the identical strand of dsDNA2 is displaced into the major groove of the nascent duplex (2, 495).

Strand exchange between DNA1 and DNA2 catalyzed by RecA filament. When homology is found (i.e., when a homologous duplex DNA2 is aligned with ssDNA1), RecA filament catalyzes the exchange of strands between the two DNA molecules. In this process, ssDNA1 forms hydrogen bonds with the complementary strand of DNA2 while the identical strand of the duplex is displaced (135) (Fig. 7). The outgoing strand of DNA2 is accommodated in the secondary ssDNA binding site at the periphery of the RecA filament to be extracted from there later by SSB (346, 420).


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FIG. 7.   The distinguishable phases of RecA-promoted reactions, as they are thought to happen in vivo. Black lines indicate a damaged duplex; white lines indicate an intact duplex; the open rectangle with rounded corners indicates a RecA filament; the irregularity in one of the black DNA strands indicates a noncoding lesion. The left side represents daughter strand gap repair; the right side represents double-strand end repair. Explanations are given in the figure.

The RecA-catalyzed strand exchange phase can be further subdivided into (i) the "initial" strand exchange, which happens concomitantly with homologous recognition, is limited in length, and does not require ATP hydrolysis by RecA, and (ii) "hybrid duplex extension," which requires ATP hydrolysis. Hybrid duplex exten