DNA Replication PDF

DNA REPLICATION  DEFINITION DNA replication is an enzymatic polymerization reaction in which DNA strand is used as a template to synthesize a copy with a base sequence that is complementary to the template. Replication is semi conservative; each parent single strand is...

DNA REPLICATION  DEFINITION DNA replication is an enzymatic polymerization reaction in which DNA strand is used as a template to synthesize a copy with a base sequence that is complementary to the template. Replication is semi conservative; each parent single strand is present in one of the double-stranded progeny molecules.  The original Watson-Crick proposal for the The proposal for the structure of DNA replication of a double-helical molecule of DNA by Watson and Crick in 1953 was accompanied by a suggested mechanism for its “self-duplication.” The two strands of the double helix are held together by hydrogen bonds between the bases. Individually, these hydrogen bonds are weak and readily broken. Watson and Crick envisioned that replication occurred by gradual separation of the strands of the double helix, much like the separation of two halves of a zipper.  Semiconservative replication Because the two strands are complementary to each other, each strand contains the information required for construction of the other strand. Thus once the strands are separated, each can act as a template to direct the synthesis of the complementary strand and restore the double-stranded state.  ALTERNATE/OLD MODELS OF DNA REPLCATION the two original strands would remain together (after serving as templates), as would the The parental strands would be broken into fragments, and the new strands would be two newly synthesized strands. As a result, one of the daughter duplexes would contain synthesized in short segments. Then the old fragments and new segments would be only parental DNA, while the other daughter duplex would contain only newly joined together to form a complete strand. As a result, the daughter duplexes would synthesized DNA. contain strands that were composites of old and new DNA.  To decide among these three possibilities, it was necessary to distinguish newly synthesized DNA strands from the original DNA strands that served as templates. This was accomplished in studies on bacteria in 1957 by Matthew Meselson and Franklin Stahl of the California Institute of Technology who used heavy (15N) and light (14N) isotopes of nitrogen to distinguish between parental and newly synthesized DNA strands. These researchers grew bacteria in medium containing 15N-ammonium chloride as the sole nitrogen source. Consequently, the nitrogen-containing bases of the DNA of these cells contained only the heavy nitrogen isotope. Cultures of “heavy” bacteria were washed free of the old medium and incubated in fresh medium with light, 14N-containing compounds, and samples were removed at increasing intervals over a period of several generations. DNA was extracted from the samples of bacteria and subjected to equilibrium density- gradient centrifugation. In this procedure, the DNA is mixed with a concentrated solution of cesium chloride and centrifuged until the double-stranded DNA molecules reach equilibrium according to their density.  Experiment demonstrating that DNA replication in bacteria is semiconservative DNA was extracted from bacteria at different stages in the experiment, mixed with a concentrated solution of the salt cesium chloride (CsCl), placed into a centrifuge tube, and centrifuged to equilibrium at high speed in an ultracentrifuge. Cesium ions have sufficient atomic mass to be affected by the centrifugal force, and they form a density gradient during the centrifugation period with the lowest concentration (lowest density) of Cs at the top of the tube and the greatest concentration (highest density) at the bottom of the tube. During centrifugation, DNA fragments within the tube become localized at a position having a density equal to their own density, which in turn depends on the ratio of 15N/14N that is present in their nucleotides. The greater the 14N content, the higher in the tube the DNA fragment is found at equilibrium. Figure: The single tube on the left indicates the position of the parental DNA and the positions at which totally light or hybrid DNA fragments would band. (b) Experimental results obtained by Meselson and Stahl. The appearance of a hybrid band and the disappearance of the heavy band after one generation eliminates conservative replication. The subsequent appearance of two bands, one light and one hybrid, eliminates the dispersive scheme.  In the Meselson-Stahl experiment, the density of a DNA molecule is directly proportional to the percentage of 15N or 14N atoms it contains. If replication is semiconservative, one would expect that the density of DNA molecules would decrease during culture in the 14N-containing medium. After one generation, all DNA molecules would be 15N-14N hybrids, and their buoyant density would be halfway between that expected for totally heavy and totally light DNA. As replication continued beyond the first generation, the newly synthesized strands would continue to contain only light isotopes, and two types of duplexes would appear in the gradients: those containing 15N–14N hybrids and those containing only 14N. As the time of growth in the light medium continued, a greater and greater percentage of the DNA molecules present would be light. However, as long as replication continued semiconservatively, the original heavy parental strands would remain intact and present in hybrid DNA molecules that occupied a smaller and smaller percentage of the total DNA. The results of the density-gradient experiments obtained by Meselson and Stahl demonstrate unequivocally that replication occurs semiconservatively.  By 1960, replication had been demonstrated to occur semiconservatively in eukaryotes as well. The original experiments were carried out by J. Herbert Taylor of Columbia University. Cultured mammalian cells were allowed to undergo replication in bromodeoxyuridine (BrdU), a compound that is incorporated into DNA in place of thymidine. Following replication, a chromosome is made up of two chromatids. After one round of replication in BrdU, both chromatids of each chromosome contained BrdU. After two rounds of replication in BrdU, one chromatid of each chromosome was composed of two BrdU-containing strands, whereas the other chromatid was a hybrid consisting of a BrdU-containing strand and a thymidine-containing strand. The thymidine-containing strand had been part of the original parental DNA molecule prior to addition of BrdU to the culture.  REPLICATION IN BACTERIAL CELLS Replication in bacterial cells is better understood than the corresponding process in eukaryotes. The early progress in bacterial research was driven by genetic and biochemical approaches including: 1. The availability of mutants that cannot synthesize one or another protein required for the replication process. The isolation of mutants unable to replicate their chromosome may seem paradoxical: how can cells with a defect in this vital process be cultured? This paradox was solved by the isolation of temperature- sensitive (ts) mutants, in which the deficiency only reveals itself at an elevated temperature, termed the nonpermissive (or restrictive) temperature. When grown at the lower (permissive) temperature, the mutant protein can function sufficiently well to carry out its required activity, and the cells can continue to grow and divide. Temperature-sensitive mutants have been isolated that affect virtually every type of physiologic activity, and they have been particularly important in the study of DNA synthesis as it occurs in replication, DNA repair, and genetic recombination. 2. The development of in vitro systems in which replication can be studied using purified cellular components. In some studies, the DNA molecule to be replicated is incubated with cellular extracts from which specific proteins suspected of being essential have been removed. In other studies, the DNA is incubated with a variety of purified proteins whose activity is to be tested. Taken together, these approaches have revealed the activity of more than 30 different proteins that are required to replicate the chromosome of E. coli. Replication in bacteria and eukaryotes occurs by very similar mechanisms, and thus most of the information presented in the discussion of bacterial replication applies to eukaryotic cells as well.  Replication Forks and Bidirectional Replication Replication begins at a specific site on the bacterial chromosome called the origin. The origin of replication on the E. coli chromosome is a specific sequence called oriC, 245 bp long, where a number of proteins bind to initiate the process of replication. Once initiated, replication proceeds outward from the origin in both directions, that is, bidirectionally. The sites where the pair of replicated segments come together and join the nonreplicated DNA are termed replication forks. Each replication fork corresponds to a site where – (1) the parental double helix is undergoing strand separation – (2) nucleotides are being incorporated into the newly synthesized complementary strands The two replication forks move in opposite directions until they meet at a point across the circle from the origin, where replication is terminated. The two newly replicated duplexes detach from one another and are ultimately directed into two different cells.  Unwinding the Duplex and Separating the Strands Separation of the strands of a circular, helical DNA duplex poses major topological problems. Separation of the two strands at the free end would generate increasing torsional stress in the rope and cause the unseparated portion to become more tightly wound. When a circular or attached DNA molecule is replicated, the DNA ahead of the replication machinery becomes overwound and accumulates positive supercoils. Consequently, movement of the replication fork generates positive supercoils in the un-replicated portion of the DNA ahead of the fork. When one considers that a complete circular chromosome of E. coli contains approximately 400,000 turns and is replicated by two forks within 40 minutes, the magnitude of the problem becomes apparent. Cells contain enzymes, called topoisomerases, that can change the state of supercoiling in a DNA molecule. One enzyme of this type, called DNA gyrase, a type II topoisomerase, relieves the mechanical strain that builds up during replication in E. coli.  DNA gyrase molecules travel along the DNA ahead of the replication fork, removing positive supercoils. DNA gyrase accomplishes this feat by cleaving both strands of the DNA duplex, passing a segment of DNA through the double-stranded break to the other side, and then sealing the cuts, a process that is driven by the energy released during ATP hydrolysis. Eukaryotic cells possess similar enzymes that carry out this required function. A model depicting the action of human topoisomerase I. Type I topoisomerases change the supercoiled state of a DNA molecule by creating a transient break in one strand of the duplex. The enzyme (yellow) cuts one of the strands of the DNA (step 1), which rotates around a phosphodiester bond in the intact strand. The cut strand is then resealed (step 2). (Note: The drawing depicts a type IB topoisomerase; type IA enzymes found in bacteria act by a different mechanism.)  Type II topoisomerases make a transient break in both strands of a DNA duplex. Topoisomerase II, a dimeric enzyme consisting of two identical halves. Step 1: the enzyme is in an “open” conformation ready to bind the G-DNA segment, so named because it will form the gate through which the T-DNA (or transported DNA) segment will pass. Step 2: the enzyme has undergone a conformational change as it binds the G-segment. Steps 3 and 4: the enzyme binds a molecule of ATP, the G-segment is cleaved, and the T-segment is passed through the open “gate.” The bracketed stage represents a hypothetical intermediate carrying out the step in which the T-segment is transported through the G- segment. At this stage, both cut ends of the G-segment are covalently bound to the enzyme. Step 5: the two ends of the G-segment are rejoined, and the T-segment is released. ATP hydrolysis and release of ADP and Pi are proposed to occur as the starting state is regenerated.  The Properties of DNA Polymerases A single-stranded DNA circle cannot serve as a template for DNA polymerase because the enzyme cannot initiate the formation of a DNA strand. Rather, it can only add nucleotides to the 3’ hydroxyl terminus of an existing strand. The strand that provides the necessary 3’ OH terminus is called a primer. All DNA polymerases—both prokaryotic and eukaryotic—have these same two basic requirements: – a template DNA strand to copy – a primer strand to which nucleotides can be added These requirements explain why certain DNA structures fail to promote DNA synthesis.  An intact, linear double helix provides the 3’ hydroxyl terminus but lacks a template. A circular single strand, on the other hand, provides a template but lacks a primer. The partially double-stranded molecule satisfies both requirements and thus promotes nucleotide incorporation. The finding that DNA polymerase cannot initiate the synthesis of a DNA strand raises a critical question: how is the synthesis of a new strand initiated in the cell?  DNA Polymerase Cannot Initiate new Strands 5’ Unable to covalently link the 2 individual nucleotides together 3’ 5’ 5’ 3’ Able to covalently link 3’ 3’ together 5’ 5’ 3’  DNA Polymerase only synthesizes DNA in a 5’-to-3’ (written 5’ → 3’) direction. Is there some other enzyme responsible for the construction of the 3’ → 5’ strand? A typical bacterial cell contains approximately 300 to 400 molecules of DNA polymerase I per cell and about 40 copies of DNA polymerase II and 10 copies of DNA polymerase III. The replicative polymerase is DNA polymerase III The discovery of other DNA polymerases did not answer the two basic questions posed above; none of the enzymes can initiate DNA chains, nor can any of them construct strands in a 3’→5’ direction.  SEMIDISCONTINUOUS REPLICATION The lack of polymerization activity in the 3’ → 5’ direction has a straightforward explanation: DNA strands cannot be synthesized in that direction. Rather, both newly synthesized strands are assembled in a 5’ → 3’ direction. During the polymerization reaction, the —OH group at the 3’ end of the primer carries out a nucleophilic attack on the 5’ α-phosphate of the incoming nucleoside triphosphate. The polymerase molecules responsible for construction of the two new strands of DNA both move in a 3’-to-5’ direction along the template, and both construct a chain that grows from its 5’-P terminus. The polymerization of a nucleotide onto the 3’ end of the primer strand A simplified model of the two-metal ion mechanism for the reaction in which nucleotides are incorporated into a growing DNA strand by a DNA polymerase. In this model, one of the magnesium ions draws the proton away from the 3’ hydroxyl group of the terminal nucleotide of the primer, facilitating the nucleophilic attack of the negatively charged 3’ oxygen atom on the α-phosphate of the incoming nucleoside triphosphate. The second magnesium ion promotes the release of the pyrophosphate. The two metal ions are bound to the enzyme by highly conserved aspartic acid residues of the active site.  Consequently, one of the newly synthesized strands grows toward the replication fork where the parental DNA strands are being separated, while the other strand grows away from the fork. Although this solves the problem concerning an enzyme that synthesizes a strand in only one direction, it creates an even more complicated dilemma. It is apparent that the strand that grows toward the fork can be constructed by the continuous addition of nucleotides to its 3’ end. But how is the other strand synthesized? The strand that grows away from the replication fork is synthesized discontinuously, that is, as fragments.  Once initiated, each fragment grows away from the replication fork toward the 5’ end of a previously synthesized fragment to which it is subsequently linked. Before the synthesis of a fragment can be initiated, a suitable stretch of template must be exposed by movement of the replication fork. Thus, the two newly synthesized strands of the daughter duplexes are synthesized by very different processes. The strand that is synthesized continuously is called the leading strand because its synthesis continues as the replication fork advances. The strand that is synthesized discontinuously is called the lagging strand because initiation of each fragment must wait for the parental strands to separate and expose additional template. Both strands are probably synthesized simultaneously, so that the terms leading and lagging may not be as appropriate as thought when they were first coined. Because one strand is synthesized continuously and the other discontinuously, replication is said to be semidiscontinuous.  A portion of the DNA is constructed in small segments (called Okazaki fragments, 1000 to 2000 nucleotides in length) that are rapidly linked to longer pieces that had been synthesized previously. The enzyme that joins the Okazaki fragments into a continuous strand is called DNA ligase. The discovery that the lagging strand is synthesized in pieces raised a new set of perplexing questions about the initiation of DNA synthesis. How does the synthesis of each of these fragments begin when none of the DNA polymerases are capable of strand initiation? Initiation is not accomplished by a DNA polymerase but, rather, by a distinct type of RNA polymerase, called primase, that constructs a short primer composed of RNA, not DNA. The leading strand, whose synthesis begins at the origin of replication, is also initiated by a primase molecule. The short RNAs synthesized by the primase at the 5’ end of the leading strand and the 5’ end of each Okazaki fragment serve as the required primer for the synthesis of DNA by a DNA polymerase. The RNA primers are subsequently removed, and the resulting gaps in the strand are filled with DNA and then sealed by DNA ligase. The formation of transient RNA primers during the process of DNA replication is a curious activity. It is thought that the likelihood of mistakes is greater during initiation than during elongation, and the use of a short removable segment of RNA avoids the inclusion of mismatched bases.  The Machinery Operating at the Replication Fork Replication involves more than incorporating nucleotides. Unwinding the duplex and separating the strands require the aid of two types of proteins that bind to the DNA, a helicase (or DNA unwinding enzyme) and single-stranded DNA binding (SSB) proteins. DNA helicases unwind a DNA duplex in a reaction that uses energy released by ATP hydrolysis to move along one of the DNA strands, breaking the hydrogen bonds that hold the two strands together and exposing the single-stranded DNA templates. E. coli has at least 12 different helicases for use in various aspects of DNA (and RNA) metabolism. One of these helicases—the product of the dnaB gene—serves as the major unwinding machine during replication. The DnaB helicase consists of six subunits arranged to form a ring-shaped protein that encircles a single DNA strand.  Initiation of replication begins in E. coli when multiple copies of the DnaA protein bind to the origin of replication (oriC) and separate (melt) the DNA strands at that site. DnaA protein binds to a 9 nucleotide sequence that is repeated 4 times within a 245-250 nucleotide sequence. Formation of this structure acts to locally denature an A-T rich region directly adjacent to it. DnaA protein binds to its binding sites in oriC , introducing a 40° bend at each site. The DnaB helicase is then loaded onto the single stranded DNA of the lagging strand of oriC, with the help of the protein DnaC. The DnaB helicase then translocates in a 5’ → 3’ direction along the lagging- strand template, unwinding the helix as it proceeds. DNA unwinding by the helicase is aided by the attachment of SSB proteins to the separated DNA strands. These proteins bind selectively to single-stranded DNA, keeping it in an extended state and preventing it from becoming rewound or damaged.  Initiation of Replication at oriC DNA replication is initiated by the binding of DnaA proteins to the DnaA box sequences  In bacteria, the primase and the helicase associate transiently to form what is called a “primosome.” Of the two members of the primosome, the helicase moves along the lagging-strand template processively (i.e., without being released from the template strand during the lifetime of the replication fork). As the helicase “motors” along the lagging-strand template, opening the strands of the duplex, the primase periodically binds to the helicase and The helicase moves along the DNA, catalyzing the ATP- synthesizes the short RNA primers that driven unwinding of the duplex. begin the formation of each Okazaki As the DNA is unwound, the strands are prevented from fragment. reforming the duplex by single-stranded DNA-binding proteins (SSBs). RNA primers are subsequently The primase associated with the helicase synthesizes the extended as DNA by a DNA RNA primers that begin each Okazaki fragment. polymerase, specifically DNA The RNA primers, which are about 10 nucleotides long, are polymerase III. subsequently removed.  A body of evidence suggests that the same DNA polymerase III molecule synthesizes successive fragments of the lagging strand. To accomplish this, the polymerase III molecule is recycled from the site where it has just completed one Okazaki fragment to the next site along the lagging-strand template closer to the site of DNA unwinding. Once at the new site, the polymerase attaches to the 3’ OH of the RNA primer that has just been laid down by a primase and begins to incorporate deoxyribonucleotides onto the end of the short RNA. How does a polymerase III molecule move from one site on the lagging-strand template to another site that is closer to the replication fork? The enzyme does this by “hitching a ride” with the DNA polymerase that is moving in that direction along the leading-strand template. Thus even though the two polymerases are moving in opposite directions with respect to the linear axis of the DNA molecule, they are, in fact, part of a single protein complex (replisome). The two tethered polymerases can replicate both strands by looping the DNA of the lagging-strand template back on itself, causing this template to have the same orientation as the leading-strand template. Both polymerases then can move together as part of a single replicative complex without violating the “5’ → 3’ rule” for synthesis of a DNA strand. Once the polymerase assembling the lagging strand reaches the 5’ end of the Okazaki fragment synthesized during the previous round, the lagging strand template is released and the polymerase begins work at the 3’ end of the next RNA primer toward the fork. Two DNA polymerases working together as part of a single complex. (a) The two DNA polymerase III molecules travel together, even though they are moving toward the opposite ends of their respective templates. This is accomplished by causing the lagging-strand template to form a loop. (b) The polymerase releases the lagging-strand template when it encounters the previously synthesized Okazaki fragment. (c) The polymerase that was involved in the assembly of the previous Okazaki fragment has now rebound the lagging-strand template farther along its length and is synthesizing DNA onto the end of RNA primer #3 that has just been constructed by the primase.  STRUCTURE AND FUNCTION OF DNA POLYMERASES There are three types of DNA polymerases namely – DNA polymerase I – DNA polymerase II – DNA polymerase III These enzymes have the same basic catalytic activity, which is to add deoxyribonucleotides onto the growing end of a single- stranded primer; however they differ in their various roles within the cell. The enzyme that acts in DNA strand formation is DNA polymerase III, which, because of its primary role in replication is often called as the replicase.  Schematic representation of DNA Polymerase III Structure resembles a human right hand Template DNA thread through the palm; Thumb and fingers wrapped around the DNA  DNA polymerase III holoenzyme is much larger than the other two polymerases, consisting of a single catalytic subunit and at least nine different associated subunits having various functions in the replication process. One of the non catalytic subunits, called the β subunit, appears responsible for keeping the polymerase associated with the DNA template. DNA polymerases have to possess two rather contrasting properties. – They have to remain associated with the template over long stretches to synthesize a continuous complementary strand. – At the same time they cannot be attached so tightly that they are unable to move from one nucleotide of the template to the next. These contrasting properties are provided by a doughnut-shaped β clamp that encircles the DNA and slides along it. As long as it is attached to a β “sliding clamp,” a DNA polymerase can move processively from one nucleotide to the next without diffusing away from the template.  The holoenzyme contains ten different subunits organized into several distinct components. Included as part of the holoenzyme are – (1) two core polymerases which replicate the DNA – (2) two or more β clamps, which allow the polymerase to remain associated with the DNA – (3) a clamp loading (γ) complex, which loads each sliding clamp onto the DNA. The clamp loader of an active replication fork contains two Ԏ subunits, which hold the core polymerases in the complex and also bind the helicase. Another term, the replisome, is often used to refer to the entire complex of proteins that are active at the replication fork, including the DNA polymerase III holoenzyme, the helicase, SSBs, and primase. DNA polymerase III holoenzyme  The polymerase on the leading-strand template remains tethered to a single β clamp during replication. In contrast, when the polymerase on the lagging-strand template completes the synthesis of an Okazaki fragment, it disengages from the β clamp and is cycled to a new β clamp that has been assembled at an RNA primer–DNA template junction located closer to the replication fork. But how does a highly elongated DNA molecule get inside of a ring-shaped clamp? The assembly of the β clamp around the DNA requires a multisubunit clamp loader that is also part of the DNA polymerase III holoenzyme. In the ATP-bound state, the clamp loader binds to a primer-template junction while holding the β clamp in an open conformation.  A model of a complex between a sliding clamp and a clamp loader from an archaean prokaryote based on electron microscopic image analysis. The clamp loader (shown with red and green subunits) is bound to the sliding clamp (blue), which is held in an open, spiral conformation resembling a lock-washer. The DNA has squeezed through the gap in the clamp. The primer strand of the DNA terminates within the clamp loader whereas the template strand extends through an opening at the top of the protein. The clamp loader has been described as a “screw- cap” that fits onto the DNA in such a way that the complex between a sliding subdomains of the protein form a spiral that can clamp and a clamp loader thread onto the helical DNA backbone. lock-washer  Once the DNA has squeezed through the opening in the clamp wall, the ATP bound to the clamp loader is hydrolyzed, causing the release of the clamp, which closes around the DNA. The β clamp is then ready to bind polymerase III. DNA polymerase I, which consists of only a single subunit, is involved primarily in DNA repair, a process by which damaged sections of DNA are corrected. DNA polymerase I also removes the RNA primers at the 5’ end of each Okazaki fragment during replication and replaces them with DNA.  All of the bacterial DNA polymerases possess exonuclease activity. Exonucleases can be divided into 5’ 3’ and 3’ 5’ exonucleases, depending on the direction in which the strand is degraded. DNA polymerase I has both 3’ 5’ and 5’ 3’ exonuclease activities, in addition to its polymerizing activity. These three activities are found in different domains of The 5’ 3’ exonuclease function removes nucleotides from the the single polypeptide. 5’ end of a single-strand nick. This activity also plays a key role Most nucleases are specific for either DNA or RNA, but in removing the RNA primers. the 5’ 3’ exonuclease of DNA polymerase I can degrade either type of nucleic acid. Initiation of Okazaki fragments by the primase leaves a stretch of RNA at the 5’ end of each fragment, which is removed by the 5’ 3’ exonuclease activity of DNA polymerase I. As the enzyme removes ribonucleotides of the primer, its polymerase activity simultaneously fills the resulting gap with deoxyribonucleotides. The last deoxyribonucleotide incorporated is subsequently joined covalently to the 5’ end of the previously The 3’ 5’ exonuclease function removes mispaired synthesized DNA fragment by DNA ligase. nucleotides from the 3 end of the growing DNA strand. This activity plays a key role in maintaining the accuracy of DNA synthesis.  Ensuring High Fidelity during DNA Replication The survival of an organism depends on the accurate duplication of the genome. A mistake made in the synthesis of a messenger RNA molecule by an RNA polymerase results in the synthesis of defective proteins, but an mRNA molecule is only one short-lived template among a large population of such molecules; therefore, little lasting damage results from the mistake. In contrast, a mistake made during DNA replication results in a permanent mutation and the possible elimination of that cell’s progeny. In E. coli, the chance that an incorrect nucleotide will be incorporated into DNA during replication and remain there is less than 10-9, or fewer than 1 out of 1 billion nucleotides. Because the genome of E. coli contains approximately 4 x 106 nucleotide pairs, this error rate corresponds to fewer than 1 nucleotide alteration for every 100 replication cycles. This represents the spontaneous mutation rate in this bacterium. Humans are thought to have a similar spontaneous mutation rate for replication of protein-coding sequences.  This job of “proofreading” is one of the most remarkable of all enzymatic activities and illustrates the sophistication to which biological molecular machinery has evolved. The 3’ 5’ exonuclease activity removes approximately 99 out of every 100 mismatched bases, raising the fidelity to about 10-7–10-8. In addition, bacteria possess a mechanism called mismatch repair that operates after replication and corrects nearly all of the mismatches that escape the proofreading step. Together these processes reduce the overall observed error rate to about 10-9. Thus the fidelity of DNA replication can be traced to three distinct activities: – accurate selection of nucleotides – immediate proofreading – post-replicative mismatch repair Another remarkable feature of bacterial replication is its rate. The replication of an entire bacterial chromosome in approximately 40 minutes at 37 °C requires that each replication fork move about 1000 nucleotides per second, which is equivalent to the length of an entire Okazaki fragment. Thus the entire process of Okazaki fragment synthesis, including formation of an RNA primer, DNA elongation and simultaneous proofreading by the DNA polymerase, excision of the RNA, its replacement with DNA, and strand ligation, occurs within a few seconds. Although it takes E. coli approximately 40 minutes to replicate its DNA, a new round of replication can begin before the previous round has been completed. Consequently, when these bacteria are growing at their maximal rate, they double their numbers in about 20 minutes.  Replication in Eukaryotic Cells Given the fact that eukaryotic cells have large genomes and complex chromosomal structure, our understanding of replication in eukaryotes has lagged behind that in bacteria. This imbalance has been addressed by the development of eukaryotic experimental systems that parallel those used for decades to study bacterial replication. These include: – The isolation of mutant yeast and animal cells unable to produce specific gene products required for various aspects of replication. – Analysis of the structure and mechanism of action of replication proteins from archaeal species. Replication in these prokaryotes begins at multiple origins and requires proteins that are homologous to those of eukaryotic cells but are less complex and easier to study. – The development of in vitro systems where replication can occur in cellular extracts or mixtures of purified proteins. The most valuable of these systems has utilized Xenopus, an aquatic frog that begins life as a huge egg stocked with all of the proteins required to carry it through a dozen or so very rapid rounds of cell division. Extracts can be prepared from these frog eggs that will replicate any added DNA, regardless of sequence. Frog egg extracts will also support the replication and mitotic division of mammalian nuclei, which has made this a particularly useful cell-free system. Antibodies can be used to deplete the extracts of particular proteins, and the replication ability of the extract can then be tested in the absence of the affected protein.  Initiation of Replication in Eukaryotic Cells Replication in E. coli begins at only one site along the single, circular chromosome. Cells of higher organisms may have a thousand times as much DNA as this bacterium, yet their polymerases incorporate nucleotides into DNA at much slower rates. To accommodate these differences, eukaryotic cells replicate their genome in small portions, termed replicons. Each replicon has its own origin from which replication forks Experimental demonstration that replication in proceed outward in both directions. eukaryotic chromosomes begins at many In a human cell, replication begins at about 10,000 to 100,000 sites along the DNA. Cells were incubated in [3H] thymidine for a brief period before different replication origins. preparation of DNA fibers for autoradiography. Approximately 10 to 15 percent of replicons are actively The lines of black silver grains indicate sites that engaged in replication at any given time during the S phase of had incorporated the radioactive DNA precursor the cell cycle. during the labeling period. It is evident that Replicons located close together in a given chromosome synthesis is occurring at separated sites along the same DNA molecule. Initiation begins in the undergo replication simultaneously. center of each site of thymidine incorporation, Moreover, those replicons active at a particular time during one forming two replication forks that travel away from round of DNA synthesis tend to be active at a comparable time each other until they meet a neighboring fork. in succeeding rounds. DNA replication initiates at many different sites simultaneously  In mammalian cells, the timing of replication of a chromosomal region is roughly correlated with the activity of the genes in the region and/or its state of compaction. The presence of acetylated histones, which is closely correlated with gene transcription, is a likely factor in determining the early replication of active gene loci. The most highly compacted, least acetylated regions of the chromosome are packaged into heterochromatin, and they are the last regions to be replicated. This difference in timing of replication is not related to DNA sequence because the inactive, heterochromatic X chromosome in the cells of female mammals is replicated late in S phase, whereas the active, euchromatic X chromosome is replicated at an earlier stage. Similarly, the β-globin locus replicates early during S phase in erythroid cells where the gene is expressed, but much later in nonerythroid cells where the gene remains silent. The mechanism by which replication is initiated in eukaryotes has been a focus of research over the past decade.  Yeast origins of replication contain a conserved sequence autonomous replicating sequences (ARSs). The core element of an ARS consists of a conserved sequence of 11 base pairs, which functions as a specific binding site for an essential multiprotein complex called the origin recognition complex (ORC). Unlike yeast, vertebrate DNA does not possess specific sequences (e.g., ARSs) at which replication is initiated. However, studies of replication of intact mammalian chromosomes in vivo suggest that replication does begin within defined regions of the DNA, rather than by random selection as occurs in the amphibian egg extract. It is thought that a DNA molecule contains many sites where DNA replication can be initiated, but only a subset of these potential sites are actually used at a given time in a given cell. Cells that reproduce via shorter cell cycles, such as those of early amphibian embryos, utilize a greater number of sites as origins of replication than cells with longer cell cycles. The actual selection of sites for initiation of replication is thought to be governed by local epigenetic factors, such as – the positions of nucleosomes, types of histone modifications, state of DNA methylation, degree of supercoiling, and the level of transcription.  Restricting Replication to Once Per Cell Cycle It is essential that each portion of the genome is replicated once, and only once, during each cell cycle. Consequently, some mechanism must exist to prevent the reinitiation of replication at a site that has already been duplicated. The initiation of replication at a particular origin requires passage of the origin through several distinct states. The basic mechanism of initiation of replication is conserved among eukaryotes. The origin of replication is bound by an ORC protein complex, which in yeast cells remains associated with the origin throughout the cell cycle. The ORC has been described as a “molecular landing pad” because of its role in binding the proteins required in subsequent steps. The next major step is the assembly of a protein– DNA complex, called the prereplication complex (pre-RC), that is “licensed” (competent) to initiate replication. Studies of the formation of the pre-RC have focused on a set of six related MCM (Minichromosome Maintenance Complex) proteins (Mcm2-Mcm7).  The MCM proteins are loaded onto the replication origin at a late stage of mitosis, or soon thereafter, with the aid of accessory factors that have previously bound to the ORC. The six Mcm2–Mcm7 proteins interact with one another to form a hexameric (6-membered) ring-shaped complex (the MCM complex) that possesses helicase activity. Evidence strongly suggests that the MCM complex is the eukaryotic replicative helicase; that is, the helicase responsible for unwinding DNA at the replication fork. At the pre-RC stage, each of the origins contains a double hexameric MCM complex, that is, two complete replicative helicases, which remain inactive at this stage of the cell cycle. Each of these replicases will travel in opposite directions away from the origin once replication begins. The assembly of a pre-RC marks that site on the genome as a potential origin of replication, but does not guarantee that it will actually be a site where replication will be initiated. During most cell cycles, many more pre-RCs are assembled than will be used and it is not clear what determines which of these potential sites of replication are subsequently selected. Regardless of the selection mechanism, just before the beginning of S phase of the cell cycle, the activation of key protein kinases leads to the phosphorylation of the MCM complex and other proteins and to the initiation of replication at selected sites in the genome.  One of these protein kinases is a cyclin-dependent kinase (Cdk). Cdk activity remains high from S phase through mitosis, which suppresses the formation of new prereplication complexes. Consequently, each origin can only be activated once per cell cycle. Cessation of Cdk activity at the end of mitosis permits the assembly pre-RCs for the next cell cycle. Once replication is initiated at the beginning of S phase, the MCM helicase moves with the replication fork, although the mechanism of action of this ring-shaped protein is debated. The fate of the MCM proteins after replication depends on the species studied. In mammalian cells, the MCM proteins are displaced from the DNA but apparently remain in the nucleus. Regardless, MCM proteins cannot reassociate with an origin of replication that has already “fired.”  Steps leading to the replication of a yeast replicon. Yeast origins of replication contain a conserved sequence (ARS) that binds the multisubunit origin recognition complex (ORC) (step 1). The presence of the bound ORC is required for initiation of replication. The ORC is bound to the origin throughout the yeast cell cycle. In step 2, a complex of six proteins (Mcm2–Mcm7) binds to the origin during or following mitosis, establishing a prereplication complex (pre-RC) that is competent to initiate replication, given the proper stimulus. Loading of MCM proteins at the origin requires additional proteins (Cdc6 and Cdt1, not shown). In step 3, DNA replication is initiated following activation of a cyclin-dependent kinase (Cdk) and a second protein kinase (DDK, DBF4-dependent kinase). Step 4 shows a stage where replication has proceeded a short distance in both directions from the origin. Each MCM complex forms a replicative DNA helicase that unwinds DNA at one of the oppositely directed replication forks. In step 5, the two strands of the original duplex have been replicated, an ORC is present at both origins, and the replication proteins, including the MCM helicases, have been displaced from the DNA. In yeast, the MCM proteins are exported from the nucleus, and reinitiation of replication cannot occur until the cell has passed through mitosis. [In vertebrate cells, several events appear to prevent reinitiation of replication, including (1) release of the ORC complex after its use in S phase, (2) continued Cdk activity from S phase into mitosis, (3) phosphorylation of Cdc6 and its subsequent export from the nucleus, and (4) degradation of Cdt1 or its inactivation by a bound inhibitor.]  The Eukaryotic Replication Fork Overall, the activities that occur at replication forks are quite similar, regardless of the type of genome being replicated—whether viral, bacterial, archaeal, or eukaryotic. All replication systems require helicases, single-stranded DNA-binding proteins, topoisomerases, primase, DNA polymerase, sliding clamp and clamp loader, and DNA ligase. When studying the initiation of eukaryotic replication in vitro, researchers often combine mammalian replication proteins with a viral helicase called the large T antigen, which is encoded by the SV40 genome. As in bacteria, the DNA of eukaryotic cells is synthesized in a semi-discontinuous manner, although the Okazaki fragments of the lagging strand are considerably smaller than in bacteria, averaging about 150 nucleotides in length. As in E. coli, the leading and lagging strands are thought to be synthesized in a coordinate manner by a single replicative complex, or replisome. To date, five “classic” DNA polymerases have been isolated from eukaryotic cells, and they are designated α, β, γ, δ, and ɛ. Of these enzymes, polymerase γ replicates mitochondrial DNA, and polymerase β functions in DNA repair. The other three polymerases have replicative functions. Polymerase α is tightly associated with the primase, and together they initiate the synthesis of each Okazaki fragment. The polymerase α-primase complex recognizes and binds to unwound as DNA that is coated by a single-stranded DNA-binding protein called RPA (Replication protein A). Primase initiates synthesis by assembly of a short RNA primer, which is then extended by the addition of about 20 deoxyribonucleotides by polymerase α.  Polymerase δ is thought to be the primary DNA-synthesizing enzyme during replication of the lagging strand, whereas polymerase ɛ is thought to be the primary DNA-synthesizing enzyme during replication of the leading strand. Like the major replicating enzyme of E. coli, both polymerase δ and ɛ require a “sliding clamp” that tethers the enzyme to the DNA, allowing it to move processively along a template. In eukaryotes, the sliding clamp is called PCNA (Proliferating cell nuclear antigen) which is very similar in structure and function to the β clamp of E. coli polymerase III. The clamp loader that loads PCNA onto the DNA is called RFC (replication factor C), and is analogous to the E. coli polymerase III clamp loader complex. After synthesizing an RNA-DNA primer, polymerase α is replaced at each template–primer junction by the PCNA–polymerase δ complex, which completes synthesis of the Okazaki fragment. When polymerase δ reaches the 5’ end of the previously synthesized Okazaki fragment, the polymerase continues along the lagging-strand template, displacing the primer. The displaced primer is cut from the newly synthesized DNA strand by a flap endonuclease (FEN-1, removes 5' overhanging flaps in DNA repair and processes the 5' ends of Okazaki fragments in lagging strand DNA) and the resulting nick in the DNA is sealed by a DNA ligase. FEN-1 and DNA ligase are thought to be recruited to the replication fork through an interaction with the PCNA sliding clamp. In fact, PCNA is thought to play a major role in orchestrating events that occur during DNA replication, repair, and recombination. Because of its ability to bind a diverse array of proteins, PCNA has been referred to as a “molecular toolbelt.”  a) A schematic view of the major components at the eukaryotic replication fork. The viral T antigen is drawn as the replicative helicase in this figure because it is prominently employed in in vitro studies of DNA replication. DNA polymerases δ and ɛ are thought to be the primary DNA synthesizing enzymes of the lagging and leading strands, respectively. PCNA acts as a sliding clamp for both polymerases δ and ɛ. The sliding clamp is loaded onto the DNA by a protein called RFC (replication factor C), which is similar in structure and function to the β-clamp loader of E. coli. RPA is a trimeric single-stranded DNA binding protein comparable in function to that of SSB utilized in E. coli replication. The RNA-DNA primers of the lagging strand that are synthesized by the polymerase α-primase complex are displaced by the continued b) movement of polymerase δ, generating a flap of RNA-DNA that is removed by the FEN-1 endonuclease. The gap is sealed by a DNA ligase. As in E. coli, a topoisomerase is required to remove the positive supercoils that develop ahead of the replication fork. A proposed version of events at the replication fork illustrating how the replicative polymerases on the leading- and lagging-strand templates might act together as part of a replisome. To date there is no firm evidence that the leading and lagging strands are replicated by a single replicative complex as in E. coli.  The various proteins in the replication “tool kit” of eukaryotic cells E. coli Eukaryotic Function protein DnaA ORC proteins Recognition of origin of replication Gyrase Topoisomerase I/II Relieves positive supercoils ahead of replication fork DnaB Mcm DNA helicase that unwinds parental duplex DnaC Cdc6, Cdt1 Loads helicase onto DNA SSB RPA (Replication protein A) Maintains DNA in single-stranded state γ-complex RFC (Replication Factor C) Subunits of the DNA polymerase holoenzyme that load the clamp onto the DNA pol III core pol δ and ɛ Primary replicating enzymes; synthesize entire leading strand and Okazaki fragments; have proofreading capability β clamp PCNA (Proliferative Cell Ring-shaped subunit of DNA polymerase holoenzyme that clamps replicating polymerase Nuclear Antigen) to DNA; works with pol III in E. coli and pol δ or ɛ in eukaryotes Primase Primase Synthesizes RNA primers _______ pol α Synthesizes short DNA oligonucleotides as part of RNA–DNA primer DNA ligase DNA ligase Seals Okazaki fragments into continuous strand pol I FEN-1 Removes RNA primers; pol I of E. coli also fills gap with DNA  Like bacterial polymerases, all of the eukaryotic polymerases elongate DNA strands in the 5’ → 3’ direction by the addition of nucleotides to a 3’ hydroxyl group, and none of them is able to initiate the synthesis of a DNA chain without a primer. Polymerases γ, δ, and ɛ possess a 3’ → 5’ exonuclease, whose proofreading activity ensures that replication occurs with very high accuracy. Several other DNA polymerases (including η, κ and ι) have a specialized function that allows cells to replicate damaged DNA. Replication forks that are active at a given time are not distributed randomly throughout the cell nucleus, but Demonstration that replication activities do not instead are localized within 50 to 250 sites, called occur randomly throughout the nucleus but are replication foci. confined to distinct sites. Prior to the onset of DNA It is estimated that each of the bright red regions synthesis at the start of S phase, various factors contains approximately 10 to 100 replication forks required for the initiation of replication are assembled at incorporating nucleotides into DNA strands discrete sites within the nucleus, forming prereplication simultaneously. centers. These sites appear as discrete red objects in The clustering of replication forks may provide a the micrograph, which has been stained with a fluorescent antibody against replication factor A (RPA), mechanism for coordinating the replication of which is a single-stranded DNA-binding protein required adjacent replicons on individual chromosomes. for replication. Other replication factors, such as PCNA and the polymerase–primase complex, are also localized to these foci.  Chromatin Structure and Replication The chromosomes of eukaryotic cells consist of DNA tightly complexed to regular arrays of histone proteins that are present in the form of nucleosomes. Movement of the replication machinery along the DNA is thought to displace nucleosomes that reside in its path. Yet, examination of a replicating DNA molecule with the electron 1.8 turns (146 base pairs) of negatively microscope reveals nucleosomes on both daughter duplexes very near the supercoiled DNA wrapped around eight core histone molecules replication fork, indicating that the reassembly of nucleosomes is a very rapid event. Collectively, the nucleosomes that form during the replication process are comprised of a roughly equivalent mixture of histone molecules that are inherited from parental chromosomes and histone molecules that have been newly synthesized. The core histone octamer of a nucleosome consists of an (H3H4)2 tetramer together with a pair of H2A/H2B dimers. The way in which parental nucleosomes are distributed during replication has been an area of recent debate. According to results from classic experiments, the (H3H4)2 tetramers present prior to replication remain intact and are distributed randomly between the two daughter duplexes. As a result, old and new (H3H4)2 tetramers are thought to be intermixed on each daughter DNA molecule.  Schematic model showing the distribution of core histones after DNA replication. Each nucleosome core particle is shown schematically to be composed of a central (H3H4)2 tetramer flanked by two H2A/H2B dimers. Histones that were present in parental nucleosomes prior to replication are indicated in blue; newly synthesized histones are indicated in red. According to this model, which is supported by a body of experimental evidence, the parental (H3H4)2 tetramers remain intact and are distributed randomly to both daughter duplexes. In contrast, the pairs of H2A/H2B dimers present in parental nucleosomes separate and recombine randomly with the (H3H4)2 tetramers on the daughter duplexes. Alternate models have also been presented in which the parental (H3H4)2 tetramer is split in half by a histone chaperone, and the two resulting H3/H4 dimers are distributed to different DNA strands.  According to this model, the two H2A/H2B dimers of each parental nucleosome fail to remain together as the replication fork moves through the chromatin. Instead, the H2A/H2B dimers of a nucleosome separate from one another and bind randomly to the new and old (H3H4)2 tetramers already present on the daughter duplexes. Results of several recent experiments have raised the possibility of another model, one in which the (H3H4)2 tetramer from parental nucleosomes is split into two H3/H4 dimers, each of which may combine with a newly synthesized H3/H4 dimer to form a “mixed” (H3H4)2 tetramer, which then assembles with H2A/H2B dimers. Regardless of the pattern by which it occurs, the stepwise assembly of nucleosomes and their orderly spacing along the DNA is facilitated by a network of accessory proteins. Included among these proteins are a number of histone chaperones that are able to accept either newly synthesized or parental histones and transfer them to the daughter strands. The best studied of these histone chaperones, CAF-1 (Chromatin Assembly Factor), is recruited to the advancing replication fork through an interaction with the sliding clamp PCNA.  For the most part, daughter cells carry out the same pattern of transcription as their parental cells; this is one of the cornerstones underlying the homeostatic functioning of tissues and organs. The transcriptional state of a cell depends to a large degree upon the epigenetic state of the cell’s chromatin, which is inherited from one cell generation to the next. Epigenetic information is not encoded within a chromosome’s DNA sequence, but rather is encoded in the pattern of methylated cytosine residues in a cell’s DNA and in the pattern of posttranslational modifications of the core histones associated with the DNA. Consequently, it is essential that these patterns be faithfully transmitted from parental chromatin to the chromatin of daughter cells, yet very little is known about how such transmission occurs. It is likely, for example, that modifications present on old histones will guide the modification of new histones within neighboring nucleosomes on the same DNA strand. DNA methylation patterns are apparently transmitted through the activities of the DNA methyltransferase DNMT1. Somehow, this enzyme appears capable of adding methyl groups to the cytosine residues of newly synthesized DNA strands using the pattern of such modifications on the parental DNA strands as a guide or template.  DNA Repair Both prokaryotic and eukaryotic cells possess a variety of proteins that patrol vast stretches of DNA, searching for subtle chemical modifications or distortions of the DNA duplex. In some cases, damage can be repaired directly. Humans, for example, possess enzymes that can directly repair damage from cancer-producing alkylating agents. Most repair systems, however, require that a damaged section of the DNA be excised, that is, selectively removed. One of the great virtues of the DNA duplex is that each strand contains the information required for constructing its partner. Consequently, if one or more nucleotides is removed from one strand, the complementary strand can serve as a template for reconstruction of the duplex. The repair of DNA damage in eukaryotic cells is complicated by the relative inaccessibility of DNA within the folded chromatin fibers of the nucleus. As in the case of transcription, DNA repair involves the participation of chromatin-reshaping machines, such as the histone modifying enzymes and nucleosome remodeling complexes. Nucleotide Excision Repair Nucleotide excision repair (NER) operates by a cut-and patch mechanism that removes a variety of bulky lesions, including pyrimidine dimers and nucleotides to which various chemical groups have become attached. Two distinct NER pathways can be distinguished: 1. A transcription-coupled pathway in which the template strands of genes that are being actively transcribed are preferentially repaired. Repair of a template strand is thought to occur as the DNA is being transcribed, and the presence of the lesion may be signaled by a stalled RNA polymerase. This preferential repair pathway ensures that those genes of greatest importance to the cell, which are the genes the cell is actively transcribing, receive the highest priority on the “repair list.” 2. A slower, less efficient global genomic pathway that corrects DNA strands in the remainder of the genome. Although recognition of the lesion is probably accomplished by different proteins in the two NER pathways (step 1, Figure 13.25), the steps that occur during repair of the lesion are thought to be very similar, as indicated in steps 2–6 of Figure 13.25. One of the key components of the NER repair machinery is TFIIH, a huge protein that also participates in the initiation of transcription. The discovery of the involvement of TFIIH established a crucial link between transcription and DNA repair, two processes that were previously assumed to be independent of one another (discussed in the Experimental Pathways, which can be accessed on the Web at www.wiley.com/college/karp). Included among the various subunits of TFIIH are two subunits (XPB and XPD) that possess helicase activity; these enzymes separate the two strands of the duplex (step 2, Figure 13.25) in preparation for removal of the lesion. The damaged strand is then cut on both sides of the lesion by a pair of endonucleases (step 3), and the segment of DNA between the incisions is released (step 4). Once excised, the gap is filled by a DNA polymerase (step 5), and the strand is sealed by DNA ligase (step 6).  Base Excision Repair A separate excision repair system operates to remove altered nucleotides generated by reactive chemicals present in the diet or produced by metabolism. The steps in this repair pathway in eukaryotes, which is called base excision repair (BER), are shown in Figure 13.26. BER is initiated by a DNA glycosylase that recognizes the alteration (step 1, Figure 13.26) and re- moves the altered base by cleavage of the glycosidic bond holding the base to the deoxyribose sugar (step 2). A number of different DNA glycosylases have been identified, each more-or-less specific for a particular type of altered base, including uracil (formed by the hydrolytic removal of the amino group of cytosine), 8-oxoguanine (caused by damage from oxygen free radicals, page 35), and 3-methyladenine (produced by transfer of a methyl group from a methyl donor, page 437). Structural studies of the DNA glycosylase that removes the highly mutagenic 8-oxoguanine (oxoG) indicate that this enzyme diffuses rapidly along the DNA “inspecting” each of the G-C base pairs within the DNA duplex (Figure 13.27, step 1). In step 2, the enzyme has come across an oxoG-C base pair. When this occurs, the enzyme inserts a specific amino acid side chain into the DNA helix, causing the nucleotide to rotate (“flip”) 180 degrees out of the DNA helix and into the body of the enzyme (step 2). If the nucleotide does, in fact, contain an oxoG, the base fits into the active site of the enzyme (step 3) and is cleaved from its associated sugar. In contrast, if the extruded nucleotide contains a normal guanine, which only differs in structure by two atoms from oxoG, it is unable to fit into the enzyme’s active site (step 4) and it is returned to its appropriate position within the stack of bases. Once an altered purine or pyrimidine is removed by a glycosylase, the “beheaded” deoxyribose phosphate remaining in the site is excised by the combined action of a specialized (AP) endonuclease and a DNA polymerase. AP endonuclease cleaves the DNA backbone (Figure 13.26, step 3) and a phosphodiesterase activity of polymerase _ removes the sugar– phosphate remnant that had been attached to the excised base (step 4). Polymerase _ then fills the gap by inserting a nucleotide complementary to the undamaged strand (step 5), and the strand is sealed by DNA ligase III (step 6). The fact that cytosine can be converted to uracil may explain why natural selection favored the use of thymine, rather than uracil, as a base in DNA, even though uracil was presumably present in RNA when it served as genetic material during the early evolution of life (page 454). If uracil had been retained as a DNA base, it would have caused difficulty for repair systems to distinguish between a uracil that “belonged” at a particular site and one that resulted from an alteration of cytosine.  Detecting damaged bases during BER. In step 1, a DNA glycosylase (named hOGG1) is inspecting a nucleotide that is paired to a cytosine. In step 2, the nucleotide is flipped out of the DNA duplex. In this case, the base is an oxidized version of guanine, 8- oxoguanine, and it is able to fit into the active site of the enzyme (step 3) where it is cleaved from its attached sugar. The subsequent steps in BER were shown in Figure 13.26. In step 4, the extruded base is a normal guanine, which is unable to fit into the active site of the glycosylase and is returned to the base stack. Failure to remove oxoG would have resulted in a G-to-T mutation.  Mismatch Repair It was noted earlier that cells can remove mismatched bases that are incorporated by the DNA polymerase and escape the enzyme’s proofreading exonuclease. This process is called mismatch repair (MMR). A mismatched base pair causes a distortion in the geometry of the double helix that can be recognized by a repair enzyme. But how does the enzyme “recognize” which member of the mismatched pair is the incorrect nucleotide? If it were to remove one of the nucleotides at random, it would make the wrong choice 50 percent of the time, creating a permanent mutation at that site. Thus, for a mismatch to be repaired after the DNA polymerase has moved past a site, it is important that the repair system distinguish the newly synthesized strand, which contains the incorrect nucleotide, from the parental strand, which contains the correct nucleotide. In E. coli, the two strands are distinguished by the presence of methylated adenosine residues on the parental strand. DNA methylation does not appear to be utilized by the MMR system in eukaryotes, and the mechanism of identification of the newly synthesized strand remains unclear.  Double-Strand Breakage Repair X-rays, gamma rays, and particles released by radioactive atoms are all described as ionizing radiation because they generate ions as they pass through matter. Millions of gamma rays pass through our bodies every minute.When these forms of radiation collide with a fragile DNA molecule, they often break both strands of the double helix. Double-strand breaks (DSBs) can also be caused by certain chemicals, including several (e.g., bleomycin) used in cancer chemotherapy, and free radicals produced by normal cellular metabolism (page 35). DSBs are also introduced during replication of damaged DNA. A single double- strand break can cause serious chromosome abnormalities, which can have grave consequences for the cell. DSBs can be repaired by several alternate pathways. The predominant pathway in mammalian cells is called nonhomologous end joining (NHEJ), in which a complex of proteins binds to the broken ends of the DNA duplex and catalyzes a series of reactions that rejoin the broken strands. The major steps that occur during NHEJ are shown in Figure 13.28a and described in the accompanying legend. Figure 13.28b shows the nucleus of a human fibroblast that had been treated with a laser to induce a localized cluster of double-strand breaks and then stained for the presence of the protein Ku at various times after laser treatment. This NHEJ repair protein is seen to localize at the site of the DSBs immediately following their appearance.  Another DSB repair pathway known as homologous recombination (HR) requires a homologous chromosome to serve as a template for repair of the broken strand. The steps that occur during homologous recombination, which include excision of the damaged DNA, are similar to those of genetic recombination depicted in Figure 14.47. A comparison between these two DSB repair pathways shows major differences. Homologous recombination is a more accurate pathway; that is, there are fewer errors in the base sequence of the repaired DNA than NHEJ. However, because it requires that a homologous chromosome be present in the nucleus, HR can only be employed during the cell cycle after DNA replication takes place (i.e., during late S or G2 phase). Defects in both repair pathways have been linked to increased cancer susceptibility.