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Cell_Biology_Alberts. Figure 5–14 The structure of a DNA helicase. (A) Diagram of the protein 5˜ as a hexameric ring drawn to scale with a replication fork. (B) Detailed structure of the bacteriophage T7 replicative helicase, as determined by x-ray diffraction. Six identical subunits bind and hydrolyze ATP in an ordered fashion to propel this molecule, like a rotary engine, along a DNA single strand that passes through the central hole. Red indicates bound ATP molecules in the structure (Movie 5.2). (PDB code: 1E0J.) finished synthesizing one Okazaki fragment on the lagging strand to be recycled quickly, so as to begin the synthesis of the next Okazaki fragment on the same strand. This rapid dissociation, however, would make it difficult for the polymerase to synthesize the long DNA strands produced at a replication fork were it not for an accessory protein (called PCNA in eukaryotes) that functions as a regulated sliding clamp. This clamp keeps the polymerase firmly on the DNA when it is moving, but


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Cell_Biology_Alberts. replication fork were it not for an accessory protein (called PCNA in eukaryotes) that functions as a regulated sliding clamp. This clamp keeps the polymerase firmly on the DNA when it is moving, but releases it as soon as the polymerase runs into a double-strand region of DNA.


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Cell_Biology_Alberts. How can a sliding clamp prevent the polymerase from dissociating without at the same time impeding the polymerase’s rapid movement along the DNA molecule? The three-dimensional structure of the clamp protein, determined by x-ray diffraction, revealed it to be a large ring around the DNA double helix. One face of the ring binds to the back of the DNA polymerase, and the whole ring slides freely along the DNA as the polymerase moves. The assembly of the clamp around the DNA requires ATP hydrolysis by a special protein complex, the clamp loader, which hydrolyzes ATP as it loads the clamp on to a primer–template junction (Figure 5–17).


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Cell_Biology_Alberts. On the leading-strand template, the moving DNA polymerase is tightly bound to the clamp, and the two remain associated for a very long time. The DNA polymerase on the lagging-strand template also makes use of the clamp, but each time the polymerase reaches the 5ʹ end of the preceding Okazaki fragment, the polymerase releases itself from the clamp and dissociates from the template. This polymerase molecule then associates with a new clamp that is assembled on the RNA primer of the next Okazaki fragment. single-stranded region of DNA template with short regions of base-paired “hairpins”


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Cell_Biology_Alberts. single-stranded region of DNA template with short regions of base-paired “hairpins” Figure 5–15 The effect of single-strand DNA-binding proteins (SSb proteins) on the structure of single-strand DNA. Because each protein molecule prefers to bind next to a previously bound molecule, long rows of this protein form on a DNA single strand. This cooperative binding straightens out the DNA template and facilitates the DNA polymerization process. The “hairpin helices” shown in the bare, single-strand DNA result from a chance matching of short regions of complementary nucleotide sequence; they are similar to the short helices that typically form in RNA molecules (see Figure 1–6).


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Cell_Biology_Alberts. Figure 5–16 Human single-strand binding protein bound to DNA. (A) Front view of the two DNA-binding domains of the protein (called RPA) which cover a total of eight nucleotides. Note that the DNA bases remain exposed in this protein–DNA complex. (B) Diagram showing the three-dimensional structure, with the DNA strand (orange) viewed end-on. (PDB code: 1JMC.)


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Cell_Biology_Alberts. Figure 5–17 The regulated sliding clamp that holds DNA polymerase on the DNA. (A) The structure of the clamp protein from E. coli, as determined by x-ray crystallography, with a DNA helix added to indicate how the protein fits around DNA (Movie 5.3). (B) Schematic illustration showing how the clamp (with red and yellow subunits) is loaded onto DNA to serve as a tether for a moving DNA polymerase molecule. The structure of the clamp loader (dark green) resembles a screw nut, (A) (B) sliding clamp clamp loader 5˜ATPADPPi ATPATP+ DNA + DNA polymerase 3˜5˜5˜3˜3˜RECYCLING OF RELEASED CLAMP LOADER ATP BINDING TO CLAMP LOADER OPENS SLIDING CLAMP DNA ENGAGED IN CLAMP ATP HYDROLYSIS LOCKS SLIDING CLAMP AROUND DNA AND RELEASES CLAMP LOADER DNA POLYMERASE BINDS TO SLIDING CLAMP + with its threads matching the grooves of double-stranded DNA. The loader binds to a free clamp molecule, forcing a gap in its ring of subunits so that this ring is able to slip around DNA. The clamp loader, thanks to


Cell_Biology_Alberts

Cell_Biology_Alberts. matching the grooves of double-stranded DNA. The loader binds to a free clamp molecule, forcing a gap in its ring of subunits so that this ring is able to slip around DNA. The clamp loader, thanks to its screw-nut structure, recognises the region of DNA that is double-stranded and latches onto it, tightening around the complex of a template strand with a freshly synthesized elongating (primer) strand. It carries the clamp along this double-stranded region until it encounters the 3ʹ end of the primer, at which point the loader hydrolyzes ATP and releases the clamp, allowing it to close around the DNA and bind to DNA polymerase. In the simplified reaction shown here, the clamp loader dissociates into solution once the clamp has been assembled. At a true replication fork, the clamp loader remains close to the polymerase so that, on the lagging strand, it is ready to assemble a new clamp at the start of each new Okazaki fragment (see Figure 5–18). (A, from X.P. Kong et al., Cell


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Cell_Biology_Alberts. remains close to the polymerase so that, on the lagging strand, it is ready to assemble a new clamp at the start of each new Okazaki fragment (see Figure 5–18). (A, from X.P. Kong et al., Cell 69:425–437, 1992. With permission from Elsevier; B, adapted from B.A. Kelch et al., Science 334:1675–1680, 2011. With permission from AAAS. PDB code: 3BEP.)


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Cell_Biology_Alberts. The Proteins at a Replication Fork Cooperate to Form a Replication Machine Although we have discussed DNA replication as though it were performed by a mix of proteins all acting independently, in reality most of the proteins are held together in a large and orderly multienzyme complex that rapidly synthesizes DNA. This complex can be likened to a tiny sewing machine composed of protein parts and powered by nucleoside triphosphate hydrolysis. Like a sewing machine, the replication complex probably remains stationary with respect to its immediate surroundings; the DNA can be thought of as a long strip of cloth being rapidly threaded through it. Although the replication complex has been most intensively studied in E. coli and several of its viruses, a very similar complex also operates in eukaryotes, as we see below.


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Cell_Biology_Alberts. Figure 5–18 summarizes the functions of the subunits of the replication machine. At the front of the replication fork, DNA helicase opens the DNA helix. Two DNA polymerase molecules work at the fork, one on the leading strand and one on the lagging strand. Whereas the DNA polymerase molecule on the leading strand can operate in a continuous fashion, the DNA polymerase molecule on the lagging strand must restart at short intervals, using a short RNA primer made by a DNA primase molecule. The close association of all these protein components increases the efficiency of replication and is made possible by a folding back of the lagging strand as shown in Figure 5–18A. This arrangement also facilitates the loading of the polymerase clamp each time that an Okazaki fragment is synthesized: the clamp loader and the lagging-strand DNA polymerase molecule are kept in place as a part of the protein machine even when they detach from their DNA template. The replication proteins are thus linked


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Cell_Biology_Alberts. clamp loader and the lagging-strand DNA polymerase molecule are kept in place as a part of the protein machine even when they detach from their DNA template. The replication proteins are thus linked together into a single large


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Cell_Biology_Alberts. Figure 5–18 A bacterial replication fork. (A) This schematic diagram shows a current view of the arrangement of replication proteins at a replication fork when DNA is being synthesized. The lagging-strand DNA is folded to bring the lagging-strand DNA polymerase molecule into a complex with the leading-strand DNA polymerase molecule. This folding also brings the 3ʹ end of each completed Okazaki fragment close to the start site for the next Okazaki fragment. Because the lagging-strand DNA polymerase molecule remains bound to the rest of the replication proteins, it can be reused to synthesize successive Okazaki fragments. In this diagram, it is about to let go of its completed DNA fragment and move to the RNA primer that is just being synthesized. Additional proteins (not shown) help to hold the different protein components of the fork together, enabling them to function as a well-coordinated protein machine (Movie 5.4 and Movie 5.5). (B) An electron micrograph showing the replication


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Cell_Biology_Alberts. the different protein components of the fork together, enabling them to function as a well-coordinated protein machine (Movie 5.4 and Movie 5.5). (B) An electron micrograph showing the replication machine from the bacteriophage T4 as it moves along a template synthesizing DNA behind it. (C) An interpretation of the micrograph is given in the sketch: note especially the DNA loop on the lagging strand. Apparently, the replication proteins became partly detached from the very front of the replication fork during the preparation of this sample for electron microscopy. (B, courtesy of Jack Griffith; see P.D. Chastain et al., J. Biol. Chem. 278:21276–21285, 2003.) unit (total molecular mass >106 daltons), enabling DNA to be synthesized on both sides of the replication fork in a coordinated and efficient manner.


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Cell_Biology_Alberts. On the lagging strand, the DNA replication machine leaves behind a series of unsealed Okazaki fragments, which still contain the RNA that primed their synthesis at their 5ʹ ends. As discussed earlier, this RNA is removed and the resulting gap is filled in by DNA repair enzymes that operate behind the replication fork (see Figure 5–11). A Strand-Directed Mismatch Repair System Removes Replication Errors That Escape from the Replication Machine


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Cell_Biology_Alberts. A Strand-Directed Mismatch Repair System Removes Replication Errors That Escape from the Replication Machine As stated previously, bacteria such as E. coli are capable of dividing once every 30 minutes, making it relatively easy to screen large populations to find a rare mutant cell that is altered in a specific process. One interesting class of mutants consists of those with alterations in so-called mutator genes, which greatly increase the rate of spontaneous mutation. Not surprisingly, one such mutant makes a defective form of the 3ʹ-to-5ʹ proofreading exonuclease that is a part of the DNA polymerase enzyme (see Figures 5–8 and 5–9). The mutant DNA polymerase no longer proofreads effectively, and many replication errors that would otherwise have been removed accumulate in the DNA.


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Cell_Biology_Alberts. The study of other E. coli mutants exhibiting abnormally high mutation rates has uncovered a proofreading system that removes replication errors made by the polymerase that have been missed by the proofreading exonuclease. This strand-directed mismatch repair system detects the potential for distortion in the DNA helix from the misfit between noncomplementary base pairs. If the proofreading system simply recognized a mismatch in newly replicated DNA and randomly corrected one of the two mismatched nucleotides, it would mistakenly “correct” the original template strand to match the error exactly half the time, thereby failing to lower the overall error rate. To be effective, such a proofreading system must be able to distinguish and remove the mismatched nucleotide only on the newly synthesized strand, where the replication error occurred.


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Cell_Biology_Alberts. The strand-distinction mechanism used by the mismatch proofreading system in E. coli depends on the methylation of selected A residues in the DNA. Methyl groups are added to all A residues in the sequence GATC, but not until some time after the A has been incorporated into a newly synthesized DNA chain. As a result, the only GATC sequences that have not yet been methylated are in the new strands just behind a replication fork. The recognition of these unmethylated GATCs allows the new DNA strands to be transiently distinguished from old ones, as required if their mismatches are to be selectively removed. The three-step process involves recognition of a newly synthesized strand, excision of the portion containing the mismatch, and resynthesis of the excised segment using the old strand as a template. This strand-directed mismatch repair system reduces the number of errors made during DNA replication by an additional factor of 100 to 1000 (see Table 5–1, p. 244).


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Cell_Biology_Alberts. A similar mismatch proofreading system functions in eukaryotic cells but uses a different strategy to distinguish the new strand from the old (Figure 5–19). Newly synthesized lagging-strand DNA transiently contains nicks (before they are sealed by DNA ligase) and such nicks (also called single-strand breaks) provide the signal that directs the mismatch proofreading system to the appropriate strand. This strategy also requires that the newly synthesized DNA on the leading strand be transiently nicked; how this occurs is uncertain.


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Cell_Biology_Alberts. The importance of mismatch proofreading in humans is seen in individuals who inherit one defective copy of a mismatch repair gene (along with a functional gene on the other copy of the chromosome). These people have a marked predisposition for certain types of cancers. For example, in a type of colon cancer called hereditary nonpolyposis colon cancer (HNPCC), spontaneous mutation of the one functional gene produces a clone of somatic cells that, because they are deficient in mismatch proofreading, accumulate mutations unusually rapidly. Most cancers arise in cells that have accumulated multiple mutations (see pp. 1096–1097), and cells deficient in mismatch proofreading therefore have a greatly enhanced chance of becoming cancerous. Fortunately, most of us inherit two good copies of each gene that encodes a mismatch proofreading protein; this protects us, because it is highly unlikely for both copies to become mutated in the same cell.


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Cell_Biology_Alberts. As a replication fork moves along double-strand DNA, it creates what has been called the “winding problem.” The two parental strands, which are wound around each other, must be unwound and separated for replication to occur. For every 10 nucleotide pairs replicated at the fork, one complete turn of the parental double helix must be unwound. In principle, this unwinding can be achieved by rapidly rotating the entire chromosome ahead of a moving fork; however, this is energetically highly unfavorable (particularly for long chromosomes) and, instead, the DNA in front of a replication fork becomes overwound (Figure 5–20). The over-winding, in turn, is continually relieved by proteins known as DNA topoisomerases. A DNA topoisomerase can be viewed as a reversible nuclease that adds itself covalently to a DNA backbone phosphate, thereby breaking a phosphodiester bond in a DNA strand. This reaction is reversible, and the phosphodiester bond re-forms as the protein leaves.


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Cell_Biology_Alberts. One type of topoisomerase, called topoisomerase I, produces a transient sin-gle-strand break; this break in the phosphodiester backbone allows the two sections of DNA helix on either side of the nick to rotate freely relative to each other, using the phosphodiester bond in the strand opposite the nick as a swivel point (Figure 5–21). Any tension in the DNA helix will drive this rotation in the direction that relieves the tension. As a result, DNA replication can occur with the rotation of only a short length of helix—the part just ahead of the fork. Because the covalent linkage that joins the DNA topoisomerase protein to a DNA phosphate retains rotate, torsional stress will build up


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Cell_Biology_Alberts. Figure 5–19 Strand-directed mismatch repair. (A) The two proteins shown are present in both bacteria and eukaryotic cells: MutS binds specifically to a mismatched base pair, while MutL scans the nearby DNA for a nick. Once MutL finds a nick, it triggers the degradation of the nicked strand all the way back through the mismatch. Because nicks are largely confined to newly replicated strands in eukaryotes, replication errors are selectively removed. In bacteria, an additional protein in the complex (MutH) nicks unmethylated (and therefore newly replicated) GATC sequences, thereby beginning the process illustrated here. In eukaryotes, MutL contains a DNA nicking activity that aids in the removal of the damaged strand.


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Cell_Biology_Alberts. (B) The structure of the MutS protein bound to a DNA mismatch. This protein is a dimer, which grips the DNA double helix as shown, kinking the DNA at the mismatched base pair. It seems that the MutS protein scans the DNA for mismatches by testing for sites that can be readily kinked, which are those with an abnormal base pair. (PDB code: 1EWQ.) Figure 5–20 The “winding problem” that arises during DNA replication.


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Cell_Biology_Alberts. Figure 5–20 The “winding problem” that arises during DNA replication. (A) For a bacterial replication fork moving at 500 nucleotides per second, the parental DNA helix ahead of the fork must rotate at 50 revolutions per second. (B) If the ends of the DNA double helix remain fixed (or difficult to rotate), tension builds up in front of the replication fork as it becomes overwound. Some of this tension can be taken up by supercoiling, whereby the DNA double helix twists around itself (see Figure 6–19). However, if the tension continues to build up, the replication fork will eventually stop because further unwinding requires more energy than the helicase can provide. Note that in (A), the dotted line represents about 20 turns of DNA. one end of the DNA double helix cannot rotate relative to the other end topoisomerase with tyrosine at the active site


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Cell_Biology_Alberts. one end of the DNA double helix cannot rotate relative to the other end topoisomerase with tyrosine at the active site CH2 OH CH2 OH DNA topoisomerase covalently attaches to a DNA phosphate, thereby breaking a phosphodiester linkage in one DNA strand the two ends of the DNA double helix can now rotate relative to each other, relieving accumulated strain Figure 5–21 The reversible DNA nicking reaction catalyzed by a eukaryotic DNA topoisomerase I enzyme. As indicated, these enzymes transiently form a single covalent bond with DNA; this allows free rotation of the DNA around the covalent backbone bonds linked to the blue phosphate.


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Cell_Biology_Alberts. CH2HO the original phosphodiester bond energy is stored in the phosphotyrosine linkage, making the reaction reversible CH2 OH spontaneous re-formation of the phosphodiester bond regenerates both the DNA helix and the DNA topoisomerase the energy of the cleaved phosphodiester bond, resealing is rapid and does not require additional energy input. In this respect, the rejoining mechanism differs from that catalyzed by the enzyme DNA ligase, discussed previously (see Figure 5–12). A second type of DNA topoisomerase, topoisomerase II, forms a covalent linkage to both strands of the DNA helix at the same time, making a transient


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Cell_Biology_Alberts. A second type of DNA topoisomerase, topoisomerase II, forms a covalent linkage to both strands of the DNA helix at the same time, making a transient Figure 5–22 The DNA-helix-passing reaction catalyzed by DNA topoisomerase II. Unlike type I topoisomerases, type II enzymes hydrolyze ATP (red), which is needed to release and reset the enzyme after each cycle. Type II topoisomerases are largely confined to proliferating cells in eukaryotes; partly for that reason, they have been effective targets for anticancer drugs. Some of these drugs inhibit topoisomerase II at the third step in the figure and thereby produce high levels of double-strand breaks that kill rapidly dividing cells. The small yellow circles represent the phosphates in the DNA backbone that become covalently bonded to the topoisomerase (see Figure 5–21).


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Cell_Biology_Alberts. double-strand break in the helix. These enzymes are activated by sites on chromosomes where two double helices cross over each other such as those generated by supercoiling in front of a replication fork (see Figure 5–20). Once a topoisomerase II molecule binds to such a crossing site, the protein uses ATP hydrolysis to perform the following set of reactions efficiently: (1) it breaks one double helix reversibly to create a DNA “gate”; (2) it causes the second, nearby double helix to pass through this opening; and (3) it then reseals the break and dissociates from the DNA. At crossover points generated by supercoiling, passage of the double helix through the gate occurs in the direction that will reduce supercoiling. In this way, type II topoisomerases can relieve the overwinding tension generated in front of a replication fork. Their reaction mechanism also allows type II DNA topoisomerases to efficiently separate two interlocked DNA circles (Figure 5–22).


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Cell_Biology_Alberts. Topoisomerase II also prevents the severe DNA tangling problems that would otherwise arise during DNA replication. This role is nicely illustrated by mutant yeast cells that produce, in place of the normal topoisomerase II, a version that is inactive above 37°C. When the mutant cells are warmed to this temperature, their daughter chromosomes remain intertwined after DNA replication and are unable to separate. The enormous usefulness of topoisomerase II for untangling chromosomes can readily be appreciated by anyone who has struggled to remove a tangle from a fishing line without the aid of scissors.


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Cell_Biology_Alberts. Much of what we know about DNA replication was first derived from studies of purified bacterial and bacteriophage multienzyme systems capable of DNA replication in vitro. The development of these systems in the 1970s was greatly facilitated by the prior isolation of mutants in a variety of replication genes; these mutants were exploited to identify and purify the corresponding replication proteins. The first mammalian replication system that accurately replicated DNA in vitro was described in the mid-1980s, and mutations in genes encoding nearly all of the replication components have now been isolated and analyzed in the yeast Saccharomyces cerevisiae. As a result, much is known about the detailed enzymology of DNA replication in eukaryotes, and it is clear that the fundamental features of DNA replication—including replication-fork geometry and the use of a multi-protein replication machine—have been conserved during the long evolutionary process that separated bacteria from


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Cell_Biology_Alberts. of DNA replication—including replication-fork geometry and the use of a multi-protein replication machine—have been conserved during the long evolutionary process that separated bacteria from eukaryotes.


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Cell_Biology_Alberts. There are more protein components in eukaryotic replication machines than there are in the bacterial analogs, even though the basic functions are the same. Thus, for example, the eukaryotic single-strand binding (SSB) protein is formed from three subunits, whereas only a single subunit is found in bacteria. Similarly, the eukaryotic DNA primase is incorporated into a multisubunit enzyme that also contains a polymerase called DNA polymerase α-primase. This protein complex begins each Okazaki fragment on the lagging strand with RNA and then extends the RNA primer with a short length of DNA. At this point, the two main eukaryotic replicative DNA polymerases, Polδand Polε, come into play: Polδcompletes each Okazaki fragment on the lagging strand and Polε extends the leading strand. The increased complexity of eukaryotic replication machinery probably reflects reversal of the covalent attachment of the topoisomerase restores an intact orange double helix more elaborate controls. For


Cell_Biology_Alberts

Cell_Biology_Alberts. increased complexity of eukaryotic replication machinery probably reflects reversal of the covalent attachment of the topoisomerase restores an intact orange double helix more elaborate controls. For example, the orderly maintenance of different cell types and tissues in animals and plants requires that DNA replication be tightly regulated. Moreover, eukaryotic DNA replication must be coordinated with the elaborate process of mitosis, as we discuss in Chapter 17.


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Cell_Biology_Alberts. As we see in the next section, the eukaryotic replication machinery has the added complication of having to replicate through nucleosomes, the repeating structural unit of chromosomes discussed in Chapter 4. Nucleosomes are spaced at intervals of about 200 nucleotide pairs along the DNA, which, as we will see, explains why new Okazaki fragments are synthesized on the lagging strand at intervals of 100–200 nucleotides in eukaryotes, instead of 1000–2000 nucleotides as in bacteria. Nucleosomes may also act as barriers that slow down the movement of DNA polymerase molecules, which may be why eukaryotic replication forks move only about one-tenth as fast as bacterial replication forks.


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Cell_Biology_Alberts. DNA replication takes place at a Y-shaped structure called a replication fork. A self-correcting DNA polymerase enzyme catalyzes nucleotide polymerization in a 5ʹ-to-3ʹ direction, copying a DNA template strand with remarkable fidelity. Since the two strands of a DNA double helix are antiparallel, this 5ʹ-to-3ʹ DNA synthesis can take place continuously on only one of the strands at a replication fork (the leading strand). On the lagging strand, short DNA fragments must be made by a “backstitching” process. Because the self-correcting DNA polymerase cannot start a new chain, these lagging-strand DNA fragments are primed by short RNA primer molecules that are subsequently erased and replaced with DNA.


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Cell_Biology_Alberts. DNA replication requires the cooperation of many proteins. These include (1) DNA polymerase and DNA primase to catalyze nucleoside triphosphate polymerization; (2) DNA helicases and single-strand DNA-binding (SSB) proteins to help in opening up the DNA helix so that it can be copied; (3) DNA ligase and an enzyme that degrades RNA primers to seal together the discontinuously synthesized lagging-strand DNA fragments; and (4) DNA topoisomerases to help to relieve helical winding and DNA tangling problems. Many of these proteins associate with each other at a replication fork to form a highly efficient “replication machine,” through which the activities and spatial movements of the individual components are coordinated.


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Cell_Biology_Alberts. We have seen how a set of replication proteins rapidly and accurately generates two daughter DNA double helices behind a replication fork. But how is this replication machinery assembled in the first place, and how are replication forks created on an intact, double-strand DNA molecule? In this section, we discuss how cells initiate DNA replication and how they carefully regulate this process to ensure that it takes place not only at the proper positions on the chromosome but also at the appropriate time in the life of the cell. We also discuss a few of the special problems that the replication machinery in eukaryotic cells must overcome. These include the need to replicate the enormously long DNA molecules found in eukaryotic chromosomes, as well as the difficulty of copying DNA molecules that are tightly complexed with histones in nucleosomes.


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Cell_Biology_Alberts. As discussed previously, the DNA double helix is normally very stable: the two DNA strands are locked together firmly by many hydrogen bonds formed between the bases on each strand. To begin DNA replication, the double helix must first be opened up and the two strands separated to expose unpaired bases. As we shall see, the process of DNA replication is begun by special initiator proteins that bind to double-strand DNA and pry the two strands apart, breaking the hydrogen bonds between the bases. leading strand of fork 1 of fork 2 leading strand lagging strand of fork 1 of fork 2 Figure 5–23 A replication bubble formed by replication-fork initiation. This diagram outlines the major steps in the initiation of replication forks at replication origins. The structure formed at the last step, in which both strands of the parental DNA helix have been separated from each other and serve as templates for DNA synthesis, is called a replication bubble.


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Cell_Biology_Alberts. Figure 5–24 DNA replication of a bacterial genome. It takes E. coli about 30 minutes to duplicate its genome of 4.6 × 106 nucleotide pairs. For simplicity, no Okazaki fragments are shown on the lagging strand. What happens as the two replication forks approach each other and collide at the end of the replication cycle is not well understood, although the replication machines are disassembled as part of the process.


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Cell_Biology_Alberts. The positions at which the DNA helix is first opened are called replication origins (Figure 5–23). In simple cells like those of bacteria or yeast, origins are specified by DNA sequences several hundred nucleotide pairs in length. This DNA contains both short sequences that attract initiator proteins and stretches of DNA that are especially easy to open. We saw in Figure 4–4 that an A-T base pair is held together by fewer hydrogen bonds than a G-C base pair. Therefore, DNA rich in A-T base pairs is relatively easy to pull apart, and regions of DNA enriched in A-T base pairs are typically found at replication origins.


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Cell_Biology_Alberts. Although the basic process of replication-fork initiation depicted in Figure 5–23 is fundamentally the same for bacteria and eukaryotes, the detailed way in which this process is performed and regulated differs between these two groups of organisms. We first consider the simpler and better-understood case in bacteria and then turn to the more complex situation found in yeasts, mammals, and other eukaryotes. Bacterial Chromosomes Typically Have a Single Origin of DNA Replication


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Cell_Biology_Alberts. The genome of E. coli is contained in a single circular DNA molecule of 4.6 × 106 nucleotide pairs. DNA replication begins at a single origin of replication, and the two replication forks assembled there proceed (at approximately 1000 nucleotides per second) in opposite directions until they meet up roughly halfway around the chromosome (Figure 5–24). The only point at which E. coli can control DNA replication is initiation: once the forks have been assembled at the origin, they synthesize DNA at relatively constant speed until replication is finished. Therefore, it is not surprising that the initiation of DNA replication is highly regulated. The process begins when initiator proteins (in their ATP-bound state) bind in multiple copies to specific DNA sites located at the replication origin, wrapping the DNA around the proteins to form a large protein–DNA complex that destabilizes the adjacent double helix. This complex then attracts two DNA helicases, each bound to a helicase loader,


Cell_Biology_Alberts

Cell_Biology_Alberts. wrapping the DNA around the proteins to form a large protein–DNA complex that destabilizes the adjacent double helix. This complex then attracts two DNA helicases, each bound to a helicase loader, and these are placed around adjacent DNA single strands whose bases have been exposed by the assembly of the initiator protein–DNA complex. The helicase loader is analogous to the clamp loader we encountered above; it has the additional job of keeping the helicase in an inactive form until it is properly loaded onto a nascent replication fork. Once the helicases are loaded, the loaders dissociate and the helicases begin to unwind DNA, exposing enough single-strand DNA for DNA primase to synthesize the first RNA primers (Figure 5–25). This quickly leads to the assembly of remaining proteins to create two replication forks, with replication machines that move, with respect to the replication origin, in opposite directions. They continue to synthesize DNA until all of the DNA template


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Cell_Biology_Alberts. to create two replication forks, with replication machines that move, with respect to the replication origin, in opposite directions. They continue to synthesize DNA until all of the DNA template downstream of each fork has been replicated.


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Cell_Biology_Alberts. In E. coli, the interaction of the initiator protein with the replication origin is carefully regulated, with initiation occurring only when sufficient nutrients are available for the bacterium to complete an entire round of replication. Initiation is also controlled to ensure that only one round of DNA replication occurs for each cell division. After replication is initiated, the initiator protein is inactivated by hydrolysis of its bound ATP molecule, and the origin of replication experiences a “refractory period.” The refractory period is caused by a delay in the methylation of newly incorporated A nucleotides in the origin (Figure 5–26). Initiation cannot occur again until the A’s are methylated and the initiator protein is restored to its ATP-bound state. Eukaryotic Chromosomes Contain Multiple Origins of Replication


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Cell_Biology_Alberts. Eukaryotic Chromosomes Contain Multiple Origins of Replication We have seen how two replication forks begin at a single replication origin in bacteria and proceed in opposite directions, moving away from the origin until all of the DNA in the single circular chromosome is replicated. The bacterial genome is sufficiently small for these two replication forks to duplicate the genome in about 30 minutes. Because of the much greater size of most eukaryotic chromosomes, a different strategy is required to allow their replication in a timely manner.


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Cell_Biology_Alberts. A method for determining the general pattern of eukaryotic chromosome replication was developed in the early 1960s. Human cells growing in culture are labeled for a short time with 3H-thymidine so that the DNA synthesized during this period becomes highly radioactive. The cells are then gently lysed, and the DNA is stretched on the surface of a glass slide coated with a photographic emulsion. Development of the emulsion reveals the pattern of labeled DNA through a technique known as autoradiography. The time allotted for radioactive labeling is chosen to allow each replication fork to move several micrometers along the DNA, so that the replicated DNA can be detected in the light microscope as lines of silver grains, even though the DNA molecule itself is too thin to be visible.


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Cell_Biology_Alberts. Figure 5–25 The proteins that initiate DNA replication in bacteria. The mechanism shown was established by studies in vitro with mixtures of highly purified proteins. For E. coli DNA replication, the major initiator protein, the helicase, and the primase are the dnaA, dnaB, and dnaG proteins, respectively. In the first step, several molecules of the initiator protein bind to specific DNA sequences at the replication origin and destabilize the double helix by forming a compact structure in which the DNA is tightly wrapped around the protein. Next, two helicases are brought in by helicaseloading proteins (the dnaC proteins), which inhibit the helicases until they are properly loaded at the replication origin. Helicase-loading proteins prevent the replicative DNA helices from inappropriately entering other single-strand stretches of DNA in the bacterial genome. Aided by single-strand binding protein (not shown), the loaded helicases open up the DNA, thereby enabling primases to enter and


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Cell_Biology_Alberts. entering other single-strand stretches of DNA in the bacterial genome. Aided by single-strand binding protein (not shown), the loaded helicases open up the DNA, thereby enabling primases to enter and synthesize initial primers. In subsequent steps, two complete replication forks are assembled at the origin and move off in opposite directions. The initiator proteins are displaced as the left-hand fork moves through them (not shown).


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Cell_Biology_Alberts. fully methylated hemimethylated origins are origin resistant to initiation initiation occurs if suffcient origins become fully resources are available to complete methylated, making them a round of DNA replication again competent for initiation In this way, both the rate and the direction of replication-fork movement can be determined (Figure 5–27). From the rate at which tracks of replicated DNA increase in length with increasing labeling time, the eukaryotic replication forks are estimated to travel at about 50 nucleotides per second. This is approximately twentyfold slower than the rate at which bacterial replication forks move, possibly reflecting the increased difficulty of replicating DNA that is packaged tightly in chromatin.


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Cell_Biology_Alberts. An average-size human chromosome contains a single linear DNA molecule of about 150 million nucleotide pairs. It would take 0.02 seconds/nucleotide × 150 × 106 nucleotides = 3.0 × 106 seconds (about 35 days) to replicate such a DNA molecule from end to end with a single replication fork moving at a rate of 50 nucleotides per second. As expected, therefore, the autoradiographic experiments just described reveal that many forks, belonging to separate replication bubbles, are moving simultaneously on each eukaryotic chromosome.


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Cell_Biology_Alberts. Much faster and more sophisticated methods now exist for monitoring DNA replication initiation and tracking the movement of DNA replication forks across whole genomes. One approach uses DNA microarrays—grids the size of a postage stamp studded with hundreds of thousands of fragments of known DNA sequence. As we will see in detail in Chapter 8, each different DNA fragment is placed at a unique position on the microarray, and whole genomes can thereby be represented in an orderly manner. If a DNA sample from a group of replicating cells is broken up and hybridized to a microarray representing that organism’s genome, the amount of each DNA sequence can be determined. Because a segment of a genome that has been replicated will contain twice as much DNA as an unreplicated segment, replication-fork initiation and fork movement can be accurately monitored across an entire genome (Figure 5–28).


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Cell_Biology_Alberts. Experiments of this type have shown the following: (1) Approximately 30,000– 50,000 origins of replication are used each time a human cell divides. (2) The human genome has many more (perhaps tenfold more) potential origins than this, and different cell types use different sets of origins. This may allow a cell to coordinate its active origins with other features of its chromosomes such as which Figure 5–26 Methylation of the E. coli replication origin creates a refractory period for DNA initiation. DNA methylation occurs at GATC sequences, 11 of which are found in the origin of replication (spanning approximately 250 nucleotide pairs). In its hemimethylated state, the origin of replication is bound by an inhibitor protein (Seq A, not shown), which blocks the ability of the initiator proteins to unwind the origin DNA. Eventually (about 15 minutes after replication is initiated), the hemimethylated origins become fully methylated by a DNA methylase enzyme; Seq A then dissociates.


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Cell_Biology_Alberts. A single enzyme, the Dam methylase, is responsible for methylating all E. coli GATC sequences. A lag in methylation after the replication of GATC sequences is also used by the E. coli mismatch proofreading system to distinguish the newly synthesized DNA strand from the parental DNA strand; in that case, the relevant GATC sequences are scattered throughout the chromosome, and they are not bound by Seq A. Figure 5–27 The experiments that demonstrated the pattern in which replication forks are formed and move on eukaryotic chromosomes. The new DNA made in human cells in culture was labeled briefly with a pulse of highly radioactive thymidine (3H-thymidine).


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Cell_Biology_Alberts. (A) In this experiment, the cells were lysed, and the DNA was stretched out on a glass slide that was subsequently covered with a photographic emulsion. After several months, the emulsion was developed, revealing a line of silver grains over the radioactive DNA. The brown DNA in this figure is shown only to help with the interpretation of the autoradiograph; the unlabeled DNA is invisible in such experiments. (B) This experiment was the same except that a further incubation in unlabeled medium allowed additional DNA, with a lower level of radioactivity, to be replicated. The pairs of dark tracks in (B) were found to have silver grains tapering off in opposite directions, demonstrating bidirectional fork movement from a central replication origin where a replication bubble forms (see Figure 5–23). A replication fork is thought to stop only when it encounters a replication fork moving in the opposite direction or when it reaches the end of the chromosome; in this way, all the DNA is


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Cell_Biology_Alberts. 5–23). A replication fork is thought to stop only when it encounters a replication fork moving in the opposite direction or when it reaches the end of the chromosome; in this way, all the DNA is eventually replicated.


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Cell_Biology_Alberts. 258 Chapter 5: DNA Replication, Repair, and Recombination culture of cells arrested before DNA replication begins allow replication to begin fragment DNA, separate strands, and fuorescently label genes are being expressed. The excess origins also provide “backups” in case a primary origin fails. (3) As in bacteria, replication forks are formed in pairs and create a replication bubble as they move in opposite directions away from a common point of origin, stopping only when they collide head-on with a replication fork moving in the opposite direction or when they reach a chromosome end. In this way, many replication forks operate independently on each chromosome and yet form two complete daughter DNA helices. In Eukaryotes, DNA Replication Takes Place During Only One Part of the Cell Cycle


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Cell_Biology_Alberts. In Eukaryotes, DNA Replication Takes Place During Only One Part of the Cell Cycle When growing rapidly, bacteria replicate their DNA nearly continuously. In contrast, DNA replication in most eukaryotic cells occurs only during a specific part of the cell-division cycle, called the DNA synthesis phase or S phase (Figure 5–29). In a mammalian cell, the S phase typically lasts for about 8 hours; in simpler eukaryotic cells such as yeasts, the S phase can be as short as 40 minutes. By its end, each chromosome has been replicated to produce two complete copies, which remain joined together at their centromeres until the M phase (M for mitosis), which soon follows. In Chapter 17, we describe the control system that runs the cell cycle, and we explain why entry into each phase of the cycle requires the cell to have successfully completed the previous phase. In the following sections, we explore how chromosome replication is coordinated within the S phase of the cell cycle.


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Cell_Biology_Alberts. In the following sections, we explore how chromosome replication is coordinated within the S phase of the cell cycle. Different Regions on the Same Chromosome Replicate at Distinct Times in S Phase In mammalian cells, the replication of DNA in the region between one replication origin and the next should normally require only about an hour to complete, given the rate at which a replication fork moves and the largest distances measured between replication origins. Yet S phase usually lasts for about 8 hours in a mammalian cell. This implies that the replication origins are not all activated simultaneously; indeed, replication origins are activated in clusters of about 50 adjacent replication origins, each of which is replicated during only a small part of the total S-phase interval.


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Cell_Biology_Alberts. Figure 5–28 Use of DNA microarrays to monitor the formation and progress of replication forks. For this experiment, a population of cells is synchronized so that they all begin replication at the same time. DNA is collected and hybridized to the microarray; DNA that has been replicated once gives a hybridization signal (dark green squares) twice as high as that of unreplicated DNA (light green squares). The spots on these microarrays represent consecutive sequences along a segment of a chromosome arranged left to right, top to bottom. Only 81 spots are shown here, but the actual arrays contain hundreds of thousands of sequences that span an entire genome. As can be seen, replication begins at an origin and proceeds bidirectionally. For simplicity, only one origin is shown here. In human cells, replication begins at 30,000–50,000 origins located throughout the genome. Using this approach it is possible to observe the formation and progress of every replication fork across a genome.


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Cell_Biology_Alberts. Figure 5–29 The four successive phases of a standard eukaryotic cell cycle. During the G1, S, and G2 phases, the cell grows continuously. During M phase growth stops, the nucleus divides, and the cell divides in two. DNA replication is confined to the part of the cell cycle known as S phase. G1 is the gap between M phase and S phase; G2 is the gap between S phase and M phase. It seems that the order in which replication origins are activated depends, in part, on the chromatin structure in which the origins reside. We saw in Chapter 4 that heterochromatin is a particularly condensed state of chromatin, while euchromatin, where most transcription occurs, has a less condensed conformation. Heterochromatin tends to be replicated very late in S phase, suggesting that the timing of replication is related to the packing of the DNA in chromatin.


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Cell_Biology_Alberts. Once initiated, however, replication forks seem to move at comparable rates throughout S phase, so the extent of chromosome condensation seems to influence the time at which replication forks are initiated, rather than their speed once formed. A Large Multisubunit Complex Binds to Eukaryotic Origins of Replication Having seen that a eukaryotic chromosome is replicated using many origins of replication, each of which “fires” at a characteristic time in S phase of the cell cycle, we turn to the nature of these origins of replication. We saw earlier in this chapter that replication origins have been precisely defined in bacteria as specific DNA sequences that attract initiator proteins, which then assemble the DNA replication machinery. We shall see that this is the case for the single-cell budding yeast S. cerevisiae, but it appears not to be strictly true for most other eukaryotes.


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Cell_Biology_Alberts. For budding yeast, the location of every origin of replication on each chromosome has been determined. The particular chromosome shown in Figure 5–30— chromosome III from S. cerevisiae—is one of the smallest chromosomes known, with a length less than 1/100 that of a typical human chromosome. Its major origins are spaced an average of 30,000 nucleotide pairs apart, but only a subset of these origins is used by a given cell. Nonetheless, this chromosome can be replicated in about 15 minutes.


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Cell_Biology_Alberts. The minimal DNA sequence required for directing DNA replication initiation in S. cerevisiae has been determined by taking a segment of DNA that spans an origin of replication and testing smaller and smaller DNA fragments for their ability to function as origins. Most DNA sequences that can serve as an origin of replication are found to contain (1) a binding site for a large, multisubunit initiator protein called ORC, for origin recognition complex; (2) a stretch of DNA that is rich in As and Ts and therefore easy to melt; and (3) at least one binding site for proteins that facilitate ORC binding, probably by adjusting chromatin structure.


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Cell_Biology_Alberts. In bacteria, once the initiator protein is properly bound to the single origin of replication, the assembly of the replication forks seems to follow more or less automatically. In eukaryotes, the situation is significantly different because of a profound problem eukaryotes have in replicating chromosomes: with so many places to begin replication, how is the process regulated to ensure that all the DNA is copied once and only once?


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Cell_Biology_Alberts. The answer lies in the sequential manner in which the replicative helicase is first loaded onto origins and is then activated to initiate DNA replication. This matter is discussed in detail in Chapter 17, where we consider the machinery that underlies the cell-division cycle. In brief, during G1 phase, the replicative helicases are loaded onto DNA next to ORC to create a prereplicative complex. Then, upon passage from G1 phase to S phase, specialized protein kinases come into play to activate the helicases. The resulting opening of the double helix allows the loading of the remaining replication proteins, including the DNA polymerases. origins of replication telomere centromere telomere


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Cell_Biology_Alberts. origins of replication telomere centromere telomere Figure 5–30 The origins of DNA replication on chromosome III of the yeast S. cerevisiae. This chromosome, one of the smallest eukaryotic chromosomes known, carries a total of 180 genes. As indicated, it contains 18 replication origins, although they are used with different frequencies. Those in red are typically used in less than 10% of cell divisions, while those in green are used about 90% of the time.


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Cell_Biology_Alberts. Figure 5–31 DNA replication initiation in eukaryotes. This mechanism ensures that each origin of replication is activated only once per cell cycle. An origin of replication can be used only if a prereplicative complex forms there in G1 phase. At the beginning of S phase, specialized kinases phosphorylate Mcm and ORC, activating the former and inactivating the latter. A new prereplicative complex cannot form at the origin until the cell progresses to the next G1 phase, when the bound ORC has been dephosphorylated. Note that the eukaryotic Mcm helicase moves along the leading-strand template, whereas the bacterial helicase moves along the lagging-strand template (see Figure 5–25). As the forks begin to move, ORC is displaced, and new ORCs rapidly bind to the newly replicated origins.


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Cell_Biology_Alberts. The protein kinases that trigger DNA replication simultaneously prevent assembly of new prereplicative complexes until the next M phase resets the entire cycle (for details, see pp. 974–975). They do this, in part, by phosphorylating ORC, rendering it unable to accept new helicases. This strategy provides a single window of opportunity for prereplicative complexes to form (G1 phase, when kinase activity is low) and a second window for them to be activated and subsequently disassembled (S phase, when kinase activity is high). Because these two phases of the cell cycle are mutually exclusive and occur in a prescribed order, each origin of replication can fire once and only once during each cell cycle. Features of the Human Genome That Specify Origins of Replication Remain to Be Discovered


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Cell_Biology_Alberts. Features of the Human Genome That Specify Origins of Replication Remain to Be Discovered Compared with the situation in budding yeast, the determinants of replication origins in other eukaryotes have been difficult to discover. It has been possible to identify specific human DNA sequences, each several thousand nucleotide pairs in length, that are sufficient to serve as replication origins. These origins continue to function when moved to a different chromosomal region by recombinant DNA methods, as long as they are placed in a region where the chromatin is relatively uncondensed. However, comparisons of such DNA sequences have not revealed specific DNA sequences that mark origins of replication.


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Cell_Biology_Alberts. Despite this, a human ORC that is very similar to the yeast ORC binds to origins of replication and initiates DNA replication in humans. Many of the other proteins that function in the initiation process in yeast likewise have central roles in humans. It therefore seems likely that the yeast and human initiation mechanisms are similar in outline, but chromatin structure, transcriptional activity, or some property of the genome other than a specific DNA sequence has the central role in attracting ORC and specifying mammalian origins of replication. These ideas could also help to explain how a given mammalian cell chooses which of the many possible origins to use when it replicates its genome and how this choice could differ from cell to cell. Clearly, we have a great deal to discover about the fundamental process of DNA replication initiation. New Nucleosomes Are Assembled Behind the Replication Fork


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Cell_Biology_Alberts. Several additional aspects of DNA replication are specific to eukaryotes. As discussed in Chapter 4, eukaryotic chromosomes are composed of roughly equal mixtures of DNA and protein. Chromosome duplication therefore requires not only the replication of DNA, but also the synthesis and assembly of new chromosomal proteins onto the DNA behind each replication fork. Although we are far from understanding this process in detail, we are beginning to learn how the fundamental unit of chromatin packaging, the nucleosome, is duplicated. The cell requires a large amount of new histone protein, approximately equal in mass to the newly synthesized DNA, to make the new nucleosomes in each cell cycle. For this reason, most eukaryotic organisms possess multiple copies of the gene for each histone. Vertebrate cells, for example, have about 20 repeated gene sets, most containing the genes that encode all five histones (H1, H2A, H2B, H3, and H4).


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Cell_Biology_Alberts. Unlike most proteins, which are made continuously, histones are synthesized mainly in S phase, when the level of histone mRNA increases about fiftyfold as a result of both increased transcription and decreased mRNA degradation. The major histone mRNAs are degraded within minutes when DNA synthesis stops at the end of S phase. The mechanism depends on special properties of the 3ʹends of these mRNAs, as discussed in Chapter 7. In contrast, the histone proteins themselves are remarkably stable and may survive for the entire life of a cell. The tight linkage between DNA synthesis and histone synthesis appears to reflect a feedback mechanism that monitors the level of free histone to ensure that the amount of histone made exactly matches the amount of new DNA synthesized.


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Cell_Biology_Alberts. As a replication fork advances, it must pass through the parental nucleosomes. In the cell, efficient replication requires chromatin remodeling complexes (discussed in Chapter 4) to destabilize the DNA–histone interfaces. Aided by such complexes, replication forks can transit even highly condensed chromatin efficiently.


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Cell_Biology_Alberts. As a replication fork passes through chromatin, the histones are transiently displaced leaving about 600 nucleotide pairs of non-nucleosomal DNA in its wake. The reestablishment of nucleosomes behind a moving fork occurs in an intriguing way. When a nucleosome is traversed by a replication fork, the histone octamer appears to be broken into an H3-H4 tetramer and two H2A-H2B dimers (discussed in Chapter 4). The H3-H4 tetramer remains loosely associated with DNA and is distributed at random to one or the other daughter duplex, but the H2A-H2B dimers are released completely from DNA. Freshly made H3-H4 tetramers are added to the newly synthesized DNA to fill in the “spaces,” and H2A-H2B dimers—half of which are old and half new—are then added at random to complete the nucleosomes (Figure 5–32). The formation of new nucleosomes behind a replication fork has an important consequence for the process of DNA replication itself. As DNA polymerase δ discontinuously synthesizes the lagging


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Cell_Biology_Alberts. 5–32). The formation of new nucleosomes behind a replication fork has an important consequence for the process of DNA replication itself. As DNA polymerase δ discontinuously synthesizes the lagging strand (see pp. 253–254), the length of each Okazaki fragment is determined by the point at which DNA polymerase δ is blocked by a newly formed nucleosome. This tight coupling between nucleosome duplication and DNA replication explains why the length of Okazaki fragments in eukaryotes (~200 nucleotides) is approximately the same as the nucleosome repeat length.


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Cell_Biology_Alberts. 262 Chapter 5: DNA Replication, Repair, and Recombination H2A-H2B dimer displaced in front of replication fork The orderly and rapid addition of new H3-H4 tetramers and H2A-H2B dimers behind a replication fork requires histone chaperones (also called chromatin assembly factors). These multisubunit complexes bind the highly basic histones and release them for assembly only in the appropriate context. The histone chaperones, along with their cargoes, are directed to newly replicated DNA through a specific interaction with the eukaryotic sliding clamp called PCNA (see Figure 5–32B). These clamps are left behind moving replication forks and remain on the DNA long enough for the histone chaperones to complete their tasks. Telomerase Replicates the Ends of Chromosomes


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Cell_Biology_Alberts. Telomerase Replicates the Ends of Chromosomes We saw earlier that synthesis of the lagging strand at a replication fork must occur discontinuously through a backstitching mechanism that produces short DNA fragments. This mechanism encounters a special problem when the replication fork reaches an end of a linear chromosome. The final RNA primer synthesized on the lagging-strand template cannot be replaced by DNA because there is no 3ʹ-OH end available for the repair polymerase. Without a mechanism to deal with this problem, DNA would be lost from the ends of all chromosomes each time a cell divides.


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Cell_Biology_Alberts. Bacteria solve this “end-replication” problem by having circular DNA molecules as chromosomes (see Figure 5–24). Eukaryotes solve it in a different way: they have specialized nucleotide sequences at the ends of their chromosomes that are incorporated into structures called telomeres (discussed in Chapter 4). Telomeres contain many tandem repeats of a short sequence that is similar in organisms as diverse as protozoa, fungi, plants, and mammals. In humans, the sequence of the repeat unit is GGGTTA, and it is repeated roughly a thousand times at each telomere.


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Cell_Biology_Alberts. Telomere DNA sequences are recognized by sequence-specific DNA-binding proteins that attract an enzyme, called telomerase, that replenishes these sequences each time a cell divides. Telomerase recognizes the tip of an existing telomere DNA repeat sequence and elongates it in the 5ʹ-to-3ʹ direction, using an RNA template that is a component of the enzyme itself to synthesize new copies of the repeat (Figure 5–33). The enzymatic portion of telomerase resembles other reverse transcriptases, proteins that synthesize DNA using an RNA template, although, in this case, the telomerase RNA also contributes functional groups to make the catalysis more efficient. After extension of the parental DNA strand by telomerase, replication of the lagging strand at the chromosome end can be completed by the conventional DNA polymerases, using these extensions as a template to synthesize the complementary strand (Figure 5–34).


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Cell_Biology_Alberts. Figure 5–32 Formation of nucleosomes behind a replication fork. Parental H3-H4 tetramers are distributed at random to the daughter DNA molecules, with roughly equal numbers inherited by each daughter. In contrast, H2A-H2B dimers are released from the DNA as the replication fork passes. This release begins just in front of the replication fork and is facilitated by chromatin remodeling complexes that move with the fork. Histone chaperones (NAP1 and CAF1) restore the full complement of histones to daughter molecules using both parental and newly synthesized histones. Although some daughter nucleosomes contain only parental histones or only newly synthesized histones, most are hybrids of old and new. For simplicity, the DNA double helix shown as a single red line. (Adapted from J.D. Watson et al., Molecular Biology of the Gene, 5th ed. Cold Spring Harbor: Cold Spring Harbor Laboratory Press, 2004.) remainder of telomerase proteintelomerase RNA region of telomerase RNA used as


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Cell_Biology_Alberts. Watson et al., Molecular Biology of the Gene, 5th ed. Cold Spring Harbor: Cold Spring Harbor Laboratory Press, 2004.) remainder of telomerase proteintelomerase RNA region of telomerase RNA used as template “palm“—active site of telomerase protein rest of synthesized chromosome telomere DNA


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Cell_Biology_Alberts. Telomeres Are Packaged Into Specialized Structures That Protect the Ends of Chromosomes The ends of chromosomes present cells with an additional problem. As we will see in the next part of this chapter, when a chromosome is accidently broken, the break is rapidly repaired (see Figure 5–45). Telomeres must clearly be distinguished from these accidental breaks; otherwise the cell will attempt to “repair” telomeres, causing chromosome fusions and other genetic abnormalities. Telomeres have several features to prevent this from happening.


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Cell_Biology_Alberts. A specialized nuclease chews back the 5ʹend of a telomere leaving a protruding single-strand end. This protruding end—in combination with the GGGTTA repeats in telomeres—attracts a group of proteins that form a protective chromosome cap known as shelterin. In particular, shelterin “hides” telomeres from the cell’s damage detectors that continually monitor DNA. When human telomeres are artificially cross-linked and viewed by electron microscopy, structures known as “t-loops” are observed in which the protruding end of the telomere loops back and tucks itself into the duplex DNA of the telomere repeat sequence (Figure 5–35). It is believed that t-loops are regulated by shelterin and provide additional protection for the ends of chromosomes. incomplete, newly synthesized lagging strand direction of AACCCC telomere 5˜ synthesisTELOMERASE EXTENDS 3˜ END telomerase with bound RNA template


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Cell_Biology_Alberts. incomplete, newly synthesized lagging strand direction of AACCCC telomere 5˜ synthesisTELOMERASE EXTENDS 3˜ END telomerase with bound RNA template Figure 5–33 Structure of a portion of telomerase. Telomerase is a large protein– RNA complex. The RNA (blue) contains a templating sequence for synthesizing new DNA telomere repeats. The synthesis reaction itself is carried out by the reverse transcriptase domain of the protein, shown in green. A reverse transcriptase is a special form of polymerase enzyme that uses an RNA template to make a DNA strand; telomerase is unique in carrying its own RNA template with it. Telomerase also has several additional protein domains (not shown) that are needed to assemble the enzyme at the ends of chromosomes. (Modified from J. Lingner and T.R. Cech, Curr. Opin. Genet. Dev. 8:226–232, 1998. With permission from Elsevier.)


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Cell_Biology_Alberts. Figure 5–34 Telomere replication. Shown here are the reactions that synthesize the repeating sequences that form the ends of the chromosomes (telomeres) of diverse eukaryotic organisms. The 3ʹ end of the parental DNA strand is extended by RNA-templated DNA synthesis; this allows the incomplete daughter DNA strand that is paired with it to be extended in its 5ʹ direction. This incomplete, lagging strand is presumed to be completed by DNA polymerase α, which carries a DNA primase as one of its subunits (Movie 5.6). The telomere sequence illustrated is that of the ciliate Tetrahymena, in which these reactions were first discovered. Because the processes that grow and shrink each telomere sequence are only approximately balanced, a chromosome end contains a variable number of telomeric repeats. Not surprisingly, many cells have homeostatic mechanisms that maintain the number of these repeats within a limited range (Figure 5–36).


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Cell_Biology_Alberts. In most of the dividing somatic cells of humans, however, telomeres gradually shorten, and it has been proposed that this provides a counting mechanism that helps prevent the unlimited proliferation of wayward cells in adult tissues. In its simplest form, this idea holds that our somatic cells start off in the embryo with a full complement of telomeric repeats. These are then eroded to different extents in different cell types. Some stem cells, notably those in tissues that must be replenished at a high rate throughout life—bone marrow or gut lining, for example— retain full telomerase activity. However, in many other types of cells, the level of telomerase is turned down so that the enzyme cannot quite keep up with chromosome duplication. Such cells lose 100–200 nucleotides from each telomere every time they divide. After many cell generations, the descendant cells will inherit chromosomes that lack telomere function, and, as a result of this defect, activate a DNA-damage response


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Cell_Biology_Alberts. every time they divide. After many cell generations, the descendant cells will inherit chromosomes that lack telomere function, and, as a result of this defect, activate a DNA-damage response causing them to withdraw permanently from the cell cycle and cease dividing—a process called replicative cell senescence (discussed in Chapter 17). In theory, such a mechanism could provide a safeguard against the uncontrolled cell proliferation of abnormal cells in somatic tissues, thereby helping to protect us from cancer.


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Cell_Biology_Alberts. fraction of chromosome ends Figure 5–35 A t-loop at the end of a mammalian chromosome. (A) Electron micrograph of the DNA at the end of an interphase human chromosome. The chromosome was fixed, deproteinated, and artificially thickened before viewing. The loop seen here is approximately 15,000 nucleotide pairs in length. (B) Structure of a t-loop. The insertion of the single-strand 3ʹ end into the duplex repeats is carried out, and the structure maintained, by specialized proteins. (From J.D. Griffith et al., Cell 97:503–514, 1999. With permission from Elsevier.)


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Cell_Biology_Alberts. Figure 5–36 A demonstration that yeast cells control the length of their telomeres. In this experiment, the telomere at one end of a particular chromosome is artificially made either longer (left) or shorter (right) than average. After many cell divisions, the chromosome recovers, showing an average telomere length and a length distribution that is typical of the other chromosomes in the yeast cell. A similar feedback mechanism for controlling telomere length has been proposed for the germ-line cells of animals.


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Cell_Biology_Alberts. The idea that telomere length acts as a “measuring stick” to count cell divisions and thereby regulate the lifetime of the cell lineage has been tested in several ways. For certain types of human cells grown in tissue culture, the experimental results support such a theory. Human fibroblasts normally proliferate for about 60 cell divisions in culture before undergoing replicative cell senescence. Like most other somatic cells in humans, fibroblasts produce only low levels of telomerase, and their telomeres gradually shorten each time they divide. When telomerase is provided to the fibroblasts by inserting an active telomerase gene, telomere length is maintained and many of the cells now continue to proliferate indefinitely.


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Cell_Biology_Alberts. It has been proposed that this type of control on cell proliferation may contribute to the aging of animals like ourselves. These ideas have been tested by producing transgenic mice that lack telomerase entirely. The telomeres in mouse chromosomes are about five times longer than human telomeres, and the mice must therefore be bred through three or more generations before their telomeres have shrunk to the normal human length. It is therefore perhaps not surprising that the first generations of mice develop normally. However, the mice in later generations develop progressively more defects in some of their highly proliferative tissues. In addition, these mice show signs of premature aging and have a pronounced tendency to develop tumors. In these and other respects these mice resemble humans with the genetic disease dyskeratosis congenita. Individuals afflicted with this disease carry one functional and one nonfunctional copy of the telomerase RNA gene; they have prematurely shortened


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Cell_Biology_Alberts. with the genetic disease dyskeratosis congenita. Individuals afflicted with this disease carry one functional and one nonfunctional copy of the telomerase RNA gene; they have prematurely shortened telomeres and typically die of progressive bone marrow failure. They also develop lung scarring and liver cirrhosis and show abnormalities in various epidermal structures including skin, hair follicles, and nails.


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Cell_Biology_Alberts. The above observations demonstrate that controlling cell proliferation by telomere shortening poses a risk to an organism, because not all of the cells that begin losing the ends of their chromosomes will stop dividing. Some apparently become genetically unstable, but continue to divide, giving rise to variant cells that can lead to cancer. Clearly, the use of telomere shortening as a regulating mechanism is not foolproof and, like many mechanisms in the cell, seems to strike a balance between benefit and risk. The proteins that initiate DNA replication bind to DNA sequences at a replication origin to catalyze the formation of a replication bubble with two outward-moving replication forks. The process begins when an initiator protein–DNA complex is formed that subsequently loads a DNA helicase onto the DNA template. Other proteins are then added to form the multienzyme “replication machine” that catalyzes DNA synthesis at each replication fork.


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Cell_Biology_Alberts. In bacteria and some simple eukaryotes, replication origins are specified by specific DNA sequences that are only several hundred nucleotide pairs long. In other eukaryotes, such as humans, the sequences needed to specify an origin of DNA replication seem to be less well defined, and the origin can span several thousand nucleotide pairs.


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Cell_Biology_Alberts. Bacteria typically have a single origin of replication in a circular chromosome. With fork speeds of up to 1000 nucleotides per second, they can replicate their genome in less than an hour. Eukaryotic DNA replication takes place in only one part of the cell cycle, the S phase. The replication fork in eukaryotes moves about 10 times more slowly than the bacterial replication fork, and the much longer eukaryotic chromosomes each require many replication origins to complete their replication in an S phase, which typically lasts for 8 hours in human cells. The different replication origins in these eukaryotic chromosomes are activated in a sequence, determined in part by the structure of the chromatin, with the most condensed regions of chromatin typically beginning their replication last. After the replication fork has passed, chromatin structure is re-formed by the addition of new histones to the old histones that are directly inherited by each daughter DNA molecule.


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Cell_Biology_Alberts. Eukaryotes solve the problem of replicating the ends of their linear chromosomes with a specialized end structure, the telomere, maintained by a special nucleotide polymerizing enzyme called telomerase. Telomerase extends one of the DNA strands at the end of a chromosome by using an RNA template that is an integral part of the enzyme itself, producing a highly repeated DNA sequence that typically extends for thousands of nucleotide pairs at each chromosome end. Telomeres have specialized structures that distinguish them from broken ends of chromosomes, ensuring that they are not mistakenly repaired.


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Cell_Biology_Alberts. Maintaining the genetic stability that an organism needs for its survival requires not only an extremely accurate mechanism for replicating DNA, but also mechanisms for repairing the many accidental lesions that DNA continually suffers. Most such spontaneous changes in DNA are temporary because they are immediately corrected by a set of processes that are collectively called DNA repair. Of the tens of thousands of random changes created every day in the DNA of a human cell by heat, metabolic accidents, radiation of various sorts, and exposure to substances in the environment, only a few (less than 0.02%) accumulate as permanent mutations in the DNA sequence. The rest are eliminated with remarkable efficiency by DNA repair.


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Cell_Biology_Alberts. The importance of DNA repair is evident from the large investment that cells make in the enzymes that carry it out: several percent of the coding capacity of most genomes is devoted solely to DNA repair functions. The importance of DNA repair is also demonstrated by the increased rate of mutation that follows the inactivation of a DNA repair gene. Many DNA repair proteins and the genes that encode them—which we now know operate in a wide range of organisms, including humans—were originally identified in bacteria by the isolation and characterization of mutants that displayed an increased mutation rate or an increased sensitivity to DNA-damaging agents.


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Cell_Biology_Alberts. Recent studies of the consequences of a diminished capacity for DNA repair in humans have linked many human diseases with decreased repair (Table 5–2). Thus, we saw previously that defects in a human gene whose product normally functions to repair the mismatched base pairs resulting from DNA replication errors can lead to an inherited predisposition to cancers of the colon and some other organs, reflecting an increased mutation rate. In another human disease, xeroderma pigmentosum (XP), the afflicted individuals have an extreme sensitivity to ultraviolet radiation because they are unable to repair certain DNA photo-products. This repair defect results in an increased mutation rate that leads to serious skin lesions and an increased susceptibility to skin cancers. Finally, mutations in the Brca1 and Brca2 genes compromise a type of DNA repair known as homologous recombination and are a cause of hereditary breast and ovarian cancer.


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Cell_Biology_Alberts. Without DNA Repair, Spontaneous DNA Damage Would Rapidly Change DNA Sequences Although DNA is a highly stable material—as required for the storage of genetic information—it is a complex organic molecule that is susceptible, even under normal cell conditions, to spontaneous changes that would lead to mutations if left unrepaired (Figure 5–37 and see Table 5–3). For example, the DNA of each Figure 5–37 A summary of spontaneous alterations that require DNA repair.


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Cell_Biology_Alberts. The sites on each nucleotide modified by spontaneous oxidative damage (red arrows), hydrolytic attack (blue arrows), and methylation (green arrows) are shown, with the width of each arrow indicating the relative frequency of each event (see Table 5–3). (After T. Lindahl, Nature 362:709–715, 1993. With permission from Macmillan Publishers Ltd.) human cell loses about 18,000 purine bases (adenine and guanine) every day because their N-glycosyl linkages to deoxyribose hydrolyze, a spontaneous reaction called depurination. Similarly, a spontaneous deamination of cytosine to uracil in DNA occurs at a rate of about 100 bases per cell per day (Figure 5–38). DNA bases are also occasionally damaged by an encounter with reactive metabolites produced in the cell, including reactive forms of oxygen and the high-energy methyl donor S-adenosylmethionine, or by exposure to chemicals in the environment. Likewise, ultraviolet radiation from the sun can produce a covalent linkage between two adjacent


Cell_Biology_Alberts

Cell_Biology_Alberts. and the high-energy methyl donor S-adenosylmethionine, or by exposure to chemicals in the environment. Likewise, ultraviolet radiation from the sun can produce a covalent linkage between two adjacent pyrimidine bases in DNA to form, for example, thymine dimers (Figure 5–39). If left uncorrected when the DNA is replicated, most of these changes would be expected to lead either to the deletion of one or more base pairs or to a base-pair substitution in the daughter DNA chain (Figure 5–40). The mutations would then be propagated throughout subsequent cell generations. Such a high rate of random changes in the DNA sequence would have disastrous consequences.


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Cell_Biology_Alberts. The DNA Double Helix Is Readily Repaired The double-helical structure of DNA is ideally suited for repair because it carries two separate copies of all the genetic information—one in each of its two strands. Thus, when one strand is damaged, the complementary strand retains an intact copy of the same information, and this copy is generally used to restore the correct nucleotide sequences to the damaged strand.


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Cell_Biology_Alberts. An indication of the importance of a double-strand helix to the safe storage of genetic information is that all cells use it; only a few small viruses use single-strand DNA or RNA as their genetic material. The types of repair processes described in this section cannot operate on such nucleic acids, and once damaged, the chance of a permanent nucleotide change occurring in these single-strand genomes of viruses is thus very high. It seems that only organisms with tiny genomes (and therefore tiny targets for DNA damage) can afford to encode their genetic information in any molecule other than a DNA double helix.


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Cell_Biology_Alberts. Figure 5–38 Depurination and deamination. These reactions are two of the most frequent spontaneous chemical reactions that create serious DNA damage in cells. Depurination can release guanine (shown here), as well as adenine, from DNA. The major type of deamination reaction converts cytosine to an altered DNA base, uracil (shown here), but deamination occurs on other bases as well. These reactions normally take place in double-helical DNA; for convenience, only one strand is shown. Figure 5–39 The most common type of thymine dimer. This type of damage occurs in the DNA of cells exposed to ultraviolet irradiation (as in sunlight). A similar dimer will form between any two neighboring pyrimidine bases (C or T residues) in DNA.


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Cell_Biology_Alberts. Cells have multiple pathways to repair their DNA using different enzymes that act upon different kinds of lesions. Figure 5–41 shows two of the most common pathways. In both, the damage is excised, the original DNA sequence is restored by a DNA polymerase that uses the undamaged strand as its template, and a remaining break in the double helix is sealed by DNA ligase (see Figure 5–12).


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Cell_Biology_Alberts. The two pathways differ in the way in which they remove the damage from DNA. The first pathway, called base excision repair, involves a battery of enzymes called DNA glycosylases, each of which can recognize a specific type of altered base in DNA and catalyze its hydrolytic removal. There are at least six types of these enzymes, including those that remove deaminated Cs, deaminated As, different types of alkylated or oxidized bases, bases with opened rings, and bases in which a carbon–carbon double bond has been accidentally converted to a carbon–carbon single bond. How is an altered base detected within the context of the double helix? A key step is an enzyme-mediated “flipping-out” of the altered nucleotide from the helix, which allows the DNA glycosylase to probe all faces of the base for damage (Figure 5–42). It is thought that these enzymes travel along DNA using base-flipping to evaluate the status of each base. Once an enzyme finds the damaged base that it recognizes, it


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Cell_Biology_Alberts. base for damage (Figure 5–42). It is thought that these enzymes travel along DNA using base-flipping to evaluate the status of each base. Once an enzyme finds the damaged base that it recognizes, it removes that base from its sugar.


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Cell_Biology_Alberts. The “missing tooth” created by DNA glycosylase action is recognized by an enzyme called AP endonuclease (AP for apurinic or apyrimidinic, endo to signify that the nuclease cleaves within the polynucleotide chain), which cuts the phosphodiester backbone, after which the resulting gap is repaired (see Figure 5–41A). Depurination, which is by far the most frequent type of damage suffered by DNA, also leaves a deoxyribose sugar with a missing base. Depurinations are directly repaired beginning with AP endonuclease, following the bottom half of the pathway in Figure 5–41A. a G has been changed to an A


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Cell_Biology_Alberts. a G has been changed to an A Figure 5–40 How chemical modifications of nucleotides produce mutations. (A) Deamination of cytosine, if uncorrected, results in the substitution of one base for another when the DNA is replicated. As shown in Figure 5–38, deamination of cytosine produces uracil. Uracil differs from cytosine in its base-pairing properties and preferentially base-pairs with adenine. The DNA replication machinery therefore adds an adenine when it encounters a uracil on the template strand. (B) Depurination can lead to the loss of a nucleotide pair. When the replication machinery encounters a missing purine on the template strand, it may skip to the next complete nucleotide as illustrated here, thus producing a nucleotide deletion in the newly synthesized strand. Many other types of DNA damage (see Figure 5–37), if left uncorrected, also produce mutations when the DNA is replicated. G C T A T C C DNA helix with missing base C G A G T A G G


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Cell_Biology_Alberts. G C T A T C C DNA helix with missing base C G A G T A G G DNA helix with single-nucleotide gap C G A G T A G G DNA POLYMERASE ADDS NEW NUCLEOTIDE, DNA LIGASE SEALS NICK G C T C A T C C DNA helix with 12nucleotide gap


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Cell_Biology_Alberts. Figure 5– 41 A comparison of two major DNA repair pathways. (A) Base excision repair. This pathway starts with a DNA glycosylase. Here, the enzyme uracil DNA glycosylase removes an accidentally deaminated cytosine in DNA. After the action of this glycosylase (or another DNA glycosylase that recognizes a different kind of damage), the sugar phosphate with the missing base is cut out by the sequential action of AP endonuclease and a phosphodiesterase. (These same enzymes begin the repair of depurinated sites directly.) The gap of a single nucleotide is then filled by DNA polymerase and DNA ligase. The net result is that the U that was created by accidental deamination is restored to a C. AP endonuclease is so-named because it recognizes any site in the DNA helix that contains a deoxyribose sugar with a missing base; such sites can arise either by the loss of a purine (apurinic sites) or by the loss of a pyrimidine (apyrimidinic sites). (B) Nucleotide excision repair. In bacteria, after


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Cell_Biology_Alberts. sugar with a missing base; such sites can arise either by the loss of a purine (apurinic sites) or by the loss of a pyrimidine (apyrimidinic sites). (B) Nucleotide excision repair. In bacteria, after a multienzyme complex has recognized a lesion such as a pyrimidine dimer (see Figure 5–39), one cut is made on each side of the lesion, and an associated DNA helicase then removes the entire portion of the damaged strand. The excision repair machinery in bacteria leaves the gap of 12 nucleotides shown. In humans, once the damaged DNA is recognized, a helicase is recruited to unwind the DNA duplex locally. Next, the excision nuclease enters and cleaves on either side of the damage, leaving a gap of about 30 nucleotides. The nucleotide excision repair machinery in both bacteria and humans can recognize and repair many different types of DNA damage.


Cell_Biology_Alberts

Cell_Biology_Alberts. The second major repair pathway is called nucleotide excision repair. This mechanism can repair the damage caused by almost any large change in the structure of the DNA double helix. Such “bulky lesions” include those created by the covalent reaction of DNA bases with large hydrocarbons (such as the carcinogen benzopyrene, found in tobacco smoke, coal tar, and diesel exhaust), as well as the various pyrimidine dimers (T-T, T-C, and C-C) caused by sunlight. In this pathway, a large multienzyme complex scans the DNA for a distortion in the double helix, rather than for a specific base change. Once it finds a lesion, it cleaves the phosphodiester backbone of the abnormal strand on both sides of the distortion, and a DNA helicase peels away the single-strand oligonucleotide containing the lesion. The large gap produced in the DNA helix is then repaired by DNA polymerase and DNA ligase (see Figure 5–41B).


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Cell_Biology_Alberts. An alternative to base and nucleotide excision repair processes is direct chemical reversal of DNA damage, and this strategy is selectively employed for the rapid removal of certain highly mutagenic or cytotoxic lesions. For example, the alkylation lesion O6-methylguanine has its methyl group removed by direct transfer to a cysteine residue in the repair protein itself, which is destroyed in the reaction. In another example, methyl groups in the alkylation lesions 1-methyladenine and 3-methylcytosine are “burnt off” by an iron-dependent demethylase, with release of formaldehyde from the methylated DNA and regeneration of the native base. Coupling Nucleotide Excision Repair to Transcription Ensures That the Cell’s Most Important DNA Is Efficiently Repaired


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Cell_Biology_Alberts. All of a cell’s DNA is under constant surveillance for damage, and the repair mechanisms we have described act on all parts of the genome. However, cells have a way of directing DNA repair to the DNA sequences that are most urgently needed. They do this by linking RNA polymerase, the enzyme that transcribes DNA into RNA as the first step in gene expression, to the nucleotide excision repair pathway. As discussed above, this repair system can correct many different types of DNA damage. RNA polymerase stalls at DNA lesions and, through the use of coupling proteins, directs the excision repair machinery to these sites. In bacteria, where genes are relatively short, the stalled RNA polymerase can be dissociated from the DNA; the DNA is repaired, and the gene is transcribed again from the beginning. In eukaryotes, where genes can be enormously long, a more complex reaction is used to “back up” the RNA polymerase, repair the damage, and then restart the polymerase.


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Cell_Biology_Alberts. The importance of transcription-coupled excision repair is seen in people with Cockayne syndrome, which is caused by a defect in this coupling. These individuals suffer from growth retardation, skeletal abnormalities, progressive neural retardation, and severe sensitivity to sunlight. Most of these problems are thought to arise from RNA polymerase molecules that become permanently stalled at sites of DNA damage that lie in important genes. The Chemistry of the DNA Bases Facilitates Damage Detection


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Cell_Biology_Alberts. The Chemistry of the DNA Bases Facilitates Damage Detection The DNA double helix seems optimal for repair. As noted above, it contains a backup copy of all genetic information. Equally importantly, the nature of the four bases in DNA makes the distinction between undamaged and damaged bases very clear. For example, every possible deamination event in DNA yields an “unnatural” base, which can be directly recognized and removed by a specific DNA glycosylase. Hypoxanthine, for example, is the simplest purine base capable of pairing specifically with C, but hypoxanthine is the direct deamination product of A (Figure 5–43A). The addition of a second amino group to hypoxanthine


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Cell_Biology_Alberts. Figure 5–42 The recognition of an unusual nucleotide in DNA by base-flipping. The DNA glycosylase family of enzymes recognizes specific inappropriate bases in the conformation shown. Each of these enzymes cleaves the glycosyl bond that connects a particular recognized base (yellow) to the backbone sugar, removing it from the DNA. (A) Stick model; (B) space-filling model.


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Cell_Biology_Alberts. Figure 5–43 The deamination of DNA nucleotides. In each case, the oxygen atom that is added in this reaction with water is colored red. (A) The spontaneous deamination products of A and G are recognizable as unnatural when they occur in DNA and thus are readily found and repaired. The deamination of C to U was also illustrated in Figure 5–38; T has no amino group to remove. (B) About 3% of the C nucleotides in vertebrate DNAs are methylated to help in controlling gene expression (discussed in Chapter 7). When these 5-methyl C nucleotides are accidentally deaminated, they form the natural nucleotide T. However, this T will be paired with a G on the opposite strand, forming a mismatched base pair. produces G, which cannot be formed from A by spontaneous deamination, and whose deamination product (xanthine) is likewise unique.


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Cell_Biology_Alberts. produces G, which cannot be formed from A by spontaneous deamination, and whose deamination product (xanthine) is likewise unique. As discussed in Chapter 6, RNA is thought, on an evolutionary time scale, to have served as the genetic material before DNA, and it seems likely that the genetic code was initially carried in the four nucleotides A, C, G, and U. This raises the question of why the U in RNA was replaced in DNA by T (which is 5-methyl U). We have seen that the spontaneous deamination of C converts it to U, but that this event is rendered relatively harmless by uracil DNA glycosylase. However, if DNA contained U as a natural base, the repair system would not be able to distinguish a deaminated C from a naturally occurring U.


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Cell_Biology_Alberts. A special situation occurs in vertebrate DNA, in which selected C nucleotides are methylated at specific CG sequences that are associated with inactive genes (discussed in Chapter 7). The accidental deamination of these methylated C nucleotides produces the natural nucleotide T (Figure 5–43B) in a mismatched base pair with a G on the opposite DNA strand. To help in repairing deaminated methylated C nucleotides, a special DNA glycosylase recognizes a mismatched base pair involving T in the sequence T-G and removes the T. This DNA repair mechanism must be relatively ineffective, however, because methylated C nucleotides are exceptionally common sites for mutations in vertebrate DNA. It is striking that, even though only about 3% of the C nucleotides in human DNA are methylated, mutations in these methylated nucleotides account for about one-third of the single-base mutations that have been observed in inherited human diseases.


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Cell_Biology_Alberts. If a cell’s DNA suffers heavy damage, the repair mechanisms that we have discussed are often insufficient to cope with it. In these cases, a different strategy is called into play, one that entails some risk to the cell. The highly accurate replicative DNA polymerases stall when they encounter damaged DNA, and in emergencies cells employ versatile, but less accurate, backup polymerases, known as translesion polymerases, to replicate through the DNA damage.


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Cell_Biology_Alberts. Human cells have seven translesion polymerases, some of which can recognize a specific type of DNA damage and correctly add the nucleotide required to restore the initial sequence. Others make only “good guesses,” especially when the template base has been extensively damaged. These enzymes are not as accurate as the normal replicative polymerases when they copy a normal DNA sequence. For one thing, the translesion polymerases lack exonucleolytic proofreading activity; in addition, many are much less discriminating than the replicative polymerase in choosing which nucleotide to incorporate initially. Presumably for this reason, each such translesion polymerase is given a chance to add only one or a few nucleotides before the highly accurate replicative polymerase resumes DNA synthesis.


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Cell_Biology_Alberts. Despite their usefulness in allowing heavily damaged DNA to be replicated, these translesion polymerases do, as noted above, pose risks to the cell. They are probably responsible for most of the base-substitution and single-nucleotide deletion mutations that accumulate in genomes; although they generally produce mutations when copying damaged DNA (see Figure 5–40), they probably also create mutations—at a low level—on undamaged DNA. Clearly, it is important for the cell to tightly regulate these polymerases, releasing them only at sites of DNA damage. Exactly how this happens for each translesion polymerase remains to be discovered, but a conceptual model is given in Figure 5–44. The principle of this model applies to many of the DNA repair processes discussed in this chapter: because the enzymes that carry out these reactions are potentially dangerous to the genome, they must be brought into play only at sites of damage.


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Cell_Biology_Alberts. An especially dangerous type of DNA damage occurs when both strands of the double helix are broken, leaving no intact template strand to enable accurate Figure 5–44 Translesion DNA polymerases can use damaged templates. According to this model, a replicative polymerase stalled at a site of DNA damage is recognized by the cell as needing rescue. Specialized enzymes covalently modify the sliding clamp (typically, it is ubiquitylated—see Figure 3–69) which releases the replicative DNA polymerase and, together with damaged DNA, attracts a translesion polymerase specific to that type of damage. Once the damaged DNA is bypassed, the covalent modification of the clamp is removed, the translesion polymerase dissociates, and the replicative polymerase is brought back into play.


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Cell_Biology_Alberts. removal of covalent modifcations, reloading of replicative DNA polymerase, DNA synthesis continues repair. Ionizing radiation, replication errors, oxidizing agents, and other metabolites produced in the cell cause breaks of this type. If these lesions were left unrepaired, they would quickly lead to the breakdown of chromosomes into smaller fragments and to loss of genes when the cell divides. However, two distinct mechanisms have evolved to deal with this type of damage (Figure 5–45). The simplest to understand is nonhomologous end joining, in which the broken ends are simply brought together and rejoined by DNA ligation, generally with the loss of nucleotides at the site of joining (Figure 5–46). This end-joining mechanism, which can be seen as a “quick and dirty” solution to the repair of double-strand breaks, is common in mammalian somatic cells. Although a change in the DNA sequence (a mutation) results at the site of breakage, so little of the mammalian genome is essential for


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Cell_Biology_Alberts. of double-strand breaks, is common in mammalian somatic cells. Although a change in the DNA sequence (a mutation) results at the site of breakage, so little of the mammalian genome is essential for life that this mechanism is apparently an acceptable solution to the problem of rejoining broken chromosomes. By the time a human reaches the age of 70, the typical somatic cell contains over 2000 such “scars,” distributed throughout its genome, representing places where DNA has been inaccurately repaired by nonhomologous end joining. But nonhomologous end joining presents another danger: because there seems to be no mechanism to ensure that two ends being joined were originally next to each other in the genome, nonhomologous end joining can occasionally generate rearrangements in which one broken chromosome becomes covalently attached to another. This can result in chromosomes with two centromeres and chromosomes lacking centromeres altogether; both processing of processing of 5˜ endsDNA