Homologous recombination

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Figure 1. During meiosis, homologous recombination can produce new combinations of genes as shown here between similar but not identical copies of human chromosome 1

Homologous recombination is a type of genetic recombination in which nucleotide sequences are exchanged between two similar or identical molecules of DNA. It is most widely used by cells to accurately repair harmful breaks that occur on both strands of DNA, known as double-strand breaks (DSB). Homologous recombination also produces new combinations of DNA sequences during meiosis, the process by which eukaryotes make gamete cells, like sperm and egg cells in animals. These new combinations of DNA represent genetic variation in offspring, which in turn enables populations to adapt during the course of evolution. Homologous recombination is also used in horizontal gene transfer to exchange genetic material between different strains and species of bacteria and viruses.

Although homologous recombination varies widely among different organisms and cell types, most forms involve the same basic steps. After a double-strand break occurs, sections of DNA around the 5' ends of the break are cut away in a process called resection. In the strand invasion step that follows, an overhanging 3' end of the broken DNA molecule then "invades" a similar or identical DNA molecule that is not broken. After strand invasion, the further sequence of events may follow either of two main pathways discussed below; the DSBR (double-strand break repair) pathway or the SDSA (synthesis-dependent strand annealing) pathway. Homologous recombination that occurs during DNA repair tends to result in non-crossover products, in effect restoring the damaged DNA molecule as it existed before the double-strand break.

Homologous recombination is conserved across all three domains of life as well as viruses, suggesting that it is a nearly universal biological mechanism. The discovery of genes for homologous recombination in protists—a diverse group of eukaryotic microorganisms—has been interpreted as evidence that meiosis emerged early in the evolution of eukaryotes. Since their dysfunction has been strongly associated with increased susceptibility to several types of cancer, the proteins that facilitate homologous recombination are topics of active research. Homologous recombination is also used in gene targeting, a technique for introducing genetic changes into target organisms. For their development of this technique, Mario Capecchi, Martin Evans and Oliver Smithies were awarded the 2007 Nobel Prize for Physiology or Medicine; Capecchi and Smithies independently discovered applications to mouse embryonic stem cells, however the highly conserved mechanisms underlying the DSB repair model, including uniform homologous integration of transformed DNA (gene therapy), were first shown in plasmid experiments by Orr-Weaver, Szostack and Rothstein. Researching the plasmid-induced DSB, using γ-irradiation in the 1970s-1980s, led to later experiments using endonucleases (e.g. I-SceI) to cut chromosomes for genetic engineering of mammalian cells, where nonhomologous recombination is more frequent than in yeast.

History and discovery

Figure 2. An early illustration of crossing over from Thomas Hunt Morgan

In the early 1900s, William Bateson and Reginald Punnett found an exception to one of the principles of inheritance originally described by Gregor Mendel in the 1860s. In contrast to Mendel's notion that traits are independently assorted when passed from parent to child—for example that a cat's hair color and its tail length are inherited independent of each other—Bateson and Punnett showed that certain genes associated with physical traits can be inherited together, or genetically linked. In 1911, after observing that linked traits could on occasion be inherited separately, Thomas Hunt Morgan suggested that "crossovers" can occur between linked genes, where one of the linked genes physically crosses over to a different chromosome. Two decades later, Barbara McClintock and Harriet Creighton demonstrated that chromosomal crossover occurs during meiosis, the process of cell division by which sperm and egg cells are made. Within the same year as McClintock's discovery, Curt Stern showed that crossing over—later called "recombination"—could also occur in somatic cells like white blood cells and skin cells that divide through mitosis.

In 1947, the microbiologist Joshua Lederberg showed that bacteria—which had been assumed to reproduce only asexually through binary fission—are capable of genetic recombination, which is more similar to sexual reproduction. This work established E. coli as a model organism in genetics, and helped Lederberg win the 1958 Nobel Prize in Physiology or Medicine. Building on studies in fungi, in 1964 Robin Holliday proposed a model for recombination in meiosis which introduced key details of how the process can work, including the exchange of material between chromosomes through Holliday junctions. In 1983, Jack Szostak and colleagues presented a model now known as the DSBR pathway, which accounted for observations not explained by the Holliday model. During the next decade, experiments in Drosophila, budding yeast and mammalian cells led to the emergence of other models of homologous recombination, called SDSA pathways, which do not always rely on Holliday junctions.

Much of the later work identifying proteins involved in the process and determining their mechanisms has been performed by a number of individuals including James Haber, Patrick Sung, Stephen Kowalczykowski, and others.

In eukaryotes

Homologous recombination (HR) is essential to cell division in eukaryotes like plants, animals, fungi and protists. In cells that divide through mitosis, homologous recombination repairs double-strand breaks in DNA caused by ionizing radiation or DNA-damaging chemicals. Left unrepaired, these double-strand breaks can cause large-scale rearrangement of chromosomes in somatic cells, which can in turn lead to cancer.

In addition to repairing DNA, homologous recombination also helps produce genetic diversity when cells divide in meiosis to become specialized gamete cells—sperm or egg cells in animals, pollen or ovules in plants, and spores in fungi. It does so by facilitating chromosomal crossover, in which regions of similar but not identical DNA are exchanged between homologous chromosomes. This creates new, possibly beneficial combinations of genes, which can give offspring an evolutionary advantage. Chromosomal crossover often begins when a protein called Spo11 makes a targeted double-strand break in DNA. These sites are non-randomly located on the chromosomes; usually in intergenic promoter regions and preferentially in GC-rich domains These double-strand break sites often occur at recombination hotspots, regions in chromosomes that are about 1,000–2,000 base pairs in length and have high rates of recombination. The absence of a recombination hotspot between two genes on the same chromosome often means that those genes will be inherited by future generations in equal proportion. This represents linkage between the two genes greater than would be expected from genes that independently assort during meiosis.

Timing within the mitotic cell cycle

Figure 3. Homologous recombination repairs DNA before the cell enters mitosis (M phase). It occurs only during and shortly after DNA replication, during the S and G₂ phases of the cell cycle

Double-strand breaks can be repaired through homologous recombination or through non-homologous end joining (NHEJ). NHEJ is a DNA repair mechanism which, unlike homologous recombination, does not require a long homologous sequence to guide repair. Whether homologous recombination or NHEJ is used to repair double-strand breaks is largely determined by the phase of cell cycle. Homologous recombination repairs DNA before the cell enters mitosis (M phase). It occurs during and shortly after DNA replication, in the S and G₂ phases of the cell cycle, when sister chromatids are more easily available. Compared to homologous chromosomes, which are similar to another chromosome but often have different alleles, sister chromatids are an ideal template for homologous recombination because they are an identical copy of a given chromosome. In contrast to homologous recombination, NHEJ is predominant in the G₁ phase of the cell cycle, when the cell is growing but not yet ready to divide. It occurs less frequently after the G₁ phase, but maintains at least some activity throughout the cell cycle. The mechanisms that regulate homologous recombination and NHEJ throughout the cell cycle vary widely between species.

Cyclin-dependent kinases (CDKs), which modify the activity of other proteins by adding phosphate groups to (that is, phosphorylating) them, are important regulators of homologous recombination in eukaryotes. When DNA replication begins in budding yeast, the cyclin-dependent kinase Cdc28 begins homologous recombination by phosphorylating the Sae2 protein. After being so activated by the addition of a phosphate, Sae2 uses its endonuclease activity to make a clean cut near a double-strand break in DNA. This allows a three-part protein known as the MRX complex to bind to DNA, and begins a series of protein-driven reactions that exchange material between two DNA molecules.

Preliminary steps

The packaging of eukaryotic DNA into chromatin presents a barrier to all DNA-based processes that require recruitment of enzymes to their sites of action. To allow HR DNA repair, the chromatin must be remodeled. In eukaryotes, ATP dependent chromatin remodeling complexes and histone-modifying enzymes are two predominant factors employed to accomplish this remodeling process.

Chromatin relaxation occurs rapidly at the site of a DNA damage. In one of the earliest steps, the stress-activated protein kinase, c-Jun N-terminal kinase (JNK), phosphorylates SIRT6 on serine 10 in response to double-strand breaks or other DNA damage. This post-translational modification facilitates the mobilization of SIRT6 to DNA damage sites, and is required for efficient recruitment of poly (ADP-ribose) polymerase 1 (PARP1) to DNA break sites and for efficient repair of DSBs. PARP1 protein starts to appear at DNA damage sites in less than a second, with half maximum accumulation within 1.6 seconds after the damage occurs. Next the chromatin remodeler Alc1 quickly attaches to the product of PARP1 action, a poly-ADP ribose chain, and Alc1 completes arrival at the DNA damage within 10 seconds of the occurrence of the damage. About half of the maximum chromatin relaxation, presumably due to action of Alc1, occurs by 10 seconds. This then allows recruitment of the DNA repair enzyme MRE11, to initiate DNA repair, within 13 seconds.

γH2AX, the phosphorylated form of H2AX is also involved in the early steps leading to chromatin decondensation after DNA double-strand breaks. The histone variant H2AX constitutes about 10% of the H2A histones in human chromatin. γH2AX (H2AX phosphorylated on serine 139) can be detected as soon as 20 seconds after irradiation of cells (with DNA double-strand break formation), and half maximum accumulation of γH2AX occurs in one minute. The extent of chromatin with phosphorylated γH2AX is about two million base pairs at the site of a DNA double-strand break. γH2AX does not, itself, cause chromatin decondensation, but within 30 seconds of irradiation, RNF8 protein can be detected in association with γH2AX. RNF8 mediates extensive chromatin decondensation, through its subsequent interaction with CHD4, a component of the nucleosome remodeling and deacetylase complex NuRD.

After undergoing relaxation subsequent to DNA damage, followed by DNA repair, chromatin recovers to a compaction state close to its pre-damage level after about 20 min.

Models

Two primary models for how homologous recombination repairs double-strand breaks in DNA are the double-strand break repair (DSBR) pathway (sometimes called the double Holliday junction model) and the synthesis-dependent strand annealing (SDSA) pathway. The two pathways are similar in their first several steps. After a double-strand break occurs, the MRX complex (MRN complex in humans) binds to DNA on either side of the break. Next a resection, in which DNA around the 5' ends of the break is cut back, is carried out in two distinct steps. In the first step of resection, the MRX complex recruits the Sae2 protein. The two proteins then trim back the 5' ends on either side of the break to create short 3' overhangs of single-strand DNA. In the second step, 5'→3' resection is continued by the Sgs1 helicase and the Exo1 and Dna2 nucleases. As a helicase, Sgs1 "unzips" the double-strand DNA, while Exo1 and Dna2's nuclease activity allows them to cut the single-stranded DNA produced by Sgs1.

Figure 4. The DSBR and SDSA pathways follow the same initial steps, but diverge thereafter. The DSBR pathway most often results in chromosomal crossover (bottom left), while SDSA always ends with non-crossover products (bottom right)

The RPA protein, which has high affinity for single-stranded DNA, then binds the 3' overhangs. With the help of several other proteins that mediate the process, the Rad51 protein (and Dmc1, in meiosis) then forms a filament of nucleic acid and protein on the single strand of DNA coated with RPA. This nucleoprotein filament then begins searching for DNA sequences similar to that of the 3' overhang. After finding such a sequence, the single-stranded nucleoprotein filament moves into (invades) the similar or identical recipient DNA duplex in a process called strand invasion. In cells that divide through mitosis, the recipient DNA duplex is generally a sister chromatid, which is identical to the damaged DNA molecule and provides a template for repair. In meiosis, however, the recipient DNA tends to be from a similar but not necessarily identical homologous chromosome. A displacement loop (D-loop) is formed during strand invasion between the invading 3' overhang strand and the homologous chromosome. After strand invasion, a DNA polymerase extends the end of the invading 3' strand by synthesizing new DNA. This changes the D-loop to a cross-shaped structure known as a Holliday junction. Following this, more DNA synthesis occurs on the invading strand (i.e., one of the original 3' overhangs), effectively restoring the strand on the homologous chromosome that was displaced during strand invasion.

DSBR pathway

After the stages of resection, strand invasion and DNA synthesis, the DSBR and SDSA pathways become distinct. The DSBR pathway is unique in that the second 3' overhang (which was not involved in strand invasion) also forms a Holliday junction with the homologous chromosome. The double Holliday junctions are then converted into recombination products by nicking endonucleases, a type of restriction endonuclease which cuts only one DNA strand. The DSBR pathway commonly results in crossover, though it can sometimes result in non-crossover products; the ability of a broken DNA molecule to collect sequences from separated donor loci was shown in mitotic budding yeast using plasmids or endonuclease induction of chromosomal events. Because of this tendency for chromosomal crossover, the DSBR pathway is a likely model of how crossover homologous recombination occurs during meiosis.

Whether recombination in the DSBR pathway results in chromosomal crossover is determined by how the double Holliday junction is cut, or "resolved". Chromosomal crossover will occur if one Holliday junction is cut on the crossing strand and the other Holliday junction is cut on the non-crossing strand (in Figure 4, along the horizontal purple arrowheads at one Holliday junction and along the vertical orange arrowheads at the other). Alternatively, if the two Holliday junctions are cut on the crossing strands (along the horizontal purple arrowheads at both Holliday junctions in Figure 4), then chromosomes without crossover will be produced.

SDSA pathway

Homologous recombination via the SDSA pathway occurs in cells that divide through mitosis and meiosis and results in non-crossover products. In this model, the invading 3' strand is extended along the recipient DNA duplex by a DNA polymerase, and is released as the Holliday junction between the donor and recipient DNA molecules slides in a process called branch migration. The newly synthesized 3' end of the invading strand is then able to anneal to the other 3' overhang in the damaged chromosome through complementary base pairing. After the strands anneal, a small flap of DNA can sometimes remain. Any such flaps are removed, and the SDSA pathway finishes with the resealing, also known as ligation, of any remaining single-stranded gaps.

During mitosis, the major homologous recombination pathway for repairing DNA double-strand breaks appears to be the SDSA pathway (rather than the DSBR pathway). The SDSA pathway produces non-crossover recombinants (Figure 4). During meiosis non-crossover recombinants also occur frequently and these appear to arise mainly by the SDSA pathway as well. Non-crossover recombination events occurring during meiosis likely reflect instances of repair of DNA double-strand damages or other types of DNA damages.

SSA pathway

Figure 5. Recombination via the SSA pathway occurs between two repeat elements (purple) on the same DNA duplex, and results in deletions of genetic material (Click to view animated diagram in Firefox, Chrome, Safari, or Opera web browsers)

The single-strand annealing (SSA) pathway of homologous recombination repairs double-strand breaks between two repeat sequences. The SSA pathway is unique in that it does not require a separate similar or identical molecule of DNA, like the DSBR or SDSA pathways of homologous recombination. Instead, the SSA pathway only requires a single DNA duplex, and uses the repeat sequences as the identical sequences that homologous recombination needs for repair. The pathway is relatively simple in concept: after two strands of the same DNA duplex are cut back around the site of the double-strand break, the two resulting 3' overhangs then align and anneal to each other, restoring the DNA as a continuous duplex.

As DNA around the double-strand break is cut back, the single-stranded 3' overhangs being produced are coated with the RPA protein, which prevents the 3' overhangs from sticking to themselves. A protein called Rad52 then binds each of the repeat sequences on either side of the break, and aligns them to enable the two complementary repeat sequences to anneal. After annealing is complete, leftover non-homologous flaps of the 3' overhangs are cut away by a set of nucleases, known as Rad1/Rad10, which are brought to the flaps by the Saw1 and Slx4 proteins. New DNA synthesis fills in any gaps, and ligation restores the DNA duplex as two continuous strands. The DNA sequence between the repeats is always lost, as is one of the two repeats. The SSA pathway is considered mutagenic since it results in such deletions of genetic material.

BIR pathway

During DNA replication, double-strand breaks can sometimes be encountered at replication forks as DNA helicase unzips the template strand. These defects are repaired in the break-induced replication (BIR) pathway of homologous recombination. The precise molecular mechanisms of the BIR pathway remain unclear. Three proposed mechanisms have strand invasion as an initial step, but they differ in how they model the migration of the D-loop and later phases of recombination.

The BIR pathway can also help to maintain the length of telomeres (regions of DNA at the end of eukaryotic chromosomes) in the absence of (or in cooperation with) telomerase. Without working copies of the enzyme telomerase, telomeres typically shorten with each cycle of mitosis, which eventually blocks cell division and leads to senescence. In budding yeast cells where telomerase has been inactivated through mutations, two types of "survivor" cells have been observed to avoid senescence longer than expected by elongating their telomeres through BIR pathways.

Maintaining telomere length is critical for cell immortalization, a key feature of cancer. Most cancers maintain telomeres by upregulating telomerase. However, in several types of human cancer, a BIR-like pathway helps to sustain some tumors by acting as an alternative mechanism of telomere maintenance. This fact has led scientists to investigate whether such recombination-based mechanisms of telomere maintenance could thwart anti-cancer drugs like telomerase inhibitors.

In bacteria

Figure 6. Crystal structure of a RecA protein filament bound to DNA.^[53] A 3' overhang is visible to the right of center.

Homologous recombination is a major DNA repair process in bacteria. It is also important for producing genetic diversity in bacterial populations, although the process differs substantially from meiotic recombination, which repairs DNA damages and brings about diversity in eukaryotic genomes. Homologous recombination has been most studied and is best understood for Escherichia coli. Double-strand DNA breaks in bacteria are repaired by the RecBCD pathway of homologous recombination. Breaks that occur on only one of the two DNA strands, known as single-strand gaps, are thought to be repaired by the RecF pathway. Both the RecBCD and RecF pathways include a series of reactions known as branch migration, in which single DNA strands are exchanged between two intercrossed molecules of duplex DNA, and resolution, in which those two intercrossed molecules of DNA are cut apart and restored to their normal double-stranded state.

RecBCD pathway

Figure 7A. Molecular model for the RecBCD pathway of recombination. This model is based on reactions of DNA and RecBCD with ATP in excess over Mg2+ ions. Step 1: RecBCD binds to a double-stranded DNA end. Step 2: RecBCD unwinds DNA. RecD is a fast helicase on the 5’-ended strand, and RecB is a slower helicase on the 3'-ended strand (that with an arrowhead) [ref 46 in current Wiki version]. This produces two single-stranded (ss) DNA tails and one ss loop. The loop and tails enlarge as RecBCD moves along the DNA. Step 3: The two tails anneal to produce a second ss DNA loop, and both loops move and grow. Step 4: Upon reaching the Chi hotspot sequence (5' GCTGGTGG 3'; red dot) RecBCD nicks the 3’-ended strand. Further unwinding produces a long 3'-ended ss tail with Chi near its end. Step 5: RecBCD loads RecA protein onto the Chi tail. At some undetermined point, the RecBCD subunits disassemble. Step 6: The RecA-ssDNA complex invades an intact homologous duplex DNA to produce a D-loop, which can be resolved into intact, recombinant DNA in two ways. Step 7: The D-loop is cut and anneals with the gap in the first DNA to produce a Holliday junction. Resolution of the Holliday junction (cutting, swapping of strands, and ligation) at the open arrowheads by some combination of RuvABC and RecG produces two recombinants of reciprocal type. Step 8: The 3' end of the Chi tail primes DNA synthesis, from which a replication fork can be generated. Resolution of the fork at the open arrowheads produces one recombinant (non-reciprocal) DNA, one parental-type DNA, and one DNA fragment.

 

Figure 7B. Beginning of the RecBCD pathway. This model is based on reactions of DNA and RecBCD with Mg²⁺ ions in excess over ATP. Step 1: RecBCD binds to a DNA double strand break. Step 2: RecBCD initiates unwinding of the DNA duplex through ATP-dependent helicase activity. Step 3: RecBCD continues its unwinding and moves down the DNA duplex, cleaving the 3' strand much more frequently than the 5' strand. Step 4: RecBCD encounters a Chi sequence and stops digesting the 3' strand; cleavage of the 5' strand is significantly increased. Step 5: RecBCD loads RecA onto the 3' strand. Step 6: RecBCD unbinds from the DNA duplex, leaving a RecA nucleoprotein filament on the 3' tail.

The RecBCD pathway is the main recombination pathway used in many bacteria to repair double-strand breaks in DNA, and the proteins are found in a broad array of bacteria.^[58][59][60] These double-strand breaks can be caused by UV light and other radiation, as well as chemical mutagens. Double-strand breaks may also arise by DNA replication through a single-strand nick or gap. Such a situation causes what is known as a collapsed replication fork and is fixed by several pathways of homologous recombination including the RecBCD pathway.

In this pathway, a three-subunit enzyme complex called RecBCD initiates recombination by binding to a blunt or nearly blunt end of a break in double-strand DNA. After RecBCD binds the DNA end, the RecB and RecD subunits begin unzipping the DNA duplex through helicase activity. The RecB subunit also has a nuclease domain, which cuts the single strand of DNA that emerges from the unzipping process. This unzipping continues until RecBCD encounters a specific nucleotide sequence (5'-GCTGGTGG-3') known as a Chi site.

Upon encountering a Chi site, the activity of the RecBCD enzyme changes drastically. DNA unwinding pauses for a few seconds and then resumes at roughly half the initial speed. This is likely because the slower RecB helicase unwinds the DNA after Chi, rather than the faster RecD helicase, which unwinds the DNA before Chi. Recognition of the Chi site also changes the RecBCD enzyme so that it cuts the DNA strand with Chi and begins loading multiple RecA proteins onto the single-stranded DNA with the newly generated 3' end. The resulting RecA-coated nucleoprotein filament then searches out similar sequences of DNA on a homologous chromosome. The search process induces stretching of the DNA duplex, which enhances homology recognition (a mechanism termed conformational proofreading). Upon finding such a sequence, the single-stranded nucleoprotein filament moves into the homologous recipient DNA duplex in a process called strand invasion. The invading 3' overhang causes one of the strands of the recipient DNA duplex to be displaced, to form a D-loop. If the D-loop is cut, another swapping of strands forms a cross-shaped structure called a Holliday junction. Resolution of the Holliday junction by some combination of RuvABC or RecG can produce two recombinant DNA molecules with reciprocal genetic types, if the two interacting DNA molecules differ genetically. Alternatively, the invading 3’ end near Chi can prime DNA synthesis and form a replication fork. This type of resolution produces only one type of recombinant (non-reciprocal).

RecF pathway

Bacteria appear to use the RecF pathway of homologous recombination to repair single-strand gaps in DNA. When the RecBCD pathway is inactivated by mutations and additional mutations inactivate the SbcCD and ExoI nucleases, the RecF pathway can also repair DNA double-strand breaks. In the RecF pathway the RecQ helicase unwinds the DNA and the RecJ nuclease degrades the strand with a 5' end, leaving the strand with the 3' end intact. RecA protein binds to this strand and is either aided by the RecF, RecO, and RecR proteins or stabilized by them. The RecA nucleoprotein filament then searches for a homologous DNA and exchanges places with the identical or nearly identical strand in the homologous DNA.

Although the proteins and specific mechanisms involved in their initial phases differ, the two pathways are similar in that they both require single-stranded DNA with a 3' end and the RecA protein for strand invasion. The pathways are also similar in their phases of branch migration, in which the Holliday junction slides in one direction, and resolution, in which the Holliday junctions are cleaved apart by enzymes. The alternative, non-reciprocal type of resolution may also occur by either pathway.

Branch migration

Immediately after strand invasion, the Holliday junction moves along the linked DNA during the branch migration process. It is in this movement of the Holliday junction that base pairs between the two homologous DNA duplexes are exchanged. To catalyze branch migration, the RuvA protein first recognizes and binds to the Holliday junction and recruits the RuvB protein to form the RuvAB complex. Two sets of the RuvB protein, which each form a ring-shaped ATPase, are loaded onto opposite sides of the Holliday junction, where they act as twin pumps that provide the force for branch migration. Between those two rings of RuvB, two sets of the RuvA protein assemble in the center of the Holliday junction such that the DNA at the junction is sandwiched between each set of RuvA. The strands of both DNA duplexes—the "donor" and the "recipient" duplexes—are unwound on the surface of RuvA as they are guided by the protein from one duplex to the other.

Resolution

In the resolution phase of recombination, any Holliday junctions formed by the strand invasion process are cut, thereby restoring two separate DNA molecules. This cleavage is done by RuvAB complex interacting with RuvC, which together form the RuvABC complex. RuvC is an endonuclease that cuts the degenerate sequence 5'-(A/T)TT(G/C)-3'. The sequence is found frequently in DNA, about once every 64 nucleotides. Before cutting, RuvC likely gains access to the Holliday junction by displacing one of the two RuvA tetramers covering the DNA there. Recombination results in either "splice" or "patch" products, depending on how RuvC cleaves the Holliday junction. Splice products are crossover products, in which there is a rearrangement of genetic material around the site of recombination. Patch products, on the other hand, are non-crossover products in which there is no such rearrangement and there is only a "patch" of hybrid DNA in the recombination product.

Facilitating genetic transfer

Homologous recombination is an important method of integrating donor DNA into a recipient organism's genome in horizontal gene transfer, the process by which an organism incorporates foreign DNA from another organism without being the offspring of that organism. Homologous recombination requires incoming DNA to be highly similar to the recipient genome, and so horizontal gene transfer is usually limited to similar bacteria. Studies in several species of bacteria have established that there is a log-linear decrease in recombination frequency with increasing difference in sequence between host and recipient DNA.

In bacterial conjugation, where DNA is transferred between bacteria through direct cell-to-cell contact, homologous recombination helps integrate foreign DNA into the host genome via the RecBCD pathway. The RecBCD enzyme promotes recombination after DNA is converted from single-strand DNA–in which form it originally enters the bacterium–to double-strand DNA during replication. The RecBCD pathway is also essential for the final phase of transduction, a type of horizontal gene transfer in which DNA is transferred from one bacterium to another by a virus. Foreign, bacterial DNA is sometimes misincorporated in the capsid head of bacteriophage virus particles as DNA is packaged into new bacteriophages during viral replication. When these new bacteriophages infect other bacteria, DNA from the previous host bacterium is injected into the new bacterial host as double-strand DNA. The RecBCD enzyme then incorporates this double-strand DNA into the genome of the new bacterial host.

Bacterial transformation

Natural bacterial transformation involves the transfer of DNA from a donor bacterium to a recipient bacterium, where both donor and recipient are ordinarily of the same species. Transformation, unlike bacterial conjugation and transduction, depends on numerous bacterial gene products that specifically interact to perform this process. Thus transformation is clearly a bacterial adaptation for DNA transfer. In order for a bacterium to bind, take up and integrate donor DNA into its resident chromosome by homologous recombination, it must first enter a special physiological state termed competence. The RecA/Rad51/DMC1 gene family plays a central role in homologous recombination during bacterial transformation as it does during eukaryotic meiosis and mitosis. For instance, the RecA protein is essential for transformation in Bacillus subtilis and Streptococcus pneumoniae, and expression of the RecA gene is induced during the development of competence for transformation in these organisms.

As part of the transformation process, the RecA protein interacts with entering single-stranded DNA (ssDNA) to form RecA/ssDNA nucleofilaments that scan the resident chromosome for regions of homology and bring the entering ssDNA to the corresponding region, where strand exchange and homologous recombination occur. Thus the process of homologous recombination during bacterial transformation has fundamental similarities to homologous recombination during meiosis.

In viruses

Homologous recombination occurs in several groups of viruses. In DNA viruses such as herpesvirus, recombination occurs through a break-and-rejoin mechanism like in bacteria and eukaryotes. There is also evidence for recombination in some RNA viruses, specifically positive-sense ssRNA viruses like retroviruses, picornaviruses, and coronaviruses. There is controversy over whether homologous recombination occurs in negative-sense ssRNA viruses like influenza.

In RNA viruses, homologous recombination can be either precise or imprecise. In the precise type of RNA-RNA recombination, there is no difference between the two parental RNA sequences and the resulting crossover RNA region. Because of this, it is often difficult to determine the location of crossover events between two recombining RNA sequences. In imprecise RNA homologous recombination, the crossover region has some difference with the parental RNA sequences – caused by either addition, deletion, or other modification of nucleotides. The level of precision in crossover is controlled by the sequence context of the two recombining strands of RNA: sequences rich in adenine and uracil decrease crossover precision.

Homologous recombination is important in facilitating viral evolution. For example, if the genomes of two viruses with different disadvantageous mutations undergo recombination, then they may be able to regenerate a fully functional genome. Alternatively, if two similar viruses have infected the same host cell, homologous recombination can allow those two viruses to swap genes and thereby evolve more potent variations of themselves.

Homologous recombination is the proposed mechanism whereby the DNA virus human herpesvirus-6 integrates into human telomeres.

When two or more viruses, each containing lethal genomic damage, infect the same host cell, the virus genomes can often pair with each other and undergo homologous recombinational repair to produce viable progeny. This process, known as multiplicity reactivation, has been studied in several bacteriophages, including phage T4. Enzymes employed in recombinational repair in phage T4 are functionally homologous to enzymes employed in bacterial and eukaryotic recombinational repair. In particular, with regard to a gene necessary for the strand exchange reaction, a key step in homologous recombinational repair, there is functional homology from viruses to humans (i. e. uvsX in phage T4; recA in E. coli and other bacteria, and rad51 and dmc1 in yeast and other eukaryotes, including humans). Multiplicity reactivation has also been demonstrated in numerous pathogenic viruses.

Effects of dysfunction

Without proper homologous recombination, chromosomes often incorrectly align for the first phase of cell division in meiosis. This causes chromosomes to fail to properly segregate in a process called nondisjunction. In turn, nondisjunction can cause sperm and ova to have too few or too many chromosomes. Down's syndrome, which is caused by an extra copy of chromosome 21, is one of many abnormalities that result from such a failure of homologous recombination in meiosis.

Deficiencies in homologous recombination have been strongly linked to cancer formation in humans. For example, each of the cancer-related diseases Bloom's syndrome, Werner's syndrome and Rothmund-Thomson syndrome are caused by malfunctioning copies of RecQ helicase genes involved in the regulation of homologous recombination: BLM, WRN and RECQL4, respectively. In the cells of Bloom's syndrome patients, who lack a working copy of the BLM protein, there is an elevated rate of homologous recombination. Experiments in mice deficient in BLM have suggested that the mutation gives rise to cancer through a loss of heterozygosity caused by increased homologous recombination. A loss in heterozygosity refers to the loss of one of two versions—or alleles—of a gene. If one of the lost alleles helps to suppress tumors, like the gene for the retinoblastoma protein for example, then the loss of heterozygosity can lead to cancer.

Decreased rates of homologous recombination cause inefficient DNA repair, which can also lead to cancer. This is the case with BRCA1 and BRCA2, two similar tumor suppressor genes whose malfunctioning has been linked with considerably increased risk for breast and ovarian cancer. Cells missing BRCA1 and BRCA2 have a decreased rate of homologous recombination and increased sensitivity to ionizing radiation, suggesting that decreased homologous recombination leads to increased susceptibility to cancer. Because the only known function of BRCA2 is to help initiate homologous recombination, researchers have speculated that more detailed knowledge of BRCA2's role in homologous recombination may be the key to understanding the causes of breast and ovarian cancer.

Tumours with a homologous recombination deficiency (including BRCA defects) are described as HRD-positive.

Evolutionary conservation

Figure 8. Protein domains in homologous recombination-related proteins are conserved across the three main groups of life: archaea, bacteria and eukaryotes.

While the pathways can mechanistically vary, the ability of organisms to perform homologous recombination is universally conserved across all domains of life. Based on the similarity of their amino acid sequences, homologs of a number of proteins can be found in multiple domains of life indicating that they evolved a long time ago, and have since diverged from common ancestral proteins.

RecA recombinase family members are found in almost all organisms with RecA in bacteria, Rad51 and DMC1 in eukaryotes, RadA in archaea, and UvsX in T4 phage.

Related single stranded binding proteins that are important for homologous recombination, and many other processes, are also found in all domains of life.

Rad54, Mre11, Rad50, and a number of other proteins are also found in both archaea and eukaryotes.

The RecA recombinase family

The proteins of the RecA recombinase family of proteins are thought to be descended from a common ancestral recombinase. The RecA recombinase family contains RecA protein from bacteria, the Rad51 and Dmc1 proteins from eukaryotes, and RadA from archaea, and the recombinase paralog proteins. Studies modeling the evolutionary relationships between the Rad51, Dmc1 and RadA proteins indicate that they are monophyletic, or that they share a common molecular ancestor. Within this protein family, Rad51 and Dmc1 are grouped together in a separate clade from RadA. One of the reasons for grouping these three proteins together is that they all possess a modified helix-turn-helix motif, which helps the proteins bind to DNA, toward their N-terminal ends. An ancient gene duplication event of a eukaryotic RecA gene and subsequent mutation has been proposed as a likely origin of the modern RAD51 and DMC1 genes.

The proteins generally share a long conserved region known as the RecA/Rad51 domain. Within this protein domain are two sequence motifs, Walker A motif and Walker B motif. The Walker A and B motifs allow members of the RecA/Rad51 protein family to engage in ATP binding and ATP hydrolysis.

Meiosis-specific proteins

The discovery of Dmc1 in several species of Giardia, one of the earliest protists to diverge as a eukaryote, suggests that meiotic homologous recombination—and thus meiosis itself—emerged very early in eukaryotic evolution. In addition to research on Dmc1, studies on the Spo11 protein have provided information on the origins of meiotic recombination. Spo11, a type II topoisomerase, can initiate homologous recombination in meiosis by making targeted double-strand breaks in DNA. Phylogenetic trees based on the sequence of genes similar to SPO11 in animals, fungi, plants, protists and archaea have led scientists to believe that the version Spo11 currently in eukaryotes emerged in the last common ancestor of eukaryotes and archaea.

Technological applications

Gene targeting

Figure 9. As a developing embryo, this chimeric mouse had the agouti coat color gene introduced into its DNA via gene targeting. Its offspring are homozygous for the agouti gene.

Main article: Gene targeting

Many methods for introducing DNA sequences into organisms to create recombinant DNA and genetically modified organisms use the process of homologous recombination. Also called gene targeting, the method is especially common in yeast and mouse genetics. The gene targeting method in knockout mice uses mouse embryonic stem cells to deliver artificial genetic material (mostly of therapeutic interest), which represses the target gene of the mouse by the principle of homologous recombination. The mouse thereby acts as a working model to understand the effects of a specific mammalian gene. In recognition of their discovery of how homologous recombination can be used to introduce genetic modifications in mice through embryonic stem cells, Mario Capecchi, Martin Evans and Oliver Smithies were awarded the 2007 Nobel Prize for Physiology or Medicine.

Advances in gene targeting technologies which hijack the homologous recombination mechanics of cells are now leading to the development of a new wave of more accurate, isogenic human disease models. These engineered human cell models are thought to more accurately reflect the genetics of human diseases than their mouse model predecessors. This is largely because mutations of interest are introduced into endogenous genes, just as they occur in the real patients, and because they are based on human genomes rather than rat genomes. Furthermore, certain technologies enable the knock-in of a particular mutation rather than just knock-outs associated with older gene targeting technologies.

Protein engineering

Protein engineering with homologous recombination develops chimeric proteins by swapping fragments between two parental proteins. These techniques exploit the fact that recombination can introduce a high degree of sequence diversity while preserving a protein's ability to fold into its tertiary structure, or three-dimensional shape. This stands in contrast to other protein engineering techniques, like random point mutagenesis, in which the probability of maintaining protein function declines exponentially with increasing amino acid substitutions. The chimeras produced by recombination techniques are able to maintain their ability to fold because their swapped parental fragments are structurally and evolutionarily conserved. These recombinable "building blocks" preserve structurally important interactions like points of physical contact between different amino acids in the protein's structure. Computational methods like SCHEMA and statistical coupling analysis can be used to identify structural subunits suitable for recombination.

Techniques that rely on homologous recombination have been used to engineer new proteins. In a study published in 2007, researchers were able to create chimeras of two enzymes involved in the biosynthesis of isoprenoids, a diverse class of compounds including hormones, visual pigments and certain pheromones. The chimeric proteins acquired an ability to catalyze an essential reaction in isoprenoid biosynthesis—one of the most diverse pathways of biosynthesis found in nature—that was absent in the parent proteins. Protein engineering through recombination has also produced chimeric enzymes with new function in members of a group of proteins known as the cytochrome P450 family, which in humans is involved in detoxifying foreign compounds like drugs, food additives and preservatives.

Cancer therapy

Cancer cells with BRCA mutations have deficiencies in homologous recombination, and drugs to exploit those deficiencies have been developed and used successfully in clinical trials. Olaparib, a PARP1 inhibitor, shrunk or stopped the growth of tumors from breast, ovarian and prostate cancers caused by mutations in the BRCA1 or BRCA2 genes, which are necessary for HR. When BRCA1 or BRCA2 is absent, other types of DNA repair mechanisms must compensate for the deficiency of HR, such as base-excision repair (BER) for stalled replication forks or non-homologous end joining (NHEJ) for double strand breaks. By inhibiting BER in an HR-deficient cell, olaparib applies the concept of synthetic lethality to specifically target cancer cells. While PARP1 inhibitors represent a novel approach to cancer therapy, researchers have cautioned that they may prove insufficient for treating late-stage metastatic cancers. Cancer cells can become resistant to a PARP1 inhibitor if they undergo deletions of mutations in BRCA2, undermining the drug's synthetic lethality by restoring cancer cells' ability to repair DNA by HR.

DNA damage (naturally occurring)

From Wikipedia, the free encyclopedia

DNA damage is distinctly different from mutation, although both are types of error in DNA. DNA damage is an abnormal chemical structure in DNA, while a mutation is a change in the sequence of standard base pairs. DNA damages cause changes in the structure of the genetic material and prevents the replication mechanism from functioning and performing properly.

 

DNA damage and mutation have different biological consequences. While most DNA damages can undergo DNA repair, such repair is not 100% efficient. Un-repaired DNA damages accumulate in non-replicating cells, such as cells in the brains or muscles of adult mammals and can cause aging. In replicating cells, such as cells lining the colon, errors occur upon replication past damages in the template strand of DNA or during repair of DNA damages. These errors can give rise to mutations or epigenetic alterations. Both of these types of alteration can be replicated and passed on to subsequent cell generations. These alterations can change gene function or regulation of gene expression and possibly contribute to progression to cancer.

Throughout the cell cycle there are various checkpoints to ensure the cell is in good condition to progress to mitosis. The three main checkpoints are at G1/s, G2/m, and at the spindle assembly checkpoint regulating progression through anaphase. G1 and G2 checkpoints involve scanning for damaged DNA. During S phase the cell is more vulnerable to DNA damage than any other part of the cell cycle. G2 checkpoint checks for damaged DNA and DNA replication completeness. DNA damage is an alteration in the chemical structure of DNA, such as a break in a strand of DNA, a base missing from the backbone of DNA, or a chemically changed base as 8-OHdG. DNA damage can occur naturally or via environmental factors. The DNA damage response (DDR) is a complex signal transduction pathway which recognizes when DNA is damaged and initiates the cellular response to the damage.

Types

Damage to DNA that occurs naturally can result from metabolic or hydrolytic processes. Metabolism releases compounds that damage DNA including reactive oxygen species, reactive nitrogen species, reactive carbonyl species, lipid peroxidation products and alkylating agents, among others, while hydrolysis cleaves chemical bonds in DNA. Naturally occurring oxidative DNA damages arise at least 10,000 times per cell per day in humans and 50,000 times or more per cell per day in rats, as documented below.

Oxidative DNA damage can produce more than 20 types of altered bases as well as single strand breaks.

Other types of endogeneous DNA damages, given below with their frequencies of occurrence, include depurinations, depyrimidinations, double-strand breaks, O6-methylguanines and cytosine deamination.

DNA can be damaged via environmental factors as well. Environmental agents such as UV light, ionizing radiation, and genotoxic chemicals. Replication forks can be stalled due to damaged DNA and double strand breaks are also a form of DNA damage.

Frequencies

The list below, from reference, shows some frequencies with which new naturally occurring DNA damages arise per day, due to endogenous cellular processes.

• Oxidative damages
  • Humans, per cell per day
    • 10,000
      11,500
      2,800 specific damages 8-oxoGua, 8-oxodG plus 5-HMUra
      2,800 specific damages 8-oxoGua, 8-oxodG plus 5-HMUra
  • Rats, per cell per day
    • 74,000
      86,000
      100,000
  • Mice, per cell per day
    • 34,000 specific damages 8-oxoGua, 8-oxodG plus 5-HMUra
      47,000 specific damages oxo8dG in mouse liver
      28,000 specific damages 8-oxoGua, 8-oxodG, 5-HMUra
• Depurinations
  • Mammalian cells, per cell per day
    • 2,000 to 10,000
      9,000
      12,000
      13,920
• Depyrimidinations
  • Mammalian cells, per cell per day
    • 600
      696
• Single-strand breaks
  • Mammalian cells, per cell per day
    • 55,200
• Double-strand breaks
  • Human cells, per cell cycle
    • 10
      50
• O6-methylguanines
  • Mammalian cells, per cell per day
    • 3,120
• Cytosine deamination
  • Mammalian cells, per cell per day
    • 192

Another important endogenous DNA damage is M1dG, short for (3-(2'-deoxy-beta-D-erythro-pentofuranosyl)-pyrimido[1,2-a]-purin-10(3H)-one). The excretion in urine (likely reflecting rate of occurrence) of M1dG may be as much as 1,000-fold lower than that of 8-oxodG. However, a more important measure may be the steady-state level in DNA, reflecting both rate of occurrence and rate of DNA repair. The steady-state level of M1dG is higher than that of 8-oxodG. This points out that some DNA damages produced at a low rate may be difficult to repair and remain in DNA at a high steady-state level. Both M1dG and 8-oxodG are mutagenic.

Steady-state levels

Steady-state levels of DNA damages represent the balance between formation and repair. More than 100 types of oxidative DNA damage have been characterized, and 8-oxodG constitutes about 5% of the steady state oxidative damages in DNA. Helbock et al. estimated that there were 24,000 steady state oxidative DNA adducts per cell in young rats and 66,000 adducts per cell in old rats. This reflects the accumulation of DNA damage with age. DNA damage accumulation with age is further described in DNA damage theory of aging.

Swenberg et al. measured average amounts of selected steady state endogenous DNA damages in mammalian cells. The seven most common damages they evaluated are shown in Table 1.

Table 1. Steady-state amounts of endogenous DNA damages

Endogenous lesions	Number per cell
Abasic sites	30,000
N7-(2-hydroxethyl)guanine (7HEG)	3,000
8-hydroxyguanine	2,400
7-(2-oxoethyl)guanine	1,500
Formaldehyde adducts	960
Acrolein-deoxyguanine	120
Malondialdehyde-deoxyguanine	60

Evaluating steady-state damages in specific tissues of the rat, Nakamura and Swenberg indicated that the number of abasic sites varied from about 50,000 per cell in liver, kidney and lung to about 200,000 per cell in the brain.

Repair of damaged DNA

In the presence of DNA damage, the cell can either repair the damage or induce cell death if the damage is beyond repair.

Types

The seven main types of DNA repair and one pathway of damage tolerance, the lesions they address, and the accuracy of the repair (or tolerance) are shown in this table. For a brief description of the steps in repair see DNA repair mechanisms or see each individual pathway.

Major pathways of DNA repair and one tolerance mechanism

Repair pathway	Lesions	Accuracy
Base excision repair	corrects DNA damage from oxidation, deamination and alkylation, also single-strand breaks	accurate
Nucleotide excision repair	oxidative endogenous lesions such as cyclopurine, sunlight induced thymine dimers (cyclobutane dimers and pyrimidine (6-4) pyrimidone photoproducts)	accurate
Homologous recombinational repair	double-strand breaks in the mid-S phase or mid-G2 phase of the cell cycle	accurate
Non-homologous end joining	double-strand breaks if cells are in the G0 phase. the G1 phase or the G2 phase of the cell cycle	somewhat inaccurate
Microhomology-mediated end joining or alt-End joining	double-strand breaks in the S phase of the cell cycle	always inaccurate
DNA mismatch repair	base substitution mismatches and insertion-deletion mismatches generated during DNA replication	accurate
Direct reversal (MGMT and AlkB)	6-O-methylguanine is reversed to guanine by MGMT, some other methylated bases are demethylated by AlkB	accurate
Translesion synthesis	DNA damage tolerance process that allows the DNA replication machinery to replicate past DNA lesions	may be inaccurate

Aging and cancer

 

DNA damage in non-replicating cells, if not repaired and

accumulated can lead to aging. DNA damage in replicating

cells, if not repaired can lead to either apoptosis or to cancer.

The schematic diagram indicates the roles of insufficient DNA repair in aging and cancer, and the role of apoptosis in cancer prevention. An excess of naturally occurring DNA damage, due to inherited deficiencies in particular DNA repair enzymes, can cause premature aging or increased risk for cancer. On the other hand, the ability to trigger apoptosis in the presence of excess un-repaired DNA damage is critical for prevention of cancer.

Apoptosis and cancer prevention

DNA repair proteins are often activated or induced when DNA has sustained damage. However, excessive DNA damage can initiate apoptosis (i.e., programmed cell death) if the level of DNA damage exceeds the repair capacity. Apoptosis can prevent cells with excess DNA damage from undergoing mutagenesis and progression to cancer.

Inflammation is often caused by infection, such as with hepatitis B virus (HBV), hepatitis C virus (HCV) or Helicobacter pylori). Chronic inflammation is also a central characteristic of obesity. Such inflammation causes oxidative DNA damage. This is due to the induction of reactive oxygen species (ROS) by various intracellular inflammatory mediators. HBV and HCV infections, in particular, cause 10,000-fold and 100,000-fold increases in intracellular ROS production, respectively. Inflammation-induced ROS that cause DNA damage can trigger apoptosis, but may also cause cancer if repair and apoptotic processes are insufficiently protective.

Bile acids, stored in the gall bladder, are released into the small intestine in response to fat in the diet. Higher levels of fat cause greater release. Bile acids cause DNA damage, including oxidative DNA damage, double-strand DNA breaks, aneuploidy and chromosome breakage. High-normal levels of the bile acid deoxycholic acid cause apoptosis in human colon cells, but may also lead to colon cancer if repair and apoptotic defenses are insufficient.

Apoptosis serves as a safeguard mechanism against tumorigenesis. It prevents the increased mutagenesis that excess DNA damage could cause, upon replication.

At least 17 DNA repair proteins, distributed among five DNA repair pathways, have a "dual role" in response to DNA damage. With moderate levels of DNA damage, these proteins initiate or contribute to DNA repair. However, when excessive levels of DNA damage are present, they trigger apoptosis.

DNA damage response

The packaging of eukaryotic DNA into chromatin is a barrier to all DNA-based processes that require enzyme action. For most DNA repair processes the chromatin must be remodeled. In eukaryotes, ATP dependent chromatin remodeling complexes and histone-modifying enzymes are two factors that act to accomplish this remodeling process after DNA damage occurs. Further DNA repair steps, involving multiple enzymes, usually follow. Some of the first responses to DNA damage, with their timing, are described below. More complete descriptions of the DNA repair pathways are presented in articles describing each pathway. At least 169 enzymes are involved in DNA repair pathways.

Base excision repair

Oxidized bases in DNA are produced in cells treated with Hoechst dye followed by micro-irradiation with 405 nm light. Such oxidized bases can be repaired by base excision repair.

When the 405 nm light is focused along a narrow line within the nucleus of a cell, about 2.5 seconds after irradiation the chromatin remodeling enzyme Alc1 achieves half-maximum recruitment onto the irradiated micro-line. The line of chromatin that was irradiated then relaxes, expanding side-to-side over the next 60 seconds.

Within 6 seconds of the irradiation with 405 nm light there is half-maximum recruitment of OGG1 to the irradiated line. OGG1 is an enzyme that removes the oxidative DNA damage 8-oxo-dG from DNA. Removal of 8-oxo-dG, during base excision repair, occurs with a half-life of 11 minutes.

Nucleotide excision repair

Ultraviolet (UV) light induces the formation of DNA damages including pyrimidine dimers (such as thymine dimers) and 6,4 photoproducts. These types of "bulky" damages are repaired by nucleotide excision repair.

After irradiation with UV light, DDB2, in a complex with DDB1, the ubiquitin ligase protein CUL4A and the RING finger protein ROC1, associates with sites of damage within chromatin. Half-maximum association occurs in 40 seconds. PARP1 also associates within this period. The PARP1 protein attaches to both DDB1 and DDB2 and then PARylates (creates a poly-ADP ribose chain) on DDB2 that attracts the DNA remodeling protein ALC1. ALC1 relaxes chromatin at sites of UV damage to DNA. In addition, the ubiquitin E3 ligase complex DDB1-CUL4A carries out ubiquitination of the core histones H2A, H3, and H4 as well as the repair protein XPC which has been attracted to the site of the DNA damage. XPC, upon ubiquitination, is activated and initiates the nucleotide excision repair pathway. Somewhat later, at 30 minutes after UV damage, the INO80 chromatin remodeling complex is recruited to the site of the DNA damage, and this coincides with the binding of further nucleotide excision repair proteins, including ERCC1.

Homologous recombinational repair

Double-strand breaks (DSBs) at specific sites can be induced by transfecting cells with a plasmid encoding I-SceI endonuclease (a homing endonuclease). Multiple DSBs can be induced by irradiating sensitized cells (labeled with 5'-bromo-2'-deoxyuridine and with Hoechst dye) with 780 nm light. These DSBs can be repaired by the accurate homologous recombinational repair or by the less accurate non-homologous end joining repair pathway. Here we describe the early steps in homologus recombinational repair (HRR).

After treating cells to introduce DSBs, the stress-activated protein kinase, c-Jun N-terminal kinase (JNK), phosphorylates SIRT6 on serine 10. This post-translational modification facilitates the mobilization of SIRT6 to DNA damage sites with half-maximum recruitment in well under a second. SIRT6 at the site is required for efficient recruitment of poly (ADP-ribose) polymerase 1 (PARP1) to a DNA break site and for efficient repair of DSBs. PARP1 protein starts to appear at DSBs in less than a second, with half maximum accumulation within 1.6 seconds after the damage occurs. This then allows half maximum recruitment of the DNA repair enzymes MRE11 within 13 seconds and NBS1 within 28 seconds. MRE11 and NBS1 carry out early steps of the HRR pathway.

γH2AX, the phosphorylated form of H2AX is also involved in early steps of DSB repair. The histone variant H2AX constitutes about 10% of the H2A histones in human chromatin. γH2AX (H2AX phosphorylated on serine 139) can be detected as soon as 20 seconds after irradiation of cells (with DNA double-strand break formation), and half maximum accumulation of γH2AX occurs in one minute. The extent of chromatin with phosphorylated γH2AX is about two million base pairs at the site of a DNA double-strand break.^[73] γH2AX does not, itself, cause chromatin decondensation, but within 30 seconds of irradiation, RNF8 protein can be detected in association with γH2AX. RNF8 mediates extensive chromatin decondensation, through its subsequent interaction with CHD4, a component of the nucleosome remodeling and deacetylase complex NuRD.

Pause for DNA repair

After rapid chromatin remodeling, cell cycle checkpoints may be activated to allow DNA repair to be completed before the cell cycle progresses. First, two kinases, ATM and ATR are activated within 5 or 6 minutes after DNA is damaged. This is followed by phosphorylation of the cell cycle checkpoint protein Chk1, initiating its function, about 10 minutes after DNA is damaged.

Role of ATR and ATM

Most damage can be repaired without triggering the damage response system, however more complex damage activates ATR and ATM, key protein kinases in the damage response system. DNA damage inhibits M-CDKs which are a key component of progression into Mitosis.

In all eukaryotic cells, ATR and ATM are protein kinases that detect DNA damage. They bind to DNA damaged sites and activate Chk1, Chk2, and, in animal cells, p53. Together, these proteins make up the DNA damage response system. Some DNA damage does not require the recruitment of ATR and ATM, it is only difficult and extensive damage that requires ATR and ATM. ATM and ATR are required for NHEJ, HR, ICL repair, and NER, as well as replication fork stability during unperturbed DNA replication and in response to replication blocks.

ATR is recruited for different forms of damage such as nucleotide damage, stalled replication forks and double strand breaks. ATM is specifically for the damage response to double strand breaks. The MRN complex (composed of Mre11, Rad50, and Nbs1) form immediately at the site of double strand break. This MRN complex recruits ATM to the site of damage. ATR and ATM phosphorylate various proteins that contribute to the damage repair system. The binding of ATR and ATM to damage sites on DNA lead to the recruitment of Chk1 and Chk2. These protein kinases send damage signals to the cell cycle control system to delay the progression of the cell cycle.

Chk1 and Chk2 functions

Chk1 leads to the production of DNA repair enzymes. Chk2 leads to reversible cell cycle arrest. Chk2 as well as ATR/ATM can activate p53 which leads to permanent cell cycle arrest or apoptosis.

p53 role in DNA damage repair system

When there is too much damage, apoptosis is triggered in order to protect the organism from potentially harmful cells.7 p53, also known as a tumor suppressor gene, is a major regulatory protein in the DNA damage response system which binds directly to the promoters of its target genes. p53 acts primarily at the G1 checkpoint (controlling the G1 to S transition), where it blocks cell cycle progression. Activation of p53 can trigger cell death or permanent cell cycle arrest. p53 can also activate certain repair pathways such was NER.

Regulation of p53

In the absence of DNA damage, p53 is regulated by Mdm2 and constantly degraded. When there is DNA damage, Mdm2 is phosphorylated, most likely caused by ATM. The phosphorylation of Mdm2 leads to a reduction in the activity of Mdm2, thus preventing the degradation of p53. Normal, undamaged cell, usually has low levels of p53 while cells under stress and DNA damage, will have high levels of p53.

p53 serves as transcription factor for bax and p21

p53 serves as a transcription factors for both bax, a proapoptotic protein as well as p21, a CDK inhibitor. CDK Inhibitors result in cell cycle arrest. Arresting the cell provides the cell time to repair the damage, and if the damage is irreparable, p53 recruits bax to trigger apoptosis.

DDR and p53 role in cancer

p53 is a major key player in the growth of cancerous cells. Damaged DNA cells with mutated p53 are at a higher risk of becoming cancerous. Common chemotherapy treatments are genotoxic. These treatments are ineffective in cancer tumor that have mutated p53 since they do not have a functioning p53 to either arrest or kill the damaged cell.

A major problem for life

One indication that DNA damages are a major problem for life is that DNA repair processes, to cope with DNA damages, have been found in all cellular organisms in which DNA repair has been investigated. For example, in bacteria, a regulatory network aimed at repairing DNA damages (called the SOS response in Escherichia coli) has been found in many bacterial species. E. coli RecA, a key enzyme in the SOS response pathway, is the defining member of a ubiquitous class of DNA strand-exchange proteins that are essential for homologous recombination, a pathway that maintains genomic integrity by repairing broken DNA. Genes homologous to RecA and to other central genes in the SOS response pathway are found in almost all the bacterial genomes sequenced to date, covering a large number of phyla, suggesting both an ancient origin and a widespread occurrence of recombinational repair of DNA damage. Eukaryotic recombinases that are homologues of RecA are also widespread in eukaryotic organisms. For example, in fission yeast and humans, RecA homologues promote duplex-duplex DNA-strand exchange needed for repair of many types of DNA lesions.

Another indication that DNA damages are a major problem for life is that cells make large investments in DNA repair processes. As pointed out by Hoeijmakers, repairing just one double-strand break could require more than 10,000 ATP molecules, as used in signaling the presence of the damage, the generation of repair foci, and the formation (in humans) of the RAD51 nucleofilament (an intermediate in homologous recombinational repair). (RAD51 is a homologue of bacterial RecA.) If the structural modification occurs during the G1 phase of DNA replication, the G1-S checkpoint arrests or postpones the furtherance of the cell cycle before the product enters the S phase.

Consequences

Differentiated somatic cells of adult mammals generally replicate infrequently or not at all. Such cells, including, for example, brain neurons and muscle myocytes, have little or no cell turnover. Non-replicating cells do not generally generate mutations due to DNA damage-induced errors of replication. These non-replicating cells do not commonly give rise to cancer, but they do accumulate DNA damages with time that likely contribute to aging. In a non-replicating cell, a single-strand break or other type of damage in the transcribed strand of DNA can block RNA polymerase II catalysed transcription. This would interfere with the synthesis of the protein coded for by the gene in which the blockage occurred.

Brasnjevic et al. summarized the evidence showing that single-strand breaks accumulate with age in the brain (though accumulation differed in different regions of the brain) and that single-strand breaks are the most frequent steady-state DNA damages in the brain. As discussed above, these accumulated single-strand breaks would be expected to block transcription of genes. Consistent with this, as reviewed by Hetman et al., 182 genes were identified and shown to have reduced transcription in the brains of individuals older than 72 years, compared to transcription in the brains of those less than 43 years old. When 40 particular proteins were evaluated in a muscle of rats, the majority of the proteins showed significant decreases during aging from 18 months (mature rat) to 30 months (aged rat) of age.

Another type of DNA damage, the double strand break, was shown to cause cell death (loss of cells) through apoptosis. This type of DNA damage would not accumulate with age, since once a cell was lost through apoptosis, its double strand damage would be lost with it. Thus, damaged DNA segments undermine the DNA replication machinery because these altered sequences of DNA cannot be utilized as true templates to produce copies of one's genetic material.

RAD genes and the cell cycle response to DNA damage in Saccharomyces cerevisiae

When DNA is damaged, the cell responds in various ways to fix the damage and minimize the effects on the cell. One such response, specifically in eukaryotic cells, is to delay cell division—the cell becomes arrested for some time in the G2 phase before progressing through the rest of the cell cycle. Various studies have been conducted to elucidate the purpose of this G2 arrest that is induced by DNA damage. Researchers have found that cells that are prematurely forced out of the delay have lower cell viability and higher rates of damaged chromosomes compared with cells that are able to undergo a full G2 arrest, suggesting that the purpose of the delay is to give the cell time to repair damaged chromosomes before continuing with the cell cycle. This ensures the proper functioning of mitosis.

Various species of animals exhibit similar mechanisms of cellular delay in response to DNA damage, which can be caused by exposure to x-irradiation. The budding yeast Saccharomyces cerevisiae has specifically been studied because progression through the cell cycle can be followed via nuclear morphology with ease. By studying Saccharomyces cerevisiae, researchers have been able to learn more about radiation-sensitive (RAD) genes, and the effect that RAD mutations may have on the typical cellular DNA damaged-induced delay response. Specifically, the RAD9 gene plays a crucial role in detecting DNA damage and arresting the cell in G2 until the damage is repaired.

Through extensive experiments, researchers have been able to illuminate the role that the RAD genes play in delaying cell division in response to DNA damage. When wild-type, growing cells are exposed to various levels of x-irradiation over a given time frame, and then analyzed with a microcolony assay, differences in the cell cycle response can be observed based on which genes are mutated in the cells. For instance, while unirradiated cells will progress normally through the cell cycle, cells that are exposed to x-irradiation either permanently arrest (become inviable) or delay in the G2 phase before continuing to divide in mitosis, further corroborating the idea that the G2 delay is crucial for DNA repair. However, rad strains, which are deficient in DNA repair, exhibit a markedly different response. For instance, rad52 cells, which cannot repair double-stranded DNA breaks, tend to permanently arrest in G2 when exposed to even very low levels of x-irradiation, and rarely end up progressing through the later stages of the cell cycle. This is because the cells cannot repair DNA damage and thus do not enter mitosis. Various other rad mutants exhibit similar responses when exposed to x-irradiation.

However, the rad9 strain exhibits an entirely different effect. These cells fail to delay in the G2 phase when exposed to x-irradiation, and end up progressing through the cell cycle unperturbed, before dying. This suggests that the RAD9 gene, unlike the other RAD genes, plays a crucial role in initiating G2 arrest. To further investigate these findings, the cell cycles of double mutant strains have been analyzed. A mutant rad52 rad9 strain—which is both defective in DNA repair and G2 arrest—fails to undergo cell cycle arrest when exposed to x-irradiation. This suggests that even if DNA damage cannot be repaired, if RAD9 is not present, the cell cycle will not delay. Thus, unrepaired DNA damage is the signal that tells RAD9 to halt division and arrest the cell cycle in G2. Furthermore, there is a dose-dependent response; as the levels of x-irradiation—and subsequent DNA damage—increase, more cells, regardless of the mutations they have, become arrested in G2.

Another, and perhaps more helpful way to visualize this effect is to look at photomicroscopy slides. Initially, slides of RAD+ and rad9 haploid cells in the exponential phase of growth show simple, single cells, that are indistinguishable from each other. However, the slides look much different after being exposed to x-irradiation for 10 hours. The RAD+ slides now show RAD+ cells existing primarily as two-budded microcolonies, suggesting that cell division has been arrested. In contrast, the rad9 slides show the rad9 cells existing primarily as 3 to 8 budded colonies, and they appear smaller than the RAD+ cells. This is further evidence that the mutant RAD cells continued to divide and are deficient in G2 arrest.

However, there is evidence that although the RAD9 gene is necessary to induce G2 arrest in response to DNA damage, giving the cell time to repair the damage, it does not actually play a direct role in repairing DNA. When rad9 cells are artificially arrested in G2 with MBC, a microtubule poison that prevents cellular division, and then treated with x-irradiation, the cells are able to repair their DNA and eventually progress through the cell cycle, dividing into viable cells. Thus, the RAD9 gene plays no role in actually repairing damaged DNA—it simply senses damaged DNA and responds by delaying cell division. The delay, then, is mediated by a control mechanism, rather than the physical damaged DNA.

On the other hand, it is possible that there are backup mechanisms that fill the role of RAD9 when it is not present. In fact, some studies have found that RAD9 does indeed play a critical role in DNA repair. In one study, rad9 mutant and normal cells in the exponential phase of growth were exposed to UV-irradiation and synchronized in specific phases of the cell cycle. After being incubated to permit DNA repair, the extent of pyrimidine dimerization (which is indicative of DNA damage) was assessed using sensitive primer extension techniques. It was found that the removal of DNA photolesions was much less efficient in rad9 mutant cells than normal cells, providing evidence that RAD9 is involved in DNA repair. Thus, the role of RAD9 in repairing DNA damage remains unclear.

Regardless, it is clear that RAD9 is necessary to sense DNA damage and halt cell division. RAD9 has been suggested to possess 3’ to 5’ exonuclease activity, which is perhaps why it plays a role in detecting DNA damage. When DNA is damaged, it is hypothesized that RAD9 forms a complex with RAD1 and HUS1, and this complex is recruited to sites of DNA damage. It is in this way that RAD9 is able to exert its effects.

Although the function of RAD9 has primarily been studied in the budding yeast Saccharomyces cerevisiae, many of the cell cycle control mechanisms are similar between species. Thus, we can conclude that RAD9 likely plays a critical role in the DNA damage response in humans as well.