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. [ citation needed ]


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. [ 103 ]


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. [ 104 ]


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. [ citation needed ]


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. [ citation needed ]


DNA mismatch repair  ( MMR ) is a system for recognizing and repairing erroneous insertion, deletion, and mis-incorporation of  bases  that can arise during  DNA replication  and  recombination , as well as  repairing  some forms of  DNA damage . [ 1 ] [ 2 ]


Mismatch repair is strand-specific. During DNA synthesis the newly synthesised (daughter) strand will commonly include errors.  In order to begin repair, the mismatch repair machinery distinguishes the newly synthesised strand from the template (parental). In gram-negative bacteria, transient  hemimethylation  distinguishes the strands (the parental is  methylated  and daughter is not). However, in other prokaryotes and eukaryotes, the exact mechanism is not clear.  It is suspected that, in eukaryotes, newly synthesized lagging-strand DNA transiently contains  nicks  (before being sealed by DNA ligase) and provides a signal that directs mismatch proofreading systems to the appropriate strand. This implies that these nicks must be present in the leading strand, and evidence for this has recently been found. [ 3 ]
Recent work [ 4 ]  has shown that nicks are sites for RFC-dependent loading of the replication sliding clamp,  proliferating cell nuclear antigen  (PCNA), in an orientation-specific manner, such that one face of the donut-shape protein is juxtaposed toward the 3'-OH end at the nick.  Loaded PCNA then directs the action of the MutLalpha endonuclease  [ 5 ]  to the daughter strand in the presence of a mismatch and MutSalpha or MutSbeta.


Any mutational event that disrupts the  superhelical structure  of  DNA  carries with it the potential to compromise the genetic stability of a cell. The fact that the damage detection and repair systems are as complex as the replication machinery itself highlights the importance evolution has attached to DNA fidelity.


Examples of mismatched bases include a G/T or A/C pairing (see  DNA repair ).  Mismatches are commonly due to  tautomerization  of bases during DNA replication. The damage is repaired by recognition of the deformity caused by the mismatch, determining the template and non-template strand, and excising the wrongly incorporated base and replacing it with the correct  nucleotide . The removal process involves more than just the mismatched nucleotide itself.  A few or up to thousands of base pairs of the newly synthesized DNA strand can be removed.


Mismatch repair is a highly conserved process from  prokaryotes  to  eukaryotes . The first evidence for mismatch repair was obtained from  S. pneumoniae  (the hexA and hexB  genes ). Subsequent work on  E. coli  has identified a number of genes that, when  mutationally  inactivated, cause hypermutable strains. The gene products are, therefore, called the "Mut" proteins, and are the major active components of the mismatch repair system. Three of these proteins are essential in detecting the mismatch and directing repair machinery to it:  MutS , MutH and MutL (MutS is a homologue of HexA and MutL of HexB).


MutS forms a dimer (MutS 2 ) that recognises the mismatched base on the daughter strand and binds the mutated DNA. MutH binds at hemimethylated sites along the daughter DNA, but its action is latent, being activated only upon contact by a MutL dimer (MutL 2 ), which binds the MutS-DNA complex and acts as a mediator between MutS 2  and MutH, activating the latter. The DNA is looped out to search for the nearest d(GATC) methylation site to the mismatch, which could be up to 1 kb away. Upon activation by the MutS-DNA complex, MutH nicks the daughter strand near the hemimethylated site. MutL recruits  UvrD   helicase  (DNA Helicase II) to separate the two strands with a specific 3' to 5' polarity. The entire MutSHL complex then slides along the DNA in the direction of the mismatch, liberating the strand to be excised as it goes. An exonuclease trails the complex and digests the ss-DNA tail. The exonuclease recruited is dependent on which side of the mismatch MutH incises the strand – 5' or 3'. If the nick made by MutH is on the 5' end of the mismatch, either RecJ or ExoVII (both 5' to 3' exonucleases) is used. If, however, the nick is on the 3' end of the mismatch,  ExoI  (a 3' to 5' enzyme) is used.


The entire process ends past the mismatch site - i.e., both the site itself and its surrounding nucleotides are fully excised. The single-strand gap created by the exonuclease can then be repaired by DNA Polymerase III (assisted by single-strand-binding protein), which uses the other strand as a template, and finally sealed by DNA ligase. DNA methylase then rapidly methylates the daughter strand.


When bound, the MutS 2  dimer bends the DNA helix and shields approximately 20 base pairs. It has weak ATPase activity, and binding of  ATP  leads to the formation of tertiary structures on the surface of the molecule. The  crystal structure  of MutS reveals that it is exceptionally asymmetric, and, while its active conformation is a dimer, only one of the two halves interacts with the mismatch site.


In eukaryotes,  M ut S   h omologs form two major heterodimers:  Msh2 /Msh6 (MutSα) and  Msh2 /Msh3 (MutSβ). The MutSα pathway is involved primarily in base substitution and small-loop mismatch repair. The MutSβ pathway is also involved in small-loop repair, in addition to large-loop (~10 nucleotide loops) repair. However, MutSβ does not repair base substitutions.


MutL also has weak ATPase activity (it uses ATP for purposes of movement). It forms a complex with MutS and MutH, increasing the MutS footprint on the DNA.


However, the processivity (the distance the enzyme can move along the DNA before dissociating) of UvrD is only ~40–50 bp. Because the distance between the nick created by MutH and the mismatch can average ~600 bp, if there is not another UvrD loaded the unwound section is then free to re-anneal to its complementary strand, forcing the process to start over. However, when assisted by MutL, the  rate  of UvrD loading is greatly increased. While the processivity (and ATP utilisation) of the individual UvrD molecules remains the same, the total effect on the DNA is boosted considerably; the DNA has no chance to re-anneal, as each UvrD unwinds 40-50 bp of DNA, dissociates, and then is immediately replaced by another UvrD, repeating the process. This exposes large sections of DNA to  exonuclease  digestion, allowing for quick excision (and later replacement) of the incorrect DNA.


Eukaryotes have five  M ut L   h omologs designated as MLH1, MLH2, MLH3, PMS1, and PMS2. They form heterodimers that mimic MutL in  E. coli . The human homologs of prokaryotic MutL form three complexes referred to as MutLα, MutLβ, and MutLγ. The MutLα complex is made of MLH1 and PMS2 subunits, the MutLβ heterodimer is made of MLH1 and PMS1, whereas MutLγ is made of MLH1 and MLH3. MutLα acts as an endonuclease that introduces strand breaks in the daughter strand upon activation by mismatch and other required proteins, MutSα and PCNA. These strand interruptions serve as entry points for an exonuclease activity that removes mismatched DNA. Roles played by MutLβ and MutLγ in mismatch repair are less-understood.


MutH is a very weak  endonuclease  that is activated once bound to MutL (which itself is bound to MutS). It nicks  unmethylated  DNA and the unmethylated strand of hemimethylated DNA but does not nick fully methylated DNA. Experiments have shown that mismatch repair is random if neither strand is methylated. [ citation needed ]  These behaviours led to the proposal that MutH determines which strand contains the mismatch.
MutH has no eukaryotic homolog. Its endonuclease function is taken up by MutL homologs, which have some specialized 5'-3' exonuclease activity. The strand bias for removing mismatches from the newly synthesized daughter strand in eukaryotes may be provided by the free 3' ends of  Okazaki fragments  in the new strand created during replication.


PCNA  and the β-sliding clamp associate with MutSα/β and MutL, respectively.  Although initial reports suggested that the PCNA-MutSα complex may enhance mismatch recognition, [ 6 ]  it has been recently demonstrated [ 7 ]  that there is no apparent change in affinity of MutSα for a mismatch in the presence or absence of PCNA.  Furthermore, mutants of MutSα that are unable to interact with PCNA  in vitro  exhibit the capacity to carry out mismatch recognition and mismatch excision to near wild type levels.  Such mutants are defective in the repair reaction directed by a 5' strand break, suggesting for the first time MutSα function in a post-excision step of the reaction.


Mutations in the human homologues of the Mut proteins affect genomic stability, which can result in  microsatellite instability  (MSI), implicated in some human cancers. In specific, the hereditary nonpolyposis colorectal cancers ( HNPCC  or Lynch syndrome) are attributed to damaging germline variants in the genes encoding the MutS and MutL homologues  MSH2  and  MLH1  respectively, which are thus classified as tumour suppressor genes. One subtype of HNPCC, the  Muir-Torre Syndrome  (MTS), is associated with skin tumors. If both inherited copies (alleles) of a MMR gene bear damaging genetic variants, this results in a very rare and severe condition: the  mismatch repair cancer syndrome  (or constitutional mismatch repair deficiency, CMMR-D), manifesting as multiple occurrences of tumors at an early age, often colon and  brain tumors . [ 8 ]


Sporadic cancers with a DNA repair deficiency only rarely have a mutation in a DNA repair gene, but they instead tend to have  epigenetic  alterations such as promoter methylation that inhibit DNA repair gene expression. [ 9 ]   About 13% of colorectal cancers are deficient in DNA mismatch repair, commonly due to loss of MLH1 (9.8%), or sometimes MSH2, MSH6 or PMS2 (all ≤1.5%). [ 10 ]   For most MLH1-deficient sporadic colorectal cancers, the deficiency was due to MLH1 promoter methylation. [ 10 ]   Other cancer types have higher frequencies of MLH1 loss (see table below), which are again largely a result of methylation of the promoter of the  MLH1  gene. A different epigenetic mechanism underlying MMR deficiencies might involve over-expression of a microRNA, for example  miR-155  levels inversely correlate with expression of MLH1 or MSH2 in colorectal cancer. [ 11 ]


A  field defect  (field cancerization) is an area of epithelium that has been preconditioned by epigenetic or genetic changes, predisposing it towards development of cancer. As pointed out by Rubin " ...there is evidence that more than 80% of the somatic mutations found in mutator phenotype human colorectal tumors occur before the onset of terminal clonal expansion." [ 20 ] [ 21 ]  Similarly, Vogelstein et al. [ 22 ]  point out that more than half of somatic mutations identified in tumors occurred in a pre-neoplastic phase (in a field defect), during growth of apparently normal cells.


MLH1 deficiencies were common in the field defects (histologically normal tissues) surrounding tumors; see Table above. Epigenetically silenced or mutated MLH1 would likely not confer a selective advantage upon a stem cell, however, it would cause increased mutation rates, and one or more of the mutated genes may provide the cell with a selective advantage. The deficient MLH1  gene could then be carried along as a selectively near-neutral passenger (hitch-hiker) gene when the mutated stem cell generates an expanded clone. The continued presence of a clone with an epigenetically repressed  MLH1  would continue to generate further mutations, some of which could produce a tumor.


MMR and mismatch repair mutations were initially observed to associate with immune checkpoint blockade efficacy in a study examining responders to anti-PD1. [ 23 ]  The association between MSI positivity and positive response to anti-PD1 was subsequently validated in a prospective clinical trial and approved by the FDA. [ 24 ]


In humans, seven DNA mismatch repair (MMR) proteins ( MLH1 ,  MLH3 ,  MSH2 ,  MSH3 ,  MSH6 ,  PMS1  and  PMS2 ) work coordinately in sequential steps to initiate repair of DNA mismatches. [ 25 ]   In addition, there are  Exo1 -dependent and Exo1-independent MMR subpathways. [ 26 ]


Other gene products involved in mismatch repair (subsequent to initiation by MMR genes) in humans include  DNA polymerase delta ,  PCNA ,  RPA ,  HMGB1 ,  RFC  and  DNA ligase I , plus  histone  and  chromatin  modifying factors. [ 27 ] [ 28 ]


In certain circumstances, the MMR pathway may recruit an error-prone DNA polymerase eta ( POLH ). This happens in B-lymphocytes during  somatic hypermutation , where POLH is used to introduce genetic variation into antibody genes. [ 29 ]  However, this error-prone MMR pathway may be triggered in other types of human cells upon exposure to genotoxins  [ 30 ]  and indeed it is broadly active in various human cancers, causing mutations that bear a signature of POLH activity. [ 31 ]


Recognizing and repairing mismatches and indels is important for cells because failure to do so results in  microsatellite instability  (MSI) and an elevated spontaneous  mutation rate  (mutator phenotype).  In comparison to other cancer types, MMR-deficient (MSI) cancer has a very high frequency of mutations, close to melanoma and lung cancer, [ 32 ]  cancer types caused by much exposure to UV radiation and mutagenic chemicals.


In addition to a very high mutation burden, MMR deficiencies result in an unusual distribution of somatic mutations across the human genome: this suggests that MMR preferentially protects the gene-rich, early-replicating euchromatic regions. [ 33 ]  In contrast, the gene-poor, late-replicating heterochromatic genome regions exhibit high mutation rates in many human tumors. [ 34 ]


The  histone modification   H3K36me3 , an  epigenetic  mark of active chromatin, has the ability to recruit the MSH2-MSH6 (hMutSα) complex. [ 35 ]  Consistently, regions of the human genome with high levels of H3K36me3 accumulate less mutations due to MMR activity. [ 31 ]


Lack of MMR often occurs in coordination with loss of other DNA repair genes. [ 9 ]  For example, MMR genes  MLH1  and   MLH3  as well as 11 other DNA repair genes (such as  MGMT  and many  NER  pathway genes) were significantly down-regulated in lower grade as well as in higher grade  astrocytomas , in contrast to normal brain tissue. [ 36 ]   Moreover, MLH1 and  MGMT  expression was closely correlated in 135 specimens of gastric cancer and loss of MLH1 and MGMT appeared to be synchronously accelerated during tumor progression. [ 37 ]


Deficient expression of multiple DNA repair genes is often found in cancers, [ 9 ]  and may contribute to the thousands of mutations usually found in cancers (see  Mutation frequencies in cancers ).


A popular idea, that has failed to gain significant experimental support, is the idea that mutation, as distinct from DNA damage, is the primary cause of aging.  Mice defective in the  mutL  homolog  Pms2  have about a 100-fold elevated mutation frequency in all tissues, but do not appear to age more rapidly. [ 38 ]   These mice display mostly normal development and life, except for early onset carcinogenesis and male infertility.


DNA repair  is a collection of processes by which a  cell  identifies and corrects damage to the  DNA  molecules that encode its  genome . [ 1 ]  In human cells, both normal  metabolic  activities and environmental factors such as  radiation  can cause DNA damage, resulting in tens of thousands of individual  molecular lesions  per cell per day. [ 2 ]  Many of these lesions cause structural damage to the DNA molecule and can alter or eliminate the cell's ability to  transcribe  the  gene  that the affected DNA encodes. Other lesions induce potentially harmful  mutations  in the cell's genome, which affect the survival of its daughter cells after it undergoes  mitosis . As a consequence, the DNA repair process is constantly active as it responds to damage in the DNA structure.  When normal repair processes fail, and when cellular  apoptosis  does not occur, irreparable DNA damage may occur. This can eventually lead to malignant tumors, or  cancer  as per the  two-hit hypothesis .


The rate of DNA repair depends on various factors, including the cell type, the age of the cell, and the extracellular environment. A cell that has accumulated a large amount of DNA damage or can no longer effectively repair its DNA may enter one of three possible states:


The DNA repair ability of a cell is vital to the integrity of its genome and thus to the normal functionality of that organism. Many genes that were initially shown to influence  life span  have turned out to be involved in DNA damage repair and protection. [ 3 ]


The 2015  Nobel Prize in Chemistry  was awarded to  Tomas Lindahl ,  Paul Modrich , and  Aziz Sancar  for their work on the molecular mechanisms of DNA repair processes. [ 4 ] [ 5 ]


DNA damage, due to environmental factors and normal  metabolic  processes inside the cell, occurs at a rate of 10,000 to 1,000,000 molecular lesions per cell per day. [ 2 ]  While this constitutes at most only 0.0003125% of the human genome's approximately 3.2 billion bases, unrepaired lesions in critical genes (such as  tumor suppressor genes ) can impede a cell's ability to carry out its function and appreciably increase the likelihood of  tumor  formation and contribute to  tumor heterogeneity .


The vast majority of DNA damage affects the  primary structure  of the double helix; that is, the bases themselves are chemically modified. These modifications can in turn disrupt the molecules' regular helical structure by introducing non-native chemical bonds or bulky  adducts  that do not fit in the standard double helix. Unlike  proteins  and  RNA , DNA usually lacks  tertiary structure  and therefore damage or disturbance does not occur at that level. DNA is, however,  supercoiled  and wound around "packaging" proteins called  histones  (in eukaryotes), and both superstructures are vulnerable to the effects of DNA damage.


DNA damage can be subdivided into two main types:


The replication of damaged DNA before cell division can lead to the incorporation of wrong bases opposite damaged ones. Daughter cells that inherit these wrong bases carry mutations from which the original DNA sequence is unrecoverable (except in the rare case of a  back mutation , for example, through  gene conversion ).


There are several types of damage to DNA due to endogenous cellular processes:


Damage caused by exogenous agents comes in many forms. Some examples are:


UV damage, alkylation/methylation, X-ray damage and oxidative damage are examples of induced damage. Spontaneous damage can include the loss of a base, deamination, sugar  ring puckering  and tautomeric shift. Constitutive (spontaneous) DNA damage caused by endogenous oxidants can be detected as a low level of histone H2AX phosphorylation in untreated cells. [ 9 ]


In human cells, and  eukaryotic  cells in general, DNA is found in two cellular locations – inside the  nucleus  and inside the  mitochondria . Nuclear DNA (n-DNA) exists as  chromatin  during non-replicative stages of the  cell cycle  and is condensed into aggregate structures known as  chromosomes  during  cell division . In either state the DNA is highly compacted and wound up around bead-like proteins called  histones . Whenever a cell needs to express the genetic information encoded in its n-DNA the required chromosomal region is unraveled, genes located therein are expressed, and then the region is condensed back to its resting conformation. Mitochondrial DNA (mtDNA) is located inside mitochondria  organelles , exists in multiple copies, and is also tightly associated with a number of proteins to form a complex known as the nucleoid. Inside mitochondria,  reactive oxygen species  (ROS), or  free radicals , byproducts of the constant production of  adenosine triphosphate  (ATP) via  oxidative phosphorylation , create a highly oxidative environment that is known to damage mtDNA. A critical enzyme in counteracting the  toxicity  of these species is  superoxide dismutase , which is present in both the mitochondria and  cytoplasm  of eukaryotic cells.


Senescence, an irreversible process in which the cell no longer  divides , is a protective response to the shortening of the chromosome ends, called  telomeres . The telomeres are long regions of repetitive  noncoding DNA  that cap chromosomes and undergo partial degradation each time a cell undergoes division (see  Hayflick limit ). [ 10 ]  In contrast,  quiescence  is a reversible state of cellular dormancy that is unrelated to genome damage (see  cell cycle ). Senescence in cells may serve as a functional alternative to apoptosis in cases where the physical presence of a cell for spatial reasons is required by the organism, [ 11 ]  which serves as a "last resort" mechanism to prevent a cell with damaged DNA from replicating inappropriately in the absence of pro-growth  cellular signaling . Unregulated cell division can lead to the formation of a tumor (see  cancer ), which is potentially lethal to an organism. Therefore, the induction of senescence and apoptosis is considered to be part of a strategy of protection against cancer. [ 12 ]


It is important to distinguish between DNA damage and mutation, the two major types of error in DNA. DNA damage and mutation are fundamentally different. Damage results in physical abnormalities in the DNA, such as single- and double-strand breaks,  8-hydroxydeoxyguanosine  residues, and polycyclic aromatic hydrocarbon adducts. DNA damage can be recognized by enzymes, and thus can be correctly repaired if redundant information, such as the undamaged sequence in the complementary DNA strand or in a homologous chromosome, is available for copying. If a cell retains DNA damage, transcription of a gene can be prevented, and thus translation into a protein will also be blocked.  Replication may also be blocked or the cell may die.


In contrast to DNA damage, a mutation is a change in the base sequence of the DNA. A mutation cannot be recognized by enzymes once the base change is present in both DNA strands, and thus a mutation cannot be repaired. At the cellular level, mutations can cause alterations in protein function and regulation. Mutations are replicated when the cell replicates. In a population of cells, mutant cells will increase or decrease in frequency according to the effects of the mutation on the ability of the cell to survive and reproduce.


Although distinctly different from each other, DNA damage and mutation are related because DNA damage often causes errors of DNA synthesis during replication or repair; these errors are a major source of mutation.


Given these properties of DNA damage and mutation, it can be seen that DNA damage is a special problem in non-dividing or slowly-dividing cells, where unrepaired damage will tend to accumulate over time. On the other hand, in rapidly dividing cells, unrepaired DNA damage that does not kill the cell by blocking replication will tend to cause replication errors and thus mutation. The great majority of mutations that are not neutral in their effect are deleterious to a cell's survival. Thus, in a population of cells composing a tissue with replicating cells, mutant cells will tend to be lost. However, infrequent mutations that provide a survival advantage will tend to clonally expand at the expense of neighboring cells in the tissue. This advantage to the cell is disadvantageous to the whole organism because such mutant cells can give rise to cancer. Thus, DNA damage in frequently dividing cells, because it gives rise to mutations, is a prominent cause of cancer. In contrast, DNA damage in infrequently-dividing cells is likely a prominent cause of aging. [ 13 ]


Cells cannot function if DNA damage corrupts the integrity and accessibility of essential information in the  genome  (but cells remain superficially functional when non-essential genes are missing or damaged). Depending on the type of damage inflicted on the DNA's double helical structure, a variety of repair strategies have evolved to restore lost information. If possible, cells use the unmodified complementary strand of the DNA or the sister  chromatid  as a template to recover the original information. Without access to a template, cells use an error-prone recovery mechanism known as  translesion synthesis  as a last resort.


Damage to DNA alters the spatial configuration of the helix, and such alterations can be detected by the cell. Once damage is localized, specific DNA repair molecules bind at or near the site of damage, inducing other molecules to bind and form a complex that enables the actual repair to take place.


Cells are known to eliminate three types of damage to their DNA by chemically reversing it. These mechanisms do not require a template, since the types of damage they counteract can occur in only one of the four bases. Such direct reversal mechanisms are specific to the type of damage incurred and do not involve breakage of the phosphodiester backbone. The formation of  pyrimidine dimers  upon irradiation with UV light results in an abnormal covalent bond between adjacent pyrimidine bases. The  photoreactivation  process directly reverses this damage by the action of the enzyme  photolyase , whose activation is obligately dependent on energy absorbed from  blue/UV light  (300–500 nm  wavelength ) to promote catalysis. [ 14 ]  Photolyase, an old enzyme present in  bacteria ,  fungi , and most  animals  no longer functions in humans, [ 15 ]  who instead use  nucleotide excision repair  to repair damage from UV irradiation. Another type of damage, methylation of guanine bases, is directly reversed by the enzyme methyl guanine methyl transferase (MGMT), the bacterial equivalent of which is called  ogt . This is an expensive process because each MGMT molecule can be used only once; that is, the reaction is  stoichiometric  rather than  catalytic . [ 16 ]  A generalized response to methylating agents in bacteria is known as the  adaptive response  and confers a level of resistance to alkylating agents upon sustained exposure by upregulation of alkylation repair enzymes. [ 17 ]  The third type of DNA damage reversed by cells is certain methylation of the bases cytosine and adenine.


When only one of the two strands of a double helix has a defect, the other strand can be used as a template to guide the correction of the damaged strand. In order to repair damage to one of the two paired molecules of DNA, there exist a number of  excision repair  mechanisms that remove the damaged nucleotide and replace it with an undamaged nucleotide complementary to that found in the undamaged DNA strand. [ 16 ]


Double-strand breaks, in which both strands in the double helix are severed, are particularly hazardous to the cell because they can lead to  genome rearrangements . In fact, when a double-strand break is accompanied by a cross-linkage joining the two strands at the same point, neither strand can be used as a template for the repair mechanisms, so that the cell will not be able to complete mitosis when it next divides, and will either die or, in rare cases, undergo a mutation. [ 22 ] [ 23 ]  Three mechanisms exist to repair double-strand breaks (DSBs):  non-homologous end joining  (NHEJ),  microhomology-mediated end joining  (MMEJ), and  homologous recombination  (HR): [ 16 ] [ 24 ]


In an  in vitro  system, MMEJ occurred in mammalian cells at the levels of 10–20% of HR when both HR and NHEJ mechanisms were also available. [ 32 ]


The  extremophile   Deinococcus radiodurans  has a remarkable ability to survive DNA damage from  ionizing radiation  and other sources. At least two copies of the genome, with random DNA breaks, can form DNA fragments through  annealing . Partially overlapping fragments are then used for synthesis of  homologous  regions through a moving  D-loop  that can continue extension until complementary partner strands are found. In the final step, there is  crossover  by means of  RecA -dependent  homologous recombination . [ 35 ]


Topoisomerases  introduce both single- and double-strand breaks in the course of changing the DNA's state of  supercoiling , which is especially common in regions near an open replication fork. Such breaks are not considered DNA damage because they are a natural intermediate in the topoisomerase biochemical mechanism and are immediately repaired by the enzymes that created them.


Another type of DNA double-strand breaks originates from the DNA heat-sensitive or heat-labile sites. These DNA sites are not initial DSBs. However, they convert to DSB after treating with elevated temperature. Ionizing irradiation can induces a highly complex form of DNA damage as clustered damage. It consists of different types of DNA lesions in various locations of the DNA helix. Some of these closely located lesions can probably convert to DSB by exposure to high temperatures. But the exact nature of these lesions and their interactions is not yet known [ 36 ]


Translesion synthesis (TLS) is a DNA damage tolerance process that allows the  DNA replication  machinery to replicate past DNA lesions such as  thymine dimers  or  AP sites . [ 37 ]  It involves switching out regular  DNA polymerases  for specialized translesion polymerases (i.e. DNA polymerase IV or V, from the Y Polymerase family), often with larger active sites that can facilitate the insertion of bases opposite damaged nucleotides. The polymerase switching is thought to be mediated by, among other factors, the post-translational modification of the replication  processivity  factor  PCNA . Translesion synthesis polymerases often have low fidelity (high propensity to insert wrong bases) on undamaged templates relative to regular polymerases. However, many are extremely efficient at inserting correct bases opposite specific types of damage. For example,  Pol η  mediates error-free bypass of lesions induced by  UV irradiation , whereas  Pol ι  introduces mutations at these sites. Pol η is known to add the first adenine across the  T^T photodimer  using  Watson-Crick base pairing  and the second adenine will be added in its syn conformation using  Hoogsteen base pairing . From a cellular perspective, risking the introduction of  point mutations  during translesion synthesis may be preferable to resorting to more drastic mechanisms of DNA repair, which may cause gross chromosomal aberrations or cell death. In short, the process involves specialized  polymerases  either bypassing or repairing lesions at locations of stalled DNA replication. For example, Human DNA polymerase eta can bypass complex DNA lesions like guanine-thymine intra-strand crosslink, G[8,5-Me]T, although it can cause targeted and semi-targeted mutations. [ 38 ]  Paromita Raychaudhury and Ashis Basu [ 39 ]  studied the toxicity and mutagenesis of the same lesion in  Escherichia coli  by replicating a G[8,5-Me]T-modified plasmid in  E. coli  with specific DNA polymerase knockouts. Viability was very low in a strain lacking pol II, pol IV, and pol V, the three SOS-inducible DNA polymerases, indicating that translesion synthesis is conducted primarily by these specialized DNA polymerases.
A bypass platform is provided to these polymerases by  Proliferating cell nuclear antigen  (PCNA). Under normal circumstances, PCNA bound to polymerases replicates the DNA. At a site of  lesion , PCNA is ubiquitinated, or modified, by the RAD6/ RAD18   proteins  to provide a platform for the specialized polymerases to bypass the lesion and resume DNA replication. [ 40 ] [ 41 ]  After translesion synthesis, extension is required. This extension can be carried out by a replicative polymerase if the TLS is error-free, as in the case of Pol η, yet if TLS results in a mismatch, a specialized polymerase is needed to extend it;  Pol ζ . Pol ζ is unique in that it can extend terminal mismatches, whereas more processive polymerases cannot. So when a lesion is encountered, the replication fork will stall, PCNA will switch from a processive polymerase to a TLS polymerase such as Pol ι to fix the lesion, then PCNA may switch to Pol ζ to extend the mismatch, and last PCNA will switch to the processive polymerase to continue replication.


Cells exposed to  ionizing radiation ,  ultraviolet light  or chemicals are prone to acquire multiple sites of bulky DNA lesions and double-strand breaks. Moreover, DNA damaging agents can damage other  biomolecules  such as  proteins ,  carbohydrates ,  lipids , and  RNA . The accumulation of damage, to be specific, double-strand breaks or adducts stalling the  replication forks , are among known stimulation signals for a global response to DNA damage. [ 42 ]  The global response to damage is an act directed toward the cells' own preservation and triggers multiple pathways of macromolecular repair, lesion bypass, tolerance, or  apoptosis . The common features of global response are induction of multiple  genes ,  cell cycle  arrest, and inhibition of  cell division .


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 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. [ 43 ]


Chromatin relaxation occurs rapidly at the site of a DNA damage. [ 44 ] [ 45 ]   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. [ 46 ]   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. [ 46 ]   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. [ 47 ]   PARP1 synthesizes  polymeric   adenosine diphosphate ribose  (poly (ADP-ribose) or PAR) chains on itself.  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. [ 45 ]   About half of the maximum chromatin relaxation, presumably due to action of ALC1, occurs by 10 seconds. [ 45 ]  This then allows recruitment of the DNA repair enzyme  MRE11 , to initiate DNA repair, within 13 seconds. [ 47 ]


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. [ 48 ]   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. [ 48 ]   The extent of chromatin with phosphorylated γH2AX is about two million base pairs at the site of a DNA double-strand break. [ 48 ]   γH2AX does not, itself, cause chromatin decondensation, but within 30 seconds of irradiation,  RNF8  protein can be detected in association with γH2AX. [ 49 ]   RNF8 mediates extensive chromatin decondensation, through its subsequent interaction with  CHD4 , [ 50 ]  a component of the nucleosome remodeling and deacetylase complex  NuRD .


DDB2  occurs in a heterodimeric complex with  DDB1 .  This complex further complexes with the  ubiquitin ligase  protein  CUL4A [ 51 ]  and with  PARP1 . [ 52 ]   This larger complex rapidly associates with UV-induced damage within chromatin, with half-maximum association completed in 40 seconds. [ 51 ]   The PARP1 protein, attached to both DDB1 and DDB2, then  PARylates  (creates a poly-ADP ribose chain) on DDB2 that attracts the DNA remodeling protein  ALC1 . [ 52 ]   Action of ALC1 relaxes the chromatin at the site of UV damage to DNA.  This relaxation allows other proteins in the  nucleotide excision repair  pathway to enter the chromatin and repair UV-induced  cyclobutane pyrimidine dimer  damages.


After rapid  chromatin remodeling ,  cell cycle   checkpoints  are activated to allow DNA repair to occur 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. [ 53 ]


After DNA damage,  cell cycle   checkpoints  are activated. Checkpoint activation pauses the cell cycle and gives the cell time to repair the damage before continuing to divide. DNA damage checkpoints occur at the  G1 / S  and  G2 / M  boundaries. An intra- S  checkpoint also exists. Checkpoint activation is controlled by two master  kinases ,  ATM  and  ATR . ATM responds to DNA double-strand breaks and disruptions in chromatin structure, [ 54 ]  whereas ATR primarily responds to stalled  replication forks . These kinases  phosphorylate  downstream targets in a  signal transduction  cascade, eventually leading to cell cycle arrest. A class of checkpoint mediator proteins including  BRCA1 ,  MDC1 , and  53BP1  has also been identified. [ 55 ]  These proteins seem to be required for transmitting the checkpoint activation signal to downstream proteins.


DNA damage checkpoint  is a  signal transduction pathway  that blocks  cell cycle  progression in G1, G2 and  metaphase  and slows down the rate of S phase progression when  DNA  is damaged. It leads to a pause in cell cycle allowing the cell time to repair the damage before continuing to divide.


Checkpoint Proteins can be separated into four groups:  phosphatidylinositol 3-kinase  (PI3K)-like  protein kinase ,  proliferating cell nuclear antigen  (PCNA)-like group, two serine/threonine(S/T) kinases and their adaptors. Central to all DNA damage induced checkpoints responses is a pair of large protein kinases belonging to the first group of PI3K-like protein kinases-the ATM ( Ataxia telangiectasia mutated ) and ATR (Ataxia- and Rad-related) kinases, whose sequence and functions have been well conserved in evolution. All DNA damage response requires either ATM or ATR because they have the ability to bind to the  chromosomes  at the site of DNA damage, together with accessory proteins that are platforms on which DNA damage response components and DNA repair complexes can be assembled.


An important downstream target of ATM and ATR is  p53 , as it is required for inducing  apoptosis  following DNA damage. [ 56 ]  The  cyclin-dependent kinase inhibitor   p21  is induced by both p53-dependent and p53-independent mechanisms and can arrest the cell cycle at the G1/S and G2/M checkpoints by deactivating  cyclin / cyclin-dependent kinase  complexes. [ 57 ]


The  SOS response  is the changes in  gene expression  in  Escherichia coli  and other bacteria in response to extensive DNA damage. The  prokaryotic  SOS system is regulated by two key proteins:  LexA  and  RecA . The LexA  homodimer  is a  transcriptional   repressor  that binds to  operator  sequences commonly referred to as SOS boxes. In  Escherichia coli  it is known that LexA regulates transcription of approximately 48 genes including the lexA and recA genes. [ 58 ]  The SOS response is known to be widespread in the Bacteria domain, but it is mostly absent in some bacterial phyla, like the  Spirochetes . [ 59 ]
The most common cellular signals activating the SOS response are regions of single-stranded DNA (ssDNA), arising from stalled  replication forks  or double-strand breaks, which are processed by  DNA helicase  to separate the two DNA strands. [ 42 ]  In the initiation step, RecA protein binds to ssDNA in an  ATP hydrolysis  driven reaction creating RecA–ssDNA filaments. RecA–ssDNA filaments activate LexA auto protease  activity, which ultimately leads to cleavage of LexA dimer and subsequent LexA degradation. The loss of LexA repressor induces transcription of the SOS genes and allows for further signal induction, inhibition of cell division and an increase in levels of proteins responsible for damage processing.


In  Escherichia coli , SOS boxes are 20-nucleotide long sequences near promoters with  palindromic  structure and a high degree of sequence conservation. In other classes and phyla, the sequence of SOS boxes varies considerably, with different length and composition, but it is always highly conserved and one of the strongest short signals in the genome. [ 59 ]  The high information content of SOS boxes permits differential binding of LexA to different promoters and allows for timing of the SOS response. The lesion repair genes are induced at the beginning of SOS response. The error-prone translesion polymerases, for example, UmuCD'2 (also called DNA polymerase V), are induced later on as a last resort. [ 60 ]  Once the DNA damage is repaired or bypassed using polymerases or through recombination, the amount of single-stranded DNA in cells is decreased, lowering the amounts of RecA filaments decreases cleavage activity of LexA homodimer, which then binds to the SOS boxes near promoters and restores normal gene expression.


Eukaryotic  cells exposed to DNA damaging agents also activate important defensive pathways by inducing multiple proteins involved in DNA repair,  cell cycle checkpoint  control, protein trafficking and degradation. Such genome wide transcriptional response is very complex and tightly regulated, thus allowing coordinated global response to damage. Exposure of  yeast   Saccharomyces cerevisiae  to DNA damaging agents results in overlapping but distinct transcriptional profiles. Similarities to environmental  shock response  indicates that a general global stress response pathway exist at the level of transcriptional activation. In contrast, different human cell types respond to damage differently indicating an absence of a common global response. The probable explanation for this difference between yeast and human cells may be in the  heterogeneity  of  mammalian  cells. In an animal different types of cells are distributed among different organs that have evolved different sensitivities to DNA damage. [ 61 ]


In general global response to DNA damage involves expression of multiple genes responsible for  postreplication repair , homologous recombination, nucleotide excision repair,  DNA damage checkpoint , global transcriptional activation, genes controlling mRNA decay, and many others. A large amount of damage to a cell leaves it with an important decision: undergo apoptosis and die, or survive at the cost of living with a modified genome. An increase in tolerance to damage can lead to an increased rate of survival that will allow a greater accumulation of mutations. Yeast Rev1 and human polymerase η are members of Y family translesion DNA  polymerases  present during global response to DNA damage and are responsible for enhanced mutagenesis during a global response to DNA damage in eukaryotes. [ 42 ]


Experimental animals with genetic deficiencies in DNA repair often show decreased life span and increased cancer incidence. [ 13 ]  For example, mice deficient in the dominant NHEJ pathway and in telomere maintenance mechanisms get  lymphoma  and infections more often, and, as a consequence, have shorter lifespans than wild-type mice. [ 62 ]  In similar manner, mice deficient in a key repair and transcription protein that unwinds DNA helices have premature onset of aging-related diseases and consequent shortening of lifespan. [ 63 ]   However, not every DNA repair deficiency creates exactly the predicted effects; mice deficient in the NER pathway exhibited shortened life span without correspondingly higher rates of mutation. [ 64 ]


The  maximum life spans  of  mice ,  naked mole-rats  and  humans  are respectively ~3, ~30 and ~129 years. [ 65 ]   Of these, the shortest lived species, mouse, expresses DNA repair genes, including core genes in several DNA repair pathways, at a lower level than do humans and naked mole rats. [ 65 ]   Furthermore several DNA repair pathways in humans and naked mole-rats are up-regulated compared to mouse.  These observations suggest that elevated DNA repair facilitates greater  longevity . [ 65 ]


If the rate of DNA damage exceeds the capacity of the cell to repair it, the accumulation of errors can overwhelm the cell and result in early senescence, apoptosis, or cancer. Inherited diseases associated with faulty DNA repair functioning result in premature aging, [ 13 ]  increased sensitivity to carcinogens and correspondingly increased cancer risk (see  below ). On the other hand, organisms with enhanced DNA repair systems, such as  Deinococcus radiodurans , the most radiation-resistant known organism, exhibit remarkable resistance to the double-strand break-inducing effects of  radioactivity , likely due to enhanced efficiency of DNA repair and especially NHEJ. [ 66 ]


A number of individual genes have been identified as influencing variations in life span within a population of organisms. The effects of these genes is strongly dependent on the environment, in particular, on the organism's diet.  Caloric restriction  reproducibly results in extended lifespan in a variety of organisms, likely via  nutrient sensing  pathways and decreased  metabolic rate . The molecular mechanisms by which such restriction results in lengthened lifespan are as yet unclear (see [ 67 ]  for some discussion); however, the behavior of many genes known to be involved in DNA repair is altered under conditions of caloric restriction. Several agents reported to have anti-aging properties have been shown to attenuate constitutive level of  mTOR  signaling, an evidence of reduction of  metabolic activity , and concurrently to reduce  constitutive level of  DNA damage  induced by endogenously generated reactive oxygen species. [ 68 ]


For example, increasing the  gene dosage  of the gene SIR-2, which regulates DNA packaging in the nematode worm  Caenorhabditis elegans , can significantly extend lifespan. [ 69 ]  The mammalian homolog of SIR-2 is known to induce downstream DNA repair factors involved in NHEJ, an activity that is especially promoted under conditions of caloric restriction. [ 70 ]  Caloric restriction has been closely linked to the rate of base excision repair in the nuclear DNA of rodents, [ 71 ]  although similar effects have not been observed in mitochondrial DNA. [ 72 ]


The  C. elegans  gene AGE-1, an upstream effector of DNA repair pathways, confers dramatically extended life span under free-feeding conditions but leads to a decrease in reproductive fitness under conditions of caloric restriction. [ 73 ]  This observation supports the  pleiotropy  theory of the  biological origins of aging , which suggests that genes conferring a large survival advantage early in life will be selected for even if they carry a corresponding disadvantage late in life.


Defects in the NER mechanism are responsible for several genetic disorders, including:


Mental retardation often accompanies the latter two disorders, suggesting increased vulnerability of developmental neurons.


Other DNA repair disorders include:


All of the above diseases are often called "segmental  progerias " (" accelerated aging diseases ") because those affected appear elderly and experience aging-related diseases at an abnormally young age, while not manifesting all the symptoms of old age.


Other diseases associated with reduced DNA repair function include  Fanconi anemia , hereditary  breast cancer  and hereditary  colon cancer .


Because of inherent limitations in the DNA repair mechanisms, if humans lived long enough, they would all eventually develop cancer. [ 74 ] [ 75 ]   There are at least 34  Inherited human DNA repair gene mutations that increase cancer risk .  Many of these mutations cause DNA repair to be less effective than normal.  In particular,  Hereditary nonpolyposis colorectal cancer  (HNPCC) is strongly associated with specific mutations in the DNA mismatch repair pathway.  BRCA1  and  BRCA2 , two important genes whose mutations confer a hugely increased risk of breast cancer on carriers, [ 76 ]  are both associated with a large number of DNA repair pathways, especially NHEJ and homologous recombination.


Cancer therapy procedures such as  chemotherapy  and  radiotherapy  work by overwhelming the capacity of the cell to repair DNA damage, resulting in cell death. Cells that are most rapidly dividing – most typically cancer cells – are preferentially affected. The side-effect is that other non-cancerous but rapidly dividing cells such as progenitor cells in the gut, skin, and hematopoietic system are also affected. Modern cancer treatments attempt to localize the DNA damage to cells and tissues only associated with cancer, either by physical means (concentrating the therapeutic agent in the region of the tumor) or by biochemical means (exploiting a feature unique to cancer cells in the body).  In the context of therapies targeting DNA damage response genes, the latter approach has been termed 'synthetic lethality'. [ 77 ]


Perhaps the most well-known of these 'synthetic lethality' drugs is the poly(ADP-ribose) polymerase 1 ( PARP1 ) inhibitor  olaparib , which was approved by the Food and Drug Administration in 2015 for the treatment in women of BRCA-defective ovarian cancer. Tumor cells with partial loss of DNA damage response (specifically,  homologous recombination  repair) are dependent on another mechanism – single-strand break repair – which is a mechanism consisting, in part, of the PARP1 gene product. [ 78 ]   Olaparib  is combined with chemotherapeutics to inhibit single-strand break repair induced by DNA damage caused by the co-administered chemotherapy. Tumor cells relying on this residual DNA repair mechanism are unable to repair the damage and hence are not able to survive and proliferate, whereas normal cells can repair the damage with the functioning homologous recombination mechanism.


Many other drugs for use against other residual DNA repair mechanisms commonly found in cancer are currently under investigation. However, synthetic lethality therapeutic approaches have been questioned due to emerging evidence of acquired resistance, achieved through rewiring of DNA damage response pathways and reversion of previously inhibited defects. [ 79 ]


It has become apparent over the past several years that the DNA damage response acts as a barrier to the malignant transformation of preneoplastic cells. [ 80 ]  Previous studies have shown an elevated DNA damage response in cell-culture models with oncogene activation [ 81 ]  and preneoplastic colon adenomas. [ 82 ]  DNA damage response mechanisms trigger cell-cycle arrest, and attempt to repair DNA lesions or promote cell death/senescence if repair is not possible. Replication stress is observed in preneoplastic cells due to increased proliferation signals from oncogenic mutations.  Replication stress  is characterized by: increased replication initiation/origin firing; increased transcription and collisions of transcription-replication complexes; nucleotide deficiency; increase in reactive oxygen species (ROS). [ 83 ]


Replication stress, along with the selection for inactivating mutations in DNA damage response genes in the evolution of the tumor, [ 84 ]  leads to downregulation and/or loss of some DNA damage response mechanisms, and hence loss of DNA repair and/or senescence/programmed cell death. In experimental mouse models, loss of DNA damage response-mediated cell senescence was observed after using a  short hairpin RNA  (shRNA) to inhibit the double-strand break response kinase ataxia telangiectasia ( ATM ), leading to increased tumor size and invasiveness. [ 82 ]  Humans born with inherited defects in DNA repair mechanisms (for example,  Li-Fraumeni syndrome ) have a higher cancer risk. [ 85 ]


The prevalence of DNA damage response mutations differs across cancer types; for example, 30% of breast invasive carcinomas have mutations in genes involved in homologous recombination. [ 80 ]  In cancer, downregulation is observed across all DNA damage response mechanisms (base excision repair (BER), nucleotide excision repair (NER), DNA mismatch repair (MMR), homologous recombination repair (HR), non-homologous end joining (NHEJ) and translesion DNA synthesis (TLS). [ 86 ]  As well as mutations to DNA damage repair genes, mutations also arise in the genes responsible for arresting the  cell cycle  to allow sufficient time for DNA repair to occur, and some genes are involved in both DNA damage repair and cell cycle checkpoint control, for example ATM and checkpoint kinase 2 (CHEK2) – a tumor suppressor that is often absent or downregulated in non-small cell lung cancer. [ 87 ]


Classically, cancer has been viewed as a set of diseases that are driven by progressive genetic abnormalities that include mutations in tumour-suppressor genes and oncogenes, and chromosomal aberrations. However, it has become apparent that cancer is also driven by  epigenetic alterations . [ 88 ]


Epigenetic alterations refer to functionally relevant modifications to the genome that do not involve a change in the nucleotide sequence. Examples of such modifications are changes in  DNA methylation  (hypermethylation and hypomethylation) and  histone modification , [ 89 ]  changes in chromosomal architecture (caused by inappropriate expression of proteins such as  HMGA2  or  HMGA1 ) [ 90 ]  and changes caused by  microRNAs . Each of these epigenetic alterations serves to regulate gene expression without altering the underlying  DNA sequence . These changes usually remain through  cell divisions , last for multiple cell generations, and can be considered to be epimutations (equivalent to mutations).


While large numbers of epigenetic alterations are found in cancers, the epigenetic alterations in DNA repair genes, causing reduced expression of DNA repair proteins, appear to be particularly important.  Such alterations are thought to occur early in progression to cancer and to be a likely cause of the  genetic  instability characteristic of cancers. [ 91 ] [ 92 ] [ 93 ]


Reduced expression of DNA repair genes causes deficient DNA repair. When DNA repair is deficient DNA damages remain in cells at a higher than usual level and these excess damages cause increased frequencies of mutation or epimutation. Mutation rates increase substantially in cells defective in  DNA mismatch repair [ 94 ] [ 95 ]  or in  homologous recombinational  repair (HRR). [ 96 ]  Chromosomal rearrangements and aneuploidy also increase in HRR defective cells. [ 97 ]


Higher levels of DNA damage not only cause increased mutation, but also cause increased epimutation. During repair of DNA double strand breaks, or repair of other DNA damages, incompletely cleared sites of repair can cause epigenetic gene silencing. [ 98 ] [ 99 ]


Deficient expression of DNA repair proteins due to an inherited mutation can cause increased risk of cancer.  Individuals with an inherited impairment in any of 34 DNA repair genes (see article  DNA repair-deficiency disorder ) have an increased risk of cancer, with some defects causing up to a 100% lifetime chance of cancer (e.g. p53 mutations). [ 100 ]  However, such  germline mutations  (which cause highly penetrant cancer syndromes) are the cause of only about 1 percent of cancers. [ 101 ]


Deficiencies in DNA repair enzymes are occasionally caused by a newly arising somatic mutation in a DNA repair gene, but are much more frequently caused by epigenetic alterations that reduce or silence expression of DNA repair genes. For example, when 113 colorectal cancers were examined in sequence, only four had a  missense mutation  in the DNA repair gene  MGMT , while the majority had reduced MGMT expression due to methylation of the MGMT promoter region (an epigenetic alteration). [ 102 ]  Five different studies found that between 40% and 90% of colorectal cancers have reduced MGMT expression due to methylation of the MGMT promoter region. [ 103 ] [ 104 ] [ 105 ] [ 106 ] [ 107 ]


Similarly, out of 119 cases of mismatch repair-deficient colorectal cancers that lacked DNA repair gene  PMS2  expression, PMS2 was deficient in 6 due to mutations in the PMS2 gene, while in 103 cases PMS2 expression was deficient because its pairing partner  MLH1  was repressed due to promoter methylation (PMS2 protein is unstable in the absence of MLH1). [ 108 ]  In the other 10 cases, loss of PMS2 expression was likely due to epigenetic overexpression of the  microRNA ,  miR-155 , which down-regulates MLH1. [ 109 ]


In a further example, epigenetic defects were found in various cancers (e.g. breast, ovarian, colorectal and head and neck). Two or three deficiencies in the expression of  ERCC1 ,  XPF  or PMS2 occur simultaneously in the majority of 49 colon cancers evaluated by Facista et al. [ 110 ]


The chart in this section shows some frequent DNA damaging agents, examples of DNA lesions they cause, and the pathways that deal with these DNA damages.  At least 169 enzymes are either directly employed in DNA repair or influence DNA repair processes. [ 111 ]  Of these, 83 are directly employed in repairing the 5 types of DNA damages illustrated in the chart. [ citation needed ]


Some of the more well studied genes central to these repair processes are shown in the chart.  The gene designations shown in red, gray or cyan indicate genes frequently epigenetically altered in various types of cancers.  Wikipedia articles on each of the genes highlighted by red, gray or cyan describe the epigenetic alteration(s) and the cancer(s) in which these epimutations are found.  Review articles, [ 112 ]  and broad experimental survey articles [ 113 ] [ 114 ]  also document most of these epigenetic DNA repair deficiencies in cancers.


Red-highlighted genes are frequently reduced or silenced by epigenetic mechanisms in various cancers.  When these genes have low or absent expression, DNA damages can accumulate.  Replication errors past these damages (see  translesion synthesis ) can lead to increased mutations and, ultimately, cancer. Epigenetic repression of DNA repair genes in  accurate  DNA repair pathways appear to be central to  carcinogenesis .


The two gray-highlighted genes  RAD51  and  BRCA2 , are required for  homologous recombinational  repair.  They are sometimes epigenetically over-expressed and sometimes under-expressed in certain cancers.  As indicated in the Wikipedia articles on  RAD51  and  BRCA2 , such cancers ordinarily have epigenetic deficiencies in other DNA repair genes. These repair deficiencies would likely cause increased unrepaired DNA damages. The over-expression of  RAD51  and  BRCA2  seen in these cancers may reflect selective pressures for compensatory  RAD51  or  BRCA2  over-expression and increased homologous recombinational repair to at least partially deal with such excess DNA damages.  In those cases where  RAD51  or  BRCA2  are under-expressed, this would itself lead to increased unrepaired DNA damages. Replication errors past these damages (see  translesion synthesis ) could cause increased mutations and cancer, so that under-expression of  RAD51  or  BRCA2  would be carcinogenic in itself.


Cyan-highlighted genes are in the  microhomology-mediated end joining  (MMEJ) pathway and are up-regulated in cancer.  MMEJ is an additional error-prone  inaccurate  repair pathway for double-strand breaks.  In MMEJ repair of a double-strand break, an homology of 5–25 complementary base pairs between both paired strands is sufficient to align the strands, but mismatched ends (flaps) are usually present. MMEJ removes the extra nucleotides (flaps) where strands are joined, and then ligates the strands to create an intact DNA double helix. MMEJ almost always involves at least a small deletion, so that it is a mutagenic pathway. [ 24 ]   FEN1 , the flap endonuclease in MMEJ, is epigenetically increased by promoter hypomethylation and is over-expressed in the majority of cancers of the breast, [ 115 ]  prostate, [ 116 ]  stomach, [ 117 ] [ 118 ]  neuroblastomas, [ 119 ]  pancreas, [ 120 ]  and lung. [ 121 ]   PARP1 is also over-expressed when its promoter region  ETS  site is  epigenetically  hypomethylated, and this contributes to progression to endometrial cancer [ 122 ]  and BRCA-mutated serous ovarian cancer. [ 123 ]  Other genes in the  MMEJ  pathway are also over-expressed in a number of cancers (see  MMEJ  for summary), and are also shown in cyan.


Differential activity of DNA repair pathways across various regions of the human genome causes mutations to be very unevenly distributed within tumor genomes. [ 124 ] [ 125 ]  In particular, the gene-rich, early-replicating regions of the human genome exhibit lower mutation frequencies than the gene-poor, late-replicating  heterochromatin . One mechanism underlying this involves the  histone modification   H3K36me3 , which can recruit  mismatch repair  proteins, [ 126 ]  thereby lowering mutation rates in  H3K36me3 -marked regions. [ 127 ]  Another important mechanism concerns  nucleotide excision repair , which can be recruited by the transcription machinery, lowering somatic mutation rates in active genes [ 125 ]  and other open chromatin regions. [ 128 ]


Damage to DNA is very common and is constantly being repaired.  Epigenetic alterations can accompany DNA repair of oxidative damage or double-strand breaks.  In human cells, oxidative DNA damage occurs about 10,000 times a day and DNA double-strand breaks occur about 10 to 50 times a cell cycle in somatic replicating cells (see  DNA damage (naturally occurring) ).  The selective advantage of DNA repair is to allow the cell to survive in the face of DNA damage.  The selective advantage of epigenetic alterations that occur with DNA repair is not clear. [ citation needed ]


In the steady state (with endogenous damages occurring and being repaired), there are about 2,400 oxidatively damaged guanines that form  8-oxo-2'-deoxyguanosine  (8-OHdG) in the average mammalian cell DNA. [ 129 ]   8-OHdG constitutes about 5% of the oxidative damages commonly present in DNA. [ 130 ]   The oxidized guanines do not occur randomly among all guanines in DNA. There is a sequence preference for the guanine at a  methylated   CpG site  (a cytosine followed by guanine along its  5' → 3' direction  and where the cytosine is methylated (5-mCpG)). [ 131 ]   A 5-mCpG site has the lowest ionization potential for guanine oxidation.


Oxidized guanine has mispairing potential and is mutagenic. [ 133 ]   Oxoguanine glycosylase  (OGG1) is the primary enzyme responsible for the excision of the oxidized guanine during DNA repair. OGG1 finds and binds to an 8-OHdG within a few seconds. [ 134 ]   However, OGG1 does not immediately excise 8-OHdG.  In HeLa cells half maximum removal of 8-OHdG occurs in 30 minutes, [ 135 ]  and in irradiated mice, the 8-OHdGs induced in the mouse liver are removed with a half-life of 11 minutes. [ 130 ]


When OGG1 is present at an oxidized guanine within a methylated  CpG site  it recruits  TET1  to the 8-OHdG lesion (see Figure). This allows TET1 to demethylate an adjacent methylated cytosine. [ 136 ]  Demethylation of cytosine is an epigenetic alteration. [ 137 ]


As an example, when human mammary epithelial cells were treated with H 2 O 2  for six hours, 8-OHdG increased about 3.5-fold in DNA and this caused about 80% demethylation of the 5-methylcytosines in the genome. [ 132 ]   Demethylation of CpGs in a gene promoter by  TET enzyme  activity increases transcription of the gene into messenger RNA. [ 138 ]   In cells treated with H 2 O 2 , one particular gene was examined,  BACE1 . [ 132 ]   The methylation level of the  BACE1   CpG island  was reduced (an epigenetic alteration) and this allowed about 6.5 fold increase of expression of  BACE1  messenger RNA. [ citation needed ]


While six-hour incubation with H 2 O 2  causes considerable demethylation of 5-mCpG sites, shorter times of H 2 O 2  incubation appear to promote other epigenetic alterations. Treatment of cells with H 2 O 2  for 30 minutes causes the mismatch repair protein heterodimer MSH2-MSH6 to recruit DNA methyltransferase 1 (DNMT1) to sites of some kinds of oxidative DNA damage. [ 139 ]  This could cause increased methylation of cytosines (epigenetic alterations) at these locations.


Jiang et al. [ 140 ]  treated  HEK 293 cells  with agents causing oxidative DNA damage, ( potassium bromate  (KBrO3) or  potassium chromate  (K2CrO4)).   Base excision repair  (BER) of oxidative damage occurred with the DNA repair enzyme  polymerase beta  localizing to oxidized guanines.  Polymerase beta is the main human polymerase in short-patch BER of oxidative DNA damage. Jiang et al. [ 140 ]  also found that polymerase beta recruited the  DNA methyltransferase  protein DNMT3b to BER repair sites. They then evaluated the methylation pattern at the single nucleotide level in a small region of DNA including the  promoter  region and the early transcription region of the  BRCA1  gene. Oxidative DNA damage from bromate modulated the DNA methylation pattern (caused epigenetic alterations) at CpG sites within the region of DNA studied. In untreated cells, CpGs located at −189, −134, −29, −19, +16, and +19 of the BRCA1 gene had methylated cytosines (where numbering is from the  messenger RNA  transcription start site, and negative numbers indicate nucleotides in the upstream  promoter  region). Bromate treatment-induced oxidation resulted in the loss of cytosine methylation at −189, −134, +16 and +19 while also leading to the formation of new methylation at the CpGs located at −80, −55, −21 and +8 after DNA repair was allowed. [ citation needed ]


At least four articles report the recruitment of  DNA methyltransferase 1 (DNMT1)  to sites of DNA double-strand breaks. [ 141 ] [ 142 ] [ 98 ] [ 143 ]   During  homologous recombinational repair (HR)  of the double-strand break, the involvement of DNMT1 causes the two repaired strands of DNA to have different levels of methylated cytosines.  One strand becomes frequently methylated at about 21  CpG sites  downstream of the repaired double-strand break.  The other DNA strand loses methylation at about six CpG sites that were previously methylated downstream of the double-strand break, as well as losing methylation at about five CpG sites that were previously methylated upstream of the double-strand break.  When the chromosome is replicated, this gives rise to one daughter chromosome that is heavily methylated downstream of the previous break site and one that is unmethylated in the region both upstream and downstream of the previous break site.  With respect to the gene that was broken by the double-strand break, half of the progeny cells express that gene at a high level and in the other half of the progeny cells expression of that gene is repressed.  When clones of these cells were maintained for three years, the new methylation patterns were maintained over that time period. [ 144 ]


In mice with a CRISPR-mediated homology-directed recombination insertion in their genome there were a large number of increased methylations of CpG sites within the double-strand break-associated insertion. [ 145 ]


Non-homologous end joining  (NHEJ) repair of a double-strand break can cause a small number of demethylations of pre-existing cytosine DNA methylations downstream of the repaired double-strand break. [ 142 ]   Further work by Allen et al. [ 146 ]  showed that NHEJ of a DNA double-strand break in a cell could give rise to some progeny cells having repressed expression of the gene harboring the initial double-strand break and some progeny having high expression of that gene due to epigenetic alterations associated with NHEJ repair.  The frequency of epigenetic alterations causing repression of a gene after an NHEJ repair of a DNA double-strand break in that gene may be about 0.9%. [ 98 ]


The basic processes of DNA repair are highly  conserved  among both  prokaryotes  and  eukaryotes  and even among  bacteriophages  ( viruses  which infect  bacteria ); however, more complex organisms with more complex genomes have correspondingly more complex repair mechanisms. [ 147 ]  The ability of a large number of protein  structural motifs  to catalyze relevant chemical reactions has played a significant role in the elaboration of repair mechanisms during evolution. For an extremely detailed review of hypotheses relating to the evolution of DNA repair, see. [ 148 ]


The  fossil record  indicates that single-cell life began to proliferate on the planet at some point during the  Precambrian  period, although exactly when recognizably modern life first emerged is unclear.  Nucleic acids  became the sole and universal means of encoding genetic information, requiring DNA repair mechanisms that in their basic form have been inherited by all extant life forms from their common ancestor. The emergence of Earth's oxygen-rich atmosphere (known as the " oxygen catastrophe ") due to  photosynthetic  organisms, as well as the presence of potentially damaging  free radicals  in the cell due to  oxidative phosphorylation , necessitated the evolution of DNA repair mechanisms that act specifically to counter the types of damage induced by  oxidative stress . The mechanism by which this came about, however, is unclear. [ citation needed ]


On some occasions, DNA damage is not repaired or is repaired by an error-prone mechanism that results in a change from the original sequence. When this occurs,  mutations  may propagate into the genomes of the cell's progeny. Should such an event occur in a  germ line  cell that will eventually produce a  gamete , the mutation has the potential to be passed on to the organism's offspring. The rate of  evolution  in a particular species (or, in a particular gene) is a function of the rate of mutation. As a consequence, the rate and accuracy of DNA repair mechanisms have an influence over the process of evolutionary change. [ 149 ]  DNA damage protection and repair does not influence the rate of adaptation by gene regulation and by recombination and selection of alleles.  On the other hand, DNA damage repair and protection does influence the rate of accumulation of irreparable, advantageous, code expanding, inheritable mutations, and slows down the evolutionary mechanism for expansion of the genome of organisms with new functionalities. The tension between evolvability and mutation repair and protection needs further investigation. [ citation needed ]


A technology named clustered regularly interspaced short palindromic repeat (shortened to  CRISPR -Cas9) was discovered in 2012.  The new technology allows anyone with molecular biology training to alter the genes of any species with precision, by inducing DNA damage at a specific point and then altering DNA repair mechanisms to insert new genes. [ 150 ]  It is cheaper, more efficient, and more precise than other technologies. With the help of CRISPR–Cas9, parts of a genome can be edited by scientists by removing, adding, or altering parts in a DNA sequence. [ citation needed ]


A  DNA repair-deficiency disorder  is a medical condition due to reduced functionality of  DNA repair .


DNA  repair defects can cause an  accelerated aging disease  or an  increased risk of cancer , or sometimes both.


DNA repair defects are seen in nearly all of the diseases described as   accelerated aging disease , in which various  tissues ,  organs  or systems of the human body  age  prematurely. Because the  accelerated aging diseases  display different aspects of aging, but never every aspect, they are often called  segmental progerias  by  biogerontologists . [ citation needed ]


Some examples of DNA repair defects causing  progeroid syndromes  in humans or mice are shown in Table 1.


Most of the DNA repair deficiency diseases show varying degrees of "accelerated aging" or cancer (often some of both). [ 37 ]  But elimination of any  gene  essential for   base excision repair  kills the  embryo —it is too lethal to display symptoms (much less symptoms of cancer or "accelerated aging"). [ 38 ]
Rothmund-Thomson syndrome and xeroderma pigmentosum display symptoms dominated by vulnerability to cancer, whereas progeria and Werner syndrome show the most features of "accelerated aging".  Hereditary nonpolyposis colorectal cancer  (HNPCC) is very often caused by a defective  MSH2  gene leading to defective  mismatch repair , but displays no symptoms of "accelerated aging". [ 39 ]  On the other hand, Cockayne Syndrome and trichothiodystrophy show mainly features of accelerated aging, but apparently without an increased risk of cancer [ 40 ]  Some DNA repair defects manifest as  neurodegeneration  rather than as cancer or "accelerated aging". [ 41 ]   (Also see the " DNA damage theory of aging " for a discussion of the evidence that DNA damage is the primary underlying cause of aging.)


Some biogerontologists question that such a thing as "accelerated aging" actually exists, at least partly on the grounds that all of the so-called accelerated aging diseases are segmental progerias. Many disease conditions such as  diabetes ,  high blood pressure , etc., are associated with increased  mortality . Without reliable  biomarkers of aging  it is hard to support the claim that a disease condition represents more than accelerated mortality. [ 42 ]


Against this position other biogerontologists argue that premature aging  phenotypes  are identifiable symptoms associated with mechanisms of molecular damage. [ 37 ]  The fact that these phenotypes are widely recognized justifies classification of the relevant diseases as "accelerated aging". [ 43 ]  Such conditions, it is argued, are readily distinguishable from  genetic diseases  associated with increased mortality, but not associated with an aging phenotype, such as  cystic fibrosis  and  sickle cell anemia . It is further argued that segmental aging phenotype is a natural part of aging insofar as genetic variation leads to some people being more disposed than others to  aging-associated diseases  such as  cancer  and  Alzheimer's disease . [ 44 ]


Individuals with an inherited impairment in  DNA repair  capability are often at increased risk of  cancer . [ 45 ]   When a  mutation  is present in a DNA repair gene, the repair gene will either not be expressed or be expressed in an altered form.  Then the repair function will likely be deficient, and, as a consequence, damages will tend to accumulate.  Such DNA damages can cause errors during  DNA synthesis  leading to mutations, some of which may give rise to cancer.  Germ-line DNA repair mutations that increase the risk of cancer are listed in the Table.


postmeiotic segregation increased 2 ( S. cerevisiae )


see also  Template:Congenital malformations and deformations of skin appendages ,  Template:Phakomatoses ,  Template:Pigmentation disorders ,  Template:DNA replication and repair-deficiency disorder


In  genetics , a  dynamic mutation  is an unstable heritable element where the probability of expression of a mutant phenotype is a function of the number of copies of the mutation. That is, the  replication  product ( progeny ) of a dynamic mutation has a different likelihood of mutation than its predecessor. These mutations, typically short sequences repeated many times,  give rise to numerous known diseases, including the  trinucleotide repeat disorders .


Robert I. Richards and Grant R. Sutherland called these phenomena, in the framework of  dynamical genetics , dynamic mutations. Triplet expansion is caused by slippage during  DNA replication . Due to the repetitive nature of the DNA sequence in these regions, 'loop out' structures may form during DNA replication while maintaining complementary base pairing between the parent strand and daughter strand being synthesized. If the loop out structure is formed from sequence on the daughter strand this will result in an increase in the number of repeats. However, if the loop out structure is formed on the parent strand a decrease in the number of repeats occurs. It appears that expansion of these repeats is more common than reduction. Generally the larger the expansion the more likely they are to cause disease or increase the severity of disease. This property results in the characteristic of  anticipation  seen in trinucleotide repeat disorders. Anticipation describes the tendency of age of onset to decrease and severity of symptoms to increase through successive generations of an affected family due to the expansion of these repeats.


Evolution by gene duplication  is an event by which a gene or part of a gene can have two identical copies that can not be distinguished from each other.   This phenomenon is understood to be an important source of novelty in evolution, providing for an expanded repertoire of molecular activities.  The underlying mutational event of duplication may be a conventional  gene duplication  mutation within a chromosome, or a larger-scale event involving whole chromosomes ( aneuploidy ) or whole genomes ( polyploidy ). A classic view, owing to  Susumu Ohno , [ 1 ]   which is known as Ohno model, he explains how duplication creates redundancy, the redundant copy accumulates beneficial mutations which provides fuel for innovation. [ 2 ]  Knowledge of evolution by gene duplication has advanced more rapidly in the past 15 years due to new genomic data, more powerful computational methods of comparative inference, and new evolutionary models.


Several models exist that try to explain how new cellular functions of genes and their encoded protein products evolve through the mechanism of duplication and divergence. Although each model can explain certain aspects of the evolutionary process, the relative importance of each aspect is still unclear. This page only presents which theoretical models are currently discussed in the literature. Review articles on this topic can be found at the bottom.


In the following, a distinction will be made between explanations for the short-term effects (preservation) of a gene duplication and its long-term outcomes.


Since a gene duplication occurs in only one cell, either in a single-celled organism or in the germ cell of a multi-cellular organism, its carrier (i.e. the organism) usually has to compete against other organisms that do not carry the duplication. If the duplication disrupts the normal functioning of an organism, the organism has a reduced reproductive success (or low  fitness ) compared to its competitors and will most likely die out rapidly. If the duplication has no effect on fitness, it might be maintained in a certain proportion of a population. In certain cases, the duplication of a certain gene might be immediately beneficial, providing its carrier with a fitness advantage.


The so-called ' dosage ' of a gene refers to the amount of mRNA transcripts and subsequently translated protein molecules produced from a gene per time and per cell.
If the amount of  gene product  is below its optimal level, there are two kinds of mutations that can increase dosage: increases in  gene expression  by  promoter  mutations and increases in gene copy number by gene duplication [ citation needed ] .


The more copies of the same (duplicated) gene a cell has in its genome, the more gene product can be produced simultaneously. Assuming that no regulatory feedback loops exist that automatically down-regulate gene expression, the amount of gene product (or gene dosage) will increase with each additional gene copy, until some upper limit is reached or sufficient gene product is available.


Furthermore, under positive selection for increased dosage, a duplicated gene could be immediately advantageous and quickly increase in frequency in a population. In this case, no further mutations would be necessary to preserve (or retain) the duplicates. However, at a later time, such mutations could still occur, leading to genes with different functions (see below).


Gene dosage effects after duplication can also be harmful to a cell and the duplication might therefore be selected against. For instance, when the  metabolic network  within a cell is fine-tuned so that it can only tolerate a certain amount of a certain gene product, gene duplication would offset this balance [ citation needed ] .


In cases of gene duplications that have no immediate fitness effect, a retention of the duplicate copy could still be possible if both copies accumulate mutations that for instance reduce the functional efficiency of the encoded proteins without inhibiting this function altogether. In such a case, the molecular function (e.g. protein/enzyme activity) would still be available to the cell to at least the extent that was available before duplication (now provided by proteins expressed from two gene loci, instead of one gene locus). However, the accidental loss of one gene copy might then be detrimental, since one copy of the gene with reduced activity would almost certainly lie below the activity that was available before duplication. [ citation needed ]


If a gene duplication is preserved, the most likely fate is that random mutations in one duplicate gene copy will eventually cause the gene to become non-functional
. [ 3 ]  Such non-functional remnants of genes, with detectable sequence  homology , can sometimes still be found in  genomes  and are called  pseudogenes .


Functional divergence  between the duplicate genes is another possible fate. There are several theoretical models that try to explain the mechanisms leading to divergence:


The term  neofunctionalization  was first coined by Force et al. 1999, [ 4 ]
but it refers to the general mechanism proposed by Ohno 1970. [ 1 ]  The long-term outcome of Neofunctionalization is that one copy retains the original (pre-duplication) function of the gene, while the second copy acquires a distinct function. It is also known as the MDN model, "mutation during non-functionality". The major criticism of this model is the high likelihood of non-functionalization, i.e. the loss of all functionality of the gene, due to random accumulation of mutations. [ 5 ] [ 6 ]


IAD stands for 'innovation, amplification, divergence' and aims to explain evolution of new gene functions while preserving its existing functions. [ 5 ]
Innovation, i.e. the establishment of a new molecular function, can occur via side-activities of genes and thus proteins this is called  Enzyme promiscuity . [ 7 ]   For example, enzymes can sometimes catalyse more than just one reaction, even though they usually are optimised for catalysing just one reaction. Such promiscuous protein functions, if they provide an advantage to the host organism, can then be amplified with additional copies of the gene. Such a rapid amplification is best known from bacteria that often carry certain genes on smaller non-chromosomal DNA molecules (called plasmids) which are capable of rapid replication. Any gene on such a plasmid is also replicated and the additional copies amplify the expression of the encoded proteins, and with it any promiscuous function. After several such copies have been made, and are also passed on to descendent bacterial cells, a few of these copies might accumulate mutations that eventually will lead to a side-activity becoming the main activity.


The IAD model have been previously tested in the lab by using bacterial enzyme with dual function as starting point. This enzyme is capable of catalyzing not only its original function, but also side function that can carried out by other enzyme.
By allowing the bacteria with this enzyme to evolve under selection to improve both activities (original and side) for several generations, it was shown that one ancestral bifunctional gene with poor activities (Innovation) evolved first by gene amplification to increase expression of the poor enzyme, and later accumulated more beneficial mutations that improved one or both of the activities that can be passed on to the next generation (divergence) [ 2 ]


"Subfunctionalization" was also first coined by Force et al. 1999. [ 4 ]  This model requires the ancestral (pre-duplication) gene to have several functions (sub-functions), which the descendant (post-duplication) genes specialise on in a complementary fashion. There are now  at least two different models that are labeled as subfunctionalization, "DDC" and "EAC".


DDC stands for "duplication-degeneration-complementation". This model was first introduced by Force et al. 1999. [ 4 ]  The first step is gene duplication. The gene duplication in itself is neither advantageous, nor deleterious, so it will remain at low frequency within a population of individuals that do not carry a duplication. According to DDC, this period of neutral drift may eventually lead to the complementary retention of sub-functions distributed over the two gene copies. This comes about by activity reducing (degenerative) mutations in both duplicates, accumulating over time periods and many generations. Taken together, the two mutated genes provide the same set of functions as the ancestral gene (before duplication). However, if one of the genes was removed, the remaining gene would not be able to provide the full set of functions and the host cell would likely suffer some detrimental consequences. Therefore, at this later stage of the process, there is a strong selection pressure against removing any of the two gene copies that arose by gene duplication. The duplication becomes permanently established in the genome of the host cell or organism.


EAC stands for "Escape from Adaptive Conflict". This name first appeared in a publication by Hittinger and Carroll 2007. [ 8 ]
The evolutionary process described by the EAC model actually begins before the gene duplication event. A singleton (not duplicated) gene evolves towards two beneficial functions simultaneously. This creates an "adaptive conflict" for the gene, since it is unlikely to execute each individual function with maximum efficiency. The intermediate evolutionary result could be a multi-functional gene and after a gene duplication its sub-functions could be carried out by specialised descendants of the gene. The result would be the same as under the DDC model, two functionally specialised genes (paralogs). In contrast to the DDC model, the EAC model puts more emphasis on the multi-functional pre-duplication state of the evolving genes and gives a slightly different explanation as to why the duplicated multi-functional genes would benefit from additional specialisation after duplication (because of the adaptive conflict of the multi-functional ancestor that needs to be resolved). Under EAC there is an assumption of a positive selection pressure driving evolution after gene duplication, whereas the DDC model only requires neutral ("undirected") evolution to take place, i.e. degeneration and complementation.


The  FET protein family  (also known as the TET protein family) [ 1 ]  consists of three similarly structured and functioning proteins. [ 2 ]  They and the  genes  in the  FET gene family  which encode them (i.e. form the  pre-messenger RNAs  that are converted to the  messenger RNAs  responsible for their production) are: [ 2 ] [ 3 ]   1)  the  EWSR1  protein encoded by the  EWSR1  gene (also termed the  Ewing sarcoma RNA  binding protein, EWS RNA binding protein 1,  or  bK984G1.4  gene) located at band 12.2 of the long (i.e. "q") arm of  chromosome 22 ; [ 4 ]   2)  the  FUS  (i.e. fused in sarcoma) protein encoded by the  FUS  gene (also termed the  FUS RNA binding protein, TLS, asTLS, ALS6, ETM4, FUS1, POMP75, altFUS , or  HNRNPP2  gene) located at  band  16 on the  short arm  of  chromosome 16 ; [ 5 ]  and  3)  the  TAF15  protein encoded by the  TAF15  gene (also termed the  TATA-box binding protein associated factor 15, Npl3, RBP56, TAF2N , or  TAFII68  gene) located at band 12 on the long arm of  chromosome 7 [ 6 ]  The FET in this protein family's name derives from the first letters of FUS, EWSR1, and TAF15. [ 7 ]


FET proteins are abundantly expressed in virtually all tissues examined. They are  RNA-binding proteins . [ 7 ]  By binding to their  RNA  targets, they contribute to the regulation of:  a)  the  transcription of genes  into  pre-messenger RNA , the  splicing  of pre-messenger RNA into  mature messenger RNA , and the transport of these RNAs between different areas of their parent cells;  b)  the processing of  micro-RNAs  that are involved in  RNA silencing  and  post-transcriptional regulation of gene expression ; and  3)  the  detection and repair of damaged DNA . Through these multiple, complex, and often incompletely understood actions, the FET family proteins regulate the  cellular expression  of diverse genes. [ 2 ]  However, the genes for FET proteins often undergo various types of  mutation . While these mutations and the diseases with which they are associated can be found in the Wikipedia pages on these diseases, this article focuses on one type of mutation, the  fusion gene  mutation. Fusion genes are formed from two previously independent genes that become united due to a  chromosome translocation ,  deletion  of some genetic material in a chromosome, or  chromosomal inversion . [ 1 ] [ 8 ]  For example, the  EWSR1-FL1 [ 9 ]  fusion gene is made by a chromosomal translocation which merges part of the  EWSR1  gene normally located on  band  12 of the  long (or "q") arm  of  chromosome 22  with part of the  FLI1   ETS transcription factor family  gene normally located on band 24 of the long arm of  chromosome 11 . The  EWSR1-FLI1  fusion gene encodes an EWS-FLI1  chimeric protein  which possesses unregulated and excessive FLI1  transcription factor  activity which it appears to contribute  to the development of  Ewing sarcomas . [ 10 ]  FET fusion genes have attracted recent interest because they have been found to be associated with, and may act to promote the development of, a wide range of  soft tissue   neoplasms  derived from  mesencyhmal tissue  cells. [ 3 ] [ 7 ] [ 11 ]  Detection of a FET gene–containing fusion gene is extremely helpful in diagnosing tumor types, [ 1 ] [ 11 ]  defining the  pathogenic mechanisms  by which these fusion proteins promote disorders, [ 3 ] [ 12 ]  and thereby identifying potential targets for treating these disorders. [ 2 ] [ 13 ]  The following are examples of these fusion genes' associations with malignant and benign neoplastic tumors. [ 1 ] [ 7 ]


In  population genetics , the  founder effect  is the loss of  genetic variation  that occurs when a new  population  is established by a very small number of individuals from a larger population. It was first fully outlined by  Ernst Mayr  in 1942, [ 1 ]  using existing theoretical work by those such as  Sewall Wright . [ 2 ]  As a result of the loss of genetic variation, the new population may be distinctively different, both  genotypically  and  phenotypically , from the parent population from which it is derived. In extreme cases, the founder effect is thought to lead to the  speciation  and subsequent  evolution  of new species. [ 3 ]


In the figure shown, the original population has nearly equal numbers of blue and red individuals. The three smaller founder populations show that one or the other color may predominate (founder effect), due to random sampling of the original population. A  population bottleneck  may also cause a founder effect, though it is not strictly a new population.


The founder effect occurs when a small group of migrants—not genetically representative of the population from which they came—establish in a new area. [ 4 ] [ 5 ]  In addition to founder effects, the new population is often  very small , so it shows increased sensitivity to  genetic drift , an increase in  inbreeding , and relatively low  genetic variation .


In  genetics , a  founder mutation  is a  mutation  that appears in the  DNA  of one or more individuals which are founders of a distinct population. Founder mutations initiate with changes that occur in the DNA and can be passed down to other generations. [ 6 ] [ 7 ]  Any organism—from a simple virus to something complex like a mammal—whose progeny carry its mutation has the potential to express the founder effect, [ 8 ]  for instance a goat [ 9 ] [ 10 ]  or a human. [ 11 ]


Founder mutations originate in long stretches of DNA on a single  chromosome ; indeed, the original  haplotype  is the whole chromosome. As the generations progress, the proportion of the haplotype that is common to all carriers of the mutation is shortened (due to  genetic recombination ). This shortening allows scientists to roughly estimate the age of the mutation. [ 12 ]