Bulky DNA adducts, increase

1162-65-8

OQIQSTLJSLGHID-WNWIJWBNSA-N

OQIQSTLJSLGHID-UHFFFAOYSA-N

Aflatoxin B1

AFB1 Cyclopenta[c]furo[3',2':4,5]furo[2,3-h][1]benzopyran-1,11-dione, 2,3,6a,9a-tetrahydro-4-methoxy-, (6aR,9aS)- (-)-Aflatoxin B1 1-Cyclopentene-1-carboxylic acid, 2-(3a,8a-dihydro-4-hydroxy-6-methoxyfuro[2,3-b]benzofuran-5-yl)-5-oxo-, delta-lactone 1H,11H-Cyclopenta[c]furo[3',2':4,5]furo[2,3-h][1]benzopyran-1,11-dione, 2,3,6a,9a-tetrahydro-4-methoxy-, (6aR,9aS)- 2,3,6A,9A-TETRAHYDRO-4-METHOXYCYCLOPENTA(C)FURO* 2,3,6aα,9aα-Tetrahydro-4-methoxycyclopenta[c]furo[2',3':4,5]furo[2,3-h]chromen-1,11-dion 2,3,6aα,9aα-tetrahydro-4-methoxycyclopenta[c]furo[2',3':4,5]furo[2,3-h]chromene-1,11-dione 2,3,6aalpha-9aalpha-tetrahidro-4-metoxiciclopenta[c]furo[2',3':4,5]furo[2,3-h]cromeno-1,11-diona Cyclopenta[c]furo[3',2':4,5]furo[2,3-h][1]benzopyran-1,11-dione, 2,3,6a,9a-tetrahydro-4-methoxy-, (6aR-cis)- Cyclopenta[c]furo[3',2':4,5]furo[2,3-h][1]benzopyran-1,11-dione, 2,3,6aalpha,9aalpha-tetrahydro-4-methoxy- NSC 529592

DTXSID9020035

313-67-7

BBFQZRXNYIEMAW-UHFFFAOYSA-N

Aristolochic acid

Aristolochic acid I Aristolochic acid A Tardolyt 8-Methoxy-3,4-methylenedioxy-10-nitrophenanthrene-1-carboxylic acid Phenanthro[3,4-d]-1,3-dioxole-5-carboxylic acid, 8-methoxy-6-nitro-

DTXSID0040969

CHEBI:16991

CHEBI deoxyribonucleic acid

FMA:74412

FMA Deoxyribonucleic acid

GO:0005694

GO chromosome

GO:0006281

GO DNA repair

NCI:C25830

NCI DNA Strand Break

D009154

MESH mutation

GO:0031052

GO chromosome breakage

WIKI increased

WIKI abnormal

Polycyclic aromatic hydrocarbons (PAHs)

2017-02-09T15:43:00

Aflatoxin B1

2016-11-29T18:42:26

Aristolochic acid

2017-05-09T15:56:55

Ionizing Radiation Ionizing radiation can vary in energy, dose, charge, and in the spatial distributions of energy transferred to other matter (linear energy transfer per unit length or LET) (ICRU 1970). At the same dose, low and high LET both generate energy deposition events, including many higher energy events (Goodhead and Nikjoo 1989). However, they differ in the spatial distribution and upper range of intensity of energy deposited. Lower LET such as gamma rays sparsely deposit many individual excitations or small clusters of excitations of low energy (Goodhead 1988). In contrast, high LET such as alpha particles have fewer tracks but readily transfer their energy to matter and therefore deposit their energy over a much smaller area (Goodhead 1994). Consequently, alpha and other high LET particles penetrate less deeply into tissue, interactions are densely focused on a narrow track, and individual energy depositions can be large (Goodhead 1988). These different energy deposition patterns can lead to differences in radiation effects including the pattern of DNA damage.

Exposure to ionizing radiation can come from natural and industrial sources. Space and terrestrial radiation includes a range of LET particles, while diagnostic radiation methods such as X-ray imaging, mammography and CT scans use low LET X-rays. Radiation therapy can use an external beam to direct radiation on a focused tissue area, or deposit solid or liquid radioactive materials in the body that release (mostly gamma) radiation internally. External radiotherapy typically uses X-rays but is moving towards higher LET charged particles such as protons and heavy ions (Durante, Orecchia et al. 2017).

2019-05-03T12:36:36

2019-05-07T12:12:13

Topoisomerase inhibitors

2019-05-19T20:21:24

Radiomimetic compounds

2019-05-19T20:21:42

WikiUser_25

Wikiuser: Cyauk human and other cells in culture

WCS_9606

common toxicological species human

10090

NCBI mouse

10116

NCBI rat

10036

NCBI Syrian golden hamster

9606

NCBI Homo sapiens

9913

NCBI cow

10090

NCBI Mus musculus

8090

NCBI medaka

4932

NCBI yeast

Bulky DNA adducts, increase

Molecular

Bulky DNA adducts are formed when activated genotoxic aromatic compounds interact with the nitrogenous bases of DNA. This occurs at various sites. The most common reactive sites for these adduct is C8, N7, N3 and N2 positions of guanine, the N7, N6, N3, and N1 positions of adenine, the N3, N4, and O2 positions of cytosine, and the N3, O2, and O4 positions of thymine (As reviewed by Hwa Yun et al., 2020). The position of the adduct depends on the chemical structure of the activated aromatic compound. Some adducts are not stable, but some can persist. For example, the most harmful adducts formed by benzo(a)pyrene are from radicals that bind to the N7 and C8 of purines (IARC., 2012). Aristolochic Acid forms adducts at N6 of adenine and Aflatoxin B1 forms adducts at the N7 of Guanine (Arlt et al., 2002). This KE describes an increase in Bulky adducts. These adducts can cause depurination, transversions which in turn cause DNA damage and chromosome aberrations.

<h3 style="text-align:justify">Quantification of Bulky DNA Adducts</h3>   <ul> <li style="text-align:justify">32P Post labelling is used for the detection of DNA adducts (for PAHs and also Aristolochic Acid) (Gupta et al., 1982; Klaene et al., 2013; Phillips and Arlt., 2014)</li> </ul> <ul style="list-style-type:circle; margin-left:40px"> <li style="text-align:justify">The DNA is isolated using the standard methods and digested into 3′-deoxynucleoside monophosphates. A 32P-orthophosphate from [gamma-32P] ATP is used to radiolabel the adducts in a reaction catalyzed by T4 polynucleotide kinase. </li> <li style="text-align:justify">The radiolabelled nucleotides are separated and detected by thin-layer chromatography. They are quantified by scintillation counting. This is usually used to detect bulky adducts. </li> <li style="text-align:justify">Nuclease P1 can be used for enrichment with PAH adducts. Using 1-Butanol to extract the adducted molecules before labelling is another optimization method and it works well with aromatic amines. </li> </ul>   <ul> <li>CometChip assay (modified by adding DNA synthesis inhibitors (Ngo et al.,2020)</li> </ul> <ul style="list-style-type:circle; margin-left:40px"> <li style="text-align:justify">This variation of the assay uses DNA synthesis inhibitors to convert bulky lesions into detectable SSBs.</li> <li style="text-align:justify">HepaCometChip uses Hydroxyurea (HU) and 1-β-d-arabinofuranosyl cytosine (AraC) to detect SSBs formed from bulky adducts in the presence of the high metabolism of HepaRG™ cells.</li> <li style="text-align:justify">HU inhibits the enzyme ribonucleotide reductase. This enzyme mediates the synthesis of deoxyribonucleotides (dNTPs). When it is inhibited dNTPs are depleted which inhibits NER. </li> <li style="text-align:justify">AraC’s structure allows it to be incorporated into DNA and interrupts DNA elongation. </li> <li style="text-align:justify">HU and AraC delay the removal of NER and SSB intermediates. The prolonged presence of NER intermediates are indicators of bulky lesions and can be observed as comet detectable SSBs.</li> <li style="text-align:justify">The number of bulky lesions is then measured by detecting the % of DNA found in the tail of the comet compared to untreated samples. Percentage DNA in the comet tail is proportional to the level of strand breaks.</li> </ul>   Other methods for adduct detection   A variety of other methods are available to measure bulky DNA adducts including Isotope dilution mass spectrometry (MS) liquid chromatography mass spectrometry (LC–MS), gas chromatography mass spectrometry (GC–MS), capillary electrophoresis mass spectrometry (CE–MS). (Long et al., 2018; Fischer et al., 2018; Chang et al., 2017; Woo et al., 2011) <ul> <li><a href="https://aopwiki.org/events/373" style="color:#0563c1; text-decoration:underline">https://aopwiki.org/events/373</a> </li> </ul>

Bulky adducts can occur in virtually any cell type or organism, as long as the organism/cell type has the xenobiotic metabolism enzymes necessary to activate pro-mutagens when required. Bulky adducts have been detected both in vitro (various cell lines) and in vivo in mammalian cells (human, mouse, rat), and can occur in males and females at any life stage.

Not Specified Unspecific

Not Specified

All life stages

Not Specified Not Specified Not Specified Not Specified

Arlt VM, Stiborova M, Schmeiser HH. Mutagenesis. 2002; 17:265–277. Barnes, J. L., Zubair, M., John, K., Poirier, M. C., & Martin, F. L. (2018). Carcinogens and DNA damage. Biochemical Society transactions, 46(5), 1213–1224. <a href="https://doi.org/10.1042/BST20180519" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1042/BST20180519</a>   Grollman, A. P., Shibutani, S., Moriya, M., Miller, F., Wu, L., Moll, U., Suzuki, N., Fernandes, A., Rosenquist, T., Medverec, Z., Jakovina, K., Brdar, B., Slade, N., Turesky, R. J., Goodenough, A. K., Rieger, R., Vukelić, M., & Jelaković, B. (2007). Aristolochic acid and the etiology of endemic (Balkan) nephropathy. Proceedings of the National Academy of Sciences of the United States of America, 104(29), 12129–12134. <a href="https://doi.org/10.1073/pnas.0701248104" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1073/pnas.0701248104</a>   Groopman, J. D., Croy, R. G., & Wogan, G. N. (1981). In vitro reactions of aflatoxin B1-adducted DNA. Proceedings of the National Academy of Sciences, 78(9), 5445-5449. Gupta, R. C., Reddy, M. V., & Randerath, K. (1982). 32 P-postlabeling analysis of non-radioactive aromatic carcinogen—DNA adducts. Carcinogenesis, 3(9), 1081-1092.   Hanahan D, Weinberg RA. Hallmarks of cancer: the next generation. Cell. 2011 Mar 4;144(5):646-74. doi: 10.1016/j.cell.2011.02.013. PMID: 21376230.   Hwa Yun, B., Guo, J., Bellamri, M., & Turesky, R. J. (2020). DNA adducts: Formation, biological effects, and new biospecimens for mass spectrometric measurements in humans. Mass spectrometry reviews, 39(1-2), 55-82.   IARC Some non-heterocyclic polycyclic aromatic hydrocarbons and some related exposures. IARC Monogr Eval Carcinog Risks Hum. 2010;92:1–853. <a href="https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4781319/" style="color:#0563c1; text-decoration:underline">https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4781319/</a>   IARC Working Group on the Evaluation of Carcinogenic Risks to Humans. Chemical Agents and Related Occupations. Lyon (FR): International Agency for Research on Cancer; 2012. (IARC Monographs on the Evaluation of Carcinogenic Risks to Humans, No. 100F.) BENZO[a]PYRENE. Available from: <a href="https://www.ncbi.nlm.nih.gov/books/NBK304415/" style="color:#0563c1; text-decoration:underline">https://www.ncbi.nlm.nih.gov/books/NBK304415/</a>   Jessica L. Barnes, Maria Zubair, Kaarthik John, Miriam C. Poirier, Francis L. Martin; Carcinogens and DNA damage. Biochem Soc Trans 19 October 2018; 46 (5): 1213–1224. doi: <a href="https://doi.org/10.1042/BST20180519" style="color:#0563c1; text-decoration:underline" target="_blank">https://doi.org/10.1042/BST20180519</a>   Li, X. L., Guo, X. Q., Wang, H. R., Chen, T., & Mei, N. (2020). Aristolochic Acid-Induced Genotoxicity and Toxicogenomic Changes in Rodents. World journal of traditional Chinese medicine, 6(1), 12–25. <a href="https://doi.org/10.4103/wjtcm.wjtcm_33_19" style="color:#0563c1; text-decoration:underline">https://doi.org/10.4103/wjtcm.wjtcm_33_19</a>   McDaniel, L. P., Elander, E. R., Guo, X., Chen, T., Arlt, V. M., & Mei, N. (2012). Mutagenicity and DNA adduct formation by aristolochic acid in the spleen of Big Blue® rats. Environmental and molecular mutagenesis, 53(5), 358-368.   Ngo, L. P., Owiti, N. A., Swartz, C., Winters, J., Su, Y., Ge, J., Xiong, A., Han, J., Recio, L., Samson, L. D., & Engelward, B. P. (2020). Sensitive CometChip assay for screening potentially carcinogenic DNA adducts by trapping DNA repair intermediates. Nucleic acids research, 48(3), e13. <a href="https://doi.org/10.1093/nar/gkz1077" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1093/nar/gkz1077</a>   Phillips, D. H., & Arlt, V. M. (2014). 32 P-Postlabeling Analysis of DNA Adducts. In Molecular Toxicology Protocols (pp. 127-138). Humana Press, Totowa, NJ.   Yun, B. H., Sidorenko, V. S., Rosenquist, T. A., Dickman, K. G., Grollman, A. P., & Turesky, R. J. (2015). New approaches for biomonitoring exposure to the human carcinogen aristolochic acid. Toxicology research, 4(4), 763-776. AOPWiki 2021-05-17T09:10:50

2021-05-17T09:48:28

Inadequate DNA repair

Cellular

DNA lesions may result from the formation of DNA adducts (i.e., covalent modification of DNA by chemicals), or by the action of agents such as radiation that may produce strand breaks or modified nucleotides within the DNA molecule. These DNA lesions are repaired through several mechanistically distinct pathways that can be categorized as follows: <ol> <li>Damage reversal acts to reverse the damage without breaking any bonds within the sugar phosphate backbone of the DNA. The most prominent enzymes associated with damage reversal are photolyases (Sancar, 2003) that can repair UV dimers in some organisms, and O6-alkylguanine-DNA alkyltransferase (AGT) (Pegg 2011) and oxidative demethylases (Sundheim et al., 2008), which can repair some types of alkylated bases.</li> <li>Excision repair involves the removal of a damaged nucleotide(s) through cleavage of the sugar phosphate backbone followed by re-synthesis of DNA within the resultant gap. Excision repair of DNA lesions can be mechanistically divided into:  a) Base excision repair (BER) (Dianov and Hübscher, 2013), in which the damaged base is removed by a damage-specific glycosylase prior to incision of the phosphodiester backbone at the resulting abasic site. This leads to an intermediate that contains a DNA strand break, whereby DNA ligase is then recruited to seal the ends of the DNA. b) Nucleotide excision repair (NER) (Schärer, 2013), in which the DNA strand containing the damaged nucleotide is incised at sites several nucleotides 5’ and 3’ to the site of damage, and a polynucleotide containing the damaged nucleotide is removed prior to DNA resynthesis within the resultant gap and sealing of the ends by DNA ligase.   c) Mismatch repair (MMR) (Li et al., 2016)  which does not act on DNA lesions but does recognize mispaired bases resulting from replication errors. In MMR the strand containing the misincorporated base is removed prior to DNA resynthesis. The major pathway that removes oxidative DNA damage is base excision repair (BER), which can be either monofunctional or bifunctional; in mammals, a specific DNA glycosylase (OGG1: 8-Oxoguanine glycosylase) is responsible for excision of 8-oxoguanine (8-oxoG) and other oxidative lesions (Hu et al., 2005; Scott et al., 2014; Whitaker et al., 2017). We note that long-patch BER is used for the repair of clustered oxidative lesions, which uses several enzymes from DNA replication pathways (Klungland and Lindahl, 1997). These pathways are described in detail in various reviews e.g., (Whitaker et al., 2017).  </li> <li>Single strand break repair (SSBR) involves different proteins and enzymes depending on the origin of the SSB (e.g., produced as an intermediate in excision repair or due to direct chemical insult) but the same general steps of repair are taken for all SSBs: detection, DNA end processing, synthesis, and ligation (Caldecott, 2014). Poly-ADP-ribose polymerase1 (PARP1) detects and binds unscheduled SSBs (i.e., not deliberately induced during excision repair) and synthesizes PAR as a signal to the downstream factors in repair. PARP1 is not required to initiate SSBR of BER intermediates. The XRCC1 protein complex is then recruited to the site of damage where a common DNA intermediate as BER was generated, and acts as a scaffold for proteins and enzymes required for repair. Depending on the nature of the damaged termini of the DNA strand, different enzymes are required for end processing to generate the substrates that DNA polymerase β (Polβ; short patch repair) or Pol δ/ε (long patch repair) can bind to synthesize over the gap, although end processing is generally done by polynucleotide kinase. Synthesis in long-patch repair displaces a single stranded flap which is excised by flap endonuclease 1 (FEN1). In short-patch repair, the XRCC1/Lig3α complex joins the two ends after synthesis. In long-patch repair, the PCNA/Lig1 complex ligates the ends. (Caldecott, 2014). </li> <li>Double strand break repair (DSBR) is necessary to preserve genomic integrity when breaks occur in both strands of a DNA molecule. There are two major pathways for DSBR: homologous recombination (HR), which operates primarily during the S phase of dividing cell types, and nonhomologous end joining (NHEJ), which can function in both dividing and non-dividing cell types. No repair occurs in the M phase (Teruaki Iyama and David M. Wilson III, 2013). DNA repair in mitosis is controversial (Mladenov et al., 2023).</li> </ol> Complex lesions can be created by a single mutagen and can be more difficult to repair, as the mechanism behind how different repair pathways cooperate to address this is still unclear (Aleksandrov et al., 2018). In higher eukaryotes such as mammals, NHEJ is usually the preferred pathway for DNA DSBR. Its use, however, is dependent on the cell type, the gene locus, and the nuclease platform (Miyaoka et al., 2016). The use of NHEJ is also dependent on the cell cycle; NHEJ is generally not the pathway of choice when the cell is in the late S or G2 phase of the cell cycle, or in mitotic cells when the sister chromatid is directly adjacent to the double-strand break (DSB) (Lieber et al., 2003). In these cases, the HR pathway is commonly used for repair of DSBs. Despite this, NHEJ is still used more commonly than HR in human cells. Classical NHEJ (C-NHEJ) is the most common NHEJ repair mechanism, but alternative NHEJ (alt-NHEJ) can also occur, especially in the absence of C-NHEJ and HR. The process of C-NHEJ in humans requires at least seven core proteins: Ku70, Ku86, DNA-dependent protein kinase complex (DNA-PKcs ), Artemis, X-ray cross-complementing protein 4 (XRCC4), XRCC4-like factor (XLF), and DNA ligase IV (Boboila et al., 2012). When DSBs occur, the Ku proteins, which have a high affinity for DNA ends, will bind to the break site and form a heterodimer. This protects the DNA from exonucleolytic attack and acts to recruit DNA-PKcs, the catalytic subunit, thus forming a trimeric complex on the ends of the DNA strands. Alternative NHEJ, or alt NHEJ, uses small similar sequences in two broken DNA ends to join them together. Unlike the usual repair method (cNHEJ), aNHEJ doesn't need specific proteins like LIG4 and KU. Instead, it relies on the MRN complex to process the breaks. However, alt NHEJ tends to cause mutations by adding or removing bits of DNA during the repair (Chaudhuri and Nussenzweig, 2017). The kinase activity of DNA-PKcs is then triggered, causing DNA-PKcs to auto-phosphorylate and thereby lose its kinase activity; the now phosphorylated DNA-PKcs dissociates from the DNA-bound Ku proteins. The free DNA-PKcs phosphorylates Artemis, an enzyme that possesses 5’-3’ exonuclease and endonuclease activity in the presence of DNA-PKcs and ATP. Artemis is responsible for ‘cleaning up’ the ends of the DNA. For 5’ overhangs, Artemis nicks the overhang, generally leaving a blunt duplex end. For 3’ overhangs, Artemis will often leave a four- or five-nucleotide single stranded overhang (Pardo et al., 2009; Fattah et al., 2010; Lieber et al., 2010). Next, the XLF and XRCC4 proteins form a complex which makes a channel to bind DNA and aligns the ends for efficient ligation via DNA ligase IV (Hammel et al., 2011). The process of alt-NHEJ is less well understood than C-NHEJ and is a lower fidelity mechanism.  Alt-NHEJ is known to involve slightly different core proteins than C-NHEJ and required microhomology repeats, but the steps of the pathway are essentially the same between the two processes (reviewed in Chiruvella et al., 2013). It is established, however, that alt-NHEJ is more error-prone in nature than C-NHEJ, which contributes to incorrect DNA repair. Alt-NHEJ is thus considered primarily to be a backup repair mechanism (reviewed in Chiruvella et al., 2013).  In contrast to NHEJ, HR takes advantage of similar or identical DNA sequences to repair DSBs and is not error-prone (Sung and Klein, 2006). The initiating step of HR is the creation of a 3’ single strand DNA (ss-DNA) overhang. Combinases such as RecA and Rad51 then bind to the ss-DNA overhang, and other accessory factors, including Rad54, help recognize and invade the homologous region on another DNA strand. From there, DNA polymerases are able to elongate the 3’ invading single strand and resynthesize the broken DNA strand using the corresponding sequence on the homologous strand.   Fidelity of DNA Repair Most DNA repair pathways are extremely efficient. However, in principal, all DNA repair pathways can be overwhelmed when the DNA lesion burden exceeds the capacity of a given DNA repair pathway to recognize and remove the lesion. Exceeded repair capacity may lead to toxicity or mutagenesis following DNA damage. Apart from extremely high DNA lesion burden, inadequate repair may arise through several different specific mechanisms. For example, during repair of DNA containing O6-alkylguanine adducts, AGT irreversibly binds a single O6-alkylguanine lesion and as a result is inactivated (this is termed suicide inactivation, as its own action causes it to become inactivated). Thus, the capacity of AGT to carry out alkylation repair can become rapidly saturated when the DNA repair rate exceeds the de novo synthesis of AGT (Pegg, 2011). A second mechanism relates to cell specific differences in the cellular levels or activity of some DNA repair proteins. For example, XPA is an essential component of the NER complex. The level of XPA that is active in NER is low in the testes, which may reduce the efficiency of NER in testes as compared to other tissues (Köberle et al., 1999). Likewise, both NER and BER have been reported to be deficient in cells lacking functional p53 (Adimoolam and Ford, 2003; Hanawalt et al., 2003; Seo and Jung, 2004). A third mechanism relates to the importance of the DNA sequence context of a lesion in its recognition by DNA repair enzymes. For example, 8-oxoguanine (8-oxoG) is repaired primarily by BER; the lesion is initially acted upon by a bifunctional glycosylase, OGG1, which carries out the initial damage recognition and excision steps of 8-oxoG repair. However, the rate of excision of 8-oxoG is modulated strongly by both chromatin components (Menoni et al., 2012) and DNA sequence context (Allgayer et al., 2013) leading to significant differences in the repair of lesions situated in different chromosomal locations. DNA repair is also remarkably error-free. However, misrepair can arise during repair under some circumstances. DSBR is notably error prone, particularly when breaks are processed through NHEJ, during which partial loss of genome information is common at the site of the double strand break (Iyama and Wilson, 2013). This is because NHEJ rejoins broken DNA ends without the use of extensive homology; instead, it uses the microhomology present between the two ends of the DNA strand break to ligate the strand back into one. When the overhangs are not compatible, however, indels (insertion or deletion events), duplications, translocations, and inversions in the DNA can occur. These changes in the DNA may lead to significant issues within the cell, including alterations in the gene determinants for cellular fatality (Moore et al., 1996). Activation of mutagenic DNA repair pathways to withstand cellular or replication stress either from endogenous or exogenous sources can promote cellular viability, albeit at a cost of increased genome instability and mutagenesis (Fitzgerald et al., 2017). These salvage DNA repair pathways including, Break-induced Replication (BIR) and Microhomology-mediated Break-induced Replication (MMBIR). BIR repairs one-ended DSBs and has been extensively studied in yeast as well as in mammalian systems. BIR and MMBIR are linked with heightened levels of mutagenesis, chromosomal rearrangements and ensuing genome instability (Deem et al., 2011; Sakofsky et al., 2015; Saini et al., 2017; Kramara et al., 2018). In mammalian genomes BIR-like synthesis has been proposed to be involved in late stage Mitotic DNA Synthesis (MiDAS) that predominantly occurs at so-called Common Fragile Sites (CFSs) and maintains telomere length under s conditions of replication stress that serve to promote cell viability (Minocherhomji et al., 2015; Bhowmick et al., 2016; Dilley et al., 2016).        Misrepair may also occur through other repair pathways. Excision repair pathways require the resynthesis of DNA and rare DNA polymerase errors during gap resynthesis will result in mutations (Brown et al., 2011). Errors may also arise during gap resynthesis when the strand that is being used as a template for DNA synthesis contains DNA lesions (Kozmin and Jinks-Robertson, 2013). In addition, it has been shown that sequences that contain tandemly repeated sequences, such as CAG triplet repeats, are subject to expansion during gap resynthesis that occurs during BER of 8-oxoG damage (Liu et al., 2009).

There is no test guideline for this event. The event is usually inferred from measuring the retention of DNA adducts or the creation of mutations as a measure of lack of repair or incorrect repair. These ‘indirect’ measures of its occurrence are crucial to determining the mechanisms of genotoxic chemicals and for regulatory applications (i.e., determining the best approach for deriving a point of departure). More recently, a fluorescence-based multiplex flow-cytometric host cell reactivation assay (FM-HCR) has been developed to directly measure the ability of human cells to repair plasmid reporters (Nagel et al., 2014). Indirect Measurement In somatic and spermatogenic cells, measurement of DNA repair is usually inferred by measuring DNA adduct formation/removal. Insufficient repair is inferred from the retention of adducts and from increasing adduct formation with dose. Insufficient DNA repair is also measured by the formation of increased numbers of mutations and alterations in mutation spectrum. The methods will be specific to the type of DNA adduct that is under study. Some EXAMPLES are given below for alkylated DNA. DOSE-RESPONSE CURVE FOR ALKYL ADDUCTS/MUTATIONS: It is important to consider that some adducts are not mutagenic at all because they are very effectively repaired. Others are effectively repaired, but if these repair processes become overwhelmed mutations begin to occur. The relationship (shape of dose-response curve) between exposure to mutagenic agents and mutations provide an indication of whether the removal of adducts occurs, and whether it is more efficient at low doses. Sub-linear dose-response curves (hockey stick or j-shape curves) for mutation induction indicates that adducts are not converted to mutations at low doses. This suggests the effective repair of adducts at low doses, followed by saturation of repair at higher doses (Clewell et al., 2019). Thus, measurement of a clear point of inflection in the dose-response curve for mutations suggests that repair does occur, at least to some extent, at low dosees but that reduced repair efficiency arises above the inflection point. A lack of increase in mutation frequencies (i.e., flat line for dose-response) for a compound showing a dose-dependent increase in adducts would imply that the adducts formed are either not mutagenic or are effectively repaired. RETENTION OF ALKYL ADDUCTS: Alkylated DNA can be found in cells long after exposure has occurred. This indicates that repair has not effectively removed the adducts. For example, DNA adducts have been measured in hamster and rat spermatogonia several days following exposure to alkylating agents, indicating lack of repair (Seiler et al., 1997; Scherer et al., 1987). MUTATION SPECTRUM: Shifts in mutation spectrum (i.e., the specific changes in the DNA sequence) following a chemical exposure (relative to non-exposed mutation spectrum) indicates that repair was not operating effectively to remove specific types of lesions. The shift in mutation spectrum is indicative of the types of DNA lesions (target nucleotides and DNA sequence context) that were not repaired. For example, if a greater proportion of mutations occur at guanine nucleotides in exposed cells, it can be assumed that the chemical causes DNA adducts on guanine that are not effectively repaired. Direct Measurement Nagel et al. (2014) we developed a fluorescence-based multiplex flow-cytometric host cell reactivation assay (FM-HCR) to measures the ability of human cells to repair plasmid reporters. These reporters contain different types and amounts of DNA damage and can be used to measure repair through by NER, MMR, BER, NHEJ, HR and MGMT. Please refer to the table below for additional details and methodologies for detecting DNA damage and repair. <table border="1" cellpadding="1" cellspacing="1" style="height:2082px; width:629px"> <tbody> <tr> <td style="background-color:#eeeeee; text-align:center">Assay Name</td> <td style="background-color:#eeeeee; text-align:center">References</td> <td style="background-color:#eeeeee; text-align:center">Description</td> <td style="background-color:#eeeeee; text-align:center">DNA Damage/Repair Being Measured</td> <td style="background-color:#eeeeee; text-align:center">OECD Approved Assay</td> </tr> <tr> <td style="text-align:center">Dose-Response Curve for Alkyl Adducts/ Mutations</td> <td> Lutz 1991   Clewell 2016 </td> <td style="text-align:center">Creation of a curve plotting the stressor dose and the abundance of adducts/mutations; Characteristics of the resulting curve can provide information on the efficiency of DNA repair</td> <td> Alkylation, oxidative damage, or DSBs </td> <td style="text-align:center">N/A</td> </tr> <tr> <td style="text-align:center">Retention of Alkyl Adducts</td> <td> Seiler 1997   Scherer 1987 </td> <td style="text-align:center">Examination of DNA for alkylation after exposure to an alkylating agent; Presence of alkylation suggests a lack of repair</td> <td style="text-align:center">Alkylation</td> <td style="text-align:center">N/A</td> </tr> <tr> <td style="text-align:center">Mutation Spectrum</td> <td style="text-align:center">Wyrick 2015</td> <td style="text-align:center">Shifts in the mutation spectrum after exposure to a chemical/mutagen relative to an unexposed subject can provide an indication of DNA repair efficiency, and can inform as to the type of DNA lesions present</td> <td> Alkylation, oxidative damage, or DSBs </td> <td style="text-align:center">N/A</td> </tr> <tr> <td style="text-align:center">DSB Repair Assay (Reporter constructs)</td> <td style="text-align:center">Mao et al., 2011</td> <td style="text-align:center">Transfection of a GFP reporter construct (and DsRed control) where the GFP signal is only detected if the DSB is repaired; GFP signal  is quantified using fluorescence microscopy or flow cytometry</td> <td style="text-align:center">DSBs</td> <td style="text-align:center">N/A</td> </tr> <tr> <td style="text-align:center">Primary Rat Hepatocyte DNA Repair Assay</td> <td> Jeffrey and Williams, 2000   Butterworth et al., 1987 </td> <td style="text-align:center">Rat primary hepatocytes are cultured with a 3H-thymidine solution in order to measure DNA synthesis in response to a stressor in non-replicating cells; Autoradiography is used to measure the amount of 3H incorporated in the DNA post-repair</td> <td style="text-align:center">Unscheduled DNA synthesis in response to DNA damage</td> <td style="text-align:center">N/A</td> </tr> <tr> <td style="text-align:center">Repair synthesis measurement by 3H-thymine incorporation</td> <td style="text-align:center">Iyama and Wilson, 2013</td> <td style="text-align:center">Measure DNA synthesis in non-dividing cells as indication of gap filling during excision repair</td> <td style="text-align:center">Excision repair</td> <td style="text-align:center">N/A</td> </tr> <tr> <td style="text-align:center">Comet Assay with Time-Course</td> <td> Olive et al., 1990   Trucco et al., 1998 - Dunkenberger et al., 2022  </td> <td style="text-align:center">Comet assay is performed with a time-course under alkaline conditions to detect SSBs and DSBs. Quantity of DNA in the tail should decrease as DNA repair progresses</td> <td style="text-align:center">DSBs</td> <td style="text-align:center"> <a href="https://read.oecd-ilibrary.org/environment/test-no-489-in-vivo-mammalian-alkaline-comet-assay_9789264264885-en">Yes</a> (No. 489)</td> </tr> <tr> <td style="text-align:center">Flow Cytometry   </td> <td>Corneo et al., 2007   </td> <td style="text-align:center">The alt-NHEJ flow cytometer method involves utilizing an extrachromosomal substrate. Green fluorescent protein (GFP) expression is indicative of successful alt-NHEJ activity, contingent on the removal of 10 nucleotides from each end of the DNA and subsequent rejoining within a 9-nucleotide microhomology region. This approach provides a quantitative and visual means to measure the efficiency of alternative non-homologous end joining in cellular processes.   </td> <td style="text-align:center">Alt NHEJ</td> <td style="text-align:center">No</td> </tr> <tr> <td style="text-align:center">Pulsed Field Gel Electro-phoresis (PFGE) with Time-Course</td> <td style="text-align:center">Biedermann et al., 1991</td> <td style="text-align:center">PFGE assay with a time-course; Quantity of small DNA fragments should decrease as DNA repair  progresses</td> <td style="text-align:center">DSBs</td> <td style="text-align:center">N/A</td> </tr> <tr> <td> Fluorescence -Based Multiplex Flow-Cytometric Host Reactivation Assay (FM-HCR) </td> <td style="text-align:center">Nagel et al., 2014</td> <td style="text-align:center">Measures the ability of human cells to repair plasma reporters, which contain different types and amounts of DNA damage; Used to measure repair processes including HR, NHEJ, BER, NER, MMR, and MGMT</td> <td style="text-align:center">HR, NHEJ, BER, NER, MMR, or MGMT</td> <td style="text-align:center">N/A</td> </tr> <tr> <td>Alkaline Unwinding Assay with Time Course </td> <td style="text-align:center">Nacci et al. 1991 </td> <td style="text-align:center">DNA is stored in alkaline solutions with DNA-specific dye and allowed to unwind following removal from tissue, increased strand damage associated with increased unwinding. Samples analyzed at different time points to compare remaining damage following repair opportunities </td> <td style="text-align:center">DSBs </td> <td style="text-align:center">Yes (No. 489) </td> </tr> <tr> <td>Sucrose Density Gradient Centrifugation with Time Course </td> <td style="text-align:center">Larsen et al. 1982 </td> <td style="text-align:center">Strand breaks alter the molecular weight of the DNA piece. DNA in alkaline solution centrifuged into sugar density gradient, repeated set time apart. The less DNA breaks identified in the assay repeats, the more repair occurred </td> <td style="text-align:center">SSBs </td> <td style="text-align:center">N/A</td> </tr> <tr> <td>y-H2AX Foci Staining with Time Course </td> <td style="text-align:center"> Mariotti et al. 2013  Penninckx et al. 2021  </td> <td style="text-align:center">Histone H2AX is phosphorylated in the presence of DNA strand breaks, the rate of its disappearance over time is used as a measure of DNA repair </td> <td style="text-align:center">DSBs </td> <td style="text-align:center">N/A</td> </tr> <tr> <td>Alkaline Elution Assay with Time Course </td> <td style="text-align:center">Larsen et al. 1982 </td> <td style="text-align:center">DNA with strand breaks elute faster than DNA without, plotted against time intervals to determine the rate at which strand breaks repair </td> <td style="text-align:center">SSBs </td> <td style="text-align:center">N/A</td> </tr> <tr> <td>53BP1 foci Detection with Time Course </td> <td style="text-align:center">Penninckx et al. 2021 </td> <td style="text-align:center">53BP1 is recruited to the site of DNA damage, the rate at which its level decreases over time is used to measure DNA repair </td> <td style="text-align:center">DSBs</td> <td style="text-align:center">N/A </td> </tr> </tbody> </table>

The retention of adducts has been directly measured in many different types of eukaryotic somatic cells (in vitro and in vivo). In male germ cells, work has been done on hamsters, rats and mice. The accumulation of mutation and changes in mutation spectrum has been measured in mice and human cells in culture. Theoretically, saturation of DNA repair occurs in every species (prokaryotic and eukaryotic). The principles of this work were established in prokaryotic models. Nagel et al. (2014) have produced an assay that directly measures DNA repair in human cells in culture. NHEJ is primarily used by vertebrate multicellular eukaryotes, but it also been observed in plants. Furthermore, it has recently been discovered that some bacteria (Matthews et al., 2014) and yeast (Emerson et al., 2016) also use NHEJ. In terms of invertebrates, most lack the core DNA-PKcs and Artemis proteins; they accomplish end joining by using the RA50:MRE11:NBS1 complex (Chen et al., 2001).  HR occurs naturally in eukaryotes, bacteria, and some viruses (Bhatti et al., 2016). Taxonomic applicability: Inadequate DNA repair is applicable to all species, as they all contain DNA (White & Vijg, 2016).   Life stage applicability: This key event is not life stage specific as any life stage can have poor repair, though as individuals age their repair process become less effective (Gorbunova & Seluanov, 2016).  Sex applicability: There is no evidence of sex-specificity for this key event, with initial rate of DNA repair not significantly different between sexes (Trzeciak et al., 2008).  Evidence for perturbation by a stressor: Multiple studies demonstrate that inadequate DNA repair can occur as a result of stressors such as ionizing and non-ionizing radiation, as well as chemical agents (Kuhne et al., 2005; Rydberg et al., 2005; Dahle et al., 2008; Seager et al., 2012; Wilhelm, 2014; O’Brien et al., 2015).

High Unspecific

High

All life stages

High Moderate Moderate High Low

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2024-03-08T12:15:51

Increase, DNA strand breaks

Molecular

DNA strand breaks are a type of damage resulting from the hydrolysis of phosphodiester groups in the backbone of DNA molecules (Gates, 2009) and can occur on a single strand (single strand breaks; SSBs) or both strands (double strand breaks; DSBs). SSBs arise when the sugar phosphate backbones connecting adjacent nucleotides in DNA are simultaneously hydrolyzed such that the hydrogen bonds between complementary bases are not able to hold the two strands together. DSBs are generated when both strands are simultaneously broken at sites that are sufficiently close to one another that base-pairing and chromatin structure are insufficient to keep the two DNA ends juxtaposed. As a consequence, the two DNA ends generated by a DSB can physically dissociate from one another, becoming difficult to repair and increasing the chance of inappropriate recombination with other sites in the genome (Jackson, 2002). SSB can turn into DSB if the replication fork stalls at the lesion leading to fork collapse. Strand breaks are intermediates in various biological events, including DNA repair (e.g., excision repair), as well as other normal cellular processes where DSBs act as genetic shufflers to generate genetic diversity for V(D)J recombination in lymphoid cells, and chromatin remodeling in both somatic cells and germ cells, and meiotic recombination in gametes.  Strand breaks are intermediates in various biological events, including DNA repair (e.g., excision repair), V(D)J recombination in developing lymphoid cells and chromatin remodeling in both somatic cells and germ cells. The spectrum of damage can be complex, particularily if the stressor is from large amounts of deposited energy which can result in complex lesions and clustered damage defined as two or more oxidized bases, abasic sites or strand breaks on opposing DNA strands within a few helical turns. These lesions are more difficult to repair and have been studied in many types of models (Barbieri et al., 2019 and Asaithamby et al., 2011). DSBs and complex lesions are of particular concern, as they are considered the most lethal and deleterious type of DNA lesion. If misrepaired or left unrepaired, DSBs may drive the cell towards genomic instability, apoptosis or tumorigenesis (Beir, 1999).

Please refer to the table below for details regarding these and other methodologies for detecting DNA DSBs.  <table border="1"> <tbody> <tr> <td> Method of Measurement   </td> <td> References   </td> <td> Description   </td> <td> OECD Approved Method?  </td> </tr> <tr> <td> Comet Assay (Single Cell Gel Eletrophoresis - Alkaline)   </td> <td> Collins, 2004; Olive and Banath, 2006; Platel et al., 2011; Nikolova et al., 2017   </td> <td> To detect SSBs or DSBs, single cells are encapsulated in agarose on a slide, lysed, and subjected to gel electrophoresis at an alkaline pH (pH >13); DNA fragments are forced to move, forming a "comet"-like appearance   </td> <td> Yes  </td> </tr> <tr> <td> γ-H2AX Foci Quantification - Flow Cytometry   </td> <td> Rothkamm and Horn, 2009; Bryce et al., 2016   </td> <td> Measurement of γ-H2AX immunostaining in cells by flow cytometry, normalized to total levels of H2AX   </td> <td> No  </td> </tr> <tr> <td> γ-H2AX Foci Quantification - Western Blot   </td> <td> Burma et al., 2001; Revet et al., 2011   </td> <td> Measurement of γ-H2AX immunostaining in cells by Western blotting, normalized to total levels of H2AX   </td> <td> No  </td> </tr> <tr> <td> γ-H2AX Foci Quantification - Microscopy   </td> <td> Redon et al., 2010; Mah et al., 2010; Garcia-Canton et al., 2013   </td> <td> Quantification of γ-H2AX immunostaining by counting γ-H2AX foci visualized with a microscope   </td> <td> No  </td> </tr> <tr> <td> γ-H2AX Foci Quantification - ELISA   </td> <td> Ji et al., 2017   </td> <td> Measurement of γ-H2AX in cells by ELISA, normalized to total levels of H2AX   </td> <td> No  </td> </tr> <tr> <td> Pulsed Field Gel Electrophoresis (PFGE)   </td> <td> Ager et al., 1990; Gardiner et al., 1985; Herschleb et al., 2007; Kawashima et al., 2017   </td> <td> To detect DSBs, cells are embedded and lysed in agarose, and the released DNA undergoes gel electrophoresis in which the direction of the voltage is periodically alternated; Large DNA fragments are thus able to be separated by size   </td> <td> No  </td> </tr> <tr> <td> The TUNEL (Terminal Deoxynucleotidyl Transferase dUTP Nick End Labeling) Assay   </td> <td> Loo, 2011   </td> <td> To detect strand breaks, dUTPs added to the 3’OH end of a strand break by the DNA polymerase terminal deoxynucleotidyl transferase (TdT) are tagged with a fluorescent dye or a reporter enzyme to allow visualization   </td> <td> No  </td> </tr> <tr> <td> In Vitro DNA Cleavage Assays using Topoisomerase   </td> <td> Nitiss, 2012   </td> <td> Cleavage of DNA can be achieved using purified topoisomerase; DNA strand breaks can then be separated and quantified using gel electrophoresis   </td> <td> No  </td> </tr> <tr> <td> PCR assay  </td> <td> Figueroa‑González & Pérez‑Plasencia, 2017  </td> <td> Assay of strand breaks through the observation of DNA amplification prevention. Breaks block Taq polymerase, reducing the number of DNA templates, preventing amplification  </td> <td> No  </td> </tr> <tr> <td> Sucrose density gradient centrifuge  </td> <td> Raschke et al. 2009  </td> <td> Division of DNA pieces by density, increased fractionation leads to lower density pieces, with the use of a sucrose cushion  </td> <td> No  </td> </tr> <tr> <td> Alkaline Elution Assay  </td> <td> Kohn, 1991  </td> <td> Cells lysed with detergent-solution, filtered through membrane to remove all but intact DNA  </td> <td> No  </td> </tr> <tr> <td> Unwinding Assay  </td> <td> Nacci et al. 1992  </td> <td> DNA is stored in alkaline solutions with DNA-specific dye and allowed to unwind following removal from tissue, increased strand damage associated with increased unwinding  </td> <td> Yes  </td> </tr> <tr> <td> STRIDE assay  </td> <td> Zilio and Ulrich, 2021  </td> <td> STRIDE (SensiTive Recognition of Individual DNA Ends) combines in situ nick translation with the proximity ligation assay (PLA) to detect single-strand breaks (sSTRIDE) or double-strand breaks (dSTRIDE). In this process, lesions labeled through nick translation with biotinylated nucleotides are identified by a PLA signal, which arises from the interaction of two anti-biotin antibodies from different species.    </td> <td> No  </td> </tr> <tr> <td> sBLISS  </td> <td> Bouwmann et al. 2020  </td> <td> sBLISS (in-suspension breaks labeling in situ and sequencing)  labels double-strand breaks (DSBs) in cells immobilized on glass coverslips, using double-stranded oligonucleotide adaptors that facilitate selective linear amplification through T7-mediated in vitro transcription (IVT), followed by next-generation sequencing (NGS) library preparation    </td> <td> No  </td> </tr> </tbody> </table>

Taxonomic applicability: DNA strand breaks are relevant to all species, including vertebrates such as humans, that contain DNA (Cannan & Pederson, 2016).   Life stage applicability: This key event is not life stage specific as all life stages display strand breaks. However, there is an increase in baseline levels of DNA strand breaks seen in older individuals though it is unknown whether this change due to increased break induction or a greater retention of breaks due to poor repair (White & Vijg, 2016).  Sex applicability: This key event is not sex specific as both sexes display evidence of strand breaks. In some cell types, such as peripheral blood mononuclear cells, males show higher levels of single strand breaks than females (Garm et al., 2012).  Evidence for perturbation by a stressor: There are studies demonstrating that increased DNA strand breaks can result from exposure to multiple stressor types including ionizing & non-ionizing radiation, chemical agents, and oxidizing agents (EPRI, 2014; Hamada, 2014; Cencer et al., 2018; Cannan & Pederson, 2016; Yang et al., 1998).

High Unspecific

High

All life stages

Not Specified

Ager, D. D., et al. (1990). Measurement of radiation-induced DNA double-strand breaks by pulsed-field gel electrophoresis. Radiation research, 122/(2), 181–187.  Anderson, D. & Laubenthal J. (2013), “Analysis of DNA Damage via Single-Cell Electrophoresis. In: Makovets S, editor. DNA Electrophoresis. Totowa.”, NJ: Humana Press. p 209-218.  Asaithamby, A., B. Hu and D.J. Chen. (2011) “Unrepaired clustered DNA lesions induce chromosome breakage in human cells.” Proc Natl Acad Sci U S A 108(20): 8293-8298 .  Barbieri, S., G. Babini, J. Morini et a l (2019). . Predicting DNA damage foci and their experimental readout with 2D microscopy: a unified approach applied to photon and neutron exposures. Scientific Reports 9(1): 14019  Bouwman, B. et al. (2020), “Genome-wide detection of DNA double-strand breaks by in-suspension BLISS”, Nature protocols,.15/12, Springer Nature, London, <a href="https://doi.org/10.1038/s41596-020-0397-2" rel="noreferrer noopener" target="_blank">https://doi.org/10.1038/s41596-020-0397-2</a>   Bryce, S. et al. (2016), “Genotoxic mode of action predictions from a multiplexed flow cytometric assay and a machine learning approach.”, Environ Mol Mutagen. 57:171-189. Doi: 10.1002/em.21996.  Burma, S. et al. (2001), “ATM phosphorylates histone H2AX in response to DNA double-strand breaks.”, J Biol Chem, 276(45): 42462-42467. doi:10.1074/jbc.C100466200  Cannan, W.J. and D.S. Pederson (2016), "Mechanisms and Consequences of Double-Strand DNA Break Formation in Chromatin.", Journal of Cellular Physiology, Vol.231(/1), Wiley, New York, https://doi.org/10.1002/jcp.25048.   Cencer, C. et al. (2018), “PARP-1/PAR Activity in Cultured Human Lens Epithelial Cells Exposed to Two Levels of UVB Light”, Photochemistry and Photobiology, Vol.(94/1), Wiley-Blackwell, Hoboken, https://doi.org/10.1111/php.12814.   Charlton, E. D. et al. (1989), “Calculation of Initial Yields of Single and Double Stranded Breaks in Cell Nuclei from Electrons, Protons, and Alpha Particles.”, Int. J. Radiat. Biol. 56(1): 1-19. doi: 10.1080/09553008914551141.  Collins, R. A. (2004), “The Comet Assay for DNA Damage and Repair. Molecular Biotechnology.”, Mol Biotechnol. 26(3): 249-61. doi:10.1385/MB:26:3:249  EPRI (2014), Epidemiology and mechanistic effects of radiation on the lens of the eye: Review and scientific appraisal of the literature, EPRI, California.  Figueroa‑González, G. and C. Pérez‑Plasencia. (2017), “Strategies for the evaluation of DNA damage and repair mechanisms in cancer”, Oncology Letters, Vol.133(/6), Spandidos Publications, Athens, https://doi.org/10.3892/ol.2017.6002.  Garcia-Canton, C. et al. (2013), “Assessment of the in vitro p-H2AX assay by High Content Screening asa novel genotoxicity test.”, Mutat Res. 757:158-166. Doi: 10.1016/j.mrgentox.2013.08.002  Gardiner, K. et al. (1986), “Fractionation of Large Mammalian DNA Restriction Fragments Using Vertical Pulsed-Field Gradient Gel Electrophoresis.”, Somatic Cell and Molecular Genetics. 12(2): 185-95.Doi: 10.1007/bf01560665.  Garm, C. et al. (2012), “Age and gender effects on DNA strand break repair in peripheral blood mononuclear cells”, Aging Cell, Vol.12/1, Blackwell Publishing Ltd, Oxford, https://doi.org/10.1111/acel.12019.  Hamada, N. (2014), “What are the intracellular targets and intratissue target cells for radiation effects?”, Radiation research, Vol. 181/1, The Radiation Research Society, Indianapolis, https://doi.org/10.1667/RR13505.1.  Herschleb, J. et al. (2007), “Pulsed-field gel electrophoresis.”, Nat Protoc. 2(3): 677-684. doi:10.1038/nprot.2007.94  Iliakis, G. et al. (2015), “Alternative End-Joining Repair Pathways Are the Ultimate Backup for Abrogated Classical Non-Homologous End-Joining and Homologous Recombination Repair: Implications for the Formation of Chromosome Translocations.”, Mutation Research/Genetic Toxicology and Environmental Mutagenesis. 2(3): 677-84. doi: 10.1038/nprot.2007.94  Jackson, S. (2002). “Sensing and repairing DNA double-strand breaks.”, Carcinogenesis. 23:687-696. Doi:10.1093/carcin/23.5.687.  Ji, J. et al. (2017), “Phosphorylated fraction of H2AX as a measurement for DNA damage in cancer cells and potential applications of a novel assay.”, PLoS One. 12(2): e0171582. doi:10.1371/journal.pone.0171582  Kawashima, Y.(2017), “Detection of DNA double-strand breaks by pulsed-field gel electrophoresis.”, Genes Cells 22:84-93. Doi: 10.1111/gtc.12457.  Khoury, L. et al. (2013), “Validation of high-throughput genotoxicity assay screening using cH2AX in-cell Western assay on HepG2 cells.”, Environ Mol Mutagen, 54:737-746. Doi: 10.1002/em.21817.  Khoury, L. et al. (2016), “Evaluation of four human cell lines with distinct biotransformation properties for genotoxic screening.”, Mutagenesis, 31:83-96. Doi: <a href="https://doi.org/10.1093/mutage/gev058" rel="noreferrer noopener" target="_blank">10.1093/mutage/gev058</a>.  Kohn, K.W. (1991), “Principles and practice of DNA filter elution”, Pharmacology & Therapeutics, Vol.49(/1), Elsevier, Amsterdam, https://doi.org/10.1016/0163-7258(91)90022-E.  Loo, DT. (2011), “In Situ Detection of Apoptosis by the TUNEL Assay: An Overview of Techniques. In: Didenko V, editor. DNA Damage Detection In Situ, Ex Vivo, and In Vivo. Totowa.”, NJ: Humana Press. p 3-13.doi: <a href="https://doi.org/10.1007/978-1-60327-409-8_1" rel="noreferrer noopener" target="_blank">10.1007/978-1-60327-409-8_1</a>.  Mah, L. J. et al. (2010), “Quantification of gammaH2AX foci in response to ionising radiation.”, J Vis Exp(38). doi:10.3791/1957.  Nacci, D. et al. (1992), “Application of the DNA alkaline unwinding assay to detect DNA strand breaks in marine bivalves”, Marine Environmental Research, Vol.33(/2), Elsevier BV, Amsterdam, https://doi.org/10.1016/0141-1136(92)90134-8.  Nikolova, T., F. et al. (2017), “Genotoxicity testing: Comparison of the γH2AX focus assay with the alkaline and neutral comet assays.”, Mutat Res 822:10-18. Doi: <a href="https://doi.org/10.1016/j.mrgentox.2017.07.004" rel="noreferrer noopener" target="_blank">10.1016/j.mrgentox.2017.07.004</a>.  Nitiss, J. L. et al. (2012), “Topoisomerase assays. ”, Curr Protoc Pharmacol. Chapter 3: Unit 3 3.  OECD. (2014). Test No. 489: “In vivo mammalian alkaline comet assay.” OECD Guideline for the Testing of Chemicals, Section 4 .  Olive, P. L., & Banáth, J. P. (2006), “The comet assay: a method to measure DNA damage in individual cells.”, Nature Protocols. 1(1): 23-29. doi:10.1038/nprot.2006.5.  Platel A. et al. (2011), “Study of oxidative DNA damage in TK6 human lymphoblastoid cells by use of the thymidine kinase gene-mutation assay and the in vitro modified comet assay: Determination of No-Observed-Genotoxic-Effect-Levels.”, Mutat Res 726:151-159. Doi: 10.1016/j.mrgentox.2011.09.003.  Raschke, S., J. Guan and G. Iliakis. (2009), “Application of alkaline sucrose gradient centrifugation in the analysis of DNA replication after DNA damage”, Methods in Molecular Biology, Vol.521, Humana Press, Totowa, https://doi.org/10.1007/978-1-60327-815-7_18.  Redon, C. et al. (2010), “The use of gamma-H2AX as a biodosimeter for total-body radiation exposure in non-human primates.”, PLoS One. 5(11): e15544. doi:10.1371/journal.pone.0015544  Revet, I. et al. (2011), “Functional relevance of the histone γH2Ax in the response to DNA damaging agents.” Proc Natl Acad Sci USA.108:8663-8667. Doi: 10.1073/pnas.1105866108  Rogakou, E.P. et al. (1998), “DNA Double-stranded Breaks Induce Histone H2AX Phosphorylation on Serine 139.” , J Biol Chem, 273:5858-5868. Doi: 10.1074/jbc.273.10.5858  Rothkamm, K. & Horn, S. (2009), “γ-H2AX as protein biomarker for radiation exposure.”, Ann Ist Super Sanità, 45(3): 265-71.  White, R.R. and J. Vijg. (2016), “Do DNA Double-Strand Breaks Drive Aging?”, Molecular Cell, Vol.63, Elsevier, Amsterdam, http://doi.org/10.1016/j.molcel.2016.08.004.  Yang, Y. et al. (1998), “The effect of catalase amplification on immortal lens epithelial cell lines”, Experimental Eye Research, Vol.67(/6), Academic Press Inc, Cambridge, https://doi.org/10.1006/exer.1998.0560.   Zilio, N. and H. D. Ulrich (2021), “Exploring the SSBreakome: genome-wide mapping of DNA single-strand breaks by next-generation sequencing”, The FEBS journal, 288(13), Wiley, Hoboken, https://doi.org/10.1111/febs.15568    AOPWiki 2019-05-19T16:33:20

2024-12-17T11:57:56

Increase, Mutations

Molecular

A mutation is a change in DNA sequence. Mutations can thus alter the coding sequence of genes, potentially leading to malformed or truncated proteins. Mutations can also occur in promoter regions, splice junctions, non-coding RNA, DNA segments, and other functional locations in the genome. These mutations can lead to various downstream consequences, including alterations in gene expression. There are several different types of mutations including missense, nonsense, insertion, deletion, duplication, and frameshift mutations, all of which can impact the genome and its expression in unique ways. Missense mutations are the substitution of one base in the codon with another. This change is significant because the three bases in a codon code for a specific amino acid and the new combination may signal for a different amino acid to be formed. Nonsense mutations also result from changes to the codon bases, but in this case, they cause the generation of a stop codon in the DNA strand where there previously was not one. This stop codon takes the place of a normal coding triplet, preventing its translation into an amino acid. This will cause the translation of the strand to prematurely stop. Both missense and nonsense mutations can result from substitutions, insertions, or deletions of bases (Chakarov et al. 2014).   Insertion and deletion mutations are the addition and removal of bases from the strand, respectively. These often accompany a frameshift mutation, as the alteration in the number of bases in the strand causes the frame of the base reader to shift by the added or reduced number, altering the amino acids that are produced if that number is not devisable by three. Codons come in specific orders, sectioned into groups of three. When the boundaries of which three bases are included in one group are changed, this can change the whole transcriptional output of the strand (Chakaroy et al. 2014).  Mutations can be propagated to daughter cells upon cellular replication. Mutations in stem cells (versus terminally differentiated non-replicating cells) are the most concerning, as these will persist in the organism. The consequence of the mutation, and thus the fate of the cell, depends on the location (e.g., coding versus non-coding) and the type (e.g., nonsense versus silent) of mutation. Mutations can occur in somatic cells or germ cells (sperm or egg).

Mutations can be measured using a variety of both OECD and non-OECD mutagenicity tests. Listed below are common methods for detecting the KE, however there may be other comparable methods that are not listed. Somatic cells: The Salmonella mutagenicity test (Ames Test) is generally used as part of a first tier screen to determine if a chemical can cause gene mutations. This well-established test has an OECD test guideline (OECD TG 471, 2020). A variety of bacterial strains are used, in the presence and absence of a metabolic activation system (e.g., rat liver microsomal S9 fraction), to determine the mutagenic potency of chemicals by dose-response analysis. A full description is found in Test No. 471: Bacterial Reverse Mutation Test (OECD, 2016). A variety of in vitro mammalian cell gene mutation tests are described in OECD’s Test Guidelines 476 (2016) and 490 (2015). TG 476 (2016) is used to identify substances that induce gene mutations at the hprt (hypoxanthine-guanine phosphoribosyl transferase) gene, or the transgenic xprt (xanthine-guanine phosphoribosyl transferase) reporter locus. The most commonly used cells for the HPRT test include the CHO, CHL and V79 lines of Chinese hamster cells, L5178Y mouse lymphoma cells, and TK6 human lymphoblastoid cells. The only cells suitable for the XPRT test are AS52 cells containing the bacterial xprt (or gpt) transgene (from which the hprt gene was deleted). The new OECD TG 490 (2015) describes two distinct in vitro mammalian gene mutation assays using the thymidine kinase (tk) locus and requiring two specific tk heterozygous cells lines: L5178Y tk+/-3.7.2C cells for the mouse lymphoma assay (MLA) and TK6 tk+/- cells for the TK6 assay. The autosomal and heterozygous nature of the thymidine kinase gene in the two cell lines enables the detection of cells deficient in the enzyme thymidine kinase following mutation from tk+/- to tk-/-. It is important to consider that different mutation spectra are detected by the different mutation endpoints assessed. The non-autosomal location of the hprt gene (X-chromosome) means that the types of mutations detected in this assay are point mutations, including base pair substitutions and frameshift mutations resulting from small insertions and deletions. Whereas, the autosomal location of the transgenic xprt, tk, or gpt locus allows the detection of large deletions not readily detected at the hemizygous hprt locus on X-chromosomes. Genetic events detected using the tk locus include both gene mutations (point mutations, frameshift mutations, small deletions) and large deletions. The transgenic rodent mutation assay (OECD TG 488, 2020) is the only assay capable of measuring gene mutation in virtually all tissues in vivo. Specific details on the rodent transgenic mutation reporter assays are reviewed in Lambert et al. (2005, 2009). The transgenic reporter genes are used for detection of gene mutations and/or chromosomal deletions and rearrangements resulting in DNA size changes (the latter specifically in the lacZ plasmid and Spi- test models) induced in vivo by test substances (OECD, 2009, OECD, 2011; Lambert et al., 2005). Briefly, transgenic rodents (mouse or rat) are exposed to the chemical agent sub-chronically. Following a manifestation period, genomic DNA is extracted from tissues, transgenes are rescued from genomic DNA, and transfected into bacteria where the mutant frequency is measured using specific selection systems. The Pig-a (phosphatidylinositol glycan, Class A) gene on the X chromosome codes for a catalytic subunit of the N-acetylglucosamine transferase complex that is involved in glycosylphosphatidyl inositol (GPI) cell surface anchor synthesis. Cells lacking GPI anchors, or GPI-anchored cell surface proteins are predominantly due to mutations in the Pig-a gene. Thus, flow cytometry of red blood cells expressing or not expressing the Pig-a gene has been developed for mutation analysis in blood cells from humans, rats, mice, and monkeys. The assay is described in detail in Dobrovolsky et al. (2010). Development of an OECD guideline for the Pig-a assay is underway. In addition, experiments determining precisely what proportion of cells expressing the Pig-a mutant phenotype have mutations in the Pig-a gene are in progress (e.g., Nicklas et al., 2015, Drobovolsky et al., 2015). A recent paper indicates that the majority of CD48 deficient cells from 7,12-dimethylbenz[a]anthracene-treated rats (78%) are indeed due to mutation in Pig-a (Drobovolsky et al., 2015). Germ cells: Tandem repeat mutations can be measured in bone marrow, sperm, and other tissues using single-molecule PCR. This approach has been applied most frequently to measure repeat mutations occurring in sperm DNA. Isolation of sperm DNA is as described above for the transgenic rodent mutation assay, and analysis of tandem repeats is done using electrophoresis for size analysis of allele length using single-molecule PCR. For expanded simple tandem repeat this involved agarose gel electrophoresis and Southern blotting, whereas for microsatellites sizing is done by capillary electrophoresis. Detailed methodologies for this approach are found in Yauk et al. (2002) and Beal et al. (2015). Mutations in rodent sperm can also be measured using the transgenic reporter model (OECD TG 488, 2020). A description of the approach is found within this published TG. Further modifications to this protocol have been made as of 2022 for the analysis of germ cells. Detailed methodology for detecting mutant frequency arising in spermatogonia is described in Douglas et al. (1995), O'Brien et al. (2013); and O'Brien et al. (2014). Briefly, male mice are exposed to the mutagen and killed at varying times post-exposure to evaluate effects on different phases of spermatogenesis. Sperm are collected from the vas deferens or caudal epididymis (the latter preferred). Modified protocols have been developed for extraction of DNA from sperm. A similar transgenic assay can be used in transgenic medaka (Norris and Winn, 2010). Please note, gene mutations that occur in somatic cells in vivo (OECD Test. No. 488, 2020) or in vitro (OECD Test No. 476: In vitro Mammalian Cell Gene Mutation Test, 2016), or in bacterial cells (i.e., OECD Test No. 471, 2020) can be used as an indicator that mutations in male pre-meiotic germ cells may occur for a particular agent (sensitivity and specificity of other assays for male germ cell effects is given in Waters et al., 1994). However, given the very unique biological features of spermatogenesis relative to other cell types, known exceptions to this rule, and the small database on which this is based, inferring results from somatic cell or bacterial tests to male pre-meiotic germ cells must be done with caution. That mutational assays in somatic cells may predict mutations in germ cells has not been rigorously tested empirically (Singer and Yauk, 2010). The IWGT working group on germ cells specifically addressed this gap in knowledge in their report (Yauk et al., 2015) and recommended that additional research address this issue. Mutations can be directly measured in humans (and other species) through the application of next-generation sequencing. Although single-molecule approaches are growing in prevalence, the most robust approach to measure mutation using next-generation sequencing today requires clonal expansion of the mutation to a sizable proportion (e.g., sequencing tumours; Shen et al., 2015), or analysis of families to identify germline derived mutations (reviewed in Campbell and Eichler, 2013; Adewoye et al., 2015). Please refer to the table below for additional details and methodologies for measuring mutations. <table border="1" cellpadding="1" cellspacing="1" style="height:2351px; width:633px"> <tbody> <tr> <td style="background-color:#eeeeee; text-align:center">Assay Name</td> <td style="background-color:#eeeeee; text-align:center">References </td> <td style="background-color:#eeeeee; text-align:center">Description </td> <td style="background-color:#eeeeee; text-align:center">OECD Approved Assay</td> </tr> <tr> <td>Assorted Gene Loci Mutation Assays</td> <td> Tindall et al., 1989; Kruger et al., 2015 </td> <td>After exposure to a chemical/mutagen, mutations can be measured by the ability of exposed cells to form colonies in the presence of specific compounds that would normally inhibit colony growth; Usually only cells -/- for the gene of interest are able to form colonies</td> <td>N/A</td> </tr> <tr> <td>TK Mutation Assay</td> <td> Yamamoto et al., 2017; Liber et al., 1982; Lloyd and Kidd, 2012 </td> <td>After exposure to a chemical/mutagen, mutations are detected at the thymidine kinase (TK) loci of L5178Y wild-type mouse lymphoma TK (+/-) cells by measuring resistance to lethaltriflurothymidine (TFT); Only TK-/- cells are able to form colonies</td> <td>Yes (No. 490)</td> </tr> <tr> <td>HPRT Mutation Assay</td> <td> Ayres et al., 2006; Parry and Parry, 2012 </td> <td>Similar to TK Mutation Assay above, X-linked HPRT mutations produced in response to chemical/mutagen exposure can be measured through colony formation in the presence of 6-TG or 8-azoguanine; Only HPRT-/- cells are able to form colonies</td> <td>Yes (No. 476)</td> </tr> <tr> <td>Salmonella Mutagenicity Test (Ames Test)</td> <td>OECD, 1997</td> <td>After exposure to a chemical/mutagen, point mutations are detected by analyzing the growth capacity of different bacterial strains in the presence and absence of various metabolic activation systems </td> <td>Yes (No. 471)</td> </tr> <tr> <td>PIG-A / PIG-O Assay</td> <td> Kruger et al., 2015; Nakamura, 2012; Chikura, 2019 </td> <td>After exposure to a chemical/mutagen, mutations  in PIG-A or PIG-O (which decrease the biosynthesis of the glycosylphosphatidylinositol (GPI) anchor protein) are assessed by the colony-forming capabilities of cells after in vitro exposure, or by flow cytometry of blood samples after in vivo exposure</td> <td>N/A</td> </tr> <tr> <td>Single Molecule PCR</td> <td> Kraytsberg & Khrapko, 2005; Yauk, 2002 </td> <td>This PCR technique uses a single DNA template, and is often employed for detection of mutations in microsatellites, recombination studies, and generation of polonies</td> <td>N/A</td> </tr> <tr> <td>ACB-PCR</td> <td> Myers et al., 2014 (Textbook, pg 345-363); Banda et al.,  2013; Banda et al.,  2015; Parsons et al., 2017 </td> <td>Using this PCR technique, single base pair substitution mutations within oncogenes or tumour suppressor genes can be detected by selectively amplifying specific point mutations within an allele and selectively blocking amplification of the wild-type allele </td> <td>N/A</td> </tr> <tr> <td>Transgenic Rodent Mutation Assay </td> <td> OECD 2013; Lambert 2005; Lambert 2009 </td> <td>This in vivo test detects gene mutations using transgenic rodents that possess transgenes and reporter genes; After in vivo exposure to a chemical/mutagen, the transgenes are analyzed by transfecting bacteria with the reporter gene and examining the resulting phenotype</td> <td>Yes (No. 488)</td> </tr> <tr> <td>Conditionally inducible transgenic mouse models</td> <td>Parsons 2018 (Review)</td> <td>Inducible mutations linked to fluorescent tags are introduced into transgenic mice; Upon exposure of the transgenic mice to an inducing agent, the presence and functional assessment of the mutations can be easily ascertained due to expression of the linked fluorescent tags </td> <td>N/A</td> </tr> <tr> <td>Error-Corrected Next Generation Sequencing (NGS)</td> <td>Salk 2018 (Review)</td> <td>This technique detects rare subclonal mutations within a pool of heterogeneous DNA samples through the application of new error-correction strategies to NGS; At present, few laboratories in the world are capable of doing this, but commercial services are becoming available (e.g., Duplex sequencing at TwinStrand BioSciences) </td> <td>N/A </td> </tr> </tbody> </table>

Taxonomic applicability: Mutations can occur in any organism and in any cell type, and are the fundamental material of evolution. The test guidelines described above range from analysis from prokaryotes, to rodents, to human cells in vitro. Mutations have been measured in virtually every human tissue sampled in vivo. Life stage applicability: This key event is not life stage specific as all stages of life have DNA that can be mutated; however, baseline levels of mutations are seen to increase with age (Slebos et al., 2004; Kirkwood, 1989).  Sex applicability: This key event is not sex specific as both sexes undergo mutations. Males have a higher mutation rate than females (Hedrick, 2007).  Evidence for perturbation by a stressor: Many studies demonstrate that increased mutations can occur as a result of ionizing radiation (Sankaranarayanan & Nikjoo, 2015; Russell et al., 1957; Winegar et al., 1994; Gossen et al., 1995).

High Unspecific

High

All life stages

High Moderate High Moderate

Adewoye, A.B. et al. (2015), "The genome-wide effects of ionizing radiation on mutation induction in the mammalian germline", Nat. Commu., 6:6684. Doi: 10.1038/ncomms7684. Ayres, M. F. et al. (2006),  “Low doses of gamma ionizing radiation increase hprt mutant frequencies of TK6 cells without triggering the mutator phenotype pathway”,  Genetics and Molecular Biology. 2(3): 558-561. Doi:10.1590/S1415-4757200600030002. Banda M, Recio L, and Parsons BL. (2013), “ACB-PCR measurement of spontaneous and furan-induced H-ras codon 61 CAA to CTA and CAA to AAA mutation in B6C3F1 mouse liver”, Environ Mol Mutagen. 54(8):659-67. Doi:10.1002/em.21808. Banda,  M. et al. (2015), “Quantification of Kras mutant fraction in the lung DNA of mice exposed to aerosolized particulate vanadium pentoxide by inhalation”,  Mutat Res Genet Toxicol Environ Mutagen. 789-790:53-60. Doi: 10.1016/j.mrgentox.2015.07.003 Campbell, C.D. & E.E. Eichler (2013), "Properties and rates of germline mutations in humans", Trends Genet., 29(10): 575-84. Doi:  10.1016/j.tig.2013.04.005 Chakarov, S. et al. (2014), “DNA damage and mutation. Types of DNA damage”, BioDiscovery, Vol.11, Pensoft Publishers, Sofia, https://doi.org/10.7750/BIODISCOVERY.2014.11.1. Chikura, S. et al. (2019), “Standard protocol for the total red blood cell Pig-a assay used in the interlaboratory trial organized by the Mammalian Mutagenicity Study Group of the Japanese Environmental Mutagen Society”,  Genes Environ.  27:41-5. Doi: 10.1186/s41021-019-0121-z. Dobrovolsky, V.N. et al. (2015), "CD48-deficient T-lymphocytes from DMBA-treated rats have de novo mutations in the endogenous Pig-a gene. CD48-Deficient T-Lymphocytes from DMBA-Treated Rats Have De Novo Mutations in the Endogenous Pig-a Gene", Environ. Mol. Mutagen., (6): 674-683. Doi: 10.1002/em.21959. Douglas, G.R. et al. (1995), "Temporal and molecular characteristics of mutations induced by ethylnitrosourea in germ cells isolated from seminiferous tubules and in spermatozoa of lacZ transgenic mice", Proceedings of the National Academy of Sciences of the United States of America, 92(16): 7485-7489. Doi: 10.1073/pnas.92.16.7485. Gossen, J.A. et al. (1995), "Spontaneous and X-ray-induced deletion mutations in a LacZ plasmid-based transgenic mouse model", Mutation Research, 331/1, Elsevier, Amsterdam, https://doi.org/10.1016/0027-5107(95)00055-N.  Hedrick, P.W. (2007), “Sex: Differences In Mutation, Recombination, Selection, Gene Flow, And Genetic Drift”, Evolution, Vol.61/12, Wiley, Hoboken, https://doi.org/10.1111/j.1558-5646.2007.00250.x.  Kirkwood, T.B.L. (1989), “DNA, mutations and aging”, Mutation Research, Vol.219/1, Elsevier B.V., Amsterdam, https://doi.org/10.1016/0921-8734(89)90035-0 Kraytsberg,Y. &  Khrapko, K. (2005),  “Single-molecule PCR: an artifact-free PCR approach for the analysis of somatic mutations”,  Expert Rev Mol Diagn. 5(5):809-15. Doi: 10.1586/14737159.5.5.809. Krüger, T. C., Hofmann, M., & Hartwig, A. (2015), “The in vitro PIG-A gene mutation assay: mutagenicity testing via flow cytometry based on the glycosylphosphatidylinositol (GPI) status of TK6 cells”, Arch Toxicol. 89(12), 2429-43. Doi: 10.1007/s00204-014-1413-5. Lambert, I.B. et al. (2005), "Detailed review of transgenic rodent mutation assays", Mutat Res., 590(1-3):1-280. Doi: 10.1016/j.mrrev.2005.04.002. Liber, L. H., & Thilly, G. W. (1982),  “Mutation assay at the thymidine kinase locus in diploid human lymphoblasts”,  Mutation Research. 94: 467-485. Doi:10.1016/0027-5107(82)90308-6. Lloyd, M., & Kidd, D. (2012), “The Mouse Lymphoma Assay. In: Parry J., Parry E. (eds) Genetic Toxicology, Methods in Molecular Biology (Methods and Protocols), 817. Springer, New York, NY. Myers, M. B. et al., (2014), “ACB-PCR Quantification of Somatic Oncomutation”,  Molecular Toxicology Protocols, Methods in Molecular Biology. DOI: 10.1007/978-1-62703-739-6_27 Nakamura, J. et al., (2012), “Detection of PIGO-deficient cells using proaerolysin: a valuable tool to investigate mechanisms of mutagenesis in the DT40 cell system”, PLoS One.7(3): e33563. Doi:10.1371/journal.pone.0033563. Nicklas, J.A., E.W. Carter and R.J. Albertini (2015), "Both PIGA and PIGL mutations cause GPI-a deficient isolates in the Tk6 cell line", Environ. Mol. Mutagen., 6(8):663-73. Doi: 10.1002/em.21953. Norris, M.B. and R.N. Winn (2010), "Isolated spermatozoa as indicators of mutations transmitted to progeny", Mutat Res., 688(1-2): 36–40. Doi: 10.1016/j.mrfmmm.2010.02.008. O'Brien, J.M. et al.(2013), "No evidence for transgenerational genomic instability in the F1 or F2 descendants of Muta™Mouse males exposed to N-ethyl-N-nitrosourea", Mutat. Res., 741-742:11-7. Doi: 10.1016/j.mrfmmm.2013.02.004. O'Brien, J.M. et al. (2014), "Transgenic rodent assay for quanitifying male germ cell mutation frequency", Journal of Visual Experimentation, Aug 6;(90). Doi: 10.3791/51576. O’Brien, J.M. et al. (2015), "Sublinear response in lacZ mutant frequency of Muta™ Mouse spermatogonial stem cells after low dose subchronic exposure to N-ethyl-N-nitrosourea", Environ. Mol. Mutagen., 6(4): 347-355. Doi: 10.1002/em.21932. OECD (2020), Test No. 471: Bacterial Reverse Mutation Test, OECD Guidelines for the Testing of Chemicals, Section 4, OECD Publishing, Paris. OECD (2016), Test No. 476: In vitro Mammalian Cell Gene Mutation Test, OECD Guidelines for the Testing of Chemicals, Section 4, OECD Publishing, Paris. OECD (2009), Detailed Review Paper on Transgenic Rodent Mutation Assays, Series on Testing and Assessment, N° 103, ENV/JM/MONO 7, OECD, Paris. OECD (2020), Test No. 488: Transgenic Rodent Somatic and Germ Cell Gene Mutation Assays, OECD Guidelines for the Testing of Chemicals, Section 4, OECD Publishing, Paris. OECD (2016), Test. No. 490: In vitro mammalian cell gene mutation mutation tests using the thymidine kinase gene, OECD Guidelines for the Testing of Chemicals, Section 4, OECD Publishing, Paris. OECD Guidelines for the Testing of Chemicals, Section 4, OECD Publishing, Paris. Parry MJ, & Parry ME. 2012. Genetic Toxicology Principles and Methods. Humana Press. Springer Protocols. Parsons BL, McKim KL, Myers MB. 2017. Variation in organ-specific PIK3CA and KRAS mutant levels in normal human tissues correlates with mutation prevalence in corresponding carcinomas. Environ Mol Mutagen. 58(7):466-476. Doi: 10.1002/em.22110. Parsons BL. Multiclonal tumor origin: Evidence and implications. Mutat Res. 2018. 777:1-18. doi: 10.1016/j.mrrev.2018.05.001. Russell, W.L. et al. (1957), "Radiation Dose Rate and Mutation Frequency.", Science, Vol.128/3338, American Association for the Advancement of Science, Washington, https://doi.org/10.1126/science.128.3338.1546. Salk JJ, Schmitt MW, &Loeb LA. (2018), “Enhancing the accuracy of next-generation sequencing for detecting rare and subclonal mutations”, Nat Rev Genet. 19(5):269-285. Doi: 10.1038/nrg.2017.117. Sankaranarayanan, K. & H. Nikjoo (2015), "Genome-based, mechanism-driven computational modeling of risks of ionizing radiation: The next frontier in genetic risk estimation?", Mutation Research, Vol.764, Elsevier, Amsterdam, https://doi.org/10.1016/j.mrrev.2014.12.003.  Shen, T., S.H. Pajaro-Van de Stadt, N.C. Yeat and J.C. Lin (2015), "Clinical applications of next generation sequencing in cancer: from panels, to exomes, to genomes" Front. Genet., 6: 215. Doi: 10.3389/fgene.2015.00215. Singer, T.M. and C.L. Yauk CL (2010), "Germ cell mutagens: risk assessment challenges in the 21st century", Environ. Mol. Mutagen., 51(8-9): 919-928. Doi: 10.1002/em.20613. Slebos, R.J.C. et al. (2004), “Mini-and microsatellite mutations in children from Chernobyl accident cleanup workers”, Mutation Research/Genetic Toxicology and Environmental Mutagenesis, Vol.559/1-2, Elsevier, Amsterdam, https://doi.org/10.1016/j.mrgentox.2004.01.003.  Tindall, R. K., & Stankowski Jr., F. L. (1989),  “Molecular analysis of spontaneous mutations at the GPT locus in Chinese hamster ovary (AS52) cells”, Mutation Research, 220, 241-53. Doi: 10.1016/0165-1110(89)90028-6. Waters, M.D. et al. (1994), "The performance of short-term tests in identifying potential germ cell mutagens: a qualitative and quantitative analysis", Mutat. Res., 341(2): 109-31. Doi: 10.1016/0165-1218(94)90093-0. Winegar, R.A. et al. (1994), "Radiation-induced point mutations, deletions and micronuclei in lacI transgenic mice", Mutation Research, Vol.307/2, Elsevier, Amsterdam, https://doi.org/10.1016/0027-5107(94)90258-5.  Yamamoto, A. et al. (2017), “Radioprotective activity of blackcurrant extract evaluated by in vitro micronucleus and gene mutation assays in TK6 human lymphoblastoid cells”, Genes and Environment. 39: 22. Doi: 10.1186/s41021-017-0082-z. Yauk, C.L. et al. (2002), "A novel single molecule analysis of spontaneous and radiation-induced mutation at a mouse tandem repeat locus", Mutat. Res., 500(1-2): 147-56. Doi: 10.1016/s0027-5107(02)00005-2. Yauk, C.L. et al. (2015), "Approaches for Identifying Germ Cell Mutagens: Report of the 2013 IWGT Workshop on Germ Cell Assays", Mutat. Res. Genet. Toxicol. Environ. Mutagen., 783: 36-54. Doi: 10.1016/j.mrgentox.2015.01.008. Yeat and J.C. Lin. 2015. Clinical applications of next generation sequencing in cancer: from panels, to exomes, to genomes. Front. Genet., 6: 215. Doi: 10.3389/fgene.2015.00215. AOPWiki 2016-11-29T18:41:23

2023-05-15T08:47:43

Increase, Chromosomal aberrations

Cellular

Structural chromosomal aberrations describe the damage to chromosomes that results from breaks along the DNA and may lead to deletion, addition, or rearrangement of sections in the chromosome. Chromosomal aberrations can be divided in two major categories: chromatid-type or chromosome-type depending on whether one or both chromatids are involved, respectively. They can be further classified as rejoined or non-rejoined aberrations. Rejoined aberrations include translocations, insertions, dicentrics and rings, while unrejoined aberrations include acentric fragments and breaks (Savage, 1976). Some of these aberrations are stable (i.e., reciprocal translocations) and can persist for many years (Tucker and Preston, 1996). Others are unstable (i.e., dicentrics, acentric fragments) and decline at each cell division because of clonogenic inactivation (Boei et al., 1996). These events may be detectable after cell division and such damage to DNA is irreversible. Chromosomal aberrations are associated with  clonogenic inactivation and carcinogenicity (Mitelman, 1982). Chromosomal aberrations (CA) refer to a missing, extra or irregular portion of chromosomal DNA. These DNA changes in the chromosome structure may be produced by different double strand break (DSB) repair mechanisms (Obe et al., 2002). There are 4 main types of CAs: deletions, duplications, translocations, and inversions. Deletions happen when a portion of the genetic material from a chromosome is lost. Terminal deletions occur when an end piece of the chromosome is cleaved. Interstitial deletions arise when a chromosome breaks in two separate locations and rejoins incorrectly, with the center piece being omitted. Duplications transpire when there is any addition or rearrangement of excess genetic material; types of duplications include transpositions, tandem duplications, reverse duplications, and displaced duplications (Griffiths et al., 2000). Translocations result from a section of one chromosome being transferred to a non-homologous chromosome (Bunting and Nussenzweig, 2013). When there is an exchange of segments on two non-homologous chromosomes, it is called a reciprocal translocation. Inversions occur in a single chromosome and involve both of the ends breaking and being ligated on the opposite ends, effectively inverting the DNA sequence.       A fifth type of CA that can occur in the genome is the copy number variant (CNV). CNVs, which may comprise greater than 10% of the human genome (Shlien et al., 2009; Zhang et al., 2016; Hastings et al., 2009),  are deletions or duplications that can vary in size from 50 base pairs (Arlt et al., 2012; Arlt et al., 2014; Liu et al., 2013) up into the megabase pair range (Arlt et al., 2012; Wilson et al., 2015; Arlt et al., 2014; Zhang et al., 2016). CNV regions are especially enriched in large genes and large active transcription units (Wilson et al., 2015), and are of particular concern when they cause deletions in tumour suppressor genes or duplications in oncogenes (Liu et al., 2013; Curtis et al., 2012). There are two types of CNVs: recurrent and non-recurrent. Recurrent CNVs are thought to be produced through a recombination process during meiosis known as non-allelic homologous recombination (NAHR) (Arlt et al., 2012; Hastings et al., 2009). These recurrent CNVs, also called germline CNVs, could be inherited and are thus common across different individuals (Shlien et al., 2009; Liu et al., 2013). Non-recurrent CNVs are believed to be produced in mitotic cells during the process of replication. Although the mechanism is not well studied, it has been suggested that stress during replication, in particular stalling replication forks, prompt microhomology-mediated mechanisms to overcome the replication stall, which often results in duplications or deletions. Two models that have been proposed to explain this mechanism include the Fork Stalling and Template Switching (FoSTeS) model, and the Microhomology-Mediated Break-Induced Replication (MMBIR) model (Arlt et al., 2012; Wilson et al., 2015; Lee et al., 2007; Hastings et al., 2009).   CAs can be classified according to whether the chromosome or chromatid is affected by the aberration. Chromosome-type aberrations (CSAs) include chromosome-type breaks, ring chromosomes, marker chromosomes, and dicentric chromosomes; chromatid-type aberrations (CTAs) refer to chromatid breaks and chromatid exchanges (Bonassi et al., 2008; Hagmar et al., 2004). When cells are blocked at the cytokinesis step, When cells are blocked at the cytokinesis step, micronuclei (MN; small nucleus-like structures that contain a chromosome or a piece of a chromosome that was lost during mitosis) can appear in the cytoplasm of binucleated cells. These micronuclei are an indication of CAs and are often related to dicentric chromosomes. Dicentric chromosomes can also cause nucleoplasmic bridges (NPBs; physical connections that exist between the two nuclei) (El-Zein et al., 2014). Other CAs can be assessed by examining the DNA sequence, as is the case when detecting copy number variants (CNVs) (Liu et al., 2013). OECD defines clastogens as ‘any substance that causes structural chromosomal aberrations in populations of cells or organisms’.

CAs can be detected before and after cell division. Widely used assays are described in the table below, however there may be other comparable methods that are not listed.  <table cellspacing="0" class="MsoTableGrid" style="border-collapse:collapse; border:none; width:0px"> <tbody> <tr> <td style="background-color:#f2f2f2; border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:1px solid black; height:37px; width:130px"> Assay </td> <td style="background-color:#f2f2f2; border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; height:37px; width:113px"> References </td> <td style="background-color:#f2f2f2; border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; height:37px; width:274px"> Description </td> <td style="background-color:#f2f2f2; border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; height:37px; width:142px"> OECD-approved assay </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:95px; text-align:center; width:130px">Premature Chromosome Condensation (PCC) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:95px; text-align:center; width:113px">Prasanna et al., 2000; Okayasu et al., 2019 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:95px; text-align:center; width:274px"> Cells are exposed to mitosis-promoting factors (MPF) following cell fusion, causing the chromosomes to condense prematurely. In another approach, cells are exposed to protein phosphatase inhibitors, such as type 1 and 2A protein phosphatases, also causing premature chromosome condensation. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:95px; text-align:center; width:142px">N/A </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:95px; text-align:center; width:130px">Chromosomal G-banding </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:95px; text-align:center; width:113px">Schwatz, 1990 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:95px; text-align:center; width:274px">Use of Giesma dye to stain chromosomal bands, abnormalities determined by the presence of altered morphology  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:95px; text-align:center; width:142px">N/A </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:95px; width:130px"> Fluorescent In Situ  Hybridization (FISH) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:95px; width:113px"> Beaton et al., 2013; Pathak et al., 2017 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:95px; width:274px"> Fluorescent assay of metaphase chromosomes that can detect CAs through chromosome painting and microscopic analysis </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:95px; width:142px"> N/A </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:95px; text-align:center; width:130px">Micronuclei (MN) Assay via Microscopy in vitro  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:95px; text-align:center; width:113px">OECD, 2016a</td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:95px; text-align:center; width:274px">Micronuclei are scored in vitro using microscopy  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:95px; text-align:center; width:142px">Yes (No. 487) </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:97px; width:130px"> Cytokinesis Block Micronucleus (CBMN) Assay with Microscopy in vitro </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:97px; width:113px"> Fenech, 2000; OECD, 2016a </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:97px; width:274px"> Cells are cultured with cytokinesis blocking agent, fixed to slides, and undergo MN quantification using microscopy. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:97px; width:142px">   Yes (No.487) </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:97px; width:130px"> Micronucleus (MN) Assay by Microscopy in vivo </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:97px; width:113px"> OECD, 2016b </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:97px; width:274px"> Cells are fixed on slides and MN are scored using microscopy. Red blood cells can also be scored for MN using flow cytometry (see below) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:97px; width:142px"> Yes (No. 474) </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:79px; width:130px"> CBMN with Imaging Flow Cytometry </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:79px; width:113px"> Rodrigues et al., 2015 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:79px; width:274px"> Cells are cultured with cytokinesis blocking agent, fixed in solution, and imaged with flow cytometry to quantify MN </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:79px; width:142px"> N/A </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:58px; width:130px"> Flow cytometry detection of MN </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:58px; width:113px"> Dertinger et al., 2004; Bryce et al., 2007; OECD 2016a, 2016b </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:58px; width:274px"> In vivo and in vitro flow cytometry-based, automated micronuclei measurements are also done without cytokinesis block. MN analysis in vivo is performed in peripheral blood cells to detect MN in erythrocytes and reticulocytes. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:58px; width:142px">     Yes (No.487; No. 474) </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:56px; width:130px"> High-throughput biomarker assays (indirect measures to confirm clastogenicity) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:56px; width:113px"> Bryce et al. 2014, 2016, 2018   Khoury et al., 2013, Khoury et al., 2016)     Hendriks et al., 2012, 2016; Wink et al., 2014 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:56px; width:274px"> Multiplexed biomarkers can be measured by flow cytometry are used to discern clastogenic and aneugenic mechanisms for MN induction. Flow cytometry-based quantification of γH2AX foci and p53 protein expression (Bryce et al., 2016).   Prediscreen Assay– In-Cell Western -based quantification of γH2AX   Green fluorescent protein reporter assay to detect the activation of stress signaling pathways, including DNA damage signaling including a reporter porter that is associated with DNA double strand breaks. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:56px; width:142px"> N/A </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:193px; width:130px"> Dicentric Chromosome Assay (DCA) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:193px; width:113px"> Abe et al., 2018 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:193px; width:274px"> Cells are fixed on microscope slides, chromosomes are stained, and the number of dicentric chromosomes are quantified </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:193px; width:142px"> N/A </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:65px; width:130px"> High content imaging </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:65px; width:113px"> Shahane et al., 2016 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:65px; width:274px"> DNA can be stained using fluorescent dyes and micronuclei can be scored high-throughput microscopy image analysis. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:65px; width:142px">   N/A </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:193px; width:130px"> Chromosomal aberration test   </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:193px; width:113px"> OECD, 2016c; 2016d; 20l16e </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:193px; width:274px"> In vitro, the cell cycle is arrested at metaphase after 1.5 cell cycle following 3-6 hour exposure   In vivo, the test chemical is administered as a single treatment and bone marrow is collected 18-24 hrs later (TG 475), while testis is collected 24-48 hrs later (TG 483). The cell cycle is arrested with a metaphase-arresting chemical (e.g., colchicine) 2-5 hours before cell collection. Once cells are fixed and stained on microscope slides, chromosomal aberrations are scored </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:193px; width:142px"> Yes. In vitro (No. 473) In vivo (No. 475 and No. 483) </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:193px; width:130px"> Array Comparative Genomic Hybridization (aCGH) or SNP Microarray </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:193px; width:113px"> Adewoye et al., 2015; Wilson et al., 2015; Arlt et al., 2014; Redon et al., 2006; Keren, 2014; Mukherjee, 2017 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:193px; width:274px"> CNVs are most commonly detected using global DNA microarray technologies; This method, however, is unable to detect balanced CAs, such as inversions </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:193px; width:142px">   N/A </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:193px; width:130px"> Next Generation Sequencing (NGS): Whole Genome Sequencing (WGS) or Whole Exome Sequencing (WES) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:193px; width:113px"> Liu, 2013; Shen, 2016; Mukherjee, 2017 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:193px; width:274px"> CNVs are detected by fragmenting the genome and using NGS to sequence either the entire genome (WGS), or only the exome (WES); Challenges with this methodology include only being able to detect CNVs in exon-rich areas if using WES, the computational investment required for the storage and analysis of these large datasets, and the lack of computational algorithms available for effectively detecting somatic CNVs </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:193px; width:142px"> N/A </td> </tr> </tbody> </table>

Taxonomic applicability: CAs are possible in nucleated cells of any species (Ferguson-Smith, 2015).   Life stage applicability: This key event is not life stage specific as subjects of all ages have chromosomes that can be improperly structured. However, older individuals have naturally higher baseline levels of CAs (Vick et al., 2017). Individuals born with stable type aberrations will retain them throughout their lifetime (Gardner et al., 2011).  Sex applicability: This key event is not sex specific, with both sexes experiencing CAs at comparable rates (Ka&scaron;uba et al., 1995).  Evidence for perturbation by a stressor: Many studies have provided evidence to support increased CAs occurring as a result of exposure to ionizing radiation (Franken et al., 2012; Cornforth et al., 2002; Loucas et al., 2013).

High Unspecific

High

All life stages

High High High

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Blakely (2000), “Induction of premature chromosome condensation by a phosphatase inhibitor and a protein kinase in unstimulated human peripheral blood lymphocytes: a simple and rapid technique to study chromosome aberrations using specific whole-chromosome DNA hybridization probes for biological dosimetry”, Mutation Research, Vol. 466/2, Elsevier B.V., Amsterdam, https://doi/org/10.1016/S1383-5718(00)00011-5  Redon, R. et al. (2006), “Global variation in copy number in the human genome”, Nature. 444(7118):444-54. 10.1038/nature05329. Rodrigues, M. A., Beaton-Green, L. A., & Wilkins, R. C. (2016), “Validation of the Cytokinesis-block Micronucleus Assay Using Imaging Flow Cytometry for High Throughput Radiation Biodosimetry”, Health Phys. 110(1): 29-36. doi:10.1097/HP.0000000000000371 Schwartz, G. G. (1990), “Chromosome aberrations. Biological Markers in Epidemiology (BS Hulka, TC Wlwosky, and JD Griffith, Eds.)”, Oxford University Press, Oxford, pp.147-172.   Shahane S, Nishihara K, Xia M. (2016), “High-Throughput and High-Content Micronucleus Assay in CHO-K1 Cells”, In: Zhu H, Xia M, editors. High-Throughput Screening Assays in Toxicology. New York, NY: Humana Press. p 77-85. Shen.TW,  (2016),”Concurrent detection of targeted copy number variants and mutations using a myeloid malignancy next generation sequencing panel allows comprehensive genetic analysis using a single testing strategy”, Br J Haematol. 173(1):49-58. doi: 10.1111/bjh.13921. Shlien A, Malkin D. (2009), “Copy number variations and cancer”, Genome Med. 1(6):62. doi: 10.1186/gm62. Tucker, J.D., Preston, R.J. (1996), “Chromosome aberrations, micronuclei, aneuploidy, sister chromatid exchanges, and cancer risk assessment”, Mutat Res, 365:147-159. Vick, E. et al. (2017), Age-related chromosomal aberrations in patients with diffuse large B-cell lymphoma, American Society of Hematology, https://doi.org/10.1182/blood.V130.Suppl_1.1571.1571  Wilson, TE. et al.  (2015), “Large transcription units unify copy number variants and common fragile sites arising under replication stress”, Genome Res. 25(2):189-200. doi: 10.1101/gr.177121.114. Wink, S. et al. (2014), “Quantitative high content imaging of cellular adaptive stress response pathways in toxicity for chemical safety assessment”, Chem Res Toxicol, 27:338-355. Zhang N, Wang M, Zhang P, Huang T. 2016. Classification of cancers based on copy number variation landscapes. Biochim Biophys Acta.  1860(11 Pt B):2750-5. doi: 10.1016/j.bbagen.2016.06.003. AOPWiki 2019-05-19T16:34:32

2024-03-08T12:20:39

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AOPWiki 2026-06-29T16:53:50

2026-06-29T16:53:50

<upstream-id>19b3e408-1b2c-42e1-8cc1-9494bd32f9de</upstream-id> <downstream-id>5e73c095-d52c-4844-9379-6a2af442e148</downstream-id>

AOPWiki 2021-05-17T09:13:57

2021-05-17T09:13:57

<upstream-id>19b3e408-1b2c-42e1-8cc1-9494bd32f9de</upstream-id> <downstream-id>17e0b8d9-bbd6-410f-be2d-f759eacefdf2</downstream-id> Bulky DNA adducts occur when aromatic compounds are metabolically activated and interact with DNA bases. Not all of these bulky adductsare stable, however some have been found to persist and cause mutations during repair or replication. The specific mutation that occurs variesby bulky DNA adduct and by chemical. Exposure to the benzo(a)pyrene (B(a)P) or its metabolite anti-benzo(a)pyrene diol epoxide (BPDE)leads to (+/-)-trans-anti-BPDE-N-2-dG adducts, these adducts are associated with G→T transversions (Chiapperino et al. 2002; Zhang et al.2000, 2002), the occurrence of these transversions has been observed both in smokers (Anna et al. 2009; Hainaut and Pfeifer 2001) and in non-smokers (DeMarini et al. 2001). Exposure to aristicholic acid (AA) leads to the persistent DNA adduct 7-(deoxyadenosin-N6-yl) aristolactamI (dA–AAI) adducts and leads to AT→TA transversions (Arlt et al., 2002). Exposure to aflatoxin B1 has been leads to 8,9-dihydro-8- (N7-guanyl)-9-hydroxyaflatoxin B1 (AFB1-N7-Gua) adducts, which can lead to the AFB1-formamidopyrimidine (FAPY) adduct and ultimately causeG→T transversions (Bailey et al. 1996; Smela et al. 2002).

There is a large body of evidence that describes the relationship between bulky DNA adducts and mutations (Alexandrov et al. 2002; Chen etal. 2008; Veglia, Matullo, and Vineis 2003; Yagi et al. 2017). The bulky DNA adducts preferentially pair with an erroneous base, resulting in amutation, the mutation that results depends on the specific bulky DNA adduct that occurs. Aristolochic acid and plants containing aristolochic acid have been found to be carcinogenic to humans due to the specific DNA adducts and theA:T to T:A transversions found in renal tissues of exposed populations (IARC 2011). Exposure to AA leads to the formation of the adduct dA-AAI. In experiments with modified bacteriophage T7 DNA polymerase and with human DNA polymerase α, dA-AAI has been found to pairequally well with adenine or tyrosine (Broschard et al. 1994; Broschard, Wiessler, and Schmeiser 1995). Pairing with tyrosine results in a non-mutagenic event, therefore mutations resulting from dA-AAI are AT→TA transversion (Arlt, Stiborova, and Schmeiser 2002; Kohara et al.2002). Aristolochic acid and plants containing aristolochic acid are considered carcinogenic to humans due to the specific DNA adducts and theA:T to T:A transversions found in renal tissues of exposed populations (IARC, 2011).  B(a)P is a known to be carcinogenic to humans due to extensive experimental evidence in many animal species along with mechanisticevidence to support the biological plausibility of bulky DNA adducts leading to mutations that cause cancer in humans (IARC 2014). Exposureto the B(a)P or its metabolite anti-benzo(a)pyrene diol epoxide (BPDE) leads to (+/-)-trans-anti-BPDE-N-2-dG adducts. Human DNApolymerase eta has been found to insert an A across from the (+/-)-trans-anti-BPDE-N-2-dG adducts, resulting in the above mentioned GvTtransversions (Chiapperino et al. 2002; Zhang et al. 2000, 2002). Polymerase eta has been found to be unlikely to extend past the lesion (Chiapperino et al. 2002)and instead polymerase kappa has been found to work as the second step in the bypass of this lesion (Zhang et al.2002). Another common mutation occurring from the bulky DNA lesion (+)-trans -anti-BPDE-N-2-dG is a G→A transversion. Through molecularmodelling, it has been suggested DNA polymerase may be more likely to insert a T if the bulk of the adduct is in the major groove and an A ifthe bulk of the adduct is in the minor groove (Kozack, Shukla, and Loechler 1999).

1. Dose concordance   <ul> <li>MutaMouse models were exposed to 11 concentrations of BaP (0, 0.10, 0.20, 0.39, 0.78, 1.56, 3.13, 6.25, 12.50, 25.00, and 50.00 mgBaP/kg body weight (BW)/day) for 28 days (Long et al. 2018) <ul style="list-style-type:circle"> <li>Bulky DNA adducts were measured using 32P post-labeling. A significant increase in bulky DNA adducts was observed in the mostsensitive tissue at 0.20 mg BaP/kg body weight (BW)/day and in all tissues at 1.56 mg BaP/kg body weight (BW)/day.</li> <li>Mutations were measured using the LacZ mutation assays. A significant increase in mutations was observed in the most sensitive tissue at 1.56 mg BaP/kg body weight (BW)/day and in all tissues 25.00 mg BaP/kg body weight (BW)/day. </li> <li>These results indicate that the formation of bulky DNA adducts occurs at lower doses than the occurrence of mutations as measured by the lacZ assay. This study also calculated BMDs for both assays. The BMD for bulky DNA adducts in bone marrow was 0.0286 (0.196 - 0.0661), the BMD for lacZ mutations in bone marrow was 2.22 (1.81 - 2.75). These BMDs support that bulky DNA adducts occur at lower doses than mutations.</li> </ul> </li> <li>TK6 cells exposed to BPDE (Akerman et al., 2004) <ul style="list-style-type:circle"> <li>After exposure to 0 uM, 0.01 uM, 0.10 uM and 1.00 uM BPDE for 4 hours with 4 hour recovery and exposure for 4 hours with 24 hour recovery, bulky DNA adducts were measured by 32P post-labeling. Samples exposed to 0.01 uM, 0.10 uM and 1.00 uM BPDE had significantly more adducts than control.</li> <li>After exposure for 4 hours and 3 day recovery (TK gene) or exposure for 4 hours and 7 day recovery (HPRT gene), mutation frequency in the TK and HPRT genes was measured by only 1.00 uM BPDE had significantly more mutations than control</li> <li>Therefore after a 4 hour exposure of TK6 cells to BPDE, the frequency of bulky DNA adducts increased at lower doses than the frequency of mutations in the TK and HPRT genes increased.</li> </ul> </li> <li>Yeast p53 cDNA samples were treated with 0, 2.5 uM, 5 uM, 10 uM, 20 uM (+/-)-Anti-BPDE for 3 hours (Park et al. 2008) <ul style="list-style-type:circle"> <li>Stable (+)-Anti-BPDE-N-dGuo Adducts were detected with LC-MS and quantified with HPLC. A linear dose response was observed, ranging from 150 adducts per 105 dGuo at 2.5 uM anti-BPDE to 940 adducts per 105 dGuo at 20 uM.</li> <li>Mutations were detected with a yeast reporter system. A linear correlation (R=0.8411) was found between the incidence of thespecific bulky DNA adduct (+)-Anti-BPDE-N-dGuo and the percentage of mutated yeast colonies.</li> <li>These results suggest that the specific bulky DNA adduct (+)-Anti-BPDE-N-dGuo is incorporated into a mutagenic lesion in a dose dependant manner.</li> </ul> </li> </ul>   <ol start="2"> <li>Temporal concordance</li> </ol>   <ul> <li>  TK6 cells exposed to BPDE (Akerman et al., 2004) <ul style="list-style-type:circle"> <li>After exposure to 0 uM, 0.01 uM, 0.10 uM and 1.00 uM BPDE for 4 hours with 4 hour recovery and exposure for 4 hours with 24 hour recovery, bulky DNA adducts were measured by 32P post-labeling. Samples exposed to 0.01 uM, 0.10 uM and 1.00 uM BPDE had significantly more adducts than control.</li> <li>After exposure for 4 hours and 3 day recovery (TK gene) or exposure for 4 hours and 7 day recovery (HPRT gene), mutation frequency in the TK and HPRT genes was measured by only 1.00 uM BPDE had significantly more mutations than control</li> <li>Therefore, after a 4 hour exposure of TK6 cells to BPDE, the frequency of bulky DNA adducts increased at earlier timepoints (4 or 24 hours) than the timepoints where mutations in the TK and HPRT genes increased (3 or 7 days).</li> </ul> </li> </ul>

There is some quantitative understanding of the amount of bulky DNA adducts that leads to mutations. <ul> <li>Broschard et al. 1994 and Broschard, Wiessler, and Schmeiser 1995 found that T7 DNA polymerase and with human DNA polymerase α,respectively, paired dA-AAI equally well with A or T, suggesting that there is a 50% chance of a dA-AAI lesion will lead to a mutation.</li> </ul>   <ul> <li>Zhang et al. 2000 found that polymerase eta predominately incorporated A opposite (+)-trans -anti-BPDE-N2-dG bulky adducts, lessfrequently a T was incorporated and least frequently a G or C was incorporated. Suggesting that it is most likely the persistence of a (+)-trans -anti-BPDE-N2 -dG bulky adduct will lead to a mutation of a G to T transversion.</li> </ul> There are also studies demonstrating the quantitative dose-response between bulky DNA adducts and mutations, see Empirical Evidence.

Not Specified Unspecific

Not Specified

All life stages

Not Specified Not Specified Not Specified Not Specified Not Specified

Bulky DNA adducts can occur in any cell type that is able to metabolically activate the stressor. Bulky adducts and resulting mutation frequencyhave been observed in various cell lines in vitro (TK6, HeLa, CHO) as well as various organisms in vivo (yeast, rat, human and mouse). This is unspecific to sex and to life stage.

Akerman, G. S. et al. 2004. “Gene Expression Profiles and Genetic Damage in Benzo(a)Pyrene Diol Epoxide-Exposed TK6 Cells.” Mutation Research - Fundamental and Molecular Mechanisms of Mutagenesis 549(1–2): 43–64. Alexandrov, Kroum et al. 2002. “CYP1A1 and GSTM1 Genotypes Affect Benzo[a]Pyrene DNA Adducts in Smokers’ Lung: Comparisonwith Aromatic/Hydrophobic Adduct Formation.” Carcinogenesis 23(12): 1969–77. Anna, Lívia et al. 2009. “Relationship between TP53 Tumour Suppressor Gene Mutations and Smoking-Related Bulky DNA Adducts in aLung Cancer Study Population from Hungary.” Mutagenesis 24(6): 475–80. Arlt, Volker M., Marie Stiborova, and Heinz H. Schmeiser. 2002. “Aristolochic Acid as a Probable Human Cancer Hazard in HerbalRemedies: A Review.” Mutagenesis 17(4): 265–77. Bailey, Elisabeth A. et al. 1996. “Mutational Properties of the Primary Aflatoxin B1-DNA Adduct.” Proceedings of the National Academy of Sciences of the United States of America 93(4): 1535–39. Broschard, Thomas H., Manfred Wiessler, Claus Wilhelm Von Der Lieth, and Heinz H. Schmeiser. 1994. “Translesional Synthesis on DNATemplates Containing Site-Specifically Placed Deoxyadenosine and Deoxyguanosine Adducts Formed by the Plant CarcinogenAristolochic Acid.” Carcinogenesis 15(10): 2331–40. Broschard, Thomas H., Manfred Wiessler, and Heinz H. Schmeiser. 1995. “Effect of Site-Specifically Located Aristolochic Acid DNAAdducts on in Vitro DNA Synthesis by Human DNA Polymerase α.” Cancer Letters 98(1): 47–56. Chen, Yanan et al. 2008. “Electronic Detection of Lectins Using Carbohydrate Functionalized Nanostructures: Graphene versus CarbonNanotubes.” Nano 6(9): 2166–71. Cho, Eunnara et al. 2020. “Oxidative DNA Damage Leading to Chromosomal Aberrations and Mutations.” AOP Wiki:https://aopwiki.org/aops/296. Chiapperino, Dominic et al. 2002. “Preferential Misincorporation of Purine Nucleotides by Human DNA Polymerase η OppositeBenzo[a]Pyrene 7,8-Diol 9,10-Epoxide Deoxyguanosine Adducts.” Journal of Biological Chemistry 277(14): 11765–71.http://dx.doi.org/10.1074/jbc.M112139200. DeMarini, David M. et al. 2001. “Lung Tumor KRAS and TP53 Mutations in Nonsmokers Reflect Exposure to PAH-Rich Coal Combustion Emissions.” Cancer Research 61(18): 6679–81. Hainaut, Pierre, and Gerd P. Pfeifer. 2001. “Patterns of P53→T Transversions in Lung Cancers Reflect the Primary Mutagenic Signature ofDNA-Damage by Tobacco Smoke.” Carcinogenesis 22(3): 367–74. IARC. 2011. “Plants Containing Aristolochic Acid.” IARC Monographs on the Evaluation of Carcinogenic Risks to Humans 100 A: 367–83.  IARC. 2014. “Benzo(a)Pyrene.” In Chemical Agents and Related Occupations IARC Monographs on the Evaluation of Carcinogenic Risksto Humans Volume 100F, 423–28.  Kohara, Arihiro et al. 2002. “Mutagenicity of Aristolochic Acid in the Lambda/LacZ Transgenic Mouse (MutaMouse).” Mutation Research- Genetic Toxicology and Environmental Mutagenesis 515(1–2): 63–72.  Kozack, Richard E., Rajiv Shukla, and Edward L. Loechler. 1999. “A Hypothesis for What Conformation of the Major Adduct of(+)-Anti-B[a]PDE (N2-DG) Causes G→T versus G→A Mutations Based upon a Correlation between Mutagenesis and Molecular Modeling Results.” Carcinogenesis 20(1): 95–102.  Long, Alexandra S. et al. 2018. “Benchmark Dose Analyses of Multiple Genetic Toxicity Endpoints Permit Robust, Cross-Tissue Comparisons of MutaMouse Responses to Orally Delivered Benzo[a]Pyrene.” Archives of Toxicology 92(2): 967–82.https://doi.org/10.1007/s00204-017-2099-2.  Mei, Nan et al. 2006. “DNA Adduct Formation and Mutation Induction by Aristolochic Acid in Rat Kidney and Liver.” Mutation Research -Fundamental and Molecular Mechanisms of Mutagenesis 602(1–2): 83–91. Park, Jong Heum et al. 2008. “Erratum: The Pattern of P53 Mutations Caused by PAH o-Quinones Is Driven by 8-Oxo-DGuo FormationWhile the Spectrum of Mutations Is Determined by Biological Selection for Dominance (Chemical Research in Toxicology (2008) 21:5(1039-1049)).” Chemical Research in Toxicology 21(9): 1907.  Smela, Maryann E. et al. 2002. “The Aflatoxin B1 Formamidopyrimidine Adduct Plays a Major Role in Causing the Types of Mutations Observed in Human Hepatocellular Carcinoma.” Proceedings of the National Academy of Sciences of the United States of America 99(10):6655–60.  Veglia, Fabrizio, Giuseppe Matullo, and Paolo Vineis. 2003. “Bulky DNA Adducts and Risk of Cancer: A Meta-Analysis.” Cancer Epidemiology Biomarkers and Prevention 12(2): 157–60.  Yagi, Takashi et al. 2017. “Error-Prone and Error-Free Translesion DNA Synthesis over Site-Specifically Created DNA Adducts of Aryl Hydrocarbons (3-Nitrobenzanthrone and 4-Aminobiphenyl).” Toxicological Research 33(4): 265–72.  Zhang, Yanbin et al. 2000. “Error-Prone Lesion Bypass by Human DNA Polymerase η.” Nucleic Acids Research 28(23): 4717–24. Zhang, Yanbin et al. 2002. “Two-Step Error-Prone Bypass of the (+)- and (-)-Trans-Anti-BPDE-N2-DG Adducts by Human DNAPolymerases η and κ.” Mutation Research - Fundamental and Molecular Mechanisms of Mutagenesis 510(1–2): 23–35. AOPWiki 2021-05-17T09:14:17

2021-09-14T16:47:57

<upstream-id>19b3e408-1b2c-42e1-8cc1-9494bd32f9de</upstream-id> <downstream-id>6f3cc035-087f-41e7-bdfa-80177f2f7b4e</downstream-id>

AOPWiki 2026-06-29T16:53:34

2026-06-29T16:53:34

<upstream-id>669b23ca-0e30-4508-ae51-b6d96fb4949d</upstream-id> <downstream-id>17e0b8d9-bbd6-410f-be2d-f759eacefdf2</downstream-id> DNA single strand breaks (SSB) are generally repaired rapidly and efficiently. However, if left unrepaired, SSBs can interfere with replication and cause the replication fork to collapse resulting in double strand breaks (DSB). Multiple SSBs in close proximity to each other can also give rise to DSBs. DSBs can be repaired virtually error-free by homologous recombination (HR), which uses DNA sequence in the homologous chromosome or sister chromatid as a template for new strand synthesis (Polo and Jackson, 2011). Alternatively, the broken ends may be joined to other sites in the genome regardless of homology via non-homologous end joining (NHEJ), irreversibly altering the DNA sequence (deletion, addition, rearrangement). Because HR is a more time-consuming and labour-intensive process, larger proportions of DSBs are repaired via NHEJ than via HR (Mao et al., 2008a; Mao et al., 2008b). Alterations in DNA sequence can also occur from structural damage to the chromosomes; observations of micronucleus indicate chromosomal aberrations and that a permanent loss of DNA segments has occurred.

The error-prone nature of DSB repair in eukaryotes has been described in numerous reviews. In mammalian and yeast cells, both HR and NHEJ can lead to alteration in DNA sequence; insertions, deletions, and translocations can arise from NHEJ and base substitutions can occur during the repair synthesis of HR (Hicks and Haber, 2010; Bunting and Nussenzweig, 2013; Byrne et al., 2014; Rodgers and McVey, 2016; Dwivedi and Haber, 2018).

The mechanisms by which strand breaks lead to mutations are very well studied and understood. Thus, we provide a small selection of empirical evidence below supporting this KER; i.e., we did not undertake and exhaustive literature search. In vitro studies <ul> <li>Strand breaks and mutation frequencies were measured in TK6 cells after exposure to bleomycin and glucose oxidase (enzyme that generates H2O2) for 1 hour (Platel et al., 2011). <ul style="list-style-type:circle"> <li>Concentration-dependent increases in strand breaks were measured using the alkaline comet assay.</li> <li>At the same concentrations, mutation frequencies measured by TK gene mutation assay also showed a concentration-dependent increasing trend.</li> <li>No Observed Genotoxic Effect Level was determined in TK assay (bleomycin: 0.6µM; GOx: 1.17x10-5 units/mL) while it couldn’t be identified in comet assay, indicating that every tested concentration induced an increase in strand breaks (First statistically significant concentration: bleomycin: 1.5 µM; GOx: 1.08x10-5 units/mL).</li> </ul> </li> </ul>   <ul> <li>Spassova et al. (2013) combined the alkaline comet assay data from Luan et al. (2007) and Tk gene mutation assay data from Harrington-Brock et al. (2003) (Spassova et al., 2013). <ul style="list-style-type:circle"> <li>Luan et al. treated TK6 cells with KBrO3 for 4 hours and performed alkaline comet assay to measure strand breaks.</li> <li>Harrington-Brock et al. treated L5178Y/Tk+/- mouse lymphoma cells with KBrO3 for 4 hours and measured the Tk mutant frequency after a 13-day incubation.</li> <li>Spassova et al. (2013) found no significant differences between the two experiments in regression analysis, thus, combined the datasets (same concentration range was used in both studies)</li> <li>In both comet assay and Tk mutation assay, concentration-dependent increase in response was observed.</li> <li>These results demonstrate the occurrence of DNA strand breaks followed by increase in mutations.</li> </ul> </li> </ul>   <ul> <li>Indirect measurement of mutations by measuring misrejoined DSBs in vitro <ul style="list-style-type:circle"> <li>Rydberg et al. (2005) exposed GM38 human primary dermal fibroblasts to increasing doses of X-rays and linear electron transfer (LET) by nitrogen, helium, and iron ions.</li> <li>DSBs were measured by pulsed field gel electrophoresis (PFGE) <ul> <li>Dose-dependent increase in DSBs was observed immediately following irradiation.</li> </ul> </li> <li>Mi<ins>s</ins>rejoining of ends was monitored using the Hybridization assay: <ul> <li>DNA is digested using a restriction enzyme and fractionated by PFGE.</li> <li>32P-labeled probe for a 3.2-Mbp NotI restriction fragment is then used in Southern blotting to detect intact restriction fragments.</li> <li>Failure to reconstitute the restriction fragment indicates incorrect joining of ends following DSBs and altered DNA sequence.</li> </ul> </li> <li>After 16 h of recovery following irradiation, Rydberg et al. observed a radiation dose-dependent increase in misrejoined DSBs in all four treatment groups.</li> <li>A similar study by Kuhne et al. (2005) reported concordant results (Kuhne et al., 2005): <ul style="list-style-type:circle"> <li>Subsequently, there was a dose-dependent increase in misrejoined DSBs 24h post irradiation.</li> <li>Increasing doses of X-rays and γ rays immediately induced DSBs in primary human fibroblasts in a dose-dependent manner.</li> <li>Alterations in the restriction fragment due to irradiation indicate changes in the DNA sequence (i.e., shorter fragments would suggest loss of DNA sequence), thus, induction of mutations (Rydberg et al., 2005; Kuhne et al., 2005).</li> <li>These results demonstrate the concentration and temporal concordance in strand breaks leading to mutations.</li> </ul> </li> </ul> </li> </ul>   <ul> <li>In a study by Kuhne et al. (2000), irradiated normal human fibroblasts were examined for both DSBs and the percentage of misrejoined DSBs (Kuhne et al., 2000). <ul> <li>Increasing doses of alpha-particle radiation from 0 – 80 Gy resulted in a linear, dose-dependent increase in the number of DSBs per mega base pair, as measured by the FAR assay.</li> <li>Using X-ray radiation, the percentage of misrejoined DSBs were found to increase approximately linearly from 0 – 40 Gy doses per fraction. By 80 Gy, the rate of misrejoining plateaued at approximately 50%, and this plateau was maintained at X-ray doses between 80 and 320 Gy.</li> <li>Overall, these results provide indirect evidence suggesting that elevated numbers of DSBs may lead to the formation of increasingly more mutations, as indicated by the corresponding increased number of misrejoined DSBs.</li> </ul> </li> </ul>   <ul> <li>Dikomey et al. (2000) performed a study using normal human skin fibroblasts that were irradiated with 200 kVp X-rays at doses ranging from 0 – 180 Gy, and then were examined for DSBs immediately following irradiation, and for non-repaired DSBs 24 hours after radiation exposure (Dikomey and Brammer, 2000). <ul> <li>As measured by constant field gel electrophoresis, there was a dose-dependent increase in the number of DSBs after exposure to X-rays doses of 0 – 80 Gy.</li> <li>The number of non-repaired DSBs also increased with increasing radiation dose from 0 – 180 Gy. After 30 Gy, there were more non-repaired DSBs when cells were exposed to radiation with a high dose-rate (4 Gy/min) relative to those exposed to radiation with a low dose-rate (0.4 Gy/min).</li> <li>These results suggest that there are increasing DSBs with increasing radiation dose, and that there are also an increasing number of DSBs that are not repaired with increasing radiation dose. This is important as non-repaired DSBs may result in mutations in the genome.</li> </ul> </li> </ul>   <ul> <li>Both lung and dermal fibroblasts were irradiated with 80 kV X-rays at 23 Gy/min, and analyzed for the number of DSBs and the percentage of correctly rejoined DSBs in a study by (Lobrich et al., 2000). <ul> <li>Results from the FAR assay showed a linear increase in the number of DSBs in all cell lines for radiation doses ranging from 0 – 80 Gy.</li> <li>After being irradiated with 80 Gy of X-rays, approximately 50% of the DSBs were correctly rejoined, as measured by the hybridization assay.</li> <li>A dose-dependent increase in the number of rearrangements per mega base pair was found in cells irradiated with 0 – 80 Gy of X-rays.</li> <li>The results of this study provide evidence of dose concordance, as the number of DSBs and the number of rearrangements both increase with increasing radiation dose.</li> </ul> </li> </ul> In vivo studies <ul> <li>Strand breaks and mutation frequencies were measured in the leaves of Nicotiana tabacum var. xanthi after the seedling plants were irradiated with 0 – 10 Gy doses of gamma-ray radiation (Ptacek et al., 2001). <ul> <li>DNA strand breaks in the leaves were measured using the Comet assay immediately following irradiation. Results of this assay showed a linear, dose-dependent increase in strand breaks, which were resolved by 24 hour post-irradiation.</li> <li>Mutations in the leaves were measured when the seedling plants put out their 6th or 7th true leaves following irradiation. Similar to results found for radiation-induced strand breaks, there was a corresponding dose-dependent increase in the number of mutations per radiation dose.</li> <li>These results demonstrate a dose concordance between DNA strands breaks and mutation frequency, and suggest a time concordance.</li> </ul> </li> </ul>

In Kuhne et al. (2005) and Rydberg et al. (2005) studies provided above, mutation was not directly measured. The PFGE and hybridization assay detects a 3.2-Mbp restriction fragment from chromosome 21. Deviation of DNA restriction fragments from the 3.2-Mbp mark during electrophoresis suggests occurrence of breakage and failed reconstruction in this segment of chromosome 21; induction of mutations can be inferred from the change in the size of the restriction fragments. The remaining 22 chromosomes are not considered. This method may not be sensitive enough to detect small base changes. Cell cycle can influence the repair pathway of DSBs and, thus, the risk of incorrect rejoining of broken ends. In G1 phase, NHEJ may be favoured, while in S, G2, or M phase, both HR and NHEJ have been observed to be active in repair (Mao et al., 2008b).

McMahon et al. (2016) compiled the data from multiple studies spanning different human and mouse cell lines (including Lobrich et al. (2000) and Rydberg et al. (2005)) to model the IR dose-dependent increase in chromosomal aberrations, misrejoined DSBs, and mutation rate (per 104 cells). The data used to model the DSB misrepair rate were generated from the hybridization assay described in the Empirical evidence section. The mutation rate model was based on HPRT gene mutation assay in Chinese hamster cells (McMahon et al., 2016). Previously, in another study, the number of γH2AX foci formed following IR irradiation was quantified over increasing doses of radiation in human fibroblasts (Rothkamm and Lobrich, 2003). Further quantitative studies that exam the relationships between the quantities of strand breaks (e.g., quantity of γH2AX foci (DSBs) formed), DSB misrepair rate, and mutation rate would provide a better quantitative understanding of this KER.

High Unspecific

High

All life stages

High High High

DNA strand breaks and subsequent mutations can occur in any eukaryotic and prokaryotic cell. Any DNA strand break has potential to cause alterations in DNA sequence (e.g., deletions and insertions), whether it is due to insufficient or faulty repair.

Bunting, S. & A. Nussenzweig (2013), "End-joining, Translocations and Cancer", Nat Rev Cancer, 13:443-454. Byrne, M. et al. (2014), "Mechanisms of oncogenic chromosomal translocations", Ann. N.Y. Acad. Sci., 1310:89-97. Dikomey, E. & I. Brammer (2000), "Relationship between cellular radiosensitivity and non-repaired double-strand breaks studied for different growth states, dose rates and plating conditions in a normal human Žfibroblast line.", Int. J. Radiat. Biol., 76:773-781. Dwivedi, G. & J.E. Haber (2018), "Assaying Mutations Associated With Gene Conversion Repair of a Double-Strand Break", Methods Enzymiol., 601:145-160. Hicks, W. & J.E. Haber (2010), "Increased Mutagenesis and Unique Mutation Signature Associated with Mitotic Gene Conversion", Nat. Rev. Cancer, 329:82-84. Kuhne, M., K. Rothkamm & M. Lobrich (2000), "No dose-dependence of DNA double-strand break misrejoining following a -particle irradiation.", Int. J. Radiat. Biol. 76(7):891-900 Kuhne, M., G. Urban & M. Lo, (2005), "DNA Double-Strand Break Misrejoining after Exposure of Primary Human Fibroblasts to C K Characteristic X Rays, 29 kVp X Rays and Co g-Rays.", Radiation Research. 164(5):669-676. doi:10.1667/RR3461.1. Lobrich, M. et al. (2000), "Joining of Correct and Incorrect DNA Double-Strand Break Ends in Normal Human and Ataxia Telangiectasia Fibroblasts.", 68(July 1999):59–68. doi: 10.1002/(SICI)1098-2264(200001)27:1<59::AID-GCC8>3.0.CO;2-9. Mao, Z. et al. (2008a), "Comparison of nonhomologous end joining and homologous recombination in human cells.", DNA Repair, 7:1765-1771. Mao, Z. et al. (2008b), "DNA repair by nonhomologous end joining and homologous recombination during cell cycle in human cells.", Cell Cycle, 7:2902-2906. McMahon, S.J. et al. (2016), "Mechanistic Modelling of DNA Repair and Cellular Survival Following Radiation-Induced DNA Damage.", Nat. Publ. Gr.(April):1–14. doi:10.1038/srep33290. Platel, A. et al. (2011), "Study of oxidative DNA damage in TK6 human lymphoblastoid cells by use of the thymidine kinase gene-mutation assay and the in vitro modified comet assay: Determination of No-Observed-Genotoxic-Effect-Levels.", Mutat. Res., 726:151-159. Polo, S. & S. Jackson (2011), "Dynamics of DNA damage response proteins at DNA breaks: a focus on protein modifications.", Genes Dev., 25:409-433. Ptacek, O. et al. (2001), "Induction and repair of DNA damage as measured by the Comet assay and the yield of somatic mutations in gamma-irradiated tobacco seedlings.", Mutat. Res., 491:17-23. Rodgers, K. & M. McVey (2016), "Error-prone repair of DNA double-strand breaks.", J. Cell. Physiol., 231:15-24. Rothkamm, K. & M. Lobrich (2003), "Evidence for a lack of DNA double-strand break repair in human cells exposed to very low x-ray doses.", PNAS, 100(9):5057-62. doi:10.1073/pnas.0830918100. Rydberg, B. et al. (2005), "Dose-Dependent Misrejoining of Radiation-Induced DNA Double-Strand Breaks in Human Fibroblasts: Experimental and Theoretical Study for High- and Low-LET Radiation.", Radiat. Res. 163(5):526–534. doi:10.1667/RR3346. Spassova, M. et al. (2013), "Dose-Response Analysis of Bromate-Induced DNA Damage andMutagenicity Is Consistent With Low-Dose Linear,Nonthreshold Processes", Environ. Mol. Mutagen., 54:19-35.   AOPWiki 2019-06-02T15:35:55

2023-01-09T21:05:00

<upstream-id>5e73c095-d52c-4844-9379-6a2af442e148</upstream-id> <downstream-id>669b23ca-0e30-4508-ae51-b6d96fb4949d</downstream-id> Inadequate repair of DNA damage includes incorrect repair (i.e., incorrect base insertion), incomplete repair (i.e., accumulation of repair intermediates such as strand breaks, stalled replications forks, and/or abasic sites), and absent repair resulting in the retention of DNA damage. It is well-established that DNA excision repair pathways require DNA strand breakage for removing the damaged sites; for example, base excision repair (BER) of oxidative lesions involves removal of oxidized bases by glycosylases followed by cleavage of the DNA strand 5’ from the abasic site. If the repair process is disrupted at this point, repair intermediates including single strand breaks (SSB) may persist in the DNA. A SSB can turn into a double strand break (DSB) if it occurs sufficiently close to another SSB on the opposite strand. SSBs can be converted into DSBs when helicase unwinds the DNA strands during replication. Furthermore, SSBs and abasic sites can act as replication blocks causing the replication fork to stall and collapse, giving rise to DSBs (Minko et al., 2016; Whitaker et al., 2017). <div> <div> The two most common DSB repair mechanisms are non-homologous end joining (NHEJ) and homologous recombination (HR). NHEJ is may favoured over HR and has also been shown to be 104 times more efficient than HR in repairing DSBs (Godwin et al., 1994; Benjamin and Little, 1992). There are two subtypes of NHEJ: canonical NHEJ (C‐NHEJ) or alternative non-homologous end joining (alt-NHEJ). During C-NHEJ, broken ends of DNA are simply ligated together. In alt‐NHEJ, one strand of the DNA on either side of the break is resected to repair the lesion (Betermeir et al., 2014). Although both repair mechanisms are error‐prone (Thurtle‐Schmidt and Lo, 2018), alt-NHEJ is considered more error-prone than C-NHEJ (Guirouil-Barbat et al., 2007; Simsek and Jasin, 2010). While NHEJ may prevent cell death due to the cytotoxicity of DSBs, it may lead to mutations and genomic instability downstream.   </div> </div>

1. DNA strand breaks generated due to faulty attempted repair Excision repair pathways require the induction of SSB as part of damage processing. Increases in DNA lesions may lead to the accumulation of intermediate SSB. Attempted excision repair of lesions on opposite strands can turn into DSBs if the two are in close proximity (Eccles et al., 2010). Generation of DSBs has been observed in both nucleotide excision repair (NER) and BER (Ma et al., 2009; Wakasugi et al., 2014). Previous studies have demonstrated that an imbalance in one of the multiple steps of BER can lead to an accumulation of repair intermediates and failed repair. It is highly likely that a disproportionate increase in oxidative DNA lesions compared to the level of available BER glycosylases leads to an imbalance between lesions and the initiating step of BER (Brenerman et al., 2014). Accumulation of oxidative lesions, abasic sites, and SSBs generated from OGG1, NTH1, and APE1 activities would be observed as a result. Moreover, studies have reported accumulation of SSB due to OGG1- and NHT1-overexpression (Yang et al., 2004; Yoshikawa et al., 2015; Wang et al., 2018). BER repair intermediates have been observed to interfere with transcription as well (Kitsera et al., 2011). While overexpression may lead to imbalanced lyase activities that generate excessive SSB intermediates, deficiency of these enzymes is also known to cause an accumulation of oxidative lesions that could lead to strand breaks downstream. Hence, both the overexpression and deficiencies of repair enzymes can lead to strand breaks due to excessive activity or inadequate repair, respectively. 2. DNA strand breaks generated due to replication stress caused by accumulated DNA lesions Retention of DNA lesions (i.e., damaged bases and SSB) can interfere with the progression of the replication fork. Thymidine glycol is an example of an oxidative DNA lesion that acts as a replication block (Dolinnaya et al., 2013). Persistent replication fork stalling and dissociation of replication machinery are known to cause the replication fork to collapse, which generates highly toxic DSBs (Zeman and Cimprich, 2014; Alexander and Orr-Weaver, 2016). Fork stalling also increases the risk of two replication forks colliding with each other, generating DSBs. In addition, the replication fork can collide with SSBs generated during BER, hindering the completion of repair and giving rise to DSBs (Ensminger et al., 2014). <div> <div> <div>   </div> </div> </div>

In vitro studies with empirical evidence are shown below for select DNA repair pathways. These studies build in elements of essentiality (modulation of DNA repair), as well as dose and incidence concordance. The primary evidence is essentiality, where repair is genetically modulated in some way. Because multiple lines of evidence are considered within individual studies, we present the data by source of evidence (in vitro versus in vivo) rather than by type of empirical evidence (dose, incidence, or temporal concordance; essentiality) to avoid repetitive use of the same studies.   Inadequate repair of oxidative lesions <ul> <li>Concentration concordance of strand breaks in repair-deficient and –proficient cells (insufficient repair) (Wu et al., 2008) <ul style="list-style-type:circle"> <li>In a study using A549 human adenocarcinoma cells, DNA strand breaks in hOGG1-proficient and hOGG1-deficient cells were compared following exposure to increasing concentrations of bleomycin.</li> <li>Strand breaks were measured as DNA migration length in alkaline comet assay after 3 hours of exposure to six increasing concentrations (0.05, 0.25, 0.5, 1, 5, and 10 mg/L).</li> <li>Concentration-dependent increase in strand breaks was observed in both cell types; however, at all concentrations significantly more strand breaks (p<0.05) were present in the hOGG1-deficient cells than in the proficient cells, demonstrating insufficient repair of oxidative lesions leading to DNA strand breaks.</li> <li>Thus, this evidence supports the essentiality of inadequate DNA repair as a modulator of the downstream KE.</li> </ul> </li> <li>Incomplete OGG1-initiated base excision repair (BER) leads to DNA strand breaks (Wang et al., 2018): <ul> <li>In a study using mouse embryonic fibroblasts (MEF), Ogg1+/+ and Ogg1-/- cells were treated with increasing concentrations of H2O2 for varying durations Higher levels of 8-oxodG were detected in Ogg1-/- cells compared to Ogg1+/+ cells after treatment with 400 µM H2O2 at all time points (5, 15, 30, 60, and 90 min) <ul> <li>Demonstrates insufficient removal of 8-oxo-dG in OGG1-deficient cells</li> </ul> </li> <li>Significantly  more strand breaks, as indicated by the higher % of TUNEL-positive cells (p<0.001), were detected in Ogg1+/+ cells compared to Ogg1-/- cells after exposure to 400 µM H2O2 for 3 hours <ul> <li>Both cell types showed a very similar increase in DNA strand breaks at lower concentrations (50, 100, and 200 µM) and there was no significant difference between Ogg1+/+ and Ogg1-/- cells at these concentrations – this suggests that up to a certain level of oxidative damage, OGG1-initiated BER does not exacerbate strand breaks but when oxidative stress is excessive (at 400µM in this study), OGG1-initiated BER is compromised and leads to increased strand breaks (incomplete repair)</li> </ul> </li> <li>Finally, DNA strand breaks in both cell types were measured using both alkaline and neutral comet assay after a 30- minute exposure to 400µM H2O2; while there was an increase in the olive tail moment (indicating DNA strand breaks) in both cell types compared to the control, the increase of strand breaks in Ogg1+/+ cells was significantly larger than in Ogg1-/- cells in both assays (p<0.001)</li> </ul> </li> </ul>   Inadequate repair of alkylated DNA <ul> <li>Interference of N-methylpurine DNA glycosylase (MPG)-initiated BER by replication leading to strand breaks (Ensminger et al., 2014) <ul style="list-style-type:circle"> <li>A549 human alveolar basal epithelial cells were exposed to increasing concentrations of methylmethane sulfonate (MMS) for 1 hour and replicating cells were labeled using a thymidine analogue, 5-ethynyl-2’-desoxyuridine (EdU).</li> <li>In S-phase cells, MMS concentration-dependent increase in γH2AX foci was detected (70 foci/cell at the highest concentration). In contrast, γH2AX foci were not detected G1- and G2-phase cells until the highest concentration (15 foci/cell).</li> <li>MPG-depleted cells in S-phase showed no significant increase in γH2AX foci, while the control cells showed significant MMS concentration-dependent increases.</li> <li>These results suggest interference of MPG-initiated BER by replication, leading to DSBs, and that the depletion of MPG decreases the probability of strand breaks in S-phase (evidence of essentiality of ‘inadequate repair’ to KEdown). </li> </ul> </li> </ul>   Inadequate mismatch repair <ul> <li>Incomplete/incorrect mismatch repair (MMR) leads to DNA strand breaks (Peterson-Roth et al., 2005): <ul style="list-style-type:circle"> <li>MLH1 (MMR protein)-deficient and -proficient HCT116 human colon cancer cells were treated with 30µM K2CrO4 (DNA crosslinking, Cr adducts, protein-DNA crosslinking, DNA oxidation) for 3, 6, and 12 hours and γH2AX foci (biomarker of DNA DSB) were scored by fluorescence microscopy</li> <li>At 6 and 12 hours, MLH1+ cells had higher percentage of γH2AX foci than MLH1- cells</li> <li>The futile repair model of MMR suggests that strand breaks arise from MMR attempting repeatedly to repair the newly synthesized strand opposite adducts in S and G2 phases; approximately 80% of the γH2AX-positive MLH1+ cells were in G2 phase 12 hours after a 3-hour exposure to 20 µM Cr(VI), while the level was five times lower in MLH1- cells, suggesting that the MMR-induced DSB occurred following DNA synthesis; this supports the futile repair model and demonstrates inadequate repair</li> </ul> </li> </ul>   Inadequate Repair of DSBs <ul> <li>Rydberg et al. [2005] exposed GM38 primary human dermal fibroblasts to increasing doses of linear electron transfer (LET) radiation of helium and iron ions (Rydberg et al., 2005). <ul style="list-style-type:circle"> <li>The cells were allowed to recover for 16 hours following irradiation.</li> <li>Unrepaired DSBs were measured after recovery using PFGE.</li> <li>There was a dose-dependent increase in unrepaired DSBs due to both ion exposures.</li> <li>Increase in persistent unrepaired DSBs with increasing dosage indicates exceeded repair capacity.</li> </ul> </li> <li>DSB repair was also monitored by measuring γH2AX foci 0.05 - 24 hours after irradiation. <ul> <li>DSBs decreased over time and less than 1 foci per cell on average remained in MRC-5 cells 24hours after 0.02, 0.2 and 2 Gy exposures.</li> <li>Repair was slower in 180BR cells, particularly for the 2 Gy exposure, where 20 foci per cell remained after 24 h. </li> <li>A follow-up study by the same group, found similar results for MRC-5 and 180BR cells exposed to 0.02 and 0.2 Gy of X-rays (Kühne et al., 2004). </li> </ul> </li> <li>Rothkamm and Löbrich (2003) exposed MRC-5 primary human lung fibroblasts (repair-proficient) and 180BR DNA ligase IV-deficient human fibroblasts to 10 and 80 Gy of X-rays (Rothkamm and Lobrich, 2003). <ul style="list-style-type:circle"> <li>DNA ligase IV deficiency results in impaired NHEJ</li> <li>DSB repair was monitored using PFGE by measuring the % of DSBs remaining after 0.25, 2, and 24 h following irradiation.</li> <li>DSBs decreased over time and, eventually, less than 10% of the DSBs remained in MRC-5 cells after 24h following both 80 and 10 Gy exposures.</li> <li>Repair was noticeably slower in 180BR cells, where the clearance of DSBs was hindered and approximately 40 and 20% of the DSBs remained at 24 hours following 80 and 10 Gy exposures, respectively.</li> <li>The above demonstrates defective DNA repair leading to persistent DSBs.</li> </ul> </li> </ul>

<ul> <li>A variety of confounding factors and genetic characteristics (i.e., SNPs) may modulate which repair pathways are invoked and the degree to which they are inadequate. These have yet to be fully defined.</li> <li>Both protective and damaging effects of OGG1 against strand breaks have been described in the literature. As demonstrated in the section above, the effect of OGG1-deficiency (BER-initiating enzyme) is observed to be different in different cell types; Wang et al. (2018) demonstrated strand breaks exacerbated by excessive OGG1 activity, while Wu et al. (2008) and Shah et al. (2018) demonstrated increased strand breaks due to lack of repair in mammalian cells in culture (Shah et al., 2018; Wu et al., 2008; Wang et al., 2018). Cell cycle and replication may influence the effect of DNA repair on exacerbating strand breaks. </li> <li>Dahle et al. (2008) exposed wild type and OGG1-overexpressing Chinese hamster ovary cells, AS52, to UVA. While OGG1-overexpression prevented the accumulation of Fpg-sensitive lesions (e.g., 8-oxo-dG and FaPyG) that were observed in wild type cells 4 hours after irradiation, there was no difference in the amount of strand breaks in the two cell types at 4h (Dahle et al., 2008). </li> <li>A recent study suggests that the NHEJ may be more accurate than previously thought (reviewed in Betermier et al., 2014). The accuracy of NHEJ may be dependent on the structure of the termini. The termini processing rather than the NHEJ itself is thus argued to be error-prone process (Betemier et al., 2014).</li> </ul>

Not Specified Unspecific

Not Specified

All life stages

Not Specified Not Specified Not Specified

This KER applies to any cell type that has DNA repair capabilities.

Alexander, J., Orr-Weaver, T. (2016), Replication fork instability and the consequences of fork collisions from rereplication, Genes Dev, 30:2241-2252. Brenerman, B., Illuzzi, J., Wilson III, D. (2014), Base excision repair capacity in informing healthspan, Carcinogenesis, 35:2643-2652. Dahle, J., Brunborg, G., Svendsrud, D., Stokke, T., Kvam, E. (2008), Overexpression of human OGG1 in mammalian cells decreases ultraviolet A induced mutagenesis, Cancer Lett, 267:18-25. Dolinnaya, N., Kubareva, E., Romanova, E., Trikin, R., Oretskaya, T. (2013), Thymidine glycol: the effect on DNA molecular structure and enzymatic processing, Biochimie, 95:134-147. Eccles, L., Lomax, M., O’Neill, P. (2010), Hierarchy of lesion processing governs the repair, double-strand break formation and mutability of three-lesion clustered DNA damage, Nucleic Acids Res, 38:1123-1134. Ensminger, M., Iloff, L., Ebel, C., Nikolova, T., Kaina, B., Lobrich, M. (2014), DNA breaks and chromosomal aberrations arise when replication meets base excision repair, J Cell Biol, 206:29. Kitsera, N., Stathis, D., Luhnsdorf, B., Muller, H., Carell, T., Epe, B., Khobta, A. (2011), 8-Oxo-7,8-dihydroguanine in DNA does not constitute a barrier to transcription, but is converted into transcription-blocking damage by OGG1, Nucleic Acids Res, 38:5926-5934. Kühne, M., E. Riballo, N. Rief, K. Rothkamm, P. Jeggo, & M. Löbrich (2004), "A Double-Strand Break Repair Defect in ATM-Deficient Cells Contributes to Radiosensitivity", Cancer Res, 64(2): 500-508. Ma, W., Panduri, V., Sterling, J., Van Houten, B., Gordenin, D., Resnick, M. (2009), The Transition of Closely Opposed Lesions to Double-Strand Breaks during Long-Patch Base Excision Repair Is Prevented by the Coordinated Action of DNA Polymerase  and Rad27/Fen1  , Mol Cell Biol, 29:1212-1221. Minko, I., Jacobs, A., de Leon, A., Gruppi, F., Donley, N., Harris, T., Rizzo, C., McCullough, A., Lloyd, R.S. (2016), Catalysts of DNA Strand Cleavage at Apurinic/Apyrimidinic Sites, Sci Rep, 6. Peterson-Roth, E., Reynolds, M., Quievryn, G., Zhitkovich, A. (2005), Mismatch Repair Proteins Are Activators of Toxic Responses to Chromium-DNA Damage, Mol Cell Biol, 25:3596-3607. Rothkamm, K., Lobrich, M. (2003), Evidence for a lack of DNA double-strand break repair in human cells exposed to very low x-ray doses, Proc Natl Acad Sci USA, 100:5057-5062. Rydberg, B., Cooper, B., Cooper, P., Holley, W., Chatterjee, A. (2005), Dose-Dependent Misrejoining of Radiation-Induced DNA Double-Strand Breaks in Human Fibroblasts: Experimental and Theoretical Study for High- and Low-LET Radiation, Radiat Res, 163:526-534. Shah, A., Gray, K., Figg, N., Finigan, A., Starks, L., Bennett, M. (2018), . Defective Base Excision Repair of Oxidative DNA Damage in Vascular Smooth Muscle Cells Promotes Atherosclerosis, Circulation, 138:1446-1462. Wakasugi, M., Sasaki, T., Matsumoto, M., Nagaoka, M., Inoue, K., Inobe, M., Horibata, K., Tanaka, K., Matsunaga, T. (2014), Nucleotide Excision Repair-dependent DNA Double-strand Break Formation and ATM Signaling Activation in Mammalian Quiescent Cells, J Biol Chem, 289:28730-28737. Wang, R., Li, C., Qiao, P., Xue, Y., Zheng, X., Chen, H., Zeng, X., Liu, W., Boldogh, I., Ba, X. (2018), OGG1-initiated base excision repair exacerbates oxidative stress-induced parthanatos, Cell Death and Disease, 9:628. Whitaker, A., Schaich, M., Smith, M.S., Flynn, T., Freudenthal, B. (2017), Base excision repair of oxidative DNA damage: from mechanism to disease, Front Biosci, 22:1493-1522. Wu, M., Zhang, Z., Che, W. (2008), Suppression of a DNA base excision repair gene, hOGG1, increases bleomycin sensitivity of human lung cancer cell line, Toxicol App Pharmacol, 228:395-402. Yang, N., Galick, H., Wallace, S. (2004), Attempted base excision repair of ionizing radiation damage in human lymphoblastoid cells produces lethal and mutagenic double strand breaks, DNA Repair, 3:1323-1334. Yoshikawa, Y., Yamasaki, A., Takatori., K., Suzuki, M., Kobayashi, J., Takao, M., Zhang-Akiyama, Q. (2015), Excess processing of oxidative damaged bases causes hypersensitivity to oxidative stress and low dose rate irradiation, Free Radic Res, 49:1239-1248. Zeman, M., Cimprich, K. (2014), Causes and Consequences of Replication Stress, Nat Cell Biol, 12:2-9.     AOPWiki 2019-05-19T16:36:02

2023-01-09T20:56:58

<upstream-id>669b23ca-0e30-4508-ae51-b6d96fb4949d</upstream-id> <downstream-id>5e73c095-d52c-4844-9379-6a2af442e148</downstream-id> The maintenance of DNA integrity is essential for genomic stability; for this reason cells have multiple response mechanisms that enable the repair of damaged DNA. Thus when DNA double strand breaks (DSBs) occur, the most detrimental type of lesion, the cell will initiate repair machinery. These mechanisms are not foolproof, and emerging evidence suggests that closely spaced lesions can compromise the repair machinery. The two most common DSB repair mechanisms are non-homologous end joining (NHEJ) and homologous recombination (HR). The latter predominates in stem cells as they are frequently in the replicative phase of the cell cycle (Choi et al., 2020). NHEJ is initiated in G1 and early S phases of the cell cycle (Lieber et al., 2003) and is preferentially used to repair DSB damage (Godwint et al., 1994), as it is rapid and more efficient than HR (Lliakis, 1991; Jeggo, 1998; Mao et al., 2008). In higher-order eukaryotes such as humans, NHEJ is the favoured DNA repair mechanism because of the large non-coding regions within the genome. However, when other repair mechanisms (e.g., NHEJ, HR) are compromised, single strand annealing, which is a low fidelity mechanism may be involved (Chang et al., 2017). NHEJ can occur through one of two subtypes: canonical NHEJ (C‐NHEJ) or alternative non-homologous end joining (alt-NHEJ). C-NHEJ, as the name suggests, simply ligates the broken ends back together. In contrast, alt‐NHEJ occurs when one strand of the DNA on either side of the break is resected to repair the lesion (Bétermier et al., 2014). All repair mechanisms are error‐prone, meaning that insertions and deletions are sometimes formed due to the DSBs being repaired imperfectly (Thurtle-Schmidt and Lo, 2018). However, alt-NHEJ is considered more error-prone than C-NHEJ, as studies have shown that it more often leads to chromosomal aberrations (Zhu et al., 2002; Guirouilh-Barbat et al., 2007; Simsek & Jasin, 2010). HR is operative during late S and G2 phases where the sister chromatid can be used as template for error-free repair (Van Gent et al 2001). Because of the reliance on the undamaged sister chromatid to repair the DSB, HR is less error-prone than NEHJ. Nevertheless, defects in HR are known to contribute to genomic instability and the formation of chromosomal aberrations (Deans et al 2000) There is extensive evidence that DNA repair capacity can be overwhelmed or saturated in the presence of high numbers of strand breaks. For example, with multiple single strand breaks (SSBs) in close proximity that can lead to DSBs (Caldecott, 2024). This is demonstrated by decades of studies showing dose-related increases in chromosomal exchanges, chromosomal breaks and micronuclei following exposure to double-strand break inducers. Additionally, the loss of heterozygosity (LOH)  is an example of how during the repair of incorrect DNA that uses HR, there may be a loss of an allele during repair (Smukowski et al., 2023). Inadequate repair not only refers to overwhelming of DNA repair machinery, but also the use of repair mechanisms that are error-prone (i.e., misrepair is considered inadequate repair).

The strategy for collating the evidence to support the relationship is described in Kozbenko et al 2022. Briefly, a scoping review methodology was used to prioritize studies based on a population, exposure, outcome, endpoint statement.

The biological rationale linking increased DNA DSB formation with inadequate DSB repair is supported strongly by literature. This is evident from the number of review articles that have been published on the subject. Of particular relevance is a recent review that focuses particularly on DSBs induced by ionizing radiation and extensively details the processes involved in repairing DSBs, including discussions of entire pathways and individual proteins involved in DNA repair (Thompson, 2012). Multiple other shorter reviews are also available on the subject, which cover such topics as: the mechanisms of DSB formation and repair, how to quantify these two events, and the biological consequences of unrepaired or misrepaired DNA damage (Lett, 1996; van Gent et al., 2001; Khanna & Jackson, 2001; Vignard et al., 2013; Moore et al., 2014; Rothkamm et al., 2015; Chang et al., 2017; Löbrich and Jeggo, 2017; Sage and Shikazono, 2017). A brief overview of the biological plausibility of this KER is given below; for more detail, please consult the above-cited reviews. When confronted with DSBs, there are two common repair pathways employed by the cell: homologous recombination (HR) and non-homologous end-joining (NHEJ). In HR, a homologous sequence on a sister chromatid is used as a template, ensuring that no sequence information is lost over the course of repair (e.g., Ferguson & Alt, 2001; van Gent et al., 2001; Jeggo & Markus, 2015; Schipler & Iliakis, 2013). Due to being inherently error-prone, NHEJ is commonly used in repairing DSBs in multicellular eukaryotic organisms, especially in humans (Feldmann et al., 2000).  Due to being inherently error-prone, this repair process is used to generate genetic variation within antigen receptor axons through VDJ recombination, a process that leads to the careful breakage and repair of DNA (Murakami & Keeney, 2008; Malu et al., 2012).  Genetic variation is also often generated during the repair of highly toxic DSB lesions. Repair to these DSB sites normally triggers cell cycle delay. NHEJ is most active in the following order of the cell cycle: G1 > S > G2/M (Mao et al., 2008). Since most somatic mammalian cells are in the G1 pre-replicative phase, DSBs also usually appear in this phase and thus are often repaired using the error-prone NHEJ (Jeggo et al., 1995). Cells in other phases of the cell cycle (S or G2) use HR (Ceccaldi et al., 2016). In addition,) and damaged cells in G0 also appear to use NHEJ repair (Frock et al., 2021). The two broken ends of DNA DSBs are bridged by overlapping single-strand microhomology termini (Anderson, 1993; Getts & Stamato, 1994; Rathmell & Chu, 1994; Jeggo et al., 1995; Miller et al., 1995; Kirchgessner et al., 1995). The microhomology termini are ligated only when complementary base pairs are overlapped and, depending on where this match is found on the termini, it can lead to deletions and other rearrangements. With increasing DSBs, the probability of insufficient or incorrect repair of these breaks increases proportionately. It has been suggested that clustered DNA damage is less easily repairable than any other form of DNA damage (United Nations, 2000; Stenerlöw et al., 2000).  With multiple lesions in close proximity within a damaged cluster, the probability of misrepair is high. This leads to an increased number of misrepaired termini (Goodhead et al., 1994; Goodhead, 1980; Tsao, 2007; Blakely, 2012), as the presence of multiple damage sites interferes with the ability of the repair enzymes to recognize and bind to the DNA accurately (Harrison et al., 1999; Tsao, 2007).

Empirical data obtained for this KER strongly supports the idea that an increase in DNA DSBs will increase the frequency of inadequate DSB repair. The evidence presented below is summarized in table 4, <a href="https://docs.google.com/spreadsheets/d/1iehBBqhFFSOhgis-0U3tasQwJ50bZJPVmenWUiR4vmA/edit?usp=sharing" target="_blank">here (click link)</a>. Much of the evidence comes from work with radiation stressors, which directly cause DNA DSBs in the genome (Pinto & Prise, 2005; Dong et al., 2017) in a dose-dependent fashion (Aufderheide, 1987; Frankenburg-Schwager et al., 1994; Rydberg et al., 1994; Durante et al., 1998; Dikomey & Brammer, 2000; Kuhne et al., 2000; Löbrich et al., 2000; Baumstark-Khan et al., 2003; Rothkamm & Lo, 2003; Kuhne et al., 2005; Asaithamby & Chen, 2009; Bracalente et al., 2013). This is a very data-rich area and it is not possible to summarize all of the evidence. However, some examples of key studies are provided below. We also direct the reader to the key event relationships 1939 (DNA strand breaks leading to chromosomal aberrations) and 1931 (DNA strand breaks leading to mutations). The formation of DSBs by ionizing radiation, the repair process, the various methods used to analyze this repair process, and the biological consequences of unrepaired or misrepaired DNA damage are reviewed in Sage & Shikazono (2017).   Dose and Incidence Concordance There is evidence in the literature suggesting a dose/incidence concordance between the occurrence of DSBs and the incidence of inadequate DNA repair upon exposure to radiation. Inadequate DNA repair appears to occur at the same radiation dose as DSBs. Visually, immunofluorescence has demonstrated a colocalization of DNA repair proteins with DSB foci in response to a radiation stressor (Paull et al., 2000; Asaithamby & Chen, 2009; Dong et al., 2017). In studies examining cellular responses to increasing doses of radiation, which is known to evoke a dose-dependent increase in DNA DSBs (Aufderheide, 1987; Durante et al., 1998; Dikomey & Brammer, 2000; Kuhne et al., 2000; Löbrich et al., 2000; Rothkamm & Löbrich, 2003; Kuhne et al., 2005; Asaithamby & Chen, 2009; Bracalente et al., 2013; Bernard et al., 2021), there were resulting dose-dependent increases in non-repaired DSBs (Aufderheide, 1987; Rydberg et al., 1994; Dikomey & Brammer, 2000; Baumstark-Khan et al., 2003), DSB misrepair rates (Mcmahon et al., 2016), and misrejoined DSBs (Durante et al., 1998; Kuhne et al., 2000; Kuhne et al., 2005; Rydberg et al., 2005), as well as a dose-dependent decrease in the total DSB rejoining (Löbrich et al., 2000). Furthermore, only 50% of the rejoined DSBs were found to be correctly repaired (Kuhne et al., 2000; Löbrich et al., 2000); 24 hours after being irradiated with an 80 Gy dose of alpha particles, this frequency of misrejoining increased to and remained constant at 80% (Kuhne et al., 2000). Furthermore, delivering radiation doses in fractionated increments also showed a dose-dependent change in the percentage of misrejoinings, such that larger fractionated doses (for example, 2 x 40 Gy) had a higher rate of DSB misrejoining than smaller fractionated doses (for example, 4 x 10 Gy) (Kuhne et al., 2000).   Temporal Concordance There is evidence suggesting a time concordance between DSBs and DNA repair. DSBs and DNA repair have both been observed within minutes to hours of radiation exposure (Paull et al., 2000; Rothkamm & Löbrich, 2003; Pinto & Prise, 2005; Asaithamby & Chen, 2009; Barnard et al., 2021; Barnard et al., 2018). Single strand breaks (SSBs) and DNA. repair has also been observed minutes to hours post-irradiation (Sidjanin et al., 1993).    Essentiality There is evidence from inhibition studies and knock-out/knock down studies suggesting that there is a strong relationship between DSBs and DNA repair. When an inhibitor of a DNA repair protein was added to cells prior to exposure to a radiation stressor, DNA repair foci were not formed post-irradiation (Paull et al., 2000), and there were significant increases in DSBs at 6 hours and 12 hours after the radiation treatment (Dong et al., 2017). Similarly, there have been several knock-out/knock-down studies in which cells lacking a DNA repair protein have been exposed to a radiation stressor. As a result, DSBs were found to persist in these cells longer than in the wild-type cells (Coquerelle et al., 1987; Rothkamm and Löbrich, 2003; Bracalente et al., 2013; Mcmahon et al., 2016; Dong et al., 2017), and there was an increase in incorrectly rejoined DSBs (Löbrich et al., 2000). In one striking example, a human cell line lacking DNA ligase IV had DSBs that were still present approximately 240 - 340 hours post-irradiation (Mcmahon et al., 2016). Interestingly, there were also increased levels of DSBs in these cells prior to being exposed to a radiation stressor (Paull et al., 2000) . Similarly, a study examining DSB repair kinetics after irradiation found that DSBs persisted for a longer time period in two repair-deficient mouse strains relative to a repair-proficient mouse strain; this pattern was found in lymphocytes, as well as tissues from the brains, lungs, hearts and intestines of these mice (Rube et al., 2008). The roles of various DNA repair proteins in the context of DSBs are highlighted in reviews by Chang et al. (2001) and Van Gent et al. (2001) with discussions focusing on the consequences of losing some of these proteins in cells, mice and humans (Van Gent et al., 2001)

Uncertainties and inconsistencies in this KER are as follows: <ul> <li>There is controversy surrounding how error-prone NHEJ truly is.  Recent studies suggest that the process may be quite accurate (reviewed in (Bétermier et al. 2014)). The accuracy of NHEJ may actually be dependent on the structure of the termini. Thus, the termini processing rather than the NHEJ mechanism itself is argued to be the error-prone process (Bétermier et al. 2014).</li> <li>There may be different cellular responses associated with low-dose radiation exposure and high-dose radiation exposure; these differences may also be dependent on a DSB threshold being exceeded prior to initiation repair. It has been suggested that DNA repair may not be activated at low doses of radiation exposure in order to prevent the risk of mutations from error-prone repair mechanisms (Marples 2004).</li> <li>DSB repair fidelity varies in terms of confounding factors and the genetic characteristics of individuals (Scott 2006). For example, individuals who smoke have a 50% reduction in the mean level of DSB repair capacity relative to the non-smokers; this is due to an increased methylation index in smokers. A higher methylation index indicates more inactivation of gene expression. It is thus possible that expression of DNA repair proteins in smokers is decreased due to increased methylation of the genes encoding for repair proteins. In terms of individual genetics, single nucleotide polymorphisms (SNPs) within the MRE11A, CHEK2, XRCC3, DNA-PKcs, and NBN repair genes have been highly associated with the methylation index (Leng et al. 2008). SNPs can critically affect the function of these core proteins, varying the fidelity of DNA repair from person to person.</li> <li> Cells containing DNA damaged may be eliminated by apoptotic pathways, therefore not undergo repair, alternatively evidence has also shown that damaged cells can propagate due to lack of detection by repair machinery (Valentin 2005).  </li> <li>The focus of this KER was on DSBs because there is lack of data to support that SSBs lead to inadequate repair. Multiple SSBs can lead to DSBs. Thus, DSBs are the focus as they can drive the cell towards genomic instability, apoptosis or tumorigenesis. Further quantitative evidence to define the extent of SSBs leading to DSBs and the relationship with repair is necessary. </li> <li>Ercc2+/- mice have a mutation in a gene involved in the nucleotide excision repair (NER) pathway, leading to DNA repair deficiency. However, when compared to wild type mice Ercc2+/- mice had fewer DNA strand breaks. This was true of both central and peripheral lens cells, as well as 4 and 24 h after irradiation (60Co γ-rays, 0.3, 0.063 Gy/min) (Barnard et al., 2021).</li> <li>DNA damage repair times can vary depending on the stressors that instigate the DNA damage. For example, it has been found that some types of radiation i.e., high linear energy transfer (LET) increases the amount of time required to repair DNA breaks (Aufderheide, 1987; Frankenburg-Schwager et al., 1994; Rydberg et al., 1994; Baumstark-Khan et al., 2003; Tsao, 2007; Blakely, 2012), however Stenerlöw et al. (2000) found that repair half-times were independent of LET.  </li> </ul>

<table border="1"> <tbody> <tr> <td> Modulating Factor  </td> <td> Details   </td> <td> Effects on the KER   </td> <td> References   </td> </tr> <tr> <td> Linear energy transfer (LET)  </td> <td> Increased LET  </td> <td> As the LET of the stressor increases, the amount of misrepaired and unrejoined DSBs also increases. One possible explanation for this is that DSB free ends are closer together at higher LETs, making it easier for misrepair to occur. Furthermore, higher LET stressors produce more complex, clustered breaks which also increasing repair difficulty. At very high LET values (over 10 000 keV/um), no significant DNA repair is detected.  </td> <td> Aufderheide, 1987; Rydberg et al., 1994; Durante et al., 1998; Kuhne et al., 2000; Stenerlöw et al., 2000; Baumstark-Khan et al., 2003; Tsao, 2007; Mukherjee et al., 2008; Blakely, 2012; Hamada, 2017  </td> </tr> <tr> <td> Oxygen   </td> <td> Decreased oxygen levels   </td> <td> Cells in an anoxic environment will rejoin DNA breaks more quickly than those in an oxic environment because oxygen can attach to the broken ends of DNA, fixing the damage and making it unrepairable.  </td> <td> Frankenburg-Schwager et al., 1994  </td> </tr> </tbody> </table>

Quantitative understanding of this linkage suggests that DSB repair can be predicted from the presence of DSBs. The following tables provide representative examples of the relationship, unless otherwise indicated, all data is significantly significant. In terms of DNA repair in response to radiation-induced DSBs, one  study suggests that complete DNA DSB repair occurs starting at a threshold dose of  5 mGy (0.005 Gy), as measured by the presence of γ-H2AX (Lobrich et al., 2005) and presence of 53BPI foci (Asaithamby & Chen, 2009). After a 10 Gy dose of radiation, approximately 10 - 15% of DSBs were found to be misrepaired (Mcmahon et al., 2016); at a dose of 80 Gy, the relative percentage of DSBs incorrectly repaired was estimated at 50 - 60% (Kuhne et al., 2000; Lobrich et al., 2000; Mcmahon et al., 2016). Twenty-four hours post-irradiation, this rate increased to approximately 80% for alpha particle irradiation at 80 Gy, and remained constant until the end of the assay (10 days) (Kuhne et al., 2000).   Dose Concordance  <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Rydberg et al., 1994  </td> <td> In vitro. Human VA13 lung fibroblast and GM38A skin fibroblast cells were exposed to neon ions (425 MeV/u, 1 – 5 Gy/min, 80 Gy), iron ions (600 MeV/u, 1 – 5 Gy/min, 50 Gy), and X rays (425 MeV/u, 1 – 2 Gy/min, 80 Gy) to induce DNA strand breaks.   Initial breaks after exposure were measured via the fraction of activity released (FAR) assay, with an increased FAR value indicating an increased number of breaks.  Repair was measured using the FAR assay after a period of incubation.  </td> <td> Exposure to X-rays, neon, and iron ions led to a 90, 70, and 50% FAR increase relative to control respectively, indicating the highest level of breaks in samples exposed to X-rays. Four h later, 15, 20, and 73% of the DNA strand breaks had not been repaired.    </td> </tr> <tr> <td> Kuhne et al., 2000   </td> <td> In vitro. Human lung fibroblast cells were exposed to X-rays (23 Gy/min) at doses from 0 - 320 Gy. Following this, both correct (measured via hybridization assay), and total (measured via FAR assay) breaks remaining were measured. Therefore, allowing for calculation of the amount of misrepaired breaks.   </td> <td> Cells exposed to 0 - 320 Gy X-rays displayed an approximately linear increase in DSBs. This led to a gradual increase in the % DSBs misrejoined, which began to plateau after 80 Gy at a misrejoining frequency of 50%.  </td> </tr> <tr> <td> Baumstark-Khan et al., 2003  </td> <td> In vitro. Bovine LECs were exposed to X-rays (5 Gy/min, 0 to 50 Gy), 16O (3.4, 8.7 MeV/u, 230.5 to 642.9 Gy), 40Ar (2.7, 6.2, 10.5, 19.3 MeV/u, 0 to 190 Gy), 132Xe (5.4, 10.1, 16.5 MeV/u, 0 to 80 Gy), 208Pb (3.0, 6.8, 15.4 MeV/u, 0 to 50 Gy), 238U (1.5, 1.9, 2.6, 4.0 MeV/u, 0 to 150 Gy). This led to the induction of both SSBs and DSBs, whose repair was measured using a method similar to the hydroxyapatite chromatography of alkaline unwound DNA.   </td> <td> Irradiation below 10 000 keV/μm led to almost 100% rejoining of SSBs and DSBs. At LETs above 10 000 keV/μm the rejoining capacity varied depending on the original level of damage. After irradiation with 238U (LET ~ 20 000 keV/μm) rejoining capacity as t -> ∞ ranged from 50 to 100%. After irradiation with 208Pb (LET ~ 18 000 keV/μm) rejoining capacity as t -> ∞ ranged from 15 to 28%.   48Ti was an exception, with an LET of 1440 keV/μm that resulted in a rejoining capacity of only 65% rather than almost 100% as t -> ∞.   </td> </tr> <tr> <td> Aufderheide, 1987  </td> <td> In vitro. Bovine lens epithelial cells (LECs) were exposed to 238U (5, 10, 20 x 106 P/cm2), 132Xe (3, 5, 7, 12, 20 x 106 P/cm2), 84Kr (9, 21 x 106 P/cm2), 40Ar (24 x 106 P/cm2), 16O (80 x 106 P/cm2), and X-rays (20, 40, 200 Gy). The radiation exposure induced DNA breaks were measured using the DNA unwinding method described by Rydberg (1975). The DNA then underwent a period of repair incubation lasting between 5 to 40 h, after which any remaining DNA damage was measured using the same method as before.  </td> <td> Bovine LECs exposed to 21 x 106 P/cm2 84Kr displayed a 1.3x increase in DNA breaks and a 5% decrease in the level of breaks repaired compared to cells exposed to 9 x 106 P/cm2.  </td> </tr> <tr> <td> Stenerlöw et al., 2000  </td> <td> In vitro. Human skin fibroblast cells were exposed to 100 Gy of photons (60Co, < 0.5 keV/um), nitrogen ions (80, 125, 175, 225 keV/um), and helium ions (40 keV/um), resulting in the formation of DSBs. Their number was calculated by fragment analysis, based upon the fraction of DNA less than 5.7 Mbp, under the assumption that the breaks were evenly distributed. DNA repair was also measured via fragment analysis.  </td> <td> Exposure to increasing LET of radiation at 100 Gy led to increasing DSBs, in general, with about 600 DSBs/Gbp after γ-ray irradiation and about 700 DSBs/Gbp after 225 keV/um nitrogen ion irradiation. A dose of 100 Gy also led to decreased repair at increased LET. About 20-22 h after γ-ray irradiation, 4% of DSBs were unrepaired, while 20-22 h after 225 keV/um nitrogen ion irradiation, 12% of DSBs were unrepaired.  </td> </tr> <tr> <td>Coquerelle et al., 1987</td> <td>In vitro. Human skin fibroblast cells were exposed to 60Co (1.5, 0.35 Gy/min) and alpha particles (120 keV/µm). Alkaline elution assay was used to detect DNA strand breaks. Repair of breaks were determined over time. The % rejoined DNA was calculated from the mean values of the entire elution profile. </td> <td>Exposure to 25 Gy gamma rays or alpha particles resulted in ~20% strand breaks. 80% of these breaks were repaired 30 mins after the exposure.</td> </tr> </tbody> </table>   Incidence Concordance  No studies were found.    Time Concordance  <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Durante et al., 1998   </td> <td> In vitro. Human, male, lymphocyte cells were exposed to either iron ions (140 keV/μm, 2 Gy), or carbon ions (42 keV/μm, 5 Gy) to induce DNA strand breaks. Misrepair was measured by producing chromosome spreads and evaluating them using a microscope and the PAINT classification code.  </td> <td> Exposure to 2 Gy iron particles resulted in about 0.45 breaks/cell, of which 50% were repaired 10 h later. However, there were 0.1 translocations/cell, 0.08 incomplete exchanges/cell, 0.075 complex exchanges/cell, and 0.07 dicentrics/cell.  Exposure to 5 Gy carbon ions resulted in 1.15 breaks/cell, of which 25% were repaired 10 h later. However, there were 0.35 translocations/cell, 0.28 incomplete exchanges/cell, 0.43 complex exchanges/cell, and 0.29 dicentrics/cell.  </td> </tr> <tr> <td> Rydberg et al., 1994  </td> <td> In vitro. Human VA13 lung fibroblast and GM38A skin fibroblast cells were exposed to neon ions (425 MeV/u, 1 – 5 Gy/min, 80 Gy), iron ions (250, 400, 600 MeV/u, 1 – 5 Gy/min, 50 Gy), and X rays (425 MeV/u, 1 – 2 Gy/min, 80 Gy) to induce DNA breaks. Their repair was measured using pulsed-field gel electrophoresis and determining the amount of DNA released from the gel plug (fraction of activity released – FAR).  </td> <td> In GM38A cells, exposure to 80 Gy of all three radiation types led to DNA breaks. Repair was observed between 0.5 and 4 h after this.  The most breaks remained after exposure to iron ions (75% of breaks remained), 25 – 42% remained after neon exposure, and only 15 – 20% remained after X ray irradiation.  </td> </tr> </tbody> </table>

There is evidence of a response-response relationship for DNA repair of radiation-induced DSBs. The frequency of DSBs has been shown to increase linearly with radiation dose (Löbrich et al., 2000; Rothkamm & Lo, 2003; Kuhne et al., 2005; Asaithamby & Chen, 2009). For DNA repair, increasing doses of a radiation stressor were found to cause a linear-quadratic relationship between the radiation dose and the number of misrejoined DSBs per cell (Kuhne et al., 2005). Interestingly, the relationships between radiation and DNA repair were found to vary depending on the type of radiation. There was a more linear response between radiation dose and the number of misrejoined DSBs for high LET particles relative to a more curvilinear relationship for lower LET particles (Rydberg et al., 2005). Additionally, a linear relationship was defined for low dose-rate radiation and the number of non-repaired DNA DSBs, but a linear-quadratic equation was described for high dose-rate radiation (Dikomey & Brammer, 2000).

Data from temporal response studies suggests that DSB repair may occur within 15 - 30 minutes of a DSB-inducing radiation stressor (Paull et al., 2000; Rothkamm & Lo, 2003; Pinto & Prise, 2005; Dong et al., 2017), with foci documented as early as 3-5 minutes post-irradiation (Asaithamby & Chen, 2009). The majority of DSB repair has been reported to occur within the first 3 - 6 hours following DSB induction (Rothkamm & Lo, 2003; Pinto & Prise, 2005; Asaithamby & Chen, 2009; Dong et al., 2017), with complete or near-complete DSB repair within 24 hours of the radiation stressor (Dikomey & Brammer, 2000; Lobrich et al., 2000; Rothkamm & Lo, 2003; Asaithamby & Chen, 2009; Mcmahon et al., 2016).  In one 48-hour time-course experiment for DSB repair using two different types of radiation, the following repair progression was found at 30 minutes, 1 hour, 3 hours, 24 hours and 48 hours, respectively: 40 - 55%, 55 - 70%, 85%, 97 - 98% and 98% repair for X-rays and 30%, 45 - 50%, 65 - 70%, 85 - 90% and 90 - 96% repair for alpha particles (Pinto & Prise, 2005). Twenty-four hours post-irradiation, the frequency of DSB misrejoining was found to remain constant at approximately 80% for the 10 days that the DSB repair was monitored (Kuhne et al., 2000).

Not identified.

High Unspecific

High

All life stages

High High High

This KER is plausible in all life stages, sexes, and organisms with DNA. The majority of the evidence is from in vivo adult mice with no specification on sex, and in vitro human models that do not specify sex.

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(2002), "Unrepaired DNA breaks in p53-deficient cells lead to oncogenic gene amplification subsequent to translocations.", Cell. 109(7):811–21. doi:10.1016/s0092-8674(02)00770-5.   AOPWiki 2019-05-19T16:36:30

2024-03-08T14:56:30

<upstream-id>5e73c095-d52c-4844-9379-6a2af442e148</upstream-id> <downstream-id>17e0b8d9-bbd6-410f-be2d-f759eacefdf2</downstream-id> The described Key Event Relationship (KER) outlines a sequence of events related to DNA repair and its consequences. The upstream event is characterized by "Inadequate DNA repair," indicating that the cellular mechanisms responsible for repairing DNA damage are compromised or insufficient. This could result from various factors, such as genetic mutations, environmental exposures, or other cellular processes. The downstream event in this KER is an "Increase in Mutations." As a consequence of inadequate DNA repair, the accumulation of unrepaired or incorrectly repaired DNA damage can lead to an elevated rate of mutations in the genome. These mutations can involve changes in the DNA sequence, structure, or arrangement, which may have various implications for cellular function, including potential disruptions to normal processes and pathways. This KER highlights the critical role of DNA repair mechanisms in maintaining genomic stability and preventing the buildup of mutations that can contribute to various biological outcomes, including disease development and other adverse effects. Insufficient repair results in the retention of damaged DNA that is then used as a template during DNA replication. During replication of damaged DNA, incorrect nucleotides may be inserted, and upon replication these become ‘fixed’ in the cell. Further replication propagates the mutation to additional cells. For example, it is well established that replication of alkylated DNA can cause insertion of an incorrect base in the DNA duplex (i.e., mutation). Replication of non-repaired O4 thymine alkylation leads primarily to A:T→G:C transitions. Retained O6 guanine alkylation causes primarily G:C→A:T transitions. For repairing DNA double strand breaks (DSBs), non-homologous end joining (NHEJ) is one of the repair mechanisms used in human somatic cells (Petrini et al., 1997; Mao et al., 2008). However, this mechanism is error-prone and may create mutations during the process of DNA repair (Little, 2000). NHEJ is considered error-prone because it does not use a homologous template to repair the DSB. The NHEJ mechanism involves many proteins that work together to bridge the DSB gap by overlapping single-strand termini that are usually less than 10 nucleotides long (Anderson, 1993; Getts & Stamato, 1994; Rathmell & Chu, 1994). Inherent in this process is the introduction of errors that may result in mutations such as insertions, deletions, inversions, or translocations. Furthermore, other repair mechanisms such as a loss in the mismatch repair (MMR) system can lead to a buildup of errors such as base-base mismatches and insertion-deletion errors in repetitive DNA sequences which are known as microsatellites. This could occur if an MMR gene (e.g. MLH1, PMS2) is inactivated through mutations or epigenetic silencing (Wang et al., 2022).

Adhering to OECD guidelines, a systematic and comprehensive evidence collection approach was executed to support the KER "Inadequate DNA repair leads to Increase, Mutations." To establish the first part of the relationship, the inadequacy of DNA repair processes, a range of molecular and cellular assays were employed. Comet assays provided direct quantification of unrepaired DNA lesions, while functional assays focusing on repair gene expression and protein activity offered mechanistic insights into repair deficiency. Crucial to establishing the link, genetic and biochemical studies delved into the interactions between the malfunctioning repair machinery and accumulating DNA lesions. The subsequent increase in mutations was substantiated by genetic analysis techniques such as whole-genome sequencing and polymerase chain reaction (PCR)-based assays. These methods not only quantified mutations but also pinpointed their specific locations, aiding in understanding their origins and patterns. Parallel assessments across different experimental conditions, strains, and species enhanced the robustness of the evidence. Additionally, case-control studies involving populations exposed to DNA damaging agents supported the real-world relevance of the KER, revealing a notable association between inadequate DNA repair and elevated mutation rates. The integration of results from diverse experimental models, mechanistic studies, and epidemiological data in line with OECD principles established a compelling and well-substantiated evidence base for the KER "Inadequate DNA repair leads to Increase, Mutations."

Overall Weight of Evidence: High

If DNA repair is able to correctly and efficiently repair DNA lesions introduced by a genotoxic stressor, then no increase in mutation frequency will occur. For example, for alkylated DNA, efficient removal by O6-alkylguanine DNA alkyltransferase will result in no increases in mutation frequency. However, above a certain dose AGT becomes saturated and is no longer able to efficiently remove the alkyl adducts. Replication of O-alkyl adducts leads to mutation. The evidence demonstrating that replication of unrepaired O-alkylated DNA causes mutations is extensive in somatic cells and has been reviewed (Basu and Essigmann 1990; Shrivastav et al. 2010); specific examples are given below. It is important to note that not all DNA lesions will cause mutations. It is well documented that many are bypassed error-free. For example, N-alkyl adducts can quite readily be bypassed error-free with no increase in mutations (Philippin et al., 2014). Inadequate repair of DSB Collective data from tumors and tumor cell lines has emerged that suggests that DNA repair mechanisms may be error-prone (reviewed in Sishc et al., 2017) (Sishc & Davis, 2017).  NHEJ, the most common pathway used to repair DSBs, has been described as error-prone. The error-prone nature of NHEJ, however, is thought to be dependent on the structure of the DSB ends being repaired, and not necessarily dependent on the NHEJ mechanism itself (Bétermier et al., 2014). Usually when perfectly cohesive ends are formed as a result of a DSB event, ligase 4 (LIG4) will have limited end processing to perform, thereby keeping ligation errors to a minimum (Waters et al., 2014). When the ends are difficult to ligate, however, the resulting repair may not be completed properly; this often leads to point mutations and other chromosomal rearrangements. It has been shown that approximately 25 - 50% of DSBs are misrejoined after exposure to ionizing radiation (Löbrich et al., 1998; Kuhne et al., 2000; Lobrich et al., 2000). Defective repair mechanisms can increase sensitivity to agents that induce DSBs and lead eventually to genomic instability (reviewed in Sishc et al., (2017)). Activation of mutagenic DNA repair pathways to withstand cellular or replication stress either from endogenous or exogenous sources can promote cellular viability, albeit at a cost of increased genome instability and mutagenesis (Fitzgerald et al., 2017). These salvage DNA repair pathways including, Break-induced Replication (BIR) and Microhomology-mediated Break-induced Replication (MMBIR). BIR repairs one-ended DSBs and has been extensively studied in yeast as well as in mammalian systems. BIR and MMBIR are linked with heightened levels of mutagenesis, chromosomal rearrangements and ensuing genome instability (Deem et al., 2011; Sakofsky et al., 2015; Saini et al., 2017; Kramara et al., 2018). In mammalian genomes BIR-like synthesis has been proposed to be involved in late-stage Mitotic DNA Synthesis (MiDAS) that predominantly occurs at so-called Common Fragile Sites (CFSs) and maintains telomere length under s conditions of replication stress that serve to promote cell viability (Minocherhomji et al., 2015; Bhowmick et al., 2016; Dilley et al., 2016).

INSUFFICIENT REPAIR OF ALKYLATED DNA Evidence in somatic cells Empirical evidence to support this KER is primarily from studies in which synthetic oligonucleotides containing well-characterized DNA lesions were genetically engineered in viral or plasmid genomes and subsequently introduced into bacterial or mammalian cells. Mutagenicity of each lesion is ascertained by sequencing, confirming that replication of alkylated DNA (i.e., unrepaired DNA) causes mutations in addition to revealing the important DNA repair pathways and polymerases involved in the process. For example, plasmids containing O6-methyl or O6-ethylguanine were introduced into AGT deficient or normal Chinese hamster ovary cells (Ellison et al. 1989). Following replication, an increase in mutant fraction to 19% for O6-methylguanine and 11% for O6-ethylguanine adducts was observed in AGT deficient cells versus undetectable levels for control plasmids. The relationship between input of alkylated DNA versus recovered mutant fractions revealed that a large proportion of alkyl adducts were converted to mutations in the AGT deficient cells (relationship slightly sublinear, with more adducts than mutations). The primary mutation occurring was G:C-A:T transitions. The results indicate that replication of the adducted DNA caused mutations and that this was more prevalent with reduced repair capacity. The number of mutations measured is less than the unrepaired alkyl adducts transfected into cells, supporting that insufficient repair occurs prior to mutation. Moreover, the alkyl adducts occur prior to mutation formation, demonstrating temporal concordance. Various studies in cultured cells and microorganisms have shown that the expression of O6-methylguanine DNA methyltransferase (AGT/MGMT) (repair machinery – i.e., decrease in DNA strand breaks) greatly reduces the incidence of mutations caused by exposure to methylating agents such as MNU and MNNG (reviewed in Kaina et al. 2007; Pegg 2011). Thomas et al. (2013) used O6-benzylguanine to specifically inhibit MGMT activity in AHH-1 cells. Inhibition was carried out for one hour prior to exposure to MNU, a potent alkylating agent. Inactivation of MGMT resulted in increased MNU-induced HPRT (hypoxanthine-guanine phosphoribosyltransferase) mutagenesis and shifted the concentrations at which induced mutations occurred to the left on the dose axis (10 fold reduction of the lowest observed genotoxic effect level from 0.01 to 0.001 µg/ml). The ratio of mutants recovered in DNA repair deficient cells was 3-5 fold higher than repair competent cells at concentrations below 0.01 µg/ml, but was approximately equal at higher concentrations, indicating that repair operated effectively to a certain concentration. Only at this concentration (above 0.01 µg/ml when repair machinery is overwhelmed and repair becomes deficient) do the induced mutations in the repair competent cells approach those of repair deficient. Thus, induced mutation frequencies in wild type cells are suppressed until repair is overwhelmed for this alkylating agent. The mutations prevented by MGMT are predominantly G:C-A:T transitions caused by O6-methylguanine. Evidence in germ cells That saturation of repair leads to mutation in spermatogonial cells is supported by work using the OECD TG488 rodent mutation reporter assay in sperm. A sub-linear dose-response was found using the lacZ MutaMouse assay in sperm exposed as spermatogonial stem cells, though the number of doses was limited (van Delft and Baan 1995). This is indirect evidence that repair occurs efficiently at low doses and that saturation of repair causes mutations at high doses. Lack of additional data motivated a dose-response study using the MutaMouse model following both acute and sub-chronic ENU exposure by oral gavage (O’Brien et al. 2015). The results indicate a linear dose-response for single acute exposures, but a sub-linear dose-response occurs for lower dose sub-chronic (28 day) exposures, during which mutation was only observed to occur at the highest dose. This is consistent with the expected pattern for dose-response based on the hypothetical AOP. Thus, this sub-linear curve for mutation at low doses following sub-chronic ENU exposure suggests that DNA repair in spermatogonia is effective in preventing mutations until the process becomes overwhelmed at higher doses. Mutation spectrum: Following exposure to alkylating agents, the most mutagenic adducts to DNA in pre-meiotic male germ cells include O6-ethylguanine, O4-ethylthymine and O2-ethylthymine (Beranek 1990; Shelby and Tindall 1997). Studies on sperm samples collected post-ENU exposure in transgenic rodents have shown that 70% of the observed mutations are at A:T sites (Douglas et al. 1995). The mutations observed at G:C base pairs are almost exclusively G:C-A:T transitions, presumably resulting from O6-ethylguanine. It is proposed that the prevalence of mutations at A:T basepairs is the result of efficient removal of O6-alkylguanine by AGT in spermatogonia, which is consistent with observation in human somatic cells (Bronstein et al. 1991; Bronstein et al. 1992). This results in the majority of O6-ethylguanine adducts being removed, leaving O4- and O2-ethylthymine lesions to mispair during replication. Thus, lack of repair predominantly at thymines and guanines at increasing doses leads to mutations in these nucleotides, consistent with the concordance expected between diminished repair capabilities at these adducts and mutation induction (i.e., concordance relates to seeing these patterns across multiple studies, species and across the data in germ cells and offspring).   Inadequate repair of oxidative DNA lesions: In vitro studies <ul> <li>AS52 Chinese hamster ovary cells (wild type and OGG1-overexpressing) were exposed to kJ/m2 UVA radiation (Dahle et al., 2008). <ul style="list-style-type:circle"> <li>Mutations in the gpt gene were quantified in both wild type and OGG1+ cells by sequencing after 13-15 days following 400 kJ/m2 UVA irradiation <ul> <li>G:C-A:T mutations in UVA-irradiated OGG1+ cells were completely eliminated</li> <li>G:C-A:T mutation frequency in wild type cells increased from 1.8 mutants/million cells to 3.8 mutants/million cells following irradiation – indicating incorrect repair or lack of repair of accumulated 8-oxo-dG</li> <li>Elevated levels of OGG1 was able to prevent G:C-A:T mutations, while the OGG1 levels in wild type cells was insufficient, leading to an increase in mutants (demonstrates inadequate repair leading to mutations)</li> </ul> </li> </ul> </li> <li>Xeroderma pigmentosum complementation group A (XPA) knockout (KO) and wild type TSCER122 human lymphoblastoid cells were transfected with TK gene-containing vectors with no adduct, a single 8-oxo-dG, or two 8-oxo-dG adducts in tandem (Sassa et al., 2015). <ul style="list-style-type:circle"> <li>XPA is a key protein in nucleotide excision repair (NER) that acts as a scaffold in the assembly the repair complex.</li> <li>Mutation frequency was determined by the number of TK-revertant colonies</li> <li>Control vector induced a mutation frequency of 1.3% in both WT and XPA KO</li> <li>Two 8-oxo-dG in tandem on the transcribed strand were most mutagenic in XPA KO, inducing 12% mutant frequency compared to 7% in WT</li> <li>For both XPA KO and WT, G:C-A:T transversion due to 8-oxo-dG was the most predominant point mutation in the mutants </li> <li>The lack of a key factor in NER leading to increased 8-oxo-dG-induced transversions demonstrates insufficient repair leading to increase in mutations </li> </ul> </li> </ul>   Inadequate repair of oxidative DNA lesions: In vivo studies in mice <ul> <li>Spontaneous mutation frequencies in the liver of Ogg1-deficient (-/-) Big Blue mice was measured at 10 weeks of age (Klungland et al., 1999). <ul style="list-style-type:circle"> <li>Mutation frequencies were 2- to 3-fold higher in the Ogg1-/- mice than in wild type</li> <li>Of the 16 base substitutions detected in Ogg1 -/- mutant plaques analyzed by sequencing, 10 indicated G:C-A:T transversions consistent with the known spectrum of mutation</li> <li>The results support that insufficient repair of oxidized bases leads to mutation.</li> </ul> </li> <li>Ogg1 knockout (Ogg1-/-) in C57BL/6J mice resulted in 4.2-fold and 12-fold increases in the amount of 8-oxo-dG in the liver compared to wild type at 9 and 14 weeks of age, respectively (Minowa et al., 2000). <ul style="list-style-type:circle"> <li>In these mice, there was an average of 2.3-fold increase in mutation frequencies in the liver (measured between 16-20 weeks) <ul> <li>57% of the observed base substitutions were G:C-A:T transversions, while 35% in wild type mice corresponded to this transversion.</li> <li>Approximately 70% of the increase in mutation frequency was due to G to T transversions.</li> </ul> </li> <li>Concordantly, KBrO3 treatment resulted in a 2.9-fold increase in mutation frequency in the kidney of Ogg1 -/- mice compared to KBrO3-treated wild type (Arai et al., 2002). <ul> <li>G:C-A:T transversions made up 50% of the base substitutions in the Ogg1-/- mice.</li> </ul> </li> <li>Heterozygous Ogg1 mutants (Ogg1+/-) retained the original repair capacity, where no increase in 8-oxo-dG lesions was observed in the liver at 9 and 14 weeks (Minowa et al., 2000). <ul> <li>This observation was consistent even after KBrO3 treatment of the mice (Arai et al., 2002).</li> </ul> </li> <li>From these results, we can infer that OGG1 proteins are present in excess and that one functional copy of the gene is sufficient in addressing endogenous and, to a certain degree, chemical-induced oxidative DNA lesions.</li> </ul> </li> </ul> Inadequate Repair of DSB Empirical data obtained for this KER moderately supports the idea that inadequate DNA repair increases the frequency of mutations. The evidence presented below related to the inadequate repair of DSBs is summarized in table 5, <a href="https://docs.google.com/spreadsheets/d/1iehBBqhFFSOhgis-0U3tasQwJ50bZJPVmenWUiR4vmA/edit?usp=sharing" target="_blank">here (click link)</a>. The review article by Sishc & Davis (2017) provides an overview of NHEJ mechanisms with a focus on the inherently error-prone nature of DSB repair mechanisms, particularly when core proteins of NHEJ are knocked-out. Although NHEJ is predominantly the preferred repair mechanism throughout the cell cycle, homologous recombination (HR) and single-stranded annealing (SSA) are favored during the S and G2 phases in scenarios where the NHEJ repair pathway is inhibited. The absence of HR leading to an increase in SSA activity is still a matter to debate (Ceccaldi et al., 2016). Another review also provides an overview of DSB induction, the repair process and how mutations may result, as well as the biological relevance of misrepaired or non-repaired DNA damage (Sage & Shikazono, 2017). Dose and Incidence Concordance There is evidence in the literature suggesting a dose/incidence concordance between inadequate DNA repair and increases in mutation frequencies. Evidence presented below related to the dose-response of mutation frequencies is summarized in table 2, <a href="https://docs.google.com/spreadsheets/d/1iehBBqhFFSOhgis-0U3tasQwJ50bZJPVmenWUiR4vmA/edit?usp=sharing" target="_blank">here (click link)</a>. In response to increasing doses from a radiation stressor, dose-dependent increases in both measures of inadequate DNA repair and mutation frequency have been found. In an analysis that amalgamated results from several different studies conducted using in vitro cell-lines, the rate of DSB misrepair was revealed to increase in a dose-dependent fashion from 0 - 80 Gy, with the mutation rate also similarly increasing from 0 - 6 Gy (Mcmahon et al., 2016). Additionally, using a plant model, it was shown that increasing radiation dose from 0 - 10 Gy resulted in increased DNA damage as a consequence of inadequate repair.  Mutations were observed 2 - 3 weeks post-irradiation (Ptácek et al., 2001). Moreover, increases in mutation densities were found in specific genomic regions of cancer samples (namely promoter DNAse I-hypersensitive sites (DHS) and 100 bp upstream of transcription start sites (TSS)) that were also found to have decreased DNA repair rates attributable to inadequate nucleotide excision repair (NER) (Perera et al., 2016).   Interestingly, mutation rates have been shown to increase as the required DNA repair becomes more complex. Upon completion of DSB repair in response to radiation and treatment with restriction enzymes, more mutations were found in cases where the ends were non-complementary and thus required more complex DNA repair (1 - 4% error-free) relative to cases where ends were complementary (34 - 38% error-free) (Smith et al., 2001). Temporal Concordance There is evidence in the literature suggesting a time concordance between the initiation of DNA repair and the occurrence of mutations. For simple ligation events, mutations were not evident until 12 - 24 hours, whereas DSB repair was evident at 6 -12 hours. For complex ligation events, however, mutations and DSB repair were both evident at 12 - 24 hours. As the relative percent of DNA repair increased over time, the corresponding percent of error-free rejoining decreased over time in both ligation cases, suggesting that overall DNA repair fidelity decreases with time ((Smith et al., 2001). Essentiality Inadequate DNA repair has been found to increase mutations above background levels. There is evidence from knock-out/knock-down studies suggesting that there is a strong relationship between the adequacy of DNA repair and mutation frequency. In all examined cases, deficiencies in proteins involved in DNA repair resulted in altered mutation frequencies relative to wild-type cases. There were significant decreases in the frequency and accuracy of DNA repair in cell lines deficient in LIG4 (DNA ligase 4, a DNA repair protein) (Smith et al., 2003) and Ku80 (Feldmann et al., 2000). Rescue experiments performed with these two cell lines further confirmed that inadequate DNA repair was the cause of the observed decreases in repair frequency and accuracy (Feldmann et al., 2000; Smith et al., 2003). In primary Nibrin-deficient mouse fibroblasts, there was increased spontaneous DNA damage relative to wild-type controls, suggestive of inadequate DNA repair. Using the corresponding Nibrin-deficient and wild-type mice, in vivo mutation frequencies were also found to be elevated in the Nibrin-deficient animals (Wessendorf et al., 2014). Furthermore, mutation densities were differentially affected in specific genomic regions in cancer patients depending on their Xeroderma pigmentosum group C (XPC) gene status. Specifically, mutation frequencies were increased in XPC-wild-type patients at DNase I-hypersensitive site (DHS) promoters and 100 bp upstream of TSS relative to cancer patients lacking functional XPC (Perera et al., 2016). Lastly, in a study using WKT1 cells with less repair capacity, radiation exposure induced four times more mutations in these cells than in TK6 cell, which had a normal repair capacity (Amundson and Chen, 1996).

Repair of alkylated DNA There were no inconsistencies in the empirical data reviewed or in the literature relating to biological plausibility. Much of the support for this KER comes predominantly from data in somatic cells and in prokaryotic organisms. We note that all of the data in germ cells used in this KER are produced exclusively from ENU exposure. Data on other chemicals are required. We consider the overall weight of evidence of this KER to be strong because of the obvious biological plausibility of the KER, and documented temporal association and incidence concordance based on studies over-expressing and repressing DNA repair in somatic cells. Repair of oxidative lesions <ul> <li>Thresholded concentration-response curve of mutation frequency was observed in AHH-1 human lymphoblastoid cells after treatment with pro-oxidants (H2O2 and  KBrO2) known to cause oxidative DNA damage (Seager et al., 2012), suggesting that cells are able to tolerate low levels of DNA damage using basal repair. However, increase in 8-oxo-dG lesions and up-regulation of DNA repair proteins were not observed under the same experimental condition.</li> <li>Mutagenicity of oxidative DNA lesions other than 8-oxo-dG, such as FaPydG and thymidine glycol, has not been as extensively studied and there are mixed results regarding the mutagenic outcome of these lesions.</li> </ul> Repair of double strand breaks  <ul> <li>One review paper found that DNA DSBs are repaired more efficiently at low dose (≤0.1 Gy) compared to high dose (>1 Gy) X-rays, but delayed mutation induction and genomic instability have also been demonstrated to occur at low doses (<1 cGy) of ionizing radiation (Preston et al., 2013).  </li> </ul> Overall <ul> <li>Mutation induction is stochastic, spontaneous, and dependent on the cell type as well as the individual’s capability to repair efficiently (NRC, 1990; Pouget & Mather, 2001).</li> </ul>

Not identified.

Thresholds for mutagenicity indicate that the response at low doses is modulated by the DNA repair machinery, which is effectively able to remove alkylated DNA at low doses [Gocke and Muller 2009; Lutz and Lutz 2009; Pozniak et al. 2009]. Kinetics of DNA repair saturation in somatic cells is described in Muller et al. [Muller et al. 2009]. For O-methyl adducts, once the primary repair process is saturated, in vitro data suggest that misreplication occurs almost every time a polymerase encounters a methylated guanine [Ellison et al. 1989; Singer et al. 1989]; however, it should be noted that this process can be modulated by flanking sequence. This conversion of adducts to mutations also appears to be reduced substantially in vivo [Ellison et al. 1989]. The probability of mutation will also depend on the type of adduct (e.g., O-alkyl adducts are more mutagenic than N-alkyl adducts; larger alkyl groups are generally more mutagenic, etc.). Overall, a substantive number of factors must be considered in developing a quantitative model. Inadequate repair of oxidative lesions The relationship between the quantity/activity of repair enzymes such as OGG1 in the cell and the quantity of oxidative lesions need to be better understood to define a threshold on the quantity of oxidative lesions exceeding basal repair capacity. Moreover, the proportion of oxidative lesions formed that lead to mutation versus strand breaks is not clearly understood. Mutations resulting from oxidative DNA damage can occur via replicative polymerases and translesion synthesis (TLS) polymerases during replication, and during attempted repair. However, an in vitro study on TLS in yeast has shown that bypass of 8-oxo-dG by TLS polymerases during replication is approximately 94-95% accurate. Therefore, the mutagenicity of 8-oxo-dG and other oxidative lesions may depend on their abundance, not on a single lesion (Rodriguez et al., 2013). Applicability of this observation in mammalian cells needs further investigation. Information on the accuracy of 8-oxo-dG bypass in mammalian cells is limited.       The most notable example of mutation arising from inadequate repair of DNA oxidation is G to T transversion due to 8-oxo-dG lesions. Previous studies have demonstrated higher mutation frequency of this lesion compared to other oxidative lesions; for example, Tan et al. (1999) compared the mutation rate of 8-oxo-dG and 8-oxo-dA in COS-7 monkey kidney cells and reported that under similar conditions, 8-oxo-dG was observed to be four times more likely to cause base substitution (Tan et al., 1999).  Inadequate Repair of DSB Quantitative understanding of this linkage is derived from the studies that examined DSB misrepair rates or mutation rates in response to a radiation stressor.  In general, combining results from these studies suggests that increased mutations can be predicted when DNA repair is inadequate. At a radiation dose of 10 Gy, the rate of DSB misrepair was found to be approximately 10 - 15% (Lobrich et al., 2000); this rate increased to 50 - 60% at a radiation exposure of 80 Gy (Kuhne et al., 2000; Lobrich et al., 2000; McMahon et al., 2016). For mutation rates in response to radiation across a variety of models and radiation doses, please refer to the example table below. <table border="1" cellpadding="1" cellspacing="1" style="height:158px; width:645px"> <tbody> <tr> <td style="text-align:center; width:150px">Reference</td> <td style="text-align:center">Summary</td> </tr> <tr> <td style="text-align:center">Matuo et al., 2018</td> <td>Yeast cells (saccharomyces cerevisiae) exposed to high LET cardbon ions (25 keV/um) and low LET carbon ions (13 keV/um) between 0-200 Gy induces a 24-fold increase overbaseline of mutations (high LET) and 11-fold increase over baseline mutations (low LET).</td> </tr> <tr> <td style="text-align:center">Nagashima et al., 2018</td> <td>Hamster cells (GM06318-10) exposed to x-rays in the 0-1 Gy. Response of 19.0 ± 6.1 mutants per 109 survivors.</td> </tr> <tr> <td style="text-align:center">Albertini et al., 1997</td> <td>T-lymphcytes isolated from human peripheral blood exposed to low LET gamma-rays (0.5-5 Gy) and high LET radon gas (0-1 Gy). Response of 7.0x10-6 mutants/Gy (Gamma-rays 0-2 Gy), 54x10-6 mutants/Gy (Gamma-rays 2-4 Gy) and 63x10-6 mutants/Gy (0-1 Gy).</td> </tr> <tr> <td style="text-align:center">Dubrova et al., 2002</td> <td>Observation of paternal ESTR mutation rates in CBAH mice following exposure to acute low LET X-rays (0-1 Gy), chronic low LET gamma-rays (0-1 Gy) and chronic high LET neutrons (0-0.5 Gy). Modelled response of y = mx + C, values of (m,C): X-rays: (0.338, 0.111), Gamma-rays: (0.373±0.082, 0.110), Neutrons: (1.135±0.202, 0.136).</td> </tr> <tr> <td style="text-align:center">McMahon et al., 2016</td> <td>Study of HPRT gene in Chinese hamster cells following exposure to radiation of 1-6 Gy. Observation of 0.2 mutations in HPRT gene per 104 cells and 0.1 point mutations per 104 cells (1 Gy). At 6 Gy, observation of 1.5 mutations in the HPRT gene per 104 cells and 0.4 point mutations per 104 cells.</td> </tr> </tbody> </table>

Inadequate Repair of DSB There is evidence of a response-response relationship between inadequate DNA repair and increased frequency of mutations. When exposed to a radiation stressor, there was a positive relationship between the radiation dose and the DSB misrepair rate, and between the mutation rate and the radiation dose (Mcmahon et al., 2016). Similarly, there was a negative correlation found between NER and the mutation densities at specific genomic regions in cancer patients. Specifically, inadequate NER resulted in more mutations in the promoter DHS and the TSS, but normal NER at DHS flanking regions resulted in fewer mutations (Perera et al., 2016).

Inadequate Repair of DSB Two studies were used to provide data regarding the time scale of DNA repair and the appearance of mutations. In a study using plants, DNA damage was evident immediately following radiation with 30 Gy of radiation; 50% of repairs were complete by 51.7 minutes, 80% by 4 hours, and repair was completed by 24 hours post-irradiation. Although no mutational analysis was performed during the period of repair, irradiated plants were found to have increased mutations when they were examined 2 - 3 weeks later (Ptácek et al., 2001). Both DNA repair and mutation frequency were examined at the same time in a study comparing simple and complex ligation of linearized plasmids. In this study, repaired plasmids were first detected between 6 - 12 hours for simple ligation events and between 12 - 24 hours for more complex ligation events; this first period was when the most error-free rejoining occurred in both cases. After this initial period of repair until its completion at 48 hr, repair became increasingly more erroneous such that mutations were found in more than half of the repaired plasmids at 48 hr regardless of the type of required ligation (Smith et al., 2001).

Not identified.

High Unspecific

High

All life stages

High High High

This KER is plausible in all life stages, sexes, and organisms with DNA. The majority of the evidence is from in vivo adult mice and male human, and mice in vitro models.  All organisms, from prokaryotes to eukaryotes, have DNA repair systems. Indeed, much of the empirical evidence on the fundamental principles described in this KER are derived from prokaryotic models. DNA adducts can occur in any cell type with DNA, and may or may not be repaired, leading to mutation. While there are differences among DNA repair systems across eukaryotic taxa, all species develop mutations following excessive burdens of DNA lesions like DNA adducts. Theoretically, any sexually reproducing organism (i.e., producing gametes) can also acquire DNA lesions that may or may not be repaired, leading to mutations in gametes.

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2024-03-08T15:00:38

<upstream-id>5e73c095-d52c-4844-9379-6a2af442e148</upstream-id> <downstream-id>6f3cc035-087f-41e7-bdfa-80177f2f7b4e</downstream-id> Cells are exposed to many insults, both endogenous and exogenous, that may cause damage to their DNA. In response to this constant threat, cells have accordingly evolved many different pathways for repairing DNA damage (Pfeiffer & Goedecke, 2000; Hoeijmakers, 2001; Jeggo & Markus, 2015; Rode et al., 2016). When confronted with double strand breaks (DSBs), there are two common repair pathways employed by the cell: homologous recombination (HR) and non-homologous end-joining (NHEJ). In HR, a homologous sequence on the sister chromatid is used as a template, ensuring that no sequence information is lost over the course of repair (Ferguson & Alt, 2001; van Gent et al., 2001; Hoeijmakers, 2001; Jeggo & Markus, 2015; Schipler & Iliakis, 2013; Venkitaraman, 2002). However, this method of DNA repair may result in a loss of an allele leading to heterozygosity. This may occur if a non-homologous chromosome with an erronous sequence is used as the template instead of the homologous chromosome, thus leading to a loss of genetic information (Ferguson & Alt, 2001). Despite this possible error, HR is generally considered to be one of the more accurate methods of DNA repair because it does make use of a template (van Gent et al., 2001; Schipler & Iliakis, 2013; Venkitaraman, 2002) .  NHEJ, however, does not use a template and is generally described as being error-prone. This repair process allows for the direct religation of broken DNA ends without using template DNA as a guide (van Gent et al., 2001; Ferguson & Alt, 2001; Hoeijmakers, 2001; Venkitaraman, 2002; Schipler & Iliakis, 2013; Jeggo & Markus, 2015; Rode et al., 2016). In lieu of a template, NHEJ utilizes rapid repair kinetics to religate the broken ends before they have time to diffuse away from each other (Schipler & Iliakis, 2013), thus fitting two ‘sticky’ DNA ends back together (Danford, 2012). There is not, however, an inherent quality control check; as such, sections of DNA may be gained or lost, or the wrong ends may be rejoined (Schipler & Iliakis, 2013). There are two versions of this error-prone DNA repair: classical or canonical NHEJ (c-NHEJ), and alternative NHEJ (alt-NHEJ) (Schipler & Iliakis, 2013). It is not well understood when or why one pathway is selected over another (Venkitaraman, 2002; Schipler & Iliakis, 2013). It has been proposed that the phase of the cell cycle may influence repair pathway choice (Ferguson & Alt, 2001; Vodicka et al., 2018); for instance, HR is generally more common than NHEJ when sister chromatids are available in S and G2 phases of the cell cycle (Hoeijmakers, 2001; Venkitaraman, 2002). If both HR and c-NHEJ are compromised, alt-NHEJ, which is slower and more error-prone than c-NHEJ, is thought to be the stand-by repair mechanism (Schipler & Iliakis, 2013). As BRCA2 is involved in both the NHEJ and HR repair pathways, it has recently been observed in BRCA2 deficient cells that single-strand annealing (SSA) may be triggered (Han et al. 2017). If these repair processes are not able to properly and adequately repair the DNA, this may lead to the formation of chromosomal aberrations (CAs). CAs are defined as abnormalities in the chromosome structure, often due to losses or gains of chromosome sections or the entire chromosomes itself (van Gent et al., 2001; Durante & Cucinotta, 2008). These abnormalities can take many different forms and can be classified according to several different schemes. CAs can be defined as breaks, which occur when DSBs are not rejoined, or as exchanges, where the presence of multiple DSBs results in misrejoining of the DNA ends (Danford, 2012; Registre et al., 2016). CA classes can be further subdivided into chromosome-type aberrations (CSAs) that affect both sister chromatids, and chromatid-type aberrations (CTAs), affecting only one chromatid (Danford, 2012) . Examples of CSAs include chromosome-type breaks, centric ring chromosomes, and dicentric chromosomes (which have two centromeres), while CTAs refer to chromatid-type breaks and chromatid exchanges (Hagmar et al., 2004; Bonassi et al., 2008). Other types of CAs that may occur include micronuclei (MN; small nucleus-like structures containing chromosome fragments enclosed by a nuclear membrane (Fenech & Natarajan, 2011; Doherty et al., 2016)), nucleoplasmic bridges (NPBs; a stretch of chromatin enclosed by a nuclear membrane that is attached to two centromeres (Fenech & Natarajan, 2011; Russo et al., 2015)), nuclear buds (NBUDs; a MN that is still connected to the nucleus by nucleoplasmic material (Fenech & Natarajan, 2011)), and copy number variants (CNVs;  base pair to megabase pair deletions or duplications of chromosomal segments (Russo et al., 2015)). CAs may also be classified as stable aberrations (translocations, inversions, insertions and deletions) and unstable aberrations (dicentric chromosomes, acentric fragments, centric rings and MN) (Hunter & Muirhead, 2009; Qian et al., 2016).

The strategy for collating the evidence to support the relationship is described in Kozbenko et al 2022.  Briefly, a scoping review methodology was used to prioritize studies based on a population, exposure, outcome, endpoint statement.

Overall Weight of Evidence: Low

There is strong biological plausibility for a relationship between inadequate repair of DNA damage and a corresponding increase in CAs. This is evident in a variety of reviews on the topic (van Gent et al., 2001; Hoeijmakers, 2001; Povirk, 2006; Weinstock et al., 2006; Lieber et al., 2010; Rode et al., 2016). The two most common methods used to repair DSBs, which are one of the most dangerous types of DNA lesions, are HR and NHEJ. Mechanisms for these two methods of DNA repair are well-established and have been thoroughly reviewed (Van Gent et al. 2001; Hoeijmakers 2001; Lieber et al. 2010; Jeggo and Markus 2015; Sishc and Davis 2017). Briefly, HR requires a template DNA strand to repair damage and thus facilitates the invasion of the damaged strand with matching sequences on homologous chromosomes or sister chromatids (Ferguson and Alt 2001; van Gent et al. 2001; Hoeijmakers 2001; Jeggo and Markus 2015; Schipler and Iliakis 2013; Venkitaraman 2002). Proteins involved in the HR pathway include the RAD50 proteins, MRE11, BRCA1, and BRCA2 (Ferguson and Alt 2001; van Gent et al. 2001; Hoeijmakers 2001; Jeggo and Markus 2015; Venkitaraman 2002). In contrast to this relatively accurate form of DNA repair ( van Gent et al. 2001; Schipler and Iliakis 2013; Venkitaraman 2002), NHEJ is more error-prone. It does not require a template to guide repair, but simply re-ligates broken DNA ends back together (Van Gent et al. 2001; Ferguson and Alt 2001; Hoeijmakers 2001; Lieber et al. 2010; Schipler and Iliakis 2013; Jeggo and Markus 2015; Rode et al. 2016; Sishc and Davis 2017) Proteins used during NHEJ include the DNA-PK complex (encompassing Ku70, Ku80 and DNA-PKcs), and the XRCC4-DNA ligase IV complex (Ferguson & Alt, 2001; van Gent et al., 2001; Hoeijmakers, 2001; Jeggo & Markus, 2015; Sishc & Davis, 2017).Interestingly, NHEJ is used in the biological V(D)J recombination process because its error-prone mechanism allows immune cells to develop a wide range of unique receptors for antigen detection (Ferguson & Alt, 2001; van Gent et al., 2001; Lieber, 2010).        Damaged DNA in the form of DSBs can follow  three possible outcomes: the DSB is rejoined accurately, with no changes made to the genome; the DSB is left unrepaired and the broken ends diffuse away from each other; or the DSB is repaired incorrectly such that the repaired version is different from the original version (Danford, 2012). These latter two errors in repair (the complete absence of repair or inaccurate repair) could arise due to interruptions to the repair process that allow time for the broken ends to move away from each other before they can be rejoined, mis-rejoining of the wrong DNA ends, or post-repair alterations that modify the junction point and lead to nucleotide losses (Schipler and Iliakis 2013). Unrepaired DSBs are the direct origin of micronuclei and unrepaired chromosomes correlated with radiosensitivity (Foray et al., 2016). Errors occurring during repair may be particularly detrimental if they interrupt or modify key genes, or if chromosome structures are created that cannot undergo proper mitosis (Schipler and Iliakis 2013).   The classic model of CA formation has centered around misrepair of DSBs. Exposing DNA to an endogenous or exogenous DSB-inducing agent directly results in DSBs, which may either persist or be misrepaired by inadequate repair mechanisms; in the event of this erroneous repair, CAs often eventually result (Bignold, 2009; Danford, 2012; Schipler & Iliakis, 2013) . Another model has been proposed that suggests CAs may actually be due to failure of enzymes that tether the DNA strands during the repair of enzyme-induced breaks in the DNA; the various pathways in the cell would likely employ assorted tethering enzymes. The numerous types of CAs would thus result from different kinds of tethering errors (Bignold 2009).   The type of CA that results may be dependent on the timing of inadequate repair. For example, DSBs may result in CSAs or CTAs depending on when during the cell cycle the DSB was incurred. DSBs that are not repaired before DNA duplication in the S-phase will be replicated and result in CTAs. If DSBs are incurred after the S-phase and are improperly repaired, CSAs  will result (Danford, 2012; Registre et al., 2016; Vodicka et al., 2018). Similarly, CNVs are thought to be induced during the DNA replication phase. DNA replication stops can also be problematic for repair. Although the mechanism is not well studied, it has been suggested that stress during replication, in particular stalling replication forks, prompt microhomology-mediated mechanisms to overcome the replication stall, which often results in duplications or deletions. Two models that have been proposed to explain this mechanism include the Fork Stalling and Template Switching (FoSTeS) model, and the Microhomology-Mediated Break-Induced Replication (MMBIR) model (Lee et al. 2007; Hastings et al. 2009; Arlt et al. 2012; Arlt et al. 2014; Wilson et al. 2015).   The type of CA may also be dependent on the type of erroneous repair that occurs. Deletions or chromosome breaks may occur when DSBs are left unrepaired (Danford 2012). Deletions may also occur when nucleotides are removed at the junctions (Schipler and Iliakis 2013) or when the wrong DNA ends are religated (Venkitaraman 2002). Ligation of the incorrect ends of DNA DSBs may also lead to translocations or dicentrics (Ferguson & Alt, 2001;  Lieber, 2010; Povirk, 2006; Venkitaraman, 2002). This type of error may occur when there are two or more DSBs in close proximity to each other that are misrejoined, thus resulting in the exchange of genetic material between two chromosomes (Ferguson and Alt 2001; Povirk 2006). NHEJ has been shown to play a significant role in the generation of chromosomal exchanges ( Lieber 2010; Povirk 2006; Weinstock et al. 2006). Evidence for this comes from analysis of breakpoint junctions, which typically have little to no chromosomal homology when NHEJ repair is used (Povirk 2006; Weinstock et al. 2006); this was demonstrated in studies using translocation reporters (reviewed in Weinstock et al., 2006). There are, however, two types of NHEJ. c-NHEJ has been shown to suppress exchanges (Simsek and Jasin 2010) , which may be due to its relatively rapid repair kinetics (Schipler and Iliakis 2013). Chromosomal exchanges are thus suggested to originate more often from alt-NHEJ (Simsek and Jasin 2010; Zhang and Jasin 2011; Schipler and Iliakis 2013) .   NHEJ is also thought to mediate the formation of other types of CAs. Based on analysis of breakpoint junctions in lung adenocarcinoma samples where reciprocal inversions were found between genes RET and KIF5B/CCDC6, the majority of the inversions were thought to be induced by NHEJ (Mizukami et al. 2014). Chromothripsis, which refers to a single event that results in a massive number of CAs localized to a single or very few chromosomes (Russo et al. 2015; Leibowitz et al. 2015; Rode et al. 2016), may also be linked to NHEJ. The single catastrophic event sparking chromothripsis likely induces a large quantity of DSBs, essentially shattering the chromosome(s). These DSBs are then processed mainly by the error-prone NHEJ, which results in a large number of CAs, including chromosomal rearrangements, CNVs, and loss of heterozygosity (Leibowitz et al. 2015; Rode et al. 2016).   Fusing two broken chromosomes may lead to the formation of dicentric chromosomes, which are characterized by the presence of two centromeres. Dicentrics may also be formed by telomere-to-telomere end fusions (Fenech and Natarajan 2011; Rode et al. 2016). Telomeres, composed of TTAGGG repeats, are important structures that protect the ends of chromosomes and ensure accurate replication (Ferguson and Alt 2001; Hoeijmakers 2001; Vodicka et al. 2018); these nucleoprotein structures are shortened (Vodicka et al. 2018) by approximately 100 base pairs after each division, and are only replenished in cell types expressing the enzyme telomerase (Hoeijmakers 2001). If the telomeres become critically short, they can be mistaken for broken DNA ends by DNA repair machinery, and thus may be ‘repaired’ by fusing the ends of two chromosomes together (Ferguson and Alt 2001; Vodicka et al. 2018).    Dicentrics can also contribute to other types of CAs. During mitosis, the two centromeres of a dicentric chromosome may be pulled to opposite ends of the cell by mitotic spindle (Ferguson and Alt 2001; Fenech and Natarajan 2011; Leibowitz et al. 2015; Rode et al. 2016). Because the ends of the chromosomes are fused, this can lead to the formation of an anaphase chromatin bridge between the daughter cells (Russo et al. 2015; Leibowitz et al. 2015; Rode et al. 2016). If this bridge persists beyond anaphase, it may become enclosed in a nucleoplasmic membrane along with the nucleus, thus generating a NPB (Fenech and Natarajan 2011). Eventually, however, these bridges do break (Ferguson and Alt 2001; Fenech and Natarajan 2011; Russo et al. 2015; Leibowitz et al. 2015; Rode et al. 2016); the break is nearly always uneven, meaning that one daughter cell will be missing genetic material and one will have extra genetic material (Fenech and Natarajan 2011). These fragments, with their ‘sticky’ ends from the break, may further propagate the formation of CAs by being ligated inappropriately to another chromosome. Thus the cycle, known as the breakage-fusion-bridge (BFB) cycle, is propagated and further contributes to chromosomal instability (Ferguson and Alt 2001; Fenech and Natarajan 2011; Russo et al. 2015; Leibowitz et al. 2015; Rode et al. 2016) .    MN may also be formed during this BFB cycle. When the anaphase bridges break, the remaining chromosome fragments may be packaged by a nuclear membrane into its own mini nucleus, thus, forming an MN. MN may also enclose acentric chromosome fragments, chromatid fragments, or even entire chromosomes that were not properly segregated during mitosis (Fenech and Natarajan 2011; Doherty et al. 2016). Similar to MN in structure are NBUDs; the only difference between these two structures is that NBUDs are still attached to the nucleus by nucleoplasmic material. A NBUD is formed if there is amplified DNA that needs to be removed; this amplified material is often segregated from the other DNA at the periphery of the nuclear membrane and excluded from the nucleus by budding, resulting in a NBUD. Additionally, NBUDs may also result from NPB breakages (Fenech and Natarajan 2011).

There is moderate empirical evidence supporting the relationship between inadequate DNA repair and the frequency of CAs. The evidence presented below is summarized in table 6, <a href="https://docs.google.com/spreadsheets/d/1iehBBqhFFSOhgis-0U3tasQwJ50bZJPVmenWUiR4vmA/edit?usp=sharing" target="_blank">here (click link)</a>. Several reviews discuss evidence that associates these two events (Ferguson and Alt 2001; van Gent et al. 2001; Sishc and Davis 2017; Venkitaraman 2002). Overall, however, there is weak empirical evidence available supporting a dose and incidence concordance, little empirical evidence supporting a temporal concordance, and strong empirical evidence supporting essentiality for this KER.    Dose and Incidence Concordance There is weak empirical evidence available that directly examines the dose and incidence concordance between DNA repair and CAs within the same study. There are, however, studies that use an ionizing radiation stressor to examine dose concordance of either inadequate DNA repair in response to radiation exposure, or CA frequencies in response to irradiation. In an analysis that amalgamated results from several different studies conducted using in vitro experiments, the rate of DSB misrepair was revealed to increase in a dose-dependent fashion from 0 - 80 Gy (Mcmahon et al. 2016). Similarly, there was a clear correlation between radiation dose (i.e., increasing amounts of energy deposition) between 0 - 10 Gy and different clastogenic endpoints (Thomas et al. 2003; Tucker et al. 2005A; George et al. 2009; Arlt et al. 2014; Balajee et al. 2014; Lin et al. 2014; Suto et al. 2015; Mcmahon et al. 2016) . Overall, this suggests that exposure to radiation may increase both inadequate repair of DNA damage and the frequency of CAs in a dose-dependent fashion. More studies, however, are required to better assess the dose and incidence concordance of this KER.   Temporal Concordance Temporal concordance between inadequate DNA repair and CA frequency is not well established. One study using cells pretreated with a DNA-PK inhibitor and irradiated with gamma rays found that DNA repair and MN were evident when they were assessed at 3 hours post-irradiation and 24 hours post-irradiation, respectively (Chernikova et al. 1999). This study does therefore suggest that there may be temporal concordance between these two events.  Other radiation-based studies examining these two events separately, however, do not provide clear evidence of temporal concordance between DNA repair and CA frequency.    Essentiality Numerous studies demonstrate that simply knocking-out one gene involved in DNA repair, without any other added stressor, is enough to increase the frequency of CAs in several types of cells (Karanjawala et al. 1999; Patel et al. 1998; Wilhelm et al. 2014). Further strengthening this relationship, addition of a DSB-inducing stressor to these DNA repair knock-out cells also significantly increases CA levels relative to wild-type cells receiving the same treatment (Cornforth and Bedford 1985; Simsek and Jasin 2010; Lin et al. 2014; Mcmahon et al. 2016). Essentiality is also supported by looking at patients with the recessive genetic disorder ataxia-telangiectasia (AT), in which mutations in the gene encoding the ATM protein results in defects in DNA damage repair signaling. One recent study showed that in comparison to control patients, patients with AT had increased levels of several types of CAs. Upon exposure to a DSB-inducing stressor such as ionizing radiation, these patients showed further increases in these aberrations as well as a significant increase in the levels of complex aberrations as compared to controls (Bucher et al. 2021).    Inhibitor studies have also found similar results. Two strains of wild-type cells that were treated with hydroxyurea, which is known to inhibit DNA repair, both had increased CAs relative to untreated wild-type cells (Wilhelm et al. 2014). Similarly, immortalized myeloid cell lines, cells from patients with myeloid leukemia, and cells from healthy donors were all found to have dose-dependent decreases in ligation efficiency after being treated with increasing doses of antibodies against various NHEJ proteins (Heterodimer et al. 2002). In addition, cells that were pretreated with DNA-PK inhibitor wortmannin prior to being irradiated were found to have not only increased levels of MN, but also decreased rates of DNA rejoining (Chernikova et al. 1999). A study by White et al. (2010) reported similar results under ATM and DNA-PK inhibition, where IR-exposed human lung cells treated for 1 hour with a reversible inhibitor of either enzyme exhibited an elevated level of CAs at all tested doses of IR, compared to the non-inhibited, IR-exposed cells 48 hours post-exposure. These findings demonstrated that even a transient inhibition of ATM or DNA-PK can sufficiently disrupt DNA damage repair and lead to CAs (White et al., 2010). Functional defects in the factors involved in NER due to mutations or knock-down/out have shown concordant results that are supportive of this KER. For example, UV61 Chinese hamster ovary cells (homogolous to human Cockayne syndrome group B cells), which have a defective ERCC6 gene, are incapable of repairing UV-induced cyclobutane pyrimidine dimers due to the compromised transcription-coupled NER (TCR). Following UV exposure, a significantly higher percentage of TCR-defective UV61 cells contained CAs than another Chinese hamster ovary cell line that is TCR-proficient (Proiettis de Santis et al., 2001). Down-regulation of xeroderma pigmentosum group A–complementing protein (XPA) by RNA interference (RNAi) in human bladder cancer cells was observed to significantly increase the baseline frequency of MN, nucleoplasmic bridges, and nuclear buds, while overexpression of XPA by transfection in the same cell line reduced these levels below that in the control cells (Zhi et al., 2017). Both studies support the essentiality of inadequate repair in the occurrence of chromosomal aberrations.    A rescue experiment provided further evidence of the essential role DNA repair plays in relation to CA frequencies. Inhibition of NHEJ through knocking out either Ku70 or Xrcc4 resulted in higher CA frequencies in the form of translocations; when Xrcc4 was transiently expressed in Xrcc4-/- cells, translocations were significantly decreased by 5-fold(Simsek and Jasin 2010) . This provides strong evidence that the NHEJ repair pathway plays an important role in the formation of CAs, specifically translocations.

Uncertainties in this KER are as follows: <ol> <li>In an experiment using both wild-type and Ku70-/- cells, knock-down of alt-NHEJ protein CtIP resulted in significantly decreased translocations in both cell types. When CtIP expression was rescued, translocation frequencies in these cells also returned to normal levels. This however, is opposite to results obtained in a similar study, where knock-out of Ku70 or Xrcc4 led to increased translocation frequency, and Xrcc4 rescue experiments resulted in decreased translocations (Simsek and Jasin 2010). It should be noted that alt-NHEJ is thought to be the major repair pathway responsible for generating translocations (Simsek and Jasin 2010; Zhang and Jasin 2011; Schipler and Iliakis 2013).  </li> <li>There is currently discussion regarding the accuracy of HR relative to NHEJ. Traditionally HR has been considered the more accurate type of DNA repair, while NHEJ is classically described as error-prone. There is emerging evidence, however, suggesting that HR may in fact be a mutagenic process. Evidence supporting HR as an error-prone repair pathway has been reviewed (Guirouilh-barbat et al. 2014).</li> </ol>

DNA repair is a modulating factor in this KER. The progression from “Inadequate DNA repair” to “Increase, Chromosomal aberrations” only occurs when "Increase, DNA strand breaks" (KE 1635) precedes "Inadequate DNA repair", which indicates that DNA strand breaks could not be repaired.

Quantitative understanding of this linkage is lacking. Most data are derived from studies that examined DSB misrepair rates or CA rates in response to a radiation stressor.  In terms of inadequate DNA repair, the rate of DSB misrepair was found to be approximately 10 - 15% at 10 Gy of radiation (Lobrich et al. 2000); this rate increased to 50 - 60% at a radiation exposure of 80 Gy (Kuhne et al. 2000; Lobrich et al. 2000; Mcmahon et al. 2016). It is not known, however, how this rate of inadequate repair directly relates to CA frequency. Overall, more studies are required that directly assess this relationship.

Studies directly examining the response-response relationship between inadequate repair and CA frequency are lacking. One study examined both DNA repair and CA frequency in cells exposed to DNA-PK inhibitor wortmannin. There was a negative, approximately linear relationship between DNA repair and increasing wortmannin dose, and a positive, approximately linear relationship between MN frequency and increasing wortmannin dose; this suggests that as adequate DNA repair declines, CA frequency increases (Chernikova et al. 1999). More studies are required, however, that directly quantify the response-response relationship between inadequate DNA repair and CAs.

The time scale between inadequate DNA repair and the increased frequency of CAs has not been well-established. Most data come from studies that assess only one of these events in relation to a radiation stressor rather than assessing the timing of the events relative to each other. More studies are thus required that directly assess this relationship.

Not identified.

Low Unspecific

Low

All life stages

Low Low Low

This KER is plausible in all life stages, sexes, and organisms with chromosomes. The majority of the evidence is from in vitro fetal human male models. No in vivo evidence was found to support the relationship.

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AOPWiki 2019-05-19T16:37:11

2024-12-17T12:37:31

<upstream-id>669b23ca-0e30-4508-ae51-b6d96fb4949d</upstream-id> <downstream-id>6f3cc035-087f-41e7-bdfa-80177f2f7b4e</downstream-id> DNA strand breaks (single and double) can arise from endogenous processes (e.g., topoisomerase reaction, excision repair, and VDJ recombination) and exogenous insults (e.g., replications stressors, ionizing radiation, and reactive oxygen species). Single strand breaks (SSBs) are generally repaired rapidly without error. However, multiple SSBs in close proximity to each other and interference of replication by unrepaired SSBs can lead to double strand breaks (DSB). DSB are more difficult to repair and are more toxic than SSB (Kuzminov, 2001). DSBs may lead to chromosomal breakages that may permanently alter the structure of chromosomes (i.e., chromosomal aberrations) and cause loss of DNA segments.

DNA strand breaks are a necessity for chromosomal aberrations to occur. However, not all strand breaks lead to clastogenic events as most of them is repaired rapidly by a variety of different repair mechanisms. DNA DSBs are the critical damage because they lead to chromosome breakage. It is well-understood that unrepaired DSBs can lead to chromosomal aberrations. Studies have demonstrated DSBs leading to irreversible structural damage; for example, treatment of cultured cells with replication stress-inducing agents such as hydroxyurea induced micronuclei that are positive for gamma-H2AX, a marker of DSBs (Xu et al., 2010). The link between DSBs and the importance of DSB repair processes, such as non-homologous end joining (NHEJ) and homologous recombination (HR), in preventing chromosomal aberrations/genomic instability is extensively discussed in literature and many reviews are available (van Gent et al., 2001; Ferguson and Alt, 2001; Hoeijmakers, 2001; Iliakis et al., 2004; Povirk, 2006; Weinstock et al., 2006; Natarajan and Palitti, 2008; Lieber et al., 2010; Mehta and Haber, 2014; Ceccaldi et al., 2016; Chang et al., 2017; Sishc and Davis, 2017; Brunet and Jasin, 2018). In addition, attempted repair of DSBs can lead to chromosomal aberrations such as translocations; NHEJ is a recognized source of oncogenic translocations in human cancers (Ferguson and Alt, 2001; Weinstock et al., 2006; Byrne et al., 2014; Brunet and Jasin, 2018), and a contributor to the carcinogenic process (Hoeijmakers, 2001; Sishc and Davis, 2017). Other types of chromosomal aberrations can serve as indicators of genomic instability that can contribute to a variety of adverse health effects including neurodegeneration (Madabhushi et al., 2014).

In vitro studies demonstrating dose and temporal concordance <ul> <li>In the 2009 and 2011 studies by Platel et al. TK6 cells were exposed to bleomycin and glucose oxidase (H2O2-generating enzyme) for 1 hour at increasing concentrations (Platel et al., 2009; Platel et al., 2011). <ul style="list-style-type:circle"> <li>Concentration-dependent increase in DNA strand breaks was measured using the alkaline comet assay 1 hr post-exposure <ul style="list-style-type:circle"> <li>First statistically significant concentration: bleomycin: 0.5 µM; GOx: 1.08x10-5 units/mL</li> <li>NOEL could not be defined, indicating that there was response at every tested concentration.</li> </ul> </li> <li>MN frequency was measured 23 hours post exposure; concentration-dependent increase in MN frequency was observed and NOEL was identified. <ul style="list-style-type:circle"> <li>NOEL: bleomycin: 0.023 µM; GOx: 1.78x10-5 units/mL</li> <li>All concentrations above the NOEL induced significant increases in MN frequency.</li> </ul> </li> <li>Thus, the data demonstrate temporal concordance for both stressors; lack of concordance in the concentration at which response for bleomycin occurs is likely due to differences in detection sensitivities between these assays.</li> </ul> </li> </ul>   <ul> <li>Strand breaks and chromosomal breakage were measured in V79 cells with the comet assay and the MN test after exposure to hyperbaric oxygen at 3 bar for different periods of time (Rothfuss et al., 1999). <ul style="list-style-type:circle"> <li>Stand breaks were observed in the comet assay after treatment of 3 bar hyperbaric oxygen starting at treatment times of 30 mins. The effect increased constantly up to 180 min.</li> <li>The MN frequency was measured 20 h post treatment and showed increasing numbers of MN starting at treatment times of 30 mins, being clearly increased at treatment times of 60 min up to 180 min. </li> <li>These data demonstrate both dose- and temporal concordance in DNA strand breaks observed by comet assay and MN frequency.</li> </ul> </li> </ul>   <ul> <li>Lymphoblastoid cell lines were investigated with the comet assay and the MN test using gamma irradiation of 1 and 2 Gy (Trenz et al., 2003). Pulsed field gel electrophoresis was used additionally to investigate the occurrence of strand breaks (Trenz et al., 2005). <ul style="list-style-type:circle"> <li>Strand breaks were shown in the comet assay in all cell lines tested, immediately after treatment with 1 and 2 Gy.  </li> <li>40 h post treatment the cell lines were prepared for MN analysis: an increase in MN frequency was shown in all cell lines after treatment with 1 and 2 Gy.</li> <li>Thus, the data demonstrate both temporal and dose concordance.</li> </ul> </li> </ul>   <ul> <li> Watters et al. (2009) treated mouse embryonic fibroblasts (MEFs) with bleomycin for 4 hours and conducted comparative investigations using the H2AX assay, the comet assay and the MN test (Watters et al., 2009). <ul style="list-style-type:circle"> <li>The occurrence of DNA DSB was shown with the gamma-H2AX assay immediately following exposure. The number of foci increased up to 0.1 µg/ml; however, it was not statistically significant until 1 µg/ml and above.</li> <li>The comet assay showed a continuous increase in tail moment immediately following exposure, showing more than 2-fold increase at 10 µg/ml, but did not reach statistical significance.</li> <li>Significant increases in MN frequency was observed 26h post exposure (~1.5 cycles) at concentrations of 0.1µg/ml and above.</li> <li>These data support temporal concordance; lack of concordance in the dose at which the endpoints reach statistical significance is likely the rest of different sensitivities of these assays.</li> </ul> </li> </ul>   <ul> <li>Using bleomycin as a stressor, Kawaguchi et al. monitored DNA strand breaks in TK6 human lymphoblastoid cells with the comet assay/modified comet assay using DNA repair inhibitors and monitored clastogenic events with the MN test after a treatment period of 2h (Kawaguchi et al., 2010). <ul style="list-style-type:circle"> <li>In the regular alkaline comet assay an increase in DNA strand breaks was observed immediately following the 2h exposure, reaching significance at 12.5 µg/mL, and in the modified AraC/HU version at 6.25 µg/ml.</li> <li>A statistically significant increase in MN frequency was observed 24 h after treatment at 5 µg/mL.</li> <li>This provides support for temporal-concordance and the lack of dose-concordance is consistent with the increased sensitivity of the MN assay relative to the comet assay.</li> </ul> </li> </ul>   <ul> <li>Wild type and N-methylpurine DNA glycosylase (MPG)-deficient (Mpg-/-) Mouse embryonic fibroblasts (MEFs) were treated with increasing concentrations of methyl methane sulfonate (MMS) (0.5, 1, 1.5, 2.5 mM) for 1 hour (Ensminger et al., 2014). <ul style="list-style-type:circle"> <li>DSBs were measured as the number of γH2AX foci immediately following the exposure.</li> <li>There was a concentration-dependent increase in DSBs in wild type MEFs, and the increase was significantly larger in wild type compared to Mpg-/- cells at every concentration.</li> <li>Chromosomal aberrations (breaks and translocations) were monitored in metaphase spreads 24h following 1h 1 mM MMS treatment.</li> <li>At 1 mM MMS, the amount of chromatid breaks and translocations was significantly larger in wild type cells, compared to Mpg-/- cells, concordant with the observations in DSBs.</li> <li> The results support that increases in DSBs lead to increases in chromosomal aberrations.</li> </ul> </li> </ul>   <ul> <li>Dertinger et al. (2019) exposed TK6 cells to 34 diverse genotoxic chemicals over a range of concentrations for 24 hrs (Dertinger et al., 2019). At 4 and 24 hr time points cell aliquots were evaluated with the MultiFlow assay, which includes the gH2AX biomarker. At the 24 hr time point, remaining cells were evaluated with the in vitro MicroFlow assay, which includes %MN measurements. <ul style="list-style-type:circle"> <li>Benchmark dose analyses were conducted to estimate Point of Departure values for MN and gamma-H2AX responses.   </li> <li>In vitro MN and gamma-H2AX BMD confidence intervals for 18 clastogens were graphed on cross system plots. Good correlations were observed for 24 hr MN and 24 hr gamma-H2AX (shown), as well as 24 hr MN and 4 hr gamma-H2AX (not shown).</li> <li>Thus, the data demonstrate both temporal and dose concordance for these endpoints.</li> </ul> </li> </ul>   <ul> <li>Isolated lymphocytes and whole blood samples taken from four healthy, adult males were exposed to gamma-ray radiation at 20 cGy/minute at doses ranging from 0 – 50 cGy. Immediately following irradiation, DNA strand breaks were assessed using the comet assay and chromosomal aberrations were examined by cytogenetic analysis (Sudprasert et al., 2006). <ul> <li>In irradiated lymphocytes, there were dose-dependent increases in the number of DNA strand breaks, with significant increases in strand breaks evident from 5 – 50 cGy doses.    </li> <li>Irradiated whole blood samples showed significantly increased strand breaks by 10 cGy, but this level stayed relatively stable from 10 - 50 cGy.</li> <li>Analysis of chromosomal aberrations in irradiated whole blood samples indicated dose-dependent increases in deletions and dicentric chromosomes across 50 cGy, with more deletions detected than dicentrics. All doses (5 – 50 cGy) showed significantly more aberrations than unirradiated controls.</li> <li>The results of this study support dose concordance and are suggestive of time concordance.</li> </ul> </li> </ul>   <ul> <li>In a study by Chernikova et al. 1999, PL61 cells were exposed to radiation sensitizer/DNA repair inhibitor wortmannin prior to gamma-ray irradiation, and then analyzed for DSBs and micronuclei (indicative of chromosomal aberrations) (Chernikova et al., 1999). <ul> <li>DSB experiments were performed with cells treated with 25 µM of wortmannin + radiation, and with cells exposed only to radiation. In both cases, there was a linear, dose-dependent increase in the number of DSBs across radiation doses ranging from 0 – 60 Gy, as measured by the FAR assay. Wortmannin treatment did not affect the number of DSBs that were formed.</li> <li>In terms of DNA repair, however, cells irradiated with 45 Gy of gamma-rays showed a dose-dependent decline in the percentage of DNA repair with increasing wortmannin concentrations from 0 – 25 µM.</li> <li>Furthermore, cells treated with wortmannin + 2 Gy of radiation demonstrated a dose-dependent increase in the number of micronuclei from 0 – 25 µM of wortmannin.</li> <li>Overall, the results of this study suggest that as the number of DSBs increase and repair processes are inhibited, there is a corresponding increase in the number of chromosomal aberrations. Thus the data demonstrate dose concordance and essentiality.</li> </ul> </li> </ul>   <ul> <li>Iliakis, et al. (2019) studied the relationship between DSB damage and chromosomal aberrations using an experimental model that mimics the clustered DNA DSB damage induced by high linear energy transfer (LET) radiation (Iliakis et al., 2019). Chinese hamster ovary cells and human retinal epithelial cells were engineered to carry I-SceI meganuclease recognition sites at specific locations in order to generate specific DSB clustered damage. Cells were then transfected with plasmids expressing I-SceI to induce the DNA breakages. Twelve hours or 24 hours post-transfection, cells were analyzed by immunofluorescence microscopy for DSBs, and by cytogenetic analysis for chromosome translocations.     <ul> <li>DSBs were increased in all cells transfected with the endonuclease relative to cells from the same cell lines that underwent a mock transfection.</li> <li>Chromosomal translocations were also elevated in cell lines transfected with an endonuclease, with increasing chromosomal translocations found in cells with increasing DSB cluster damage.</li> <li>This study shows an association between DSB cluster damage and chromosomal translocation incidence.</li> </ul> </li> </ul>   In vivo studies <ul> <li>Sprague-Dawley rats were dosed with different genotoxic compounds at select concentrations (methotrexate, cisplatin, chlorambucil, and cyclophosphamide) and blood samples were collected at different time points following the dosing (6, 12, 24, 36, 48, 72, and 96 hours post dosing) (Mughal et al., 2010). <ul style="list-style-type:circle"> <li>Peripheral blood lymphocytes were isolated for comet assay and peripheral blood erythrocytes were used to measure MN at each time point. <ul> <li>Different comet assay parameters such as tail length, moment, olive tail moment, and % tail DNA were compared to MN frequency</li> <li>All comet assay parameters had a positive correlation to MN frequency demonstrated in all chemical treatments.</li> <li>DNA tail length and % tail DNA showed visible increases in strand breaks at early time points (6 and 12h), while the increase in MN frequency was not observed until after 12-24 h.</li> <li>This early response at 6 h was not observed in tail moment or olive tail moment; these two paramenters did not show as strong of a response as tail length and % tail DNA to all four chemical treatments.</li> </ul> </li> <li>The results suggest temporal concordance in strand breaks measured by comet assay and induction of MN, where strand breaks are observed earlier than MN.</li> </ul> </li> </ul>   <ul> <li>C57BL/6 mice were irradiated with increasing doses of X-rays (1.1, 2.2, 4.4 Gy) at rate of 1.03 Gy/min (acute high dose) and 0.31 cGy/min (low dose rate). Lymphocytes were isolated and collected 24h and 7 days from the start of irradiation (different mice were used for each time point) (Turner et al., 2015). <ul style="list-style-type:circle"> <li>γH2AX measured at 24h showed a dose-dependent increase in DSBs in both acute and low dose rate exposed mice. <ul> <li>The level of DSBs due to the acute dose treatment was significantly higher than due to the low dose rate treatment at 1.1 and 2.2 Gy.</li> </ul> </li> <li>MN frequency was also measured 24h and 7 days post exposure; <ul> <li>At both time points and in both treatment groups, MN frequency increased with dose from 1.1 and 2.2 Gy. However, there was no further increase at 4.4 Gy</li> <li>There was no statistical difference in the two treatment groups</li> </ul> </li> </ul> </li> </ul> Overall, the above data demonstrate that when strand breaks occur there is an increase in MN frequency, which is indicative of chromosomal aberrations. There is a clear temporal-concordance but dose-concordance is not always consistent due to differences in assay sensitivity.

As described above, statistically significant increases in MN occur, in some cases, at lower concentrations than strand breaks measured with the comet assay (Platel et al., 2001; Watters et al., 2009; Kawaguchi et al., 2010). The two assays measure different endpoints at different time points; the MN test may appear to be more sensitive than the comet assay but it is difficult to directly compare these two assays. Mughal et al. (2010) study compared different parameters of comet assay (tail moment, length, and % tail DNA) to MN frequency. Depending on the parameter, the observation of increase in strand breaks varied. For example, % tail DNA would show a visible increase in strand breaks at one concentration; however, no change would be observed in the tail moment calculated using the same data. Use of different parameters in presenting comet assay data may add subjectivity to the results that are reported in certain papers. Rossner Jr. et al. exposed human embryonic lung fibroblasts (HEL12469) to 1, 10, and 25 µM of benzo[a]pyrene (B[a]P) for 24 hours and measured DSB (γH2AX immunodetection by Western blotting) and translocations (by fluorescence in situ hybridization of chromosomes 1, 2, 4, 5, 7, 17) (Rossner Jr. et al., 2014). <ul style="list-style-type:circle"> <li>Increases in γH2AX were observed only at 25 µM B[a]P (~2.5 fold increase) after the 24h exposure.</li> <li>Translocations were quantified and expressed as the genomic frequency of translocations per 100 cells (FG/100) <ul> <li>All concentrations of B[a]P induced an elevated frequency of translocations compared to the DMSO control (DMSO: ~0.19/100; 1 µM: ~0.53/100 cells; 10 µM: ~0.33/100; 25 µM: ~0.39/100)</li> </ul> </li> </ul> In this study, the increase in translocations was detected at concentrations that did not induce an increase in γH2AX signal. This observation of the discordant relationship between γH2AX and translocations may be due to the differences in assay sensitivity. In addition, immunodetection by Western blotting cannot precisely measure small changes in protein content.

As with the regularly used alkaline comet assay a variety of DNA damage is detected – SSBs, DSBs, alkaline labile sites, as well as sites of DNA repair; thus, a quantitative understanding for specific types of damage is rather difficult. There exists the possibility to quantify the amount of DNA breaks by comparing the induced damage with Gy equivalents (Collins et al., 2008), however, this is not the standard. DSBs can be measured more specific with the neutral version of the Comet assay, however, this version is not that regularly used. As reviewed in Takahashi et al, 2005 the efficiency of DSB detection measured with the PFGE, the neutral Comet assay and the DNA elution assay has a lower detection limit of 100 DSB per cell (Collins et al., 2008). In contrast to this, each Gamma-H2AX focus seems to represent a DNA DSB in vivo (Rogakou et al., 1999).

High Unspecific

High

All life stages

High High High

DNA strand breaks and subsequent chromosomal aberrations can occur in any eukaryotic and prokaryotic cell.

Brunet, E. & M. Jasin (2018), "Induction of chromosomal translocations with CRISPR-Cas9 and other nucleases: Understanding the repair mechanisms that give rise to translocations.", Adv. Exp. Med. Biol. 1044:15-25. Byrne, M. et al. (2014), "Mechanisms of oncogenic chromosomal translocations.", Ann. N.Y. Acad. Sci., 1310:89-97. Ceccaldi, R., B. Rondinelli & A.D. D'Andrea (2016), "Repair Pathway Choices and Consequences at the Double-Strand Break.", Trends Cell Biol. 26(1):52-64. Chang, H. et al. (2017), "Non-homologous DNA end joining and alternative pathways to double‑strand break repair.", Nature Rev. Mol. Cell. Biol., 18:495-506. Chernikova, S.B., R.L. Wells & M. Elkind (1999), "Wortmannin Sensitizes Mammalian Cells to Radiation by Inhibiting the DNA-Dependent Protein Kinase-Mediated Rejoining of Double-Strand Breaks.", Radiat. Res., 151:159-166. Collins, A.R. et al. (2008), "The comet assay: topical issues.", Mutagenesis, 23:143-151. Dertinger, S.D. et al. (2019), "Predictions of genotoxic potential, mode of action, molecular targets, and potency via a tiered multiflow® assay data analysis strategy.", Environ. Mol. Mutagen., 60(6):513-533 Ensminger, M. et al. (2014), "DNA breaks and chromosomal aberrations arise when replication meets base excision repair.", J. Cell Biol.,  206:29. Ferguson, D.O. & F.W. Alt (2001), "DNA double strand break repair and chromosomal translocation: Lessons from animal models.", Oncogene 20(40):5572–5579. Hoeijmakers, J.H. (2001), "Genome maintenance mechanisms for preventing cancer.", Nature, 411:366-374. Iliakis, G. et al. (2019), "Defined Biological Models of High-LET Radiation Lesions.", Radiat. Protect Dosimet., 183:60-68. Iliakis, G. et al. (2004), "Mechanisms of DNA double strand break repair and chromosome aberration formation.", Cytogenet. Genome Res. 104:14-20. Kawaguchi, S. et al. (2010), "Is the comet assay a sensitive procedure for detecting genotoxicity?.", J. Nucleic Acids, 2010:541050. 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2023-01-09T21:05:56

Bulky DNA adducts leading to chromosomal aberrations and mutations

Bulky DNA adducts, chromosomal aberrations, and mutations

Beckner Andersano

BY-SA

2.8

A variety of stressors can induce bulky adducts. The most well studied are Benzo(a)pyrene, Aflatoxin B1 and Aristolochic acid. These compounds have been demonstrated to induce bulky adducts in various in vivo and in vitro methods (). Many studies use 32-P post labelling and other methods to observe the presence of bulky adducts when different doses of a carcinogenic compound are administered. A dose-dependent increase of bulky adducts measured by 32P post-labelling and comet assay was observed by Long et al., 2018 and Lemieux et al., 2011 from Benzo(a)pyrene doses as low as 0.20 mg/kg BW/day in mammalian cells from various tissues in vivo. AFB1-induced adducts were observed by Woo et al. (year) by isotope dilution mass spectrometry (MS) in mice. A dose-dependent increase in Aristolochic Acid induced adducts have been demonstrated by 32-P post labelling in in rodent models (Mei et al., 2006 and McDaniel et al., 2012). The authors noted an increase in adducts from a dose as low as 0.1 mg/kg bodyweight (Li et al., 2020).

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AOPWiki 2026-06-25T11:56:40

2026-06-29T16:54:03