SNAPSHOT

Event: 1634: Increase, Oxidative damage to DNA

Short Name: Increase, Oxidative DNA damage

AOPs Including This Key Event

Stressors

Biological Context

Cell term

Taxonomic Applicability

Life Stage Applicability

Sex Applicability

Theoretically, DNA oxidation can occur in any cell type, in any organism. Oxidative DNA lesions have been measured in mammalian cells (human, mouse, calf, rat) in vitro and in vivo, and in prokaryotes.

Event: 155: N/A, Inadequate DNA repair

Short Name: N/A, Inadequate DNA repair

AOPs Including This Key Event

Stressors

Biological Context

Taxonomic Applicability

Life Stage Applicability

Sex Applicability

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-PK_cs 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).

Event: 1635: Increase, DNA strand breaks

Short Name: Increase, DNA strand breaks

AOPs Including This Key Event

Stressors

Biological Context

Taxonomic Applicability

Life Stage Applicability

Sex Applicability

DNA strand breaks can occur in any eukaryotic or prokaryotic cell.

Event: 185: Increase, Mutations

Short Name: Increase, Mutations

AOPs Including This Key Event

Stressors

Biological Context

Taxonomic Applicability

Life Stage Applicability

Sex 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.

Event: 1636: Increase, Chromosomal aberrations

Short Name: Increase, Chromosomal aberrations

AOPs Including This Key Event

Stressors

Biological Context

Taxonomic Applicability

Life Stage Applicability

Sex Applicability

Chromosomal aberrations indicating clastogenicity can occur in any eukaryotic or prokaryotic cell. However, dose-response curves can differ depending on the cell cycle stage when the DSB agent was introduced (Obe et al., 2002).

Relationship: 1909: Increase, Oxidative DNA damage leads to N/A, Inadequate DNA repair

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

Oxidative DNA damage can occur and overwhelm the repair mechanisms in any eukaryotic and prokaryotic cell. Observation of this KER has been described in mammalian cells, yeast, and bacteria.

Key Event Relationship Description

Oxidative DNA lesions are present in the cell at steady state due to low levels of reactive oxygen species (ROS) and other free radicals generated by endogenous processes involving redox reactions. The most prominent examples of oxidative DNA lesions include 7, 8-dihydro-8oxo-deoxyGuanine (8-oxo-dG), 2,6-diamino-4-hydroxy-5-formamidopyrimidine (FaPydG), and thymidine glycol (Tg). Under homeostatic conditions, cells are able to regulate the level of free radicals and readily repair oxidized DNA bases using basal repair mechanisms to prevent irreversible damage (Swenberg et al., 2011). Oxidative DNA lesions are mainly repaired by base excision repair (BER) initiated by DNA glycosylases such as oxoguanine glycosylase 1 (OGG1), endonuclease III homologue 1 (NTH1), and Nei-like DNA glycosylases (NEIL 1/2), which detect and remove damaged bases. Abasic sites are then cleaved by endonucleases or lyases, resulting in transient single-strand breaks (SSB) that enter either short-patch or long-patch repair. Nucleotide excision repair (NER) is involved in repairing oxidized bases to a lesser extent (Shafirovich et al., 2016). Increase in free radicals or exposure to oxidizing agents can increase the level of oxidative DNA lesions and overwhelm the repair pathways, compromising the quality of repair. If the repair mechanisms are compromised, oxidative lesions may accumulate (insufficient repair) and cause incorrect base pairing during replication or incomplete repair (indicated by accumulation of repair intermediates) (Markkanen, 2017).

Evidence Supporting this KER

Inadequate repair of oxidative lesions is indicated by an increase in oxidative lesions above background, activation of repair enzymes, increase in repair intermediates (abasic sites and single strand breaks), and incorrect base insertion opposite lesion during replication (lesion bypass by translesion DNA synthesis).

Biological Plausibility

The mechanism of repair of oxidative DNA lesions in humans is well-established and numerous literature reviews are available on this topic (Berquist and Wilson III, 2012; Cadet and Wagner, 2013). As described above, oxidative DNA lesions are mostly repaired via BER and, to a lesser extent, NER. Previous studies have reported thresholded dose-response curves in oxidative DNA damage and attributed these observations to exceeded repair capacity at the inflection point on the curve (Gagne et al., 2012; Seager et al., 2012). In vivo, increase and accumulation of oxidative DNA lesions despite the activation of BER have been observed following chemical exposures, demonstrating insufficient repair of oxidative DNA lesions past a certain level (Ma et al., 2008).

OGG1 and NTH1, the glycosylases that initiate the BER of 8-oxo-dG and thymine glycol (Tg) lesions, respectively, are bifunctional, containing both glycosylase and lyase activities. The glycosylase removes the oxidized guanine by cleaving the glycosidic bond, giving rise to an apurinic site. The lyase then cleaves the phosphodiester bond 5’ to the AP site; a transient SSB is created for further processing in BER (Delaney et al., 2012). Abasic sites created by OGG1 and other glycosylases are also processed by apuric/apyrimidinic endonucleases (APE1) to create the 5’ nick (Allgayer et al., 2016).

Previous studies have demonstrated that an imbalance in any one of the multiple steps of BER can lead to an accumulation of repair intermediates and failed repair. Given that OGG1 is relatively slower in releasing its catalytic product than other glycosylases, it is highly likely that a disproportionate increase in oxidative DNA lesions compared to the level of available OGG1 would lead to an imbalance between lesions and the initiating step of BER (Brenerman et al., 2014). Accumulation of oxidative lesions would be observed as a result. Moreover, studies have reported accumulation of SSB due to OGG1 and NTH1 overexpression, demonstrating that the imbalanced lyase activity generates excessive SSB intermediates (Yang et al., 2004; Yoshikawa et al., 2015; Wang et al., 2018). 

Increases in oxidative lesions may produce more lesions and repair intermediates in close proximity to each other. Previous studies in mammalian cell extracts have reported reduction in repair efficiency when oxidative lesions are in tandem or opposite each other. For example, OGG1 showed reduced binding to 8-oxo-dG near an AP site incision. Furthermore, the OGG1-8-oxo-dG complex has been observed to hinder the repair of neighbouring AP site incision, delaying the completion of BER; this interaction between BER enzymes has been suggested to cause an accumulation of oxidative lesions and repair intermediates (Pearson et al., 2004; Budworth et al., 2005; Bellon et al., 2009; Yoshikawa et al., 2015; Sharma et al., 2016).

If oxidative lesions persist in the genome due to insufficient repair, incorrect base insertion opposite unrepaired oxidative DNA lesions may occur during replication. This is a well-established event. For example, 8-oxo-dG and FaPydG, the two most prominent oxidative DNA lesions, are able to form base pairs with dATP, giving rise to G:C→T:A transversions after subsequent DNA synthesis (Freudenthal et al., 2013; Gehrke et al., 2013; Markkanen, 2017). Replicative DNA polymerases such as DNA polymerase α, δ, and ε (pol α, δ, ε) have a poor ability to extend the DNA strand past 8-oxo-dG:dCTP base pairs and may cause replication to stall or incorrectly insert dATP opposite 8-oxo-dG (Hashimoto et al., 2004; Markkanen et al., 2012). In stalled replication forks, repair polymerases may be recruited to perform translesion DNA synthesis (TLS). Human Y-family DNA polymerases (Rev 1, pol κ, ι, and η) are DNA repair polymerases mainly involved in TLS in stalled replication forks. However, TLS is not free of error and its accuracy differs for each repair polymerase. For example, it is known that pol κ and η perform TLS across 8-oxo-dG and preferentially insert dATP opposite the lesion, generating G:C→T:A transversions. The error-prone nature of bypassing unrepaired oxidative lesions has been described in many previous studies and reviews (Greenberg, 2012; Maddukuri et al., 2014; Taggart et al., 2014; Shah et al., 2018).

Repair by OGG1 requires 8-oxo-dG:dC base pairing, thus, it is unable to repair 8-oxo-dG:dA mispairing in newly synthesized strands. The repair of 8-oxo-dG:dA base pairs post-replication is performed by MUT Y homologue, MYH, an adenine DNA glycosylase. However, the removal of dA instead of the damaged guanine may lead to futile cycles of BER because: 1) another dA is often inserted opposite the lesion, or 2) BER ligases have a poor ability of ligating the 3’end of dC opposite 8-oxo-dG (Hashimoto et al., 2004; Caglayan and Wilson, 2015). Accumulated 8-oxo-dG may be more resistant to repair post-replication due to this futile BER.         

Empirical Evidence

Example in vitro studies demonstrating dose and temporal concordance, or essentiality

• Human normal hepatocytes (HL-7702) were exposed to N,N-dimethylformamide for 24 hours at increasing concentrations (C. Wang et al., 2016)
  • Concentration-dependent increase in ROS was observed; the increase was statistically significant compared to control at all concentrations (6.4, 16, 40, 100 mM)
  • No significant increase in 8-oxodG was observed until the highest two concentrations (40 and 100 mM) indicating insufficient repair at these concentrations
  • Significant up-regulation of excision repair genes (XRCC2 and XRCC3) occurred at 6.4 and 16 mM, below the concentrations that significantly induced 8-oxodG, supporting sufficient DNA repair at these low concentrations.
  • These results demonstrate that repair is sufficient at low concentrations (rapidly removing 8-oxodG) and not until higher concentrations is repair overwhelmed (i.e., insufficient), where 8-oxo-dG significantly increases.

 

• AS52 Chinese hamster ovary cells (wild type and OGG1-overexpressing) were exposed to varying doses of ultraviolet A (UVA) radiation (Dahle et al., 2008)
  • Formamidopyrimidine glycosylase (Fpg)-sensitive sites were quantified using alkaline elution after increasing repair times (0, 1, 2, 3, 4 h) following 100 kJ/m² UVA irradiation
  • OGG1-overexpressing AS52 cells (OGG1+): Fpg-sensitive sites reduced to 71% within half an hour and down to background levels at 4h
  • Wild type AS52 cells: at 4h, 70% of the Fpg-sensitive sites remained, indicating accumulation of oxidative lesions
  • The above results demonstrated that excess OGG1 was able to prevent the accumulation of oxidative lesions, while the amount of OGG1 in wild type was insufficient to handle the amount of lesions induced by the same magnitude of UVA irradiation.
  • Mutations in the Gpt gene was quantified in both wild type and OGG1+ cells by sequencing after 13-15 days following 400 kJ/m² UVA irradiation
    • G:C→T:A mutations in UVA-irradiated OGG1+ cells were completely eliminated (thus, repair was sufficient when repair overexpressed).
    • G:C→T:A 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.
    • The above result also demonstrates the essentiality of 8-oxo-dG formation in the oxidative DNA damage-induced G to T transversion mutations.

 

• HL-60 human leukemia cells were irradiated with X-rays at a rate of 0.5 Gy/min for increasing durations (i.e., increasing doses). 8-OHdG levels were quantified by HPLC as number of 8-OHdG per 10⁶ deoxyguanosine (Li et al., 2013)
  • No increase in 8-OHdG was observed up to 2 Gy (sufficient repair at low doses), above which the level of lesions increased linearly up to 20 Gy (insufficient repair)
  • This thresholded dose-response curve, indicative of overwhelmed repair processes, was also observed in mouse liver in the same study described below.

 

In vivo studies demonstrating dose concordance

• Two groups of 5-week-old C57BL/6J mice were exposed to increasing doses of X-rays at a rate of 0.5 Gy/min (200 kV, 12 mA). The livers were collected from one group immediately after exposure and urine samples were collected over 24 hours following irradiation in the second group of mice (Li et al., 2013).
  • 8-OHdG in the mouse liver DNA were quantified by HPLC and expressed as 8-OHdG per 10⁶ deoxyguanosine
  • Between 0 and 0.5 Gy, no increase in lesions was observed
  • Between 0.5 and 30 Gy, a linear dose-response in 8-OHdG was observed
  • The thresholded dose-response curve was concordant in the urine samples; no increase in urinary 8-OHdG (8-OHdG/creatinine (ng/mg)) was observed between 0 and 0.1 Gy but between 0.1 and 5 Gy, the number of lesions increased linearly with dose

 

• Male Sprague-Dawley rats were fed 0.5 mmol aniline/kg/day for 30 days. Genomic DNA, nuclear extracts, and mitochondrial extracts were collected from spleen tissues (Ma et al., 2008).
  • 8-OHdG was quantified using enzyme-linked immunosorbent assay (ELISA) on digested genomic DNA. There was a significant 2.8-fold increase in lesions in aniline-fed rats than in control rats.
  • Both the nuclear extracts and mitochondrial extracts were tested for OGG1 activity, where 1.32-fold and 1.15-fold increase in enzyme activity (both significant; p<0.05) were observed in the respective extracts of aniline-treated rats. 
  • The OGG1 enzyme content in the extracts was detected using Western blotting; the increase in OGG1 content in aniline-treated rats was consistent with the OGG1 activity assay.
  • Despite the increase in OGG1 enzyme content and activity, the quantity of 8-OHdG increased.
  • Together, these results demonstrate that the level of OGG1 capacity was exceeded for repair of 8-oxo-dG, leading to accumulation of these lesions during this sub-chronic exposure. 

Uncertainties and Inconsistencies

Although the dual functionality of OGG1 as a glycosylase and lyase has been widely accepted and demonstrated experimentally, there are studies showing that the cleavage of phosphodiester bond 5’ to the lesion is mainly performed by apurinic endonuclease 1 (APE1) (Allgayer et al., 2016; R. Wang et al., 2018). In some cases, APE1 may be the main factor driving the accumulation of BER intermediates. Some studies suggest that OGG1 is involved in the repair of non-transcribed strands and is not required for transcription-coupled repair of 8-oxo-dG; Le Page et al. reported efficient repair of 8-oxo-dG in the transcribed sequence in Ogg1 knockout mouse cells (Le Page et al., 2000). Moreover, the repair of 8-oxo-dG is also affected by the neighbouring sequence; the position of the lesions may have a negative effect on repair efficiency (Pastoriza-Gallego et al., 2007). We note that the study by Allgayer et al. was investigating the fate and effect of 8-oxo-dG during transcription; repair mechanism may vary by situation and availability of repair enzymes at the time.

References

Allgayer, J., Kitsera, N., Bartelt, S., Epe, B., Khobta, A. (2016), Widespread transcriptional gene inactivation initiated by a repair intermediate of 8-oxoguanine, Nucleic Acids Res, 44:7267-7280.

Bellon, S., Shikazono, N., Cunniffe, S., Lomax, M., O’Neill, P. (2009), Processing of thymine glycol in a clustered DNA damage site: mutagenic or cytotoxic, Nucleic Acids Res, 37:4430-4440.

Berquist, B., Wilson III, D. (2012), Pathways for Repairing and Tolerating the Spectrum of Oxidative DNA Lesions, Cancer Lett, 327:61-72.

Brenerman, B., Illuzzi, J., Wilson III, D. (2014), Base excision repair capacity in informing healthspan, Carcinogenesis, 35:2643-2652.

Budworth, H., Matthewman, G., O’Neill, P., Dianov, G. (2005), Repair of Tandem Base Lesions in DNA by Human Cell Extracts Generates Persisting Single-strand Breaks, J Mol Biol, 351:1020-1029.

Cadet, J., Wagner, J.R. (2013), DNA Base Damage by Reactive Oxygen Species, Oxidizing Agents, and UV Radiation, Cold Spring Harb Perspect Biol, 5:a012559.

Caglayan, M., Wilson, S. (2015), Oxidant and environmental toxicant-induced effects compromise DNA ligation during base excision DNA repair, DNA Repair, 35:85-89.

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.

Delaney, S., Jarem, D., Volle, C., Yennie, C. (2012), Chemical and Biological Consequences of Oxidatively Damaged Guanine in DNA, Free Radic Res, 46:420-441.

Freudenthal, B., Beard, W., Wilson, S. (2013), DNA polymerase minor groove interactions modulate mutagenic bypass of a templating 8-oxoguanine lesion., Nucleic Acids Res, 41:1848-1858.

Gagne, J., Rouleau, M., Poirier, G. (2012), PARP-1 Activation— Bringing the Pieces Together, Science, 336:678-279.

Gehrke, T., Lischke, U., Gasteiger, K., Schneider, S., Arnold, S., Muller, H., Stephenson, D., Zipse, H., Carell, T. (2013), Unexpected non-Hoogsteen–based mutagenicity mechanism of FaPy-DNA lesions, Nat Chem Biol, 9:455-461.

Greenberg, M. (2012), Purine Lesions Formed in Competition With 8-Oxopurines From Oxidative Stress, Acc Chem Res, 45:588-597.

Hashimoto, K., Tominaga, Y., Nakabeppu, Y., Moriya, M. (2004), Futile short-patch DNA base excision repair of adenine:8-oxoguanine mispair, Nucleic Acids Res, 32:5928-5934.

Le Page, F., Klunglund, A., Barnes, D., Sarasin, A., Boiteux, S. (2000), Transcription coupled repair of 8-oxoguanine in murine cells: The Ogg1 protein is required for repair in nontranscribed sequences but not in transcribed sequences, Proc Natl Acad Sci USA, 97:8397-8402.

Li, Y., Song, M., Kasai, H., Kawai, K. (2013), Generation and threshold level of 8-OHdG as oxidative DNA damage elicited by low dose ionizing radiation, Genes Environ, 35:88-92.

Ma, H., Wang, J., Abdel-Rahman, S., Boor, P., Firoze, M. (2008), Oxidative DNA damage and its repair in rat spleen following subchronic exposure to aniline, Toxicol Appl Pharmacol, 233:247-253.

Maddukuri, L., Ketkar, A., Eddy, S., Zafar, M., Eoff, R. (2014), The Werner syndrome protein limits the error-prone 8-oxo-dG lesion bypass activity of human DNA polymerase kappa, Nucleic Acids Res, 42:12027-12040.

Markkanen, E. (2017), Not breathing is not an option: How to deal with oxidative DNA damage, DNA Repair, 59:82-105.

Markkanen, E., Castrec, B., Vilani, G., Hubscher, U. (2012), A switch between DNA polymerases δ and λ promotes error-free bypass of 8-oxo-G lesions, Proc Natl Acad Sci USA, 27:931-940.

Pastoriza-Gallego, M., Armier, J., Sarasin, A. (2007), Transcription through 8-oxoguanine in DNA repair-proficient and Csb−/Ogg1− DNA repair-deficient mouse embryonic fibroblasts is dependent upon promoter strength and sequence context, Mutagenesis, 22:343-351.

Pearson, C., Shikazono, N., Thacker, J., O’Neill, P. (2004), Enhanced mutagenic potential of 8-oxo-7,8-dihydroguanine when present within a clustered DNA damage site, Nucleic Acids Res, 32:263-270.

Seager, A., Shah, U., Mikhail, J., Nelson, B., Marquis, B., Doak, S., Johnson, G., Griffiths, S., Carmichael, P., Scott, S., Scott, A., Jenkins, G. (2012), Pro-oxidant Induced DNA Damage in Human Lymphoblastoid Cells: Homeostatic Mechanisms of Genotoxic Tolerance, Toxicol Sci, 128:387-397.

Shafirovich, V., Kropachev, K., Anderson, T., Li, Z., Kolbanovskiy, M., Martin, B., Sugden, K., Shim, Y., Min, J., Ceacintov, N. (2016), Base and Nucleotide Excision Repair of Oxidatively Generated Guanine Lesions in DNA, J Biol Chem, 291:5309-5319.

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.

Sharma, V., Collins, L., Chen, T., Herr, N., Takeda, S., Sun, W., Swenberg, J., Nakamura, J. (2016), Oxidative stress at low levels can induce clustered DNA lesions leading to NHEJ mediated mutations, Oncotarget, 7:25377-25390.

Sokhansanj, B., Wilson III, D. (2004), Oxidative DNA damage background estimated by a system model of base excision repair, Free Rad Biol Med, 37:433-427.

Swenberg, J., Lu, K., Moeller, B., Gao, L., Upton, P., Nakamura, J., Starr, T. (2011), Endogenous versus Exogenous DNA Adducts: Their Role in Carcinogenesis, Epidemiology, and Risk Assessment, Toxicol Sci, 120:S130-S145.

Taggart, D., Fredrickson, S., Gadkari, V., Suo, Z. (2014), Mutagenic Potential of 8-Oxo-7,8-dihydro-2′-deoxyguanosine Bypass Catalyzed by Human Y-Family DNA Polymerases, Chem Res Toxicol, 27:931-940.

Wang, C., Yang, J., Lu, D., Fan, Y., Zhao, M., Li, Z. (2016), Oxidative stress-related DNA damage and homologous recombination repairing induced by N,N-dimethylformamide , J Appl Toxicol, 36:936-945.

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.

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.

 

Relationship: 1910: N/A, Inadequate DNA repair leads to Increase, DNA strand breaks

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

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

Key Event Relationship Description

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).

Evidence Supporting this KER

Biological Plausibility

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).

Empirical Evidence

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

• Concentration concordance of strand breaks in repair-deficient and –proficient cells (insufficient repair) (Wu et al., 2008)
  • 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.
  • 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).
  • 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.
  • Thus, this evidence supports the essentiality of inadequate DNA repair as a modulator of the downstream KE.
• Incomplete OGG1-initiated base excision repair (BER) leads to DNA strand breaks (Wang et al., 2018):
  • 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)
    • Demonstrates insufficient removal of 8-oxo-dG in OGG1-deficient cells
  • Significantly 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
    • 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)
  • 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)

 

Inadequate repair of alkylated DNA

• Interference of N-methylpurine DNA glycosylase (MPG)-initiated BER by replication leading to strand breaks (Ensminger et al., 2014)
  • 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).
  • 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).
  • MPG-depleted cells in S-phase showed no significant increase in γH2AX foci, while the control cells showed significant MMS concentration-dependent increases.
  • 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). 

 

Inadequate mismatch repair

• Incomplete/incorrect mismatch repair (MMR) leads to DNA strand breaks (Peterson-Roth et al., 2005):
  • 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
  • At 6 and 12 hours, MLH1+ cells had higher percentage of γH2AX foci than MLH1- cells
  • 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

 

Inadequate Repair of DSBs

• 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).
  • The cells were allowed to recover for 16 hours following irradiation.
  • Unrepaired DSBs were measured after recovery using PFGE.
  • There was a dose-dependent increase in unrepaired DSBs due to both ion exposures.
  • Increase in persistent unrepaired DSBs with increasing dosage indicates exceeded repair capacity.
• 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).
  • DNA ligase IV deficiency results in impaired NHEJ
  • DSB repair was monitored using PFGE by measuring the % of DSBs remaining after 0.25, 2, and 24 h following irradiation.
  • 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.
  • 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.
  • The above demonstrates defective DNA repair leading to persistent DSBs.

 

Uncertainties and Inconsistencies

• 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.
• 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. 
• 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). 
• 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).

References

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.

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.

 

 

Relationship: 1911: Increase, DNA strand breaks leads to N/A, Inadequate DNA repair

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

The domain of applicability is multicellular eukaryotes (Lieber, 2008; Hartlerode & Scully, 2009) , plants (Gorbunova, 1997; Puchta, 2005), certain strains of bacteria such as Mycobacteria, Pseudomonas, Bacillus and Agrobacterium (Shuman & Glickman, 2007), and yeast (Wilson & Lieber, 1999).

Key Event Relationship Description

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). 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. 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). Both repair mechanisms are error‐prone, meaning 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).

Evidence Supporting this KER

Biological Plausibility

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 which focussed particularly on DSBs induced by ionizing radiation and extensively detailed 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 (van Gent et al., 2001; Khanna & Jackson, 2001; Vignard et al., 2013; Moore et al., 2014; Rothkamm et al., 2015; Chang et al., 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.

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).

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).  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), 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).

Empirical Evidence

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, here (click link). 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 (Dikomey & Brammer, 2000; Kuhne et al., 2000; Lobrich et al., 2000; Rothkamm & Lo, 2003; Kuhne et al., 2005; Asaithamby & Chen, 2009; Bracalente et al., 2013).

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 (Dikomey & Brammer, 2000; Kuhne et al., 2000; Lobrich et al., 2000; Rothkamm & Lo, 2003; Kuhne et al., 2005; Asaithamby & Chen, 2009; Bracalente et al., 2013), there were resulting dose-dependent increases in non-repaired DSBs (Dikomey & Brammer, 2000), DSB misrepair rates (Mcmahon et al., 2016), and misrejoined DSBs (Kuhne et al., 2000; Kuhne et al., 2005; Rydberg et al., 2005), as well as a dose-dependent decrease in the total DSB rejoining (Lobrich et al., 2000). Furthermore, only 50% of the rejoined DSBs were found to be correctly repaired (Kuhne et al., 2000; Lobrich 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 & Lo, 2003; Pinto & Prise, 2005; Asaithamby & Chen, 2009). 

 

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 (Rothkamm and Lo, 2003; Bracalente et al., 2013; Mcmahon et al., 2016; Dong et al., 2017), and there was an increase in incorrectly rejoined DSBs (Lobrich 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 focussing on the consequences of losing some of these proteins in cells, mice and humans (Van Gent et al., 2001)

Uncertainties and Inconsistencies

Uncertainties and inconsistencies in this KER are as follows:

1. 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).
2. 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).
3. 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.
4.  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).  

Quantitative Understanding of the Linkage

Quantitative understanding of this linkage suggests that DSB repair can be predicted from the presence of DSBs. 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 phosphorylation of gamma-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).

Response-response relationship

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 (Lobrich 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).

Time-scale

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).

Known modulating factors

Not identified.

Known Feedforward/Feedback loops influencing this KER

Not identified.

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Relationship: 164: N/A, Inadequate DNA repair leads to Increase, Mutations

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

The domain of applicability is multicellular eukaryotes (Lieber, 2008; Hartlerode & Scully, 2009), plants (Gorbunova, 1997; Puchta, 2005), certain strains of bacteria such as Mycobacteria, Pseudomonas, Bacillus and Agrobacterium (Shuman & Glickman, 2007), and yeast (Wilson & Lieber, 1999).

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, 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.

Key Event Relationship Description

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.

Evidence Supporting this KER

Biological Plausibility

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 AGT 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).       

Empirical Evidence

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 AGT/MGMT (repair machinery – i.e., decrease in KE1) 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 hypothesized 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

• AS52 Chinese hamster ovary cells (wild type and OGG1-overexpressing) were exposed to kJ/m² UVA radiation (Dahle et al., 2008).
  • 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
    • G:C-A:T mutations in UVA-irradiated OGG1+ cells were completely eliminated
    • 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
    • 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)
• 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).
  • XPA is a key protein in nucleotide excision repair (NER) that acts as a scaffold in the assembly the repair complex.
  • Mutation frequency was determined by the number of TK-revertant colonies
  • Control vector induced a mutation frequency of 1.3% in both WT and XPA KO
  • 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
  • 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 
  • The lack of a key factor in NER leading to increased 8-oxo-dG-induced transversions demonstrates insufficient repair leading to increase in mutations 

 

Inadequate repair of oxidative DNA lesions: In vivo studies in mice

• Spontaneous mutation frequencies in the liver of Ogg1-deficient (-/-) Big Blue mice was measured at 10 weeks of age (Klungland et al., 1999).
  • Mutation frequencies were 2- to 3-fold higher in the Ogg1-/- mice than in wild type
  • 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
  • The results support that insufficient repair of oxidized bases leads to mutation.
• 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).
  • In these mice, there was an average of 2.3-fold increase in mutation frequencies in the liver (measured between 16-20 weeks)
    • 57% of the observed base substitutions were G:C-A:T transversions, while 35% in wild type mice corresponded to this transversion.
    • Approximately 70% of the increase in mutation frequency was due to G to T transversions.
  • 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).
    • G:C-A:T transversions made up 50% of the base substitutions in the Ogg1-/- mice.
  • 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).
    • This observation was consistent even after KBrO3 treatment of the mice (Arai et al., 2002).
  • 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.

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, here (click link). 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. 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. 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

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 (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 XPC status. Specifically, mutation frequencies were increased in XPC-wild-type patients at 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).

Uncertainties and Inconsistencies

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

• 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.
• 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.

Overall

• 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).

Quantitative Understanding of the Linkage

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 table below.

Reference	Summary
Matuo et al., 2018	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).
Nagashima et al., 2018	Hamster cells (GM06318-10) exposed to x-rays in the 0-1 Gy. Response of 19.0 ± 6.1 mutants per 109 survivors.
Albertini et al., 1997	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).
Dubrova et al., 2002	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).
McMahon et al., 2016	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.

 

Response-response relationship

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).

Time-scale

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).

Known modulating factors

Not identified.

Known Feedforward/Feedback loops influencing this KER

Not identified.

References

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Relationship: 1912: N/A, Inadequate DNA repair leads to Increase, Chromosomal aberrations

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

The domain of applicability for this KER is multicellular eukaryotes at any stage of development, including plants (Varga & Aplan 2005; Schipler & Iliakis 2013; Manova & Gruszka 2015).

Key Event Relationship Description

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 a 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 a 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). 

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). 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 (CTAs) that affect both sister chromatids, and chromatid-type aberrations (CSAs), affecting only one chromatid (Danford, 2012) . Examples of CTAs include chromosome-type breaks, centric ring chromosomes, and dicentric chromosomes (which have two centromeres), while CSAs 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). 

Evidence Supporting this KER

Biological Plausibility

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-PK_cs), 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 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). 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 centred 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. 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 (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 and a translocated chromosome (Ferguson and Alt 2001; Povirk 2006). NHEJ has been shown to play a significant role in the generation of translocations ( 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 translocations (Simsek and Jasin 2010) , which may be due to its relatively rapid repair kinetics (Schipler and Iliakis 2013). Translocations 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, dicentric chromosomes 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).

Empirical Evidence

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, here (click link). 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

There is strong evidence for 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 fortifying 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) .

 

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). Lastly, 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 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 and Inconsistencies

Uncertainties in this KER are as follows:

1. 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).  
2. 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).

Quantitative Understanding of the Linkage

Quantitative understanding of this linkage is lacking. Most data is derived from the 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.

Response-response 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 assess the quantitative response-response relationship between inadequate DNA repair and CAs. 

Time-scale

The time scale between inadequate DNA repair and the increased frequency of CAs has not been well-established. Most data comes 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.

Known modulating factors

Not identified.

Known Feedforward/Feedback loops influencing this KER

Not identified.

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Relationship: 1913: Increase, Oxidative DNA damage leads to Increase, DNA strand breaks

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

DNA in any cell type is susceptible to oxidative damage due to endogenous (e.g., aerobic respiration) and exogenous (i.e., exposure to oxidants) oxidative insults.

 

Key Event Relationship Description

The repair of oxidative DNA lesions produced by exposure to reactive oxygen species (ROS) involves excision repair, where damaged base is removed by glycosylases, a strand break is generated 5’ to the apurinic/apyrimidinic (AP) site by lyases and endonucleases, and finally, a new strand is synthesized across the break. Although these strand breaks are mostly transient under normal conditions, elevated levels of oxidative DNA lesions can increase the early AP lyase activities generating a higher number of SSBs that can be more persistent (Yang et al., 2004; Yang et al., 2006). These SSBs can exacerbate the DNA damage by interfering with the replication fork causing it to collapse, and ultimately becoming double strand breaks (DSBs).      

Evidence Supporting this KER

Biological Plausibility

The mechanism of repair of oxidative DNA damage in humans is well-established and numerous literature reviews are available on this topic (Berquist and Wilson III, 2012; Cadet and Wagner, 2013). Oxidative DNA damage is mostly repaired via base excision repair (BER) and via nucleotide excision repair (NER) to a lesser extent. With an increase in oxidative DNA lesions, the more glycosylase and lyase activities occur, introducing SSBs at a higher rate than at homeostasis. It is highly plausible that an increase SSBs also increases the risk for DSBs, which are more difficult to repair accurately. Previous studies have reported thresholded dose-response curves in oxidative DNA damage and attributed these observations to failed repair at the inflection point on the curve, thus allowing strand breaks to accumulate (Gagne et al., 2012; Seager et al., 2012).

Empirical Evidence

The studies collected frequently address both dose and temporal concordance within a single study. Thus, we have not split out these types of empirical data by sub-headings. Instead, we indicate what evidence is available both in vitro and in vivo.

In vitro studies

• Concentration concordance in the formation of oxidative DNA lesions and strand breaks in  HepG2 cells treated with nodularin (ROS-inducing substance (Bouaicha and Maatouk, 2004)) (Lankoff et al., 2006):
  • A concentration-dependent increase in oxidative lesions and strand breaks was observed after 6, 12, and 24h of treatment using Fpg-modified and regular comet assays, respectively.
    • At 6h, the increase in oxidative lesions was significant at 2.5, 5, and 10 µg/mL, while the increase strand breaks was significant at 5 and 10 µg/mL.
    • At 12 and 24 h, the increase in lesions was significant from 1 µg/mL and above, while significant increase in strand breaks occurred from 2.5 µg/mL and above.  
  • At all time points, significant increase in oxidative DNA lesions occurred at a lower concentration than DNA strand breaks.
  • These results demonstrate the concentration concordance in the formation of oxidative DNA lesions and DNA strand breaks.
• Concentration and temporal concordance in human glioblastoma LN-229 cells treated with artesunate, a ROS inducing agent (Berdelle et al., 2011).
  • Concentration and time dependent increases in oxidative lesions were observed using the +Fpg comet test and immunofluorescence staining of 8-oxo-dG.
    • Significant increases in oxidative lesions were observed in cells treated with 25 µg/ml after 6 and 24 hours of treatment, but not 2 and 4 hours, using the + Fpg comet. No increases were observed using -Fpg comet.
    • Concentration-dependent increases in oxidative lesions were observed at the 24 hour timepoint using the +Fpg comet (50 and 75 µg/ml).
    • Oxidative lesions were also measured using immunofluorescence staining of 8-OxodG. Significant increases in oxidative lesions were observed at 6 and 8 hours of continuous treatment with 15 ug/ml artesunate, but not 1 and 4 hours.
      • Upon removal of test chemical, 8-OxodG levels decreased, returning to negative control level after 6 hours.
  • Significant increases in strand breaks as measured by ɣH2AX were observed 2 and 10 hours after treatment (15 µg/ml).
• Deferme et al. (2013) exposed HepG2 cells to 100 µM menadione, 200 µM tert butylhydroperoxide, and 50 µM hydrogen peroxide for increasing durations (30 min, 1, 2, 4, 6, 8, 24 h). The temporal profiles of strand breaks and oxidative lesions were analyzed. The results shown below demonstrate incidence and temporal concordance in oxidative lesion formation and strand breaks (Deferme et al., 2013).
  • Strand breaks were measured by alkaline comet assay.
  • Oxidative DNA lesions were measured by Fpg-modified comet assay
  • Menadione: strand breaks and oxidative lesions increased in a time-dependent manner from 30 min to 4h, when both reached their maximum. The tail moment values of fpg-digested comets were significantly higher than those of no-fpg comets at 1, 2, and 4h, indicating that the induction of oxidative lesions was significant at these time points. After 4h, both strand breaks and oxidative lesions gradually decreased.
  • Tert butylhydroperoxide: From 30 min to 1h, both strand breaks and oxidative lesions increased and gradually decreased from 2 to 24h. Oxidative lesion induction was significant at both 30min and 1h.
  • Hydrogen peroxide: The highest amount of strand breaks and oxidative lesions occurred at 30 min. From 1h onward, the levels of both decreased. Notably, the induction of oxidative lesions was significant at 30min and also at 1h, despite the decrease from 30min.

In vivo studies

• (Trouiller et al., 2009)Concentration concordance in Wistar rats orally exposed to ochratoxin A (OTA) and fumonisin B1 (FB1), ROS inducing agents (Domijan et al., 2006).
  • Kidney cells of male Wistar rats were examined using the comet assay +/- Fpg after oral exposure to OTA for 15 days (5ng, 0.05 mg, 0.5 mg/kg b.w.) or FB1 for 5 days (200 ng, 0.05 mg, 0.5 mg/kg b.w.).
    • Significant increases in oxidative lesions were observed using +Fpg comet at all concentrations tested of both OTA and FB1
    • Significant increases were observed in strand breaks using the standard comet assay at all concentrations of both OTA and FB1.
• Concentration concordance in mice exposed to ROS inducing titanium dioxide nanoparticles (Trouiller et al., 2009).
  • Mice were exposed to 50, 100, 250 or 500 mg/kg of 21 nm P25 TiO2 particles via drinking water for 5 days.
    • A significant increase in 8-oxodG in liver was observed at the 500 mg/kg concentration as measured by high performance liquid chromatography (other concentrations were not examined).
    • Significant increases in gamma-H2AX positive bone marrow cells were observed at all concentrations tested.
    • A significant increase in micronuclei in peripheral blood erythrocytes was observed only at the top concentration tested.

Uncertainties and Inconsistencies

As demonstrated by the Domijan et al paper, results can be complicated by mixed MOA’s. The comet results were positive with and without Fpg suggesting oxidative stress is not the only mechanism.

References

Berdelle, N., Nikolova, T., Quiros, S., Efferth, T., Kaina, B. (2011), Artesunate Induces Oxidative DNA Damage, Sustained DNA Double-Strand Breaks, and the ATM/ATR Damage Response in Cancer Cells, Mol Cancer Ther, 10:2224-2233.

Berquist, B., Wilson III, D. (2012), Pathways for Repairing and Tolerating the Spectrum of Oxidative DNA Lesions, Cancer Lett, 327:61-72.

Bouaicha, N., Maatouk, I. (2004), Microcystin-LR and nodularin induce intracellular glutathione alteration, reactive oxygene species production and lipid peroxidation in primary cultured rat hepatocytes, Toxicol Lett, 148:53-63.

Cadet, J., Wagner, J.R. (2013), DNA Base Damage by Reactive Oxygen Species, Oxidizing Agents, and UV Radiation, Cold Spring Harb Perspect Biol, 5:a012559.

Deferme, L., Briede, J.J., Claessen, S.M., Jennen, D.G., Cavill, R., Kleinjans, J.C. (2013), Time series analysis of oxidative stress response patterns in HepG2: A toxicogenomics approach  , Toxicol, 306:24-34.

Domijan, A., Zeljezic, D., Kopjar, D., Peraica, M. (2006), Standard and Fpg-modified comet assay in kidney cells of ochratoxin A- and fumonisin B(1)-treated rats, Toxciol, 222:53-59.

Gagne, J., Rouleau, M., Poirier, G. (2012), PARP-1 Activation— Bringing the Pieces Together, Science, 336:678-279.

Lankoff, A., Wojcik, A., Fessard, V., Meriluoto, J. (2006), Nodularin-induced genotoxicity following oxidative DNA damage and aneuploidy in HepG2 cells, Toxicol Lett, 164:239-248.

Seager, A., Shah, U., Mikhail, J., Nelson, B., Marquis, B., Doak, S., Johnson, G., Griffiths, S., Carmichael, P., Scott, S., Scott, A., Jenkins, G. (2012), Pro-oxidant Induced DNA Damage in Human Lymphoblastoid Cells: Homeostatic Mechanisms of Genotoxic Tolerance, Toxicol Sci, 128:387-397.

Spassova, M., Miller, D., Nikolov, A. (2015), Kinetic Modeling Reveals the Roles of Reactive Oxygen Species Scavenging and DNA Repair Processes in Shaping the Dose-Response Curve of KBrO3-Induced DNA Damage, Oxid Med Cell Longev, 2015:764375.

Trouiller, B., Reliene, R., Westbrook, A., Solaimani, P., Schiestl, R. (2009), Titanium dioxide nanoparticles induce DNA damage and genetic instability in vivo in mice, Cancer Res, 69:8784-8789.

Yang, N., Chaudry, A., Wallace, S. (2006), Base excision repair by hNTH1 and hOGG1: A two edged sword in the processing of DNA damage in gamma-irradiated human cells, DNA Repair, 5:43-51.

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.

Relationship: 1914: Increase, Oxidative DNA damage leads to Increase, Mutations

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

DNA in any cell type is susceptible to oxidative damage due to endogenous (e.g., aerobic respiration) and exogenous (i.e., exposure to oxidants) oxidative insults. Resulting increase in mutation frequency has been described in both eukaryotic and prokaryotic cells.

Key Event Relationship Description

Oxidative DNA lesions such as 7, 8-dihydro-8oxo-deoxyGuanine (8-oxo-dG) and 2,6-diamino-4-hydroxy-5-formamidopyrimidine (FaPydG) are mutagenic because they are able to form base pairs with dATP instead dCTP during replication. This can lead to permanent changes in the DNA sequence that is inherited by daughter cells with subsequent replication. G:C→T:A transversions are the most abundant base substitution attributed to oxidative DNA lesions (Cadet and Wagner, 2013).

Evidence Supporting this KER

Biological Plausibility

Mutagenicity of oxidative DNA lesions has been extensively studied; incorrect base insertion opposite unrepaired oxidative DNA lesions during replication is a well-established event.

For example, 8-oxo-dG and FaPydG, the two most prominent oxidative DNA lesions, are able to form base pairs with dATP, giving rise to G:C→T:A transversions with subsequent DNA synthesis (Gehrke et al., 2013; Freudenthal et al., 2013; Markkanen, 2017). Replicative DNA polymerases such as DNA polymerase α, δ, and ε (pol α, δ, ε) have a poor ability to extend the DNA strand past 8-oxo-dG:dCTP base pairs and may cause replication to stall or incorrectly insert dATP opposite 8-oxo-dG (Hashimoto et al., 2004; Markkanen et al., 2012). In stalled replication forks, repair polymerases may be recruited to perform translesion DNA synthesis (TLS). Human Y-family DNA polymerases (Rev 1, pol κ, ι, and η) are DNA repair polymerases mainly involved in TLS for stalled replication forks. However, TLS is not free of error and its accuracy differs for each repair polymerase. For example, it is known that pol κ and η perform TLS across 8-oxo-dG and often insert dATP opposite the lesion, generating G:C→T:A transversions. The error-prone nature of bypassing unrepaired oxidative lesions has been described in many previous studies and reviews (Greenberg, 2012; Maddukuri et al., 2014; Taggart et al., 2014; Sha et al., 2017).

Empirical Evidence

In vitro studies

• Concentration-dependent increase in oxidative lesions observed in TK6 human lymphoblastoid cells exposed to KBrO₃ and glucose oxidase (GOx; enzyme that produces H₂O₂) for 1 hour; increase in mutation frequency measured in TK assay after 14 days (3 days in non-selective medium and 11 days in selective medium) following 1 hour exposure corresponded with the concentration-response observed in oxidative lesions (Platel et al., 2011).
  • NOGEL could be determined in TK assay (KBrO3: 1.75 mM; bleomycin: 0.6µM; GOx: 1.17x10-5 units/mL) but not in the Fpg-modified comet assays (First statistically significant concentrations: KBrO3: 1 mM; bleomycin: 0.5µM; GOx: 1.08x10-5 units/mL)
    • These results indicate that statistically significant increases in oxidative lesions (measured in Fpg comet assay) occur at lower concentrations of the above three stressors than mutations measured by the Tk gene mutation assay at a later time point (after 14-day recovery)
    • Demonstrates concentration concordance in oxidative DNA lesions and mutation

In vivo studies

• Klungland et al. (1999) measured and compared the level of 8-oxodG in the liver of OGG1-null Big Blue mice and Ogg1+/+ Blue Blue mice at 13-15 weeks of age. (Klungland et al., 1999).
  • The amount of 8-oxodG in the OGG1-null mice was 1.7-fold higher than in wild-type mice at the time of measurement.
  • Spontaneous mutation frequencies in the liver of OGG1 null (Ogg1 -/-) Big Blue mice and wild type (Ogg1 +/+) Big Blue mice were measured at 10 and 20 weeks of age:
• At 10 weeks, mutation frequency increased by 2- to 3-fold in OGG1 -/- mice compared to the wild type mice. No further increase was observed at 20 weeks.
• Of the 16 base substitutions detected in Ogg1 -/- mutant plaques analyzed by sequencing, 10 indicated G→T transversions.
• This study demonstrates that increased levels of oxidative DNA damage in the null mice was concordant with increased incidence of mutations.
• Unfried et al. (2002) measured the level of 8-oxodG and mutations in the omenta of rats exposed to crocidolite for various durations (Unfried et al., 2002).
  • Statistically significant increases in 8-oxodG were observed compared to control after 10 and 20 weeks of exposure.
  • The number of G→T transversions after 4, 12, and 24 weeks of exposure was significantly higher compared to control and G→T transversions were the most prominent base substitution in these samples.
  • This mutation spectrum supports that oxidative DNA lesions were the source of mutations.
• Five-week-old male gpt delta mice were given drinking water containing 85 ppm sodium arsenite for 3 weeks and sacrificed 2 weeks after administration was stopped (Takumi et al., 2014).
  • The gpt mutation assay and 8-OHdG quantification was performed using genomic DNA isolated from the liver
  •  Significantly higher levels of 8-OHdG were observed in the arsenite group (1.15/10⁶ dG) compared to the control group (0.86/10⁶ dG)
  •  Elevated mutation frequency was observed in the arsenite group with an average of 1.10 x10^-5, compared to that of the control group (0.71x10^-5 )
  • G:C→T:A made up 46% of the mutations in the arsenite-fed mice
  • These data demonstrating a positive correlation between incidence of oxidative lesions in DNA and elevation in mutation frequency support that these events are associated, and the mutation spectrum further suggest that mutations were the result of oxidative lesions.

Uncertainties and Inconsistencies

The provided empirical evidence examined only the quantities of 8-oxo-dG and related the observed mutations to this oxidative lesion; the level of overall DNA oxidation is inferred from the level of 8-oxo-dG present. It is unclear how other oxidative DNA lesions such as FapyG, FapyA, and thymidine glycol contribute to the mutation spectra and frequencies.

References

Cadet, J., Wagner, J.R. (2013), DNA Base Damage by Reactive Oxygen Species, Oxidizing Agents, and UV Radiation, Cold Spring Harb Perspect Biol, 5:a012559.

Freudenthal, B., Beard, W., Wilson, S. (2013), DNA polymerase minor groove interactions modulate mutagenic bypass of a templating 8-oxoguanine lesion., Nucleic Acids Res, 41:1848-1858.

Gehrke, T., Lischke, U., Gasteiger, K., Schneider, S., Arnold, S., Muller, H., Stephenson, D., Zipse, H., Carell, T. (2013), Unexpected non-Hoogsteen–based mutagenicity mechanism of FaPy-DNA lesions, Nat Chem Biol, 9:455-461.

Greenberg, M. (2012), Purine Lesions Formed in Competition With 8-Oxopurines From Oxidative Stress, Acc Chem Res, 45:588-597.

Hashimoto, K., Tominaga, Y., Nakabeppu, Y., Moriya, M. (2004), Futile short-patch DNA base excision repair of adenine:8-oxoguanine mispair, Nucleic Acids Res, 32:5928-5934.

Klungland, A., Rosewell, I., Hollenbach, S., Larsen, E., Daly, G., Epe, B., Seeberg, E., Lindahl, T., Barnes, D. (1999), Accumulation of premutagenic DNA lesions in mice defective in removal of oxidative base damage, Proc Natl Acad Sci USA, 96:13300-13305.

Maddukuri, L., Ketkar, A., Eddy, S., Zafar, M., Eoff, R. (2014), The Werner syndrome protein limits the error-prone 8-oxo-dG lesion bypass activity of human DNA polymerase kappa, Nucleic Acids Res, 42:12027-12040.

Markkanen, E. (2017), Not breathing is not an option: How to deal with oxidative DNA damage, DNA Repair, 59:82-105.

Markkanen, E., Castrec, B., Vilani, G., Hubscher, U. (2012), A switch between DNA polymerases δ and λ promotes error-free bypass of 8-oxo-G lesions, Proc Natl Acad Sci USA, 27:931-940.

Platel, A., Nesslany, F., Gervais, V., Claude, N., Marzin, D. (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.

Sha, Y., Minko, I., Malik, C., Rizzo, C., Lloyd, R.S. (2017), Error-Prone Replication Bypass of the Imidazole Ring-Opened Formamidopyrimidine Deoxyguanosine Adduct, Envrion Mol Mutatgen, 58:182-189.

Taggart, D., Fredrickson, S., Gadkari, V., Suo, Z. (2014), Mutagenic Potential of 8-Oxo-7,8-dihydro-2′-deoxyguanosine Bypass Catalyzed by Human Y-Family DNA Polymerases, Chem Res Toxicol, 27:931-940.

Takumi, S., Aoki, Y., Sano, T., Suzuki, T., Nohmi, T., Nohara, K. (2014), In vivo mutagenicity of arsenite in the livers of gpt delta transgenic mice  , Mutat Res, 760:42-47.

Unfried, K., Schurkes, C., Abel, J. (2002), Distinct Spectrum of Mutations Induced by Crocidolite Asbestos: Clue for 8-Hydroxydeoxyguanosine-dependent Mutagenesis in Vivo, Cancer Res, 62:104.

Relationship: 1931: Increase, DNA strand breaks leads to Increase, Mutations

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

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.

Key Event Relationship Description

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.

Evidence Supporting this KER

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.

Biological Plausibility

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).

                                                                                                                            

Empirical Evidence

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

• 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).
  • Concentration-dependent increases in strand breaks were measured using the alkaline comet assay.
  • At the same concentrations, mutation frequencies measured by TK gene mutation assay also showed a concentration-dependent increasing trend.
  • 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).

 

• 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).
  • Luan et al. treated TK6 cells with KBrO₃ for 4 hours and performed alkaline comet assay to measure strand breaks.
  • 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.
  • 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)
  • In both comet assay and Tk mutation assay, concentration-dependent increase in response was observed.
  • These results demonstrate the occurrence of DNA strand breaks followed by increase in mutations.

 

• Indirect measurement of mutations by measuring misrejoined DSBs in vitro
  • 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.
  • DSBs were measured by pulsed field gel electrophoresis (PFGE)
    • Dose-dependent increase in DSBs was observed immediately following irradiation.
  • Misrejoining of ends was monitored using the Hybridization assay:
    • DNA is digested using a restriction enzyme and fractionated by PFGE.
    • ³²P-labeled probe for a 3.2-Mbp NotI restriction fragment is then used in Southern blotting to detect intact restriction fragments.
    • Failure to reconstitute the restriction fragment indicates incorrect joining of ends following DSBs and altered DNA sequence.
  • After 16 h of recovery following irradiation, Rydberg et al. observed a radiation dose-dependent increase in misrejoined DSBs in all four treatment groups.
  • A similar study by Kuhne et al. (2005) reported concordant results (Kuhne et al., 2005):
    • Subsequently, there was a dose-dependent increase in misrejoined DSBs 24h post irradiation.
    • Increasing doses of X-rays and γ rays immediately induced DSBs in primary human fibroblasts in a dose-dependent manner.
    • 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).
    • These results demonstrate the concentration and temporal concordance in strand breaks leading to mutations.

 

• 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).
  • 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.
  • 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.
  • 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.

 

• 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).
  • 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.
  • 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).
  • 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.

 

• 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).
  • 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.
  • After being irradiated with 80 Gy of X-rays, approximately 50% of the DSBs were correctly rejoined, as measured by the hybridization assay.
  • 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.
  • 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.

In vivo studies

• 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).
  • 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.
  • Mutations in the leaves were measured when the seedling plants put out their 6^th or 7^th 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.
  • These results demonstrate a dose concordance between DNA strands breaks and mutation frequency, and suggest a time concordance.

Uncertainties and Inconsistencies

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). 

References

Bunting, S., Nussenzweig, A. (2013), End-joining, Translocations and Cancer, Nat Rev Cancer, 13:443-454.

Byrne, M., Wray, J., Reinert, B., Wu, Y., Nickoloff, J., Lee, S.H., Hromas, R., Williamson, E. (2014), Mechanisms of oncogenic chromosomal translocations, Ann NY Acad Sci, 1310:89-97.

Dikomey, E., Brammer, I. (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., Haber, J. (2018), Assaying Mutations Associated With Gene Conversion Repair of a Double-Strand Break  , Methods Enzymiol, 601:145-160.

Hicks, W., Haber, J. (2010), Increased Mutagenesis and Unique Mutation Signature Associated with Mitotic Gene Conversion  , Nat Rev Cancer, 329:82-84.

Kuhne, M., Rothkamm, K., Lobrich, M. (2000), No dose-dependence of DNA double-strand break misrejoining following a-particle irradiation, Int J Radiat Biol, 76:891-900.

Kuhne, M., Urban, G., Frankenberg, D., Lobrich, M. (2005), DNA double-strand break misrejoining after exposure of primary human fibroblasts to CK characteristic X rays, 29 kVp X rays and 60Co gamma rays, Radiat Res, 164:669-676.

Lobrich, M., Kuhne, M., Wetzel, J., Rothkamm, K. (2000), Joining of Correct and Incorrect DNA Double-Strand Break Ends in Normal Human and Ataxia Telangiectasia Fibroblasts, Genes Chromosomes Cancer, 27:59-68.

Mao, Z., Bozzella, M., Seluanov, A., Gorbunova, V. (2008a), Comparison of nonhomologous end joining and homologous recombination in human cells, DNA Repair, 7:1765-1771.

Mao, Z., Bozzella, M., Seluanov, A., Gorbunova, V. (2008b), DNA repair by nonhomologous end joining and homologous recombination during cell cycle in human cells, Cell Cycle, 7:2902-2906.

McMahon, S., Schuemann, J., Paganetti, H., Prise, K. (2016), Mechanistic Modelling of DNA Repair and Cellular Survival Following Radiation-Induced DNA Damage, Sci Rep, 6:33290.

Platel, A., Nesslany, F., Gervais, V., Claude, N., Marzin, D. (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., Jackson, S. (2011), Dynamics of DNA damage response proteins at DNA breaks: a focus on protein modifications, Genes Dev, 25:409-433.

Ptacek, O., Stavreva, D., Kim, J.K., Gichner, T. (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., McVey, M. (2016), Error-prone repair of DNA double-strand breaks, J Cell Physiol, 231:15-24.

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.

Spassova, M., Miller, D.J., Eastmond, D., Nikolova, N., Vulimiri, S.V., Caldwell, J., Chen, C., White, P.D. (2013), Dose-Response Analysis of Bromate-Induced DNA Damage andMutagenicity Is Consistent With Low-Dose Linear,Nonthreshold Processes, Environ Mol Mutagen, 54:19-35.

 

Relationship: 1939: Increase, DNA strand breaks leads to Increase, Chromosomal aberrations

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

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

Key Event Relationship Description

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.

Evidence Supporting this KER

Biological Plausibility

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 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).

Empirical Evidence

In vitro studies demonstrating dose and temporal concordance

• 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).
  • Concentration-dependent increase in DNA strand breaks was measured using the alkaline comet assay 1 hr post-exposure
    • First statistically significant concentration: bleomycin: 0.5 µM; GOx: 1.08x10^-5 units/mL
    • NOEL could not be defined, indicating that there was response at every tested concentration.
  • MN frequency was measured 23 hours post exposure; concentration-dependent increase in MN frequency was observed and NOEL was identified.
    • NOEL: bleomycin: 0.023 µM; GOx: 1.78x10^-5 units/mL
    • All concentrations above the NOEL induced significant increases in MN frequency.
  • 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.

 

• 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).
  • 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.
  • 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. 
  • These data demonstrate both dose- and temporal concordance in DNA strand breaks observed by comet assay and MN frequency.

 

• 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).
  • Strand breaks were shown in the comet assay in all cell lines tested, immediately after treatment with 1 and 2 Gy.  
  • 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.
  • Thus, the data demonstrate both temporal and dose concordance.

 

•  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).
  • 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.
  • 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.
  • Significant increases in MN frequency was observed 26h post exposure (~1.5 cycles) at concentrations of 0.1µg/ml and above.
  • 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.

 

• 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).
  • 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.
  • A statistically significant increase in MN frequency was observed 24 h after treatment at 5 µg/mL.
  • 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.

 

• 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).
  • DSBs were measured as the number of γH2AX foci immediately following the exposure.
  • 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.
  • Chromosomal aberrations (breaks and translocations) were monitored in metaphase spreads 24h following 1h 1 mM MMS treatment.
  • 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.
  •  The results support that increases in DSBs lead to increases in chromosomal aberrations.

 

• 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.
  • Benchmark dose analyses were conducted to estimate Point of Departure values for MN and gamma-H2AX responses.   
  • 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).
  • Thus, the data demonstrate both temporal and dose concordance for these endpoints.

 

• 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).
  • 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.    
  • Irradiated whole blood samples showed significantly increased strand breaks by 10 cGy, but this level stayed relatively stable from 10 - 50 cGy.
  • 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.
  • The results of this study support dose concordance and are suggestive of time concordance.

 

• 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).
  • 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.
  • 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.
  • 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.
  • 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.

 

• 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.    
  • DSBs were increased in all cells transfected with the endonuclease relative to cells from the same cell lines that underwent a mock transfection.
  • Chromosomal translocations were also elevated in cell lines transfected with an endonuclease, with increasing chromosomal translocations found in cells with increasing DSB cluster damage.
  • This study shows an association between DSB cluster damage and chromosomal translocation incidence.

 

In vivo studies

• 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).
  • Peripheral blood lymphocytes were isolated for comet assay and peripheral blood erythrocytes were used to measure MN at each time point.
    • Different comet assay parameters such as tail length, moment, olive tail moment, and % tail DNA were compared to MN frequency
    • All comet assay parameters had a positive correlation to MN frequency demonstrated in all chemical treatments.
    • 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.
    • 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.
  • The results suggest temporal concordance in strand breaks measured by comet assay and induction of MN, where strand breaks are observed earlier than MN.

 

• 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).
  • γH2AX measured at 24h showed a dose-dependent increase in DSBs in both acute and low dose rate exposed mice.
    • 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.
  • MN frequency was also measured 24h and 7 days post exposure;
    • 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
    • There was no statistical difference in the two treatment groups

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.

Uncertainties and Inconsistencies

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).

• Increases in γH2AX were observed only at 25 µM B[a]P (~2.5 fold increase) after the 24h exposure.
• Translocations were quantified and expressed as the genomic frequency of translocations per 100 cells (F_G/100)
  • 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)

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.

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