Alkylation, DNA

55-18-5

WBNQDOYYEUMPFS-UHFFFAOYSA-N

N-Nitrosodiethylamine

DEN Ethanamine, N-ethyl-N-nitroso- Diethylamine, N-nitroso- Diethylnitrosamide Diethylnitrosoamin Diethylnitrosoamine dietilnitrosoamina N,N-Diethylnitrosoamine N-Ethyl-N-nitrosoethanamine Nitrosodiethylamine N-NITROSODIAETHYLAMIN N-Nitroso-N,N-diethylamine NSC 132 diethylnitrosamine

DTXSID2021028

64-67-5

DENRZWYUOJLTMF-UHFFFAOYSA-N

Diethyl sulfate

Sulfuric acid, diethyl ester DIAETHYLSULFAT diethyl sulphate Diethylsulfat NSC 56380 Sulfate de diethyle sulfato de dietilo Sulfuric acid diethyl ester UN 1594

DTXSID1024045

62-75-9

UMFJAHHVKNCGLG-UHFFFAOYSA-N

N-Nitrosodimethylamine

DMN Methanamine, N-methyl-N-nitroso- DIMETHYLAMINE, N-NITROSO- Dimethylnitrosoamin Dimethylnitrosoamine dimetilnitrosoamina Nitrosodimethylamine Nitrosodimetilamina N-Methyl-N-nitrosomethanamine N-NITROSODIMETHYLAMIN N-Nitroso-N,N-dimethylamine NSC 23226 Dimethylnitrosamine

DTXSID7021029

77-78-1

VAYGXNSJCAHWJZ-UHFFFAOYSA-N

Dimethyl sulfate

Sulfuric acid, dimethyl ester Dimethyl monosulfate dimethyl sulphate Dimethylsulfat DIMETHYLSULFATE NSC 56194 Sulfate de dimethyle sulfato de dimetilo Sulfuric acid dimethyl ester SULFURIC ACID DIMETHYLESTER UN 1595

DTXSID5024055

62-50-0

PLUBXMRUUVWRLT-UHFFFAOYSA-N

Ethyl methanesulfonate

EMS Methanesulfonic acid, ethyl ester Ethyl mesylate Ethyl methane sulfonate ethyl methanesulphonate Ethylmethansulfonat metanosulfonato de etilo Methanesulfonate d'ethyle Methanesulfonic acid ethyl ester METHANSULFONSAEURE-AETHYLESTER METHYLSULFONATE, ETHYL NSC 26805 O-Ethyl methylsulfonate

DTXSID6025309

759-73-9

FUSGACRLAFQQRL-UHFFFAOYSA-N

1-Ethyl-1-nitrosourea

ENU Urea, N-ethyl-N-nitroso- N-AETHYL-N-NITROSO-HARNSTOFF N-Ethyl-N-nitrosoharnstoff N-ethyl-N-nitrosourea N-Ethyl-N-nitrosouree N-etil-N-nitrosourea N-Nitroso-N-ethylurea NSC 45403 Urea, 1-ethyl-1-nitroso-

DTXSID8020593

63885-23-4

ZGONASGBWOJHDD-UHFFFAOYSA-N

N′-Ethyl-N-nitro-N-nitrosoguanidine

DTXSID3020592

926-06-7

SWWHCQCMVCPLEQ-UHFFFAOYSA-N

Isopropyl methanesulfonate

DTXSID8031497

66-27-3

MBABOKRGFJTBAE-UHFFFAOYSA-N

Methyl methanesulfonate

MMS Methanesulfonic acid, methyl ester metanosulfonato de metilo Methanesulfonate de methyle methyl methanesulphonate Methyl methylsulfonate Methylmethansulfonat METHYLSULFONATE, METHYL NSC 50256

DTXSID7020845

70-25-7

VZUNGTLZRAYYDE-UHFFFAOYSA-N

Methylnitronitrosoguanidine

1-Methyl-3-nitro-1-nitroso-guanidine Guanidine, N-methyl-N'-nitro-N-nitroso- 1-Methyl-1-nitroso-2-nitroguanidine 1-Methyl-1-nitroso-3-nitroguanidine 1-Methyl-3-nitro-1-nitrosoguanidin 1-METHYL-3-NITRO-1-NITROSO-GUANIDIN 1-Methyl-3-nitro-1-nitrosoguanidine 1-metil-3-nitro-1-nitrosoguanidina 1-Nitroso-3-nitro-1-methylguanidine Guanidine, 1-methyl-3-nitro-1-nitroso- Methylnitronitrosoguanidine N-Methyl-N1-nitro-N-nitrosoguanidine N-Methyl-nitroso-N'-nitroguanidine N-Methyl-N'-nitro-N-nitrosoguanadine N-Methyl-N'-nitro-N-nitrosoquanidine N-Methyl-N-nitroso-N'-nitroguanidine N-Nitroso-N-methylnitroguanidine N-Nitroso-N-methyl-N'-nitroguanidine N-Nitroso-N'-nitro-N-methylguanidine NSC 9369

DTXSID2020846

CHEBI:16991

CHEBI deoxyribonucleic acid

D009369

MESH Neoplasms

GO:0006305

GO DNA alkylation

GO:0006281

GO DNA repair

D009154

MESH mutation

WIKI increased

WIKI functional change

Diethyl nitrosamine

2016-11-29T18:42:09

Diethyl sulfate

2016-11-29T18:42:11

Dimethyl nitrosamine

2016-11-29T18:42:11

2016-11-29T21:19:02

Dimethyl sulfate

2016-11-29T18:42:13

Ethyl methanesulfonate

2016-11-29T18:42:14

Ethyl nitrosourea

2016-11-29T18:42:14

Ethyl-N'-nitro-N-nitrosoguanidine

2016-11-29T18:42:15

Isopropyl methanesulfonate

2016-11-29T18:42:17

Methyl methanesulfonate

2016-11-29T18:42:19

Methyl-l-N'-nitro-N-nitroguanidine

2016-11-29T18:42:19

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

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

2019-05-03T12:36:36

2019-05-07T12:12:13

10090

NCBI mouse

10036

NCBI Syrian golden hamster

10116

NCBI rat

9606

NCBI Homo sapiens

9913

NCBI cow

10090

NCBI Mus musculus

8090

NCBI medaka

10116

NCBI Rattus norvegicus

WCS_9606

common toxicological species human

Alkylation, DNA

Molecular

The event involves DNA alkylation to form a variety of different DNA adducts (i.e., alkylated nucleotides). Alkylation occurs at various sites in DNA and can include alkylation of adenine- Nl, - N3, - N7, guanine- N3, - O6, - N7, thymine-O2, - N3, - O4, cytosine- O2, -N3, and the phosphate (diester) group (reviewed in detail in Beranek 1990). In addition, alkylation can involve modification with different sizes of alkylation groups (e.g., methyl, ethyl, propyl). It should be noted that many of these adducts are not stable or are readily repaired (discussed in more detail below). A small proportion of adducts are stable and remain bound to DNA for long periods of time.

There is no OECD guideline for measurement of alkylated DNA, although technologies for their detection are established. Reviews of modern methods to measure DNA adducts include Himmelstein et al,. 2009 and Philips et al., 2000. High performance liquid chromatography (HPLC) methods can be used to measure whether an agent is capable of alkylating DNA in somatic cells. Alkyl adducts in somatic cells can be measured using immunological methods (described in Nehls et al. 1984), as well as HPLC (methods in de Groot et al. 1994) or a combination of 32P post-labeling, HPLC and immunologic detection (Kang et al. 1992). We note that mass spectrometry provides structural specificity and confirmation of the structure of DNA adducts. DNA alkylation can also be measured using a modified comet assay. This method involves the digestion of alkylated DNA bases with 3–methyladenine DNA glycosylase (Collins et al., 2001; Hasplova et al., 2012) followed by the standard comet assay to detect where alkyl adducts occur. The advantage of this method is that the alkaline version of the comet assay, as a core method, has an in vivo OECD guideline. Finally, structure-activity relationships (SARs) have been developed to predict the possibility that a chemical will alkylate DNA (e.g., Vogel and Ashby, 1994; Benigni, 2005; Dai et al., 1989; Lewis and Griffith, 1987). Measurement of alkylation in male germ cells: In rodent testes, studies have detected adducts in situ by immuhistocytological staining. For example, fixed testes are incubated with O6-EtGua -specific mouse monoclonal antibody and subsequently with a labeled anti-mouse IgG F antibody. Nuclear DNA is counterstained with DAPI 4,6-diamidino- 2-phenylindole. Fluorescence signals from immunostained O6-EtGua residues in DNA are visualized by fluorescence microscopy and quantitated using an image analysis system. Methods are described in (Seiler et al. 1997). An immunoslot blot assay for detection of O6-EtGua has been described previously in (Mientjes et al. 1996). Alternatively, rodents have also been exposed to radio-labeled alkylating agents. Examples from the literature include [2-3H] ENU, [1-3H]di-ethyl sulfate, or [1-3H]ethyl-methane sulfonate. Following treatment with the labeled chemical, testis and other tissues of interest are removed. Germ cells are released from tubuli by pushing out the contents with forceps. Using this procedure all germ-cell stages are liberated from the tubuli, with the possible exception of part of the population of stem-cell spermatogonia that remain attached to the walls of the tubuli. DNA is then extracted from germ cells, empty testis tubuli and other tissues of interest. DNA adduct formation is determined after neutral and acid hydrolysis of DNA followed by separation of the various ethylation products using HPLC (described in van Zeeland et al. 1990).

Alkylated DNA has been measured in somatic cells in a variety of species, from prokaryotic organisms, to rodents in vivo, to human cells in culture. Theoretically, DNA alkylation can occur in any cell type in any organism.

CL:0000255

CL eukaryotic cell

High Mixed High High High High

Benigni, R. (2005), "Structure-activity relationship studies of chemical mutagens and carcinogens: mechanistic investigations and prediction and approaches", Chem. Rev., 105: 1767-1800. Beranek, D.T. (1990), "Distribution of methyl and ethyl adducts following alkylation with monofunctional alkylating agents", Mutation Res., 231: 11-30. Collins, A.R., M. Dusinská and A. Horská (2001), "A Detection of alkylation damage in human lymphocyte DNA with the comet assay". Acta Biochim Pol., 48: 611-4. Dai, Q.H. and R.G. Zhong (1989), "Quantitative pattern recognition for structure-carcinogenic activity relationship of N-nitroso compounds based upon Di-region theory", Sci China B., 32:776-790. de Groot, A.J., J.G. Jansen, C.F. van Valkenburg and A.A. van Zeeland (1994), "Molecular dosimetry of 7-alkyl- and O6-alkylguanine in DNA by electrochemical detection", Mutat Res., 307: 61-6. Hašplová, K., A. Hudecová, Z. Magdolénová, M. Bjøras, E. Gálová, E. Miadoková and M. Dušinská (2012), "DNA alkylation lesions and their repair in human cells: modification of the comet assay with 3-methyladenine DNA glycosylase (AlkD)", Toxicol Lett., 208: 76-81. Himmelstein, M.W., P.J. Boogaard, J. Cadet, P.B. Farmer, J.J. Kim, E.A. Martin, R. Persaud and D.E. Shuker (2009), "Creating context for the use of DNA adduct data in cancer risk assessment: II. Overview of methods of identification and quantitation of DNA damage", Crit. Rev. Toxicol., 39: 679-94. Kamino, K., F. Seiler, M. Emura, J. Thomale, M.F. Rajewsky and U. Mohr (1995), "Formation of O6-ethylguanine in spermatogonial DNA of adult Syrian golden hamster by intraperitoneal injection of diethylnitrosamine", Exp. Toxicol. Pathol., 47: 443-445. Kang, H.I., C. Konishi, G. Eberle, M.F. Rajewsky, T. Kuroki and N.H. Huh (1992), "Highly sensitive, specific detection of O6-methylguanine, O4-methylthymine, and O4-ethylthymine by the combination of high-performance liquid chromatography prefractionation, 32P postlabeling, and immunoprecipitation", Cancer Res., 52: 5307-5312. Lewis, D.F. and V.S. Griffiths (1987), "Molecular electrostatic potential energies and methylation of DNA bases: a molecular orbital-generated quantitative structure-activity relationship", Xenobiotica, 17: 769-776. Mientjes, E.J., K. Hochleitner, A. Luiten-Schuite, J.H. van Delft, J. Thomale, F. Berends, M.F. Rajewsky, P.H. Lohman and R.A. Baan (1996), "Formation and persistence of O6-ethylguanine in genomic and transgene DNA in liver and brain of lambda(lacZ) transgenic mice treated with N-ethyl-N-nitrosourea", Carcinogenesis, 17: 2449-2454. Nehls, P., M.F. Rajewsky, E. Spiess, D. Werner (1984), "Highly sensitive sites for guanine-O6 ethylation in rat brain DNA exposed to N-ethyl-N-nitrosourea in vivo", EMBO J., 3:327-332. Phillips, D.H., P.B. Farmer, F.A. Beland, R.G. Nath, M.C. Poirier, M.V. Reddy and K.W. Turteltaub (2000), "Methods of DNA adduct determination and their application to testing compounds for genotoxicity", Environ. Mol. Mutagen., 35: 222-233. Scherer, E., A.A. Jenner and L. den Engelse (1987), "Immunocytochemical studies on the formation and repair of O6-alkylguanine in rat tissues", IARC Sci. Publ., 84: 55-58. Sega, G.A., C.R. Rohrer, H.R. Harvey and A.E. Jetton (1986), "Chemical dosimetry of ethyl nitrosourea in the mouse testis", Mutat. Res., 159: 65-74. Seiler, F., K. Kamino, M. Emura, U. Mohr and J. Thomale (1997), "Formation and persistence of the miscoding DNA alkylation product O6-ethylguanine in male germ cells of the hamster", Mutat. Res., 385: 205-211. van Zeeland, A.A., A. de Groot and A. Neuhäuser-Klaus (1990), "DNA adduct formation in mouse testis by ethylating agents: a comparison with germ-cell mutagenesis", Mutat. Res. 231: 55-62. Vogel, E.W., Ashby, J. (1994), "Structure-activity relationships: experimental approaches." In: Methods to asses DNA Damage and repair: Interspecies comparisons. Edited by R.T. Tardiff, P.H.M. Lohman and G.N. Wogan, SCOPE, Wiley and Sons LTD. AOPWiki 2016-11-29T18:41:22

2017-09-16T10:14:29

Inadequate DNA repair

Cellular

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

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

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

High Unspecific

High

All life stages

High Moderate Moderate High Low

Adimoolam, S. & J.M. Ford (2003), "p53 and regulation of DNA damage recognition during nucleotide excision repair" DNA Repair (Amst), 2(9): 947-54. Allgayer, J. et al. (2013), "Modulation of base excision repair of 8-oxoguanine by the nucleotide sequence", Nucleic Acids Res, 41(18): 8559-8571. Doi: <a href="https://doi.org/10.1093/nar/gkt620" target="_blank">10.1093/nar/gkt620</a>. Beranek, D.T. (1990), "Distribution of methyl and ethyl adducts following alkylation with monofunctional alkylating agents", Mutation Research, 231(1): 11-30. Doi: 10.1016/0027-5107(90)90173-2. Bhatti, A. et al., (2016), “Homologous Recombination Biology.”, Encyclopedia Britannica. Bhowmick, R., S. et al. (2016), "RAD52 Facilitates Mitotic DNA Synthesis Following Replication Stress", Mol Cell, 64:1117-1126. Doi: 10.1016/j.molcel.2016.10.037. Biedermann, A. K. et al. (1991), “SCID mutation in mice confers hypersensitivity to ionizing radiation and a deficiency in DNA double-strand break repair”, Cell Biology, 88(4): 1394-7. Doi: 10.1073/pnas.88.4.1394. Boboila, C., F. W. Alt & B. Schwer. (2012), “Classical and alternative end-joining pathways for repair of lymphocyte-specific and general DNA double-strand breaks.” Adv Immunol, 116, 1-49. doi:10.1016/B978-0-12-394300-2.00001-6 Bronstein, S.M. et al. (1991), "Toxicity, mutagenicity, and mutational spectra of N-ethyl-N-nitrosourea in human cell lines with different DNA repair phenotypes", Cancer Research, 51(19): 5188-5197. Bronstein, S.M. et al. (1992), "Efficient repair of O6-ethylguanine, but not O4-ethylthymine or O2-ethylthymine, is dependent upon O6-alkylguanine-DNA alkyltransferase and nucleotide excision repair activities in human cells", Cancer Research, 52(7): 2008-2011.  Brown, J.A. et al. (2011), "Efficiency and fidelity of human DNA polymerases λ and β during gap-filling DNA synthesis", DNA Repair (Amst)., 10(1):24-33. Butterworth, E. B. et al., (1987), A protocol and guide for the in vitro rat hepatocyte DNA-repair assay. Mutation Research. 189, 113-21. Doi: 10.1016/0165-1218(87)90017-6. Caldecott, K. W. (2014), "DNA single-strand break repair", Exp Cell Res, 329(1): 2-8. Chen, L. et al., (2001), Promotion of DNA ligase IV-catalyzed DNA end-joining by the Rad50/Mre11/Xrs2 and Hdf1/Hdf2 complexes. Mol Cell. 8(5), 1105-15. Chiruvella, K. K., Z. Liang & T. E. Wilson, (2013), Repair of Double-Strand Breaks by End Joining. Cold Spring Harbor Perspectives in Biology, 5(5):127-57. Doi: 10.1101/cshperspect.a012757. Dahle, J., et al. (2008), “Overexpression of human OGG1 in mammalian cells decreases ultraviolet A induced mutagenesis”, Cancer Letters, Vol.267, Elsevier, Amsterdam, https://doi.org/10.1016/j.canlet.2008.03.002.  Deem, A. et al. (2011), "Break-Induced Replication Is Highly Inaccurate.", PLoS Biol.  9:e1000594. Doi: 10.1371/journal.pbio.1000594. Dianov, G.L. & U. Hübscher (2013), "Mammalian base excision repair: the forgotten archangel", Nucleic Acids Res., 41(6):3483-90. Doi: 10.1093/nar/gkt076. Dilley, R.L. et al.  Greenberg (2016), "Break-induced telomere synthesis underlies alternative telomere maintenance", Nature, 539:54-58. Doi: 10.1038/nature20099. Douglas, G.R. et al.  (1995), "Temporal and molecular characteristics of mutations induced by ethylnitrosourea in germ cells isolated from seminiferous tubules and in spermatozoa of lacZ transgenic mice", Proceedings of the National Academy of Sciences of the United States of America, 92(16):7485-7489. Doi: 10.1073/pnas.92.16.7485. Fattah, F. et al., (2010), Ku regulates the non-homologous end joining pathway choice of DNA double-strand break repair in human somatic cells. PLoS Genet, 6(2), doi:10.1371/journal.pgen.1000855 Fitzgerald, D.M., P.J. Hastings, and S.M. Rosenberg (2017), "Stress-Induced Mutagenesis: Implications in Cancer and Drug Resistance", Ann Rev Cancer Biol, 1:119-140. Doi: 10.1146/annurev-cancerbio-050216-121919. Gorbunova, V. and A. Seluanov. (2016), “DNA double strand break repair, aging and the chromatin connection”, Mutation Research/Fundamental and Molecular Mechanisms of Mutagenesis, Vol.788/1-2, Elsevier, Amsterdam, http://dx.doi.org/10.1016/j.mrfmmm.2016.02.004.  Hammel, M. et al., (2011), XRCC4 protein interactions with XRCC4-like factor (XLF) create an extended grooved scaffold for DNA ligation and double strand break repair. J Biol Chem, 286(37), 32638-32650. doi:10.1074/jbc.M111.272641. Hanawalt, P.C., J.M. Ford and D.R. Lloyd (2003), "Functional characterization of global genomic DNA repair and its implications for cancer", Mutation Research, 544(2-3): 107–114. Harbach, P. R. et al., (1989), “The in vitro unscheduled DNA synthesis (UDS) assay in rat primary hepatocytes”, Mutation Research, 216(2):101-10. Doi:10.1016/0165-1161(89)90010-1. Iyama, T. and D.M. Wilson III (2013), "DNA repair mechanisms in dividing and non-dividing cells", DNA Repair, 12(8): 620– 636. Jeffrey, M. A.& M. G. Williams, (2000), “Lack of DNA-damaging Activity of Five Non-nutritive Sweeteners in the Rat Hepatocyte/DNA Repair Assay”,  Food and Chemical Toxicology, 38: 335-338. Doi: 10.1016/S0278-6915(99)00163-5. Köberle, B. et al. (1999), "Defective repair of cisplatin-induced DNA damage caused by reduced XPA protein in testicular germ cell tumours", Curr. Biol., 9(5):273-6. Doi: <a href="https://doi.org/10.1016/s0960-9822(99)80118-3" target="_blank">10.1016/s0960-9822(99)80118-3</a>. Kozmin, S.G. & S. Jinks-Robertson S. (2013), “The mechanism of nucleotide excision repair-mediated UV-induced mutagenesis in nonproliferating cells”, Genetics, 193(3): 803-17. Doi: 10.1534/genetics.112.147421. Kramara, J., B. Osia, and A. Malkova (2018), "Break-Induced Replication: The Where, The Why, and The How", Trends Genet, 34:518-531. Doi: 10.1016/j.tig.2018.04.002. Kuhne, M., G. Urban and M. Lo (2005), "DNA Double-Strand Break Misrejoining after Exposure of Primary Human Fibroblasts to CK Characteristic X Rays, 29 kVp X Rays and 60Co γ Rays", Radiation. Research, Vol.164/5, Radiation Research Society, Indianapolis, https://doi.org/10.1667/RR3461.1.  Larsen, K.H. et al. (1982), “DNA repair assays as tests for environmental mutagens: A report of the U.S. EPA gene-tox program”, Mutation Research, Vol.98/3, Elsevier, Amsterdam, https://doi.org/10.1016/0165-1110(82)90037-9.  Li Z, A. H. Pearlman, and P. Hsieh (2016), "DNA mismatch repair and the DNA damage response", DNA Repair (Amst), 38:94-101. Lieber, M. R., (2010), “The mechanism of double-strand DNA break repair by the nonhomologous DNA end-joining pathway.” Annu Rev Biochem. 79:181-211. doi:10.1146/annurev.biochem.052308.093131. Lieber, M. R. et al., (2003), “Mechanism and regulation of human non-homologous DNA end-joining”, Nat Rev Mol Cell Biol. 4(9):712-720. doi:10.1038/nrm1202. Liu, Y. et al. (2009), "Coordination between polymerase beta and FEN1 can modulate CAG repeat expansion", J. Biol. Chem., 284(41): 28352-28366. Doi: 10.1074/jbc.M109.050286. Mao, Z. et al., (2011), “SIRT6 promotes DNA repair under stress by activating PARP1”, Science. 332(6036): 1443-1446. doi:10.1126/science.1202723. Mariotti, L.G. et al. (2013), “Use of the γ-H2AX Assay to Investigate DNA Repair Dynamics Following Multiple Radiation Exposures”, PLoS ONE, Vol.8/11, PLoS, San Francisco, https://doi.org/10.1371/journal.pone.0079541.  Matthews, L. A., & L. A. Simmons, (2014), “Bacterial nonhomologous end joining requires teamwork”, J Bacteriol. 196(19): 3363-3365. doi:10.1128/JB.02042-14. Menoni, H. et al. (2012), "Base excision repair of 8-oxoG in dinucleosomes", Nucleic Acids Res. ,40(2): 692-700. Doi: <a href="https://doi.org/10.1093/nar/gkr761" target="_blank">10.1093/nar/gkr761</a>. Minocherhomji, S. et al. (2015), "Replication stress activates DNA repair synthesis in mitosis", Nature, 528:286-290. Doi: 10.1038/nature16139. Miyaoka, Y. et al., (2016), “Systematic quantification of HDR and NHEJ reveals effects of locus, nuclease, and cell type on genome-editing”, Sci Rep, 6, 23549. doi:10.1038/srep23549/. Moore, J. K., & J. E. Haber, (1996), “Cell cycle and genetic requirements of two pathways of nonhomologous end-joining repair of double-strand breaks in Saccharomyces cerevisiae”, Molecular and Cellular Biology, 16(5), 2164–73.  Doi: 10.1128/MCB.16.5.2164. Nacci, D. et al. (1992), “Application of the DNA alkaline unwinding assay to detect DNA strand breaks in marine bivalves”, Marine Environmental Research, Vol.33/2, Elsevier BV, Amsterdam, https://doi.org/10.1016/0141-1136(92)90134-8.  Nagel, Z.D. et al. (2014), "Multiplexed DNA repair assays for multiple lesions and multiple doses via transcription inhibition and transcriptional mutagenesis", Proc. Natl. Acad. Sci. USA, 111(18):E1823-32. Doi: 10.1073/pnas.1401182111. O’Brien, J.M. et al. (2015), "Sublinear response in lacZ mutant frequency of Muta™ Mouse spermatogonial stem cells after low dose subchronic exposure to N-ethyl-N-nitrosourea", Environ. Mol. Mutagen., 56(4): 347-55. Doi: 10.1002/em.21932. Olive, L. P., J. P. Bnath & E. R. Durand, (1990), “Heterogeneity in Radiation-Induced DNA Damage and Repairing Tumor and Normal Cells Measured Using the "Comet" Assay”, Radiation Research. 122: 86-94. Doi: 10.1667/rrav04.1. Pardo, B., B. Gomez-Gonzalez & A. Aguilera, (2009), “DNA repair in mammalian cells: DNA double-strand break repair: how to fix a broken relationship”, Cell Mol Life Sci, 66(6), 1039-1056. doi:10.1007/s00018-009-8740-3. Pegg, A.E. (2011), "Multifaceted roles of alkyltransferase and related proteins in DNA repair, DNA damage, resistance to chemotherapy, and research tools", Chem. Res. Toxicol., 4(5): 618-39. Doi: 10.1021/tx200031q. Penninckx, S. et al. (2021), “Quantification of radiation-induced DNA double strand break repair foci to evaluate and predict biological responses to ionizing radiation”, NAR Cancer, Vol.3/4, Oxford University Press, Oxford, https://doi.org/10.1093/narcan/zcab046.  Rydberg, B. et al. (2005), "Dose-Dependent Misrejoining of Radiation-Induced DNA Double-Strand Breaks in Human Fibroblasts: Experimental and Theoretical Study for High- and Low-LET Radiation", Radiation Research, Vol.163/5, Radiation Research Society, Indianapolis, https://doi.org/10.1667/RR3346.   Sancar, A. (2003), "Structure and function of DNA photolyase and cryptochrome blue-light photoreceptors", Chem Rev., 103(6): 2203-37. Doi: 10.1021/cr0204348. Saini, N. et al. (2017), "Migrating bubble during break-induced replication drives conservative DNA synthesis", Nature, 502:389-392. Doi: 10.1038/nature12584. Sakofsky, C.J. et al. (2015), "Translesion Polymerases Drive Microhomology-Mediated Break-Induced Replication Leading to Complex Chromosomal Rearrangements", Mol Cell, 60:860-872. Doi: 10.1016/j.molcel.2015.10.041. Schärer, O.D. (2013), "Nucleotide excision repair in eukaryotes", Cold Spring Harb. Perspect. Biol., 5(10): a012609. Doi: 10.1101/cshperspect.a012609. Scherer, E., A.A. Jenner and L. den Engelse (1987), "Immunocytochemical studies on the formation and repair of O6-alkylguanine in rat tissues", IARC Sci Publ., 84: 55-8. Seiler, F., K. Kamino, M. Emura, U. Mohr and J. Thomale (1997), "Formation and persistence of the miscoding DNA alkylation product O6-ethylguanine in male germ cells of the hamster", Mutat Res., 385(3): 205-211. Doi: 10.1016/s0921-8777(97)00043-8. Shelby, M.D. and K.R. Tindall (1997), "Mammalian germ cell mutagenicity of ENU, IPMS and MMS, chemicals selected for a transgenic mouse collaborative study", Mutation Research, 388(2-3): 99-109. Doi: 10.1016/s1383-5718(96)00106-4. Seo, Y.R. and H.J. Jung (2004), "The potential roles of p53 tumor suppressor in nucleotide excision repair (NER) and base excision repair (BER)", Exp. Mol. Med., 36(6): 505-509. Doi: 10.1038/emm.2004.64. Sundheim, O. et al. (2008), "AlkB demethylases flip out in different ways", DNA Repair (Amst)., 7(11): 1916-1923. Doi: <a href="https://doi.org/10.1016/j.dnarep.2008.07.015" target="_blank">10.1016/j.dnarep.2008.07.015</a>. Sung, P., & H. Klein, (2006), “Mechanism of homologous recombination: mediators and helicases take on regulatory functions”,  Nat Rev Mol Cell Biol, 7(10), 739-750. Doi:10. 1038/nrm2008. Trucco, C., et al., (1998), “DNA repair defect i poly(ADP-ribose) polymerase-deficient cell lines”, Nucleic Acids Research. 26(11): 2644–2649. Doi: 10.1093/nar/26.11.2644. Trzeciak, A.R. et al. (2008), “Age, sex, and race influence single-strand break repair capacity in a human population”, Free Radical Biology & Medicine, Vol. 45, Elsevier, Amsterdam, https://doi.org/10.1016/j.freeradbiomed.2008.08.031.  White, R.R. and J. Vijg. (2016), “Do DNA Double-Strand Breaks Drive Aging?”, Molecular Cell, Vol.63, Elsevier, Amsterdam, http://doi.org/10.1016/j.molcel.2016.08.004.  Wyrick, J.J. & S. A. Roberts, (2015), “Genomic approaches to DNA repair and mutagenesis”, DNA Repair (Amst). 36:146-155. doi: 10.1016/j.dnarep.2015.09.018. van Zeeland, A.A., A. de Groot and A. Neuhäuser-Klaus (1990), "DNA adduct formation in mouse testis by ethylating agents: a comparison with germ-cell mutagenesis", Mutat. Res., 231(1): 55-62. AOPWiki 2016-11-29T18:41:23

2023-05-15T08:41:34

Increase, Mutations

Molecular

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

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

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

High Unspecific

High

All life stages

High Moderate High Moderate

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

2023-05-15T08:47:43

Increase, Cancer

Tissue

Cancer is a general key event for related diseases each exhibiting uncontrolled proliferation of abnormal cells (for review see Hanahan and Weinberg 2011).  A cancer often is initially associated with a specific organ, with malignant tumors developing ability to metastasize, or travel to other areas of the body.  Most cancers develop from genetic mutations in normal cells, although a minority of cancers are hereditary.   Exposure to chemical stressors, radiation, tobacco smoke, or viruses can increase the likelihood that cancer will develop. Cancer cells proliferate due to capabilities summarized by Hanahan and Weinberg (2011): <ol> <li>Sustained proliferation signaling – by deregulating normal cell signals, cancer cells can sustain chronic proliferation.</li> <li>Evading growth suppressors – by evading activities of tumor suppressor genes, cancer cells continue to proliferate.</li> <li>Activating invasion and metastasis – by altering shape and attachment to cells in the extracellular matrix, cancer cells gain ability to move to other locations.</li> <li>Enabling replicative immortality – by disabling senescence pathways, cancer cells have extended lifespans.</li> <li>Inducing angiogenesis – by enabling neovasculature, cancer cells receive nutrients and oxygen and get rid of waste products.</li> <li>Resisting cell death – by evading apotosis and necrosis defense pathways, cancer cells avoid elimination.</li> </ol>

Most carcinogenicity studies are conducted with rodents (see OECD 2018 for methods) or in-vitro with mammalian cell lines (see OECD 2023 for methods).  Cancer is usually detected by biopsy or histopathological examination of tissue.  Gene expression levels can also be assessed, as increased transcription of known genes have been associated with specific cancers (ex. Tumor Necrosis Factor (Pavet et al. 2014); Heat Shock Factors (Vihervaara and Sistonen 2014; Androgen Receptor (Heinlein and Chang 2004)).

Life Stage: All life stages.  Older individuals are more likely to manifest this key event (adults > juveniles > embryos). Sex: Applies to both males and females. Taxonomic: Appears to be present broadly, with representative studies including mammals (humans, lab mice, lab rats), teleost fish, and invertebrates (cladocerans, mussels).

High Unspecific

High

All life stages

High High High

Abraha, A.M. and Ketema, E.B.  2016.  Apoptotic pathways as a therapeutic target for colorectal cancer treatment.  World Journal of Gastrointestinal Oncology 8 (8): 583-491 European Parliament.  2022.  Directive 2004/37/EC of the European Parliament on the protection of workers from the risks related to exposure to carcinogens, mutagens or reprotoxic substances at work.  Retrieved 3 August 2023 from http://data.europa.eu/eli/dir/2004/37/2022-04-05 Hanahan, D. and Weinberg, R.A.  2011.  Hallmarks of cancer: the next generation.  Cell 144(5): 646-674. Heinlein, C.A. and Chang, C.  2004.  Androgen receptor in prostate cancer.  Endocrine Reviews 25: 276-308. OECD.  2018.  Test no. 451: OECD Guideline for the Testing of Chemicals: Carcinogenicity Studies.  OECD Publishing, Paris.  Retrieved 3 August 2023 from <a href="https://www.oecd.org/env/test-no-451-carcinogenicity-studies-9789264071186-en.htm" style="color:#0563c1; text-decoration:underline">https://www.oecd.org/env/test-no-451-carcinogenicity-studies-9789264071186-en.htm</a> OECD.  2023. Test No. 487: In Vitro Mammalian Cell Micronucleus Test, OECD Guidelines for the Testing of Chemicals, Section 4, OECD Publishing, Paris.  Retrieved 3 August 2023 from  <a href="https://doi.org/10.1787/9789264264861-en.htm" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1787/9789264264861-en.htm</a> OSHA. 2023.  Carcinogens.  Retrieved 3 August 2023 from <a href="https://www.osha.gov/carcinogens/standards" style="color:#0563c1; text-decoration:underline">https://www.osha.gov/carcinogens/standards</a> Pavet, V., Shlyakhtina, Y., He, T., Ceschin, D.G., Kohonen, P., Perala, M., Kallioniemi, O., and Gronemeyer, H.  2014.  Plasminogen activator urokinase expression reveals TRAIL responsiveness and support fractional survival of cancer cells.  Cell Death and Disease 5: e1043. Vihervaara, A. and Sistonen, L.  2014.  HSF1 at a glance.  Journal of Cell Scientce 127: 261-266. Zhou, Y., Xia, J., Xu, S., She, T., Zhang, Y., Sun, Y., Wen, M., Jiang, T., Xiong, Y., and Lei, J.  2023.  Experimental mouse models for translational human cancer research.  Frontiers in Immunology 14: 1095388. AOPWiki 2016-11-29T18:41:27

2023-08-10T14:59:50

<upstream-id>02d1eee7-71a4-4d86-818c-7c9527f05b8b</upstream-id> <downstream-id>fb2b9f6b-28b0-4a81-8daa-592de2e2332c</downstream-id> Alkylated DNA may be tolerated and/or repaired error-free by a variety of DNA repair pathways. However, at high doses, it is established that the primary DNA repair pathway (O6-Alkylguanine-DNA alkyltransferase: AGT) responsible for removing alkylated DNA becomes saturated. This may lead to several potential adduct fates: (i) error-free repair of the DNA adduct using alternative DNA repair mechanisms; (ii) no repair (DNA damage is retained); or (iii) instability in the DNA duplex leading to DNA strand breaks and possibly activation of DNA damage signaling. For repair of alkyl adducts it is well established that the O6-alkylguanine-DNA alkyltransferase pathway becomes saturated at high doses leading to insufficient repair at high doses.

General details: The weight of evidence for this KER is strong. It is widely accepted that damaged DNA is subject to repair, and that in the absence of DNA repair, mutations will ensue. Specifically, AGT (Damage Reversal DNA repair: pathway #1 in KE155), also known as O6-methylguanine-DNA methyltransferase (MGMT), reverses alkylation damage by directly transferring alkyl groups from the O6 position of guanine to a cysteine residue on the AGT (or MGMT) molecule, restoring the DNA in a single step. However, transfer of the alky group to AGT results in concomitant inactivation of AGT (Pegg 2011). The mammalian protein is also active on O6-ethylguanine and can remove only one ethyl group from DNA, following which the protein is degraded. Thus, high levels of alkylation damage overwhelm the cellular AGT capacity to remove lesions. In mammalian cells, O4-ethylthymine and O2-ethylthymine are poor substrates for AGT (Fang et al. 2010) and no other DNA repair pathway has been identified that is able to efficiently repair these lesions; consequently, these lesions are extremely persistent in cells. Reviews on this topic have been published (Kaina et al. 2007; Pegg 2011). In the absence of the AGT/MGMT pathway, other DNA repair pathways may be invoked, but the relative efficiency of these pathways is not well understood (further details described below). The role of nucleotide excision repair (NER; excision repair pathways: #2 in KE155) in alkylation damage repair in mammalian cells remains unclear. Earlier studies using human cell lines suggested that both AGT and NER may be involved in the repair of O6-ethylguanine (Bronstein et al. 1991; Bronstein et al. 1992). Very recently, an alkyltransferase like protein (ATL1) that has homology to AGT has been identified in a range of prokaryotes and lower eukaryotes. This protein has no alkyltransferase activity but can couple O6-alkylguanine damage to NER (Latypov et al. 2012). ATL1 proteins have not yet been identified in mammals. Some alkyl adducts, such as N7-ethylguanine and N3-ethyladenine, are inherently unstable and may depurinate (i.e., hydrolytic cleavage of the glycosidic bond, which releases adenine or guanine). The resultant abasic sites are normally repaired through error-free pathways although they may occasionally be transformed to DNA strand breaks. In mammals, N-methylpurine DNA glycosylases, such as alkyladenine DNA glycosylase (AAG), have a wide range of substrates including N7-alkylguanine and N3-alkyladenine derivatives (Wyatt et al. 1999). However, there are no specific reports in the literature that the ethylated derivatives are AAG substrates. Glycosylases such as AAG yield abasic sites that are processed as described above. An alternative repair mechanism for repairing minor lesions such as N3-ethylcytosine and N1-ethyladenine is through oxidative dealkylation catalyzed by AlkB and mammalian homologs (Drabløs et al. 2004). This pathway is an error-free damage reversal pathway that releases the oxidized ethyl group as acetaldehyde (Duncan et al. 2002). A final mechanism through which DNA repair pathways may influence the fate of alkylation damage is through futile cycling of the mismatch repair (MMR; excision repair pathways: #2 in KE155) system at an O6-alkyl G:T mispair. In this scenario, unrepaired O6-alkylguanine is able to mispair with T, and the mispair is recognized by MMR enzymes resulting in the removal of the newly incorporated thymine from the nascent strand opposite the O6-alkyguanine adduct. During DNA repair synthesis, O6-alkylguanine preferentially pairs once again with thymine, reinitiating the repair/synthesis cycle. This iteration of excision and synthesis may produce strand breaks and activate damage signaling pathways (York and Modrich 2006).   If the pathways described above become saturated or do not operate properly, the alkylated DNA will not be repaired and will provide a template for replication of this damaged DNA. This is widely understood and accepted. Many studies have demonstrated that the introduction of plasmids or vectors with alkylated DNA (i.e., unrepaired lesions) into prokaryotic and eukaryotic cells, followed by replication, results in the formation of mutations at the alkylated sites, and that the probability of a mutation occurring at the alkylated site is modified by specific DNA repair genes/pathways (reviewed in Basu and Essigmann 1990; Shrivastav et al. 2010).

Insufficient repair is inferred from the formation and retention of adducts, and the formation of increased numbers of mutations above background (i.e., KE185 - methodologies described therein). A variety of studies show that alkylated DNA persists for prolonged periods of time post-exposure. For example, persistence of different alkylated nucleotides was shown in livers and brains of C57BL mice exposed to N-methyl-N-nitrosourea, N-ethyl-N-nitrosourea and ethyl methanesulfonate using high-performance liquid chromatography several days post-exposure (Frei et al., 1978). The stability of methyl and ethyl adducts in somatic tissues for various adduct types is summarized in Beranuk, 1990. The in vivo liver half life of methyl adducts ranges from 0-3 days, and liver ethyl adduct half lives can be up to 17 days, indicating poorer repair of oxygen-bound ethyl adducts. This prolonged retention of adducts indicates that there is insufficient repair by AGT or other DNA repair pathways of these adducts. Studies in both hamsters and rats show persistence of alkylated nucleotides several days post-exposure, indicating lack of DNA repair of some adducts (Scherer et al. 1987; Seiler et al. 1997). For example, 101xC3H mouse hybrid testes exhibited DNA adducts within 1 hour of exposure to ENU (10 or 100 mg/kg by i.p.), but some adducts remained unrepaired six days post-exposure (Sega et al. 1986). O6-ethylguanine adducts were also found in hamster spermatogonia DNA up to four days after exposure to DEN (100 µg/g body weight) (Seiler et al. 1997). O6-ethylguanine adducts were found in spermatogonia 1.5 hours post-exposure to ENU in Syrian Golden hamsters (Seiler et al. 1997). Approximately 30% persisted in spermatogonia four days post-exposure. Moreover, the amount of O6-ethylguanine recovered after a 100 mg ENU/kg exposure was 40% greater than predicted from a linear extrapolation of the amount of O6-ethylguanine recovered after exposure to 10 mg/kg. The data suggest that the high dose exposure to ENU results in depletion of AGT within the testis and permits O6-ethylguanine to persist at higher levels than would be predicted from lower exposure. The relationship between dose and formation of DNA adducts in tubular germ cells is non-linear, indicating relatively rapid repair at low doses that becomes saturated at higher doses (van Zeeland et al. 1990). Thus, with increasing dose, increasing incidence of KE1 (insufficient repair) occurs. This implies that mouse spermatogonia are capable of repairing a major part of the DNA damage at low doses. However, at higher doses the repair process is saturated and mutations begin to occur. Indeed, the dose-response curve for mutations in spermatogonia measured in sperm of exposed males is sub-linear with a clear point of inflection at low sub-chronic doses of ENU (O’Brien et al. 2015). Finally, both alkyl adducts and mutations increase with increasing doses of alkylating agents in somatic cells and in male germ cells, indicating that DNA repair processes are not operating to remove all of the damage (ability to remove adducts and prevent mutations).

DNA repair is not generally measured directly; thus, insufficient repair is inferred from the retention of adducts or the induction of increases in mutation frequencies post-exposure. In addition, various sizes of alkylation groups (e.g., methyl, ethyl, propyl) can be involved. Although it appears that the larger alkyl adducts tend to be more mutagenic (Beranek, 1990), this is not completely established and there are insufficient data to establish that this is true for germ cells. However, in general, this KER is biologically plausible, broadly accepted for alkyl adducts and has few uncertainties. The direct measurement of insufficient repair can be considered a data gap.

Moderate Moderate

DNA adducts can occur in any cell type. While there are differences across taxa, all species have some DNA repair systems in place and it is common to extrapolate conclusions across eukaryotic species.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00005eb8a24f1fc8> AOPWiki 2016-11-29T18:41:33

2019-12-10T10:43:25

<upstream-id>fb2b9f6b-28b0-4a81-8daa-592de2e2332c</upstream-id> <downstream-id>88691209-9194-4245-86ee-8fd2f185cd1f</downstream-id> 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.

Overall Weight of Evidence: High

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

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

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

Not identified.

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

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

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

Not identified.

High Unspecific

High

All life stages

High High High

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

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00005eb8a27d2238> AOPWiki 2016-11-29T18:41:33

2023-05-15T13:37:37

<upstream-id>88691209-9194-4245-86ee-8fd2f185cd1f</upstream-id> <downstream-id>74dd9841-19fb-45a2-96a0-36a43cb5ae6a</downstream-id>

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00005eb8a27e8330> AOPWiki 2016-11-29T18:41:35

2016-12-03T16:38:01

<upstream-id>02d1eee7-71a4-4d86-818c-7c9527f05b8b</upstream-id> <downstream-id>88691209-9194-4245-86ee-8fd2f185cd1f</downstream-id> Alkylated DNA may be ‘misread’ during DNA replication, leading to insertion of incorrect nucleotides. Upon replication, these changes become fixed as mutations. Subsequent replication propagates these mutations to daughter cells. Mutations in stem cells are of the greatest concern, as these will persist throughout the organism’s lifetime. Thus, increased mutations will be found in the cells of organisms that possess alkylated DNA.

Alkylating agents can cause a variety of adducts and DNA damage (e.g., alkali labile sites, DNA strand breaks, etc.) that are potentially mutagenic and clastogenic. This KER focuses on the probability that an alkyl DNA adduct will lead to a mutation. Not all adducts are equally mutagenic. Very generally, chemicals that preferentially cause O-alkylation in DNA induce DNA sequence changes, whereas chemicals that cause N-alkylation of DNA are more efficient inducers of structural chromosomal aberrations (reviewed in Beranek 1990). Indeed, a review of the biological significance of N7 alkyl-guanine adducts concluded that these adducts simply be used to confirm exposure to target tissue (Boysen et al., 2009), because the vast majority of studies shows that these adducts do not cause mispairing. A variety of work has demonstrated that N7-alkylguanine adducts can be bypassed essentially error free (e.g., Philippin et al., 2014; Shrivastav et al., 2010). Moreover, alkylation can involve modification with different sizes of alkylation groups (e.g., methyl, ethyl, propyl). Although response to these is qualitatively similar with respect to the key events, in general, larger alkylating groups tend to be more mutagenic (Beranek, 1990). It is widely known that chemicals that preferentially cause O-alkylation in DNA induce mutations. ENU (N-ethyl-N-nitrosourea) is a prototypical O-alkylating agent and the most studied male germ cell mutagen. Alkylating agents are prototypical somatic and male germ cell mutagens.

Evidence in somatic cells It is well established that transfection of cells with alkylated DNA leads to mutation at the sites of alkyl damage. The design of these experiments requires waiting for cellular replication in order to produce the mutation, confirming the temporal concordance of DNA alkylation and subsequent mutation. A summary of the empirical data to support this is reviewed in Shrivastav et al. (2010). Various studies have examined the dose-response of DNA adduct levels and mutations. These studies demonstrate that alkyl adducts can be seen at lower doses in the absence of increased mutations both in vitro and in vivo, or demonstrate equal or increased incidence of adducts relative to mutations at the same doses. For example, following exposure of AHH-1 cells to increasing concentrations of MMS, a linear increase in alkylated DNA is measured with significant increases occurring in adduct levels at 0.25 µg/ml (Swenberg et al. 2008). Significant increases in mutations in the HPRT gene in the same cells are not measurable until 1.25 µg MMS/ml (Swenberg et al. 2008) (Figure 1). In vivo, time-series analysis of λlacZ transgenic mice exposed to a single dose of either ENU or DEN, demonstrate that global and lacZ-specific O6-EtGua adducts occur within hours of exposure in the liver, with the bulk of adducts removed by three days post-exposure (Mientjes et al. 1996 and 1998). In contrast, mutant frequency does not begin to significantly increase until three days post-exposure, demonstrating temporal concordance of adduct and mutation formation (see Figure 1 in Mientjes et al. 1998). Levels of O6-EtGua adducts are also consistently higher than the induced mutation frequency per nucleotide. In the bone marrow, DEN is not metabolized and thus is unable to create O6-EtGua adducts. The finding of lack of O6-alkyl adducts in bone marrow is consistent with the lack of an increase in mutations observed in this tissue (Mientjes et al. 1998). This is in contrast to ENU exposure (a direct acting mutagen that does not require metabolic activation), where both adducts and mutations increase in a concordant fashion in the bone marrow. This pattern of adduct incidence versus mutation incidence is consistent for somatic tissues in rodents in vivo for other types of adducts (we were not able to find suitable dose-response studies to compare oxygen-alkyl adducts to mutation frequencies in vivo). For example, MutaMouse males were exposed to increasing doses of the polycyclic aromatic hydrocarbon dibenzo[a,l]pyrene (forms mutagenic bulky DNA adducts) for 28 days (followed by a 3 day break for mutation fixation) following OECD protocol TG488 (Malik et al. 2013). Significant increases in hepatic DNA adducts were found at 25 mg/kg, but increases in lacZ mutant frequency in liver did not occur until 50 mkg/kg. Bulky DNA adducts in both the livers and lungs of MutaMouse males exhibit an order of magnitude greater incidence per nucleotide than mutations in the lacZ gene using a similar experimental design following exposure to benzo[a]pyrene (Labib et al. 2012; Malik et al. 2012). Lower tissue adduct burden is correlated with lower tissue-specific gene mutation frequencies in Big Blue mice exposed to benzo[a]pyrene (Skopek et al. 1996). Thus, adducts in DNA occurs at lower doses than mutations and are correlated with mutation burden in somatic tissues for different types of DNA adducts. Evidence in germ cells: No study has compared dose-response for adduct formation and mutation in a single experiment on germ cells. However, it is possible to look across experiments. It is important to note that adducts are measured immediately following exposure because they are relatively quickly repaired. However, analysis of lacZ mutation requires collection of mature sperm from the caudal epididymis. Thus, sperm is collected 42 or 49 days post-exposure (OECD TG488). This is because spermatogonia can not be sampled directly for these purposes. Therefore, comparison of adducts to mutations in pre-meiotic male germ cells requires sampling at different time points for these endpoints (early for adducts, much later for mutations), which is consistent with the expected temporal order of events, with adducts occurring before mutations. Dose-response for alkyl adduct levels has been very well characterized in mouse testicular DNA for ENU, EMS and DES (van Zeeland et al. 1990). A summary of dose-response data for mouse exposure to ENU and mutation analysis using the transgenic rodent mutation assay in sperm is given in the attached Table I and Figure 2. These studies involve acute injections or oral gavage studies only. Alkyl adducts are evident in gonadal tissues within 2 hours of exposure (van Zeeland et al. 1990) and are fairly efficiently removed within days of exposure in the absence of continued exposure. For this analysis, transgene mutant frequencies were converted to per nucleotide mutation frequency by dividing mutant frequency by the length of the lacZ gene (3096 bp). The data demonstrate that alkyl adduct incidence at low doses is much greater per nucleotide than transgene mutations in the lacZ locus. Adducts are observed to increase substantially at the lowest exposure dose (10 mg/kg), whereas mutation increases in lacZ are marginal at 25 mg/kg. Alkyl adducts in mouse testes following ENU exposure were in the range of approximately 40 in 10E7 nucleotides for 80 mg/kg ENU exposure (van Zeeland et al. 1990). Conversion of the data in O’Brien et al. (2015) to mutations per nucleotide (by dividing mutant frequency by the length of the lacZ locus, which is 3096 bp) produces an estimated induced mutation frequency in spermatogonia of approximately 4 mutations per 10E7 nucleotides for the highest dose (100 mg/kg ENU), an increase of approximately 2 in 10E7 above controls. This suggests that the incidence of adducts is an order of magnitude greater than incidence of mutations. Similarly, exposure to a single dose of 250 mg/kg EMS leads to an over 10-fold increase in the number of alkyl adducts (although the majority are on nitrogen atoms, with only a small proportion on oxygen) (van Zeeland et al. 1990), but only a marginal 2-fold increase in lacZ mutation frequency [van Delft et al. 1997]. Indeed, Van Zeeland et al. (1990) estimate that approximately 10 O6-ethylguanine adducts are required in the gene-coding region to generate a mutation. Analysis of germ cell mutation during a sub-chronic exposure was carried out by O’Brien et al. (2015). The lower doses used in that study revealed that significant increases in mutations occurred only after 28 days of exposure to the highest dose of 5 mg/kg ENU (cumulative dose of 140 mg/kg), again supporting that higher adduct burdens are required to lead to mutations. In general, many studies in different mouse strains have used similar experimental designs to conclusively demonstrate that exposure to a variety of alkylating agents causes mutations in spermatogonia (Brooks and Dean 1997; Douglas et al. 1995; Katoh et al. 1997; Katoh et al. 1994; Liegibel and Schmezer 1997; Mattison et al. 1997; O'Brien et al. 2014; O’Brien et al. 2015; Renault et al. 1997; Suzuki et al. 1997; Swayne et al. 2012; Tinwell et al. 1997; van Delft et al. 1997). These studies have been done using a single dose and thus do not enable further comparison of the concordance of dose-response. We also note that ENU exposure of pre-meiotic male germ cells in fish (transgenic medaka) also causes significant increases in mutations observed in spermatozoa (Norris and Winn 2010), supporting the effects of alkylating agents on mutations in male pre-meiotic germs across taxa using similar experimental designs.

As described above, not all alkyl adducts are mutagenic. The proportion of oxygen-alkylation and the type of mutation (with ethylation > methylation) will govern mutagenicity, but there are few empirical data to support this. There are no inconsistencies or uncertainties for ENU or iPMS; other alkylating agents (EMS, MMS) have yielded some discrepancies in the transgenic rodent mutation assay. However, the experimental protocols applied were sub-standard (the OECD TG for this analysis was revised and published in 2013). Overall, more work is needed on alkylating agents other than ENU to fill important data gaps.

Is it known how much change in the first event is needed to impact the second? Are there known modulators of the response-response relationships? Are there models or extrapolation approaches that help describe those relationships? The shape of the dose-response curve for alkyl adduct formation versus mutation demonstrates that a threshold exists whereby alkyl adducts can be seen at low doses in the absence of increased mutations occurring. For example, following exposure of AHH-1 cells to increasing concentrations of MMS, a linear increase in alkylated DNA is measured. However, a hockey-stick shaped curve was found for mutations at HPRT in the same cells (Thomas et al. 2013). Thus, alkylation of DNA occurs at lower doses than mutation, and above a certain dose (where repair is saturated), mutation frequencies increase. That DNA alkylation leads to mutation in spermatogonia in a similar hockey stick-shaped response (implying that a minimal dose must be exceeded) is supported by work using the LacZ Muta™Mouse assay. Exposure of male mice to the prototypical agent ENU was used to examine effects on spermatogonial stem cells, though the number of doses was limited (van Delft and Baan 1995). This analysis revealed that mutations did not occur at the lowest dose, where adducts are known to be measurable in other studies (van Zeeland et al., 1990). This data gap motivated a dose-response study using the Muta™Mouse model following both acute and sub-chronic ENU exposure by oral gavage at Health Canada (O’Brien et al., 2015). These data indicate a clear dose-response for single acute exposures, whereas a hockey stick-shaped dose-response occurs for lower dose sub-chronic (28 day) exposures. At the single acute high doses where the DNA repair machinery is expected to be overwhelmed (and thus higher levels of alkylation occur), significantly more mutations occur relative to the same dose spread out over 28 daily oral gavage exposures (O’Brien et al., 2015). Additional contributors to the probability that an adduct will cause mutation include the site of alkylation, with agents that cause O-alkyl lesions being the primary mutagens, and the size of the alkyl group, with larger alkyl groups generally being more mutagenic. A computational model to describe the mutational efficiency of different alkyl adducts has not yet been developed to our knowledge.

High Moderate

Alkylating agents are well-established to cause mutation in virtually any cell type in any organism.

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2016-11-29T19:53:41

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#<Reference::ActiveRecord_Associations_CollectionProxy:0x00005eb8a2764ad0> AOPWiki 2016-11-29T18:41:35

2016-12-03T16:38:01

Alkylation of DNA leading to cancer 1

DNA alkylation -> cancer 1

Arthur Author

Carole Yauk Environmental Health Centre Health Canada 50 Colombine Driveway Ottawa, ON Canada K1S 0K1 email: Carole.Yauk@canada.ca

Open for adoption

1.0

Alkylating agents are prototypical DNA-reactive compounds and have been extensively studied for decades (reviewed in Beranek 1990). The chemicals can be direct-acting electrophiles, or can be converted from non-reactive substances to reactive metabolites via metabolism. A prototypical alkylating agent is N-ethyl-N-nitrosourea (chemical formula C3H7N3O2) (ENU). ENU is rapidly absorbed following oral exposure and intraperitoneal injections and distributed widely across the tissues. ENU is unstable and readily reacts with somatic and germ cell DNA in mice, rats, flies and hamsters, to alkylate DNA. Very generally, mono-functional (referring to the transfer of a single alkyl group) alkylating agents include: 1. Alkyl sulfates: e.g., diethyl (DES) and dimethyl sulfate (DMS); 2. Alkyl alkanesulfonates: e.g., methyl (MMS) and ethyl methanesulfonate (EMS); 3. Nitrosamides: e.g., methyl (MNU) and ethyl nitrosourea (ENU), methyl- (MNNG) and ethyl-N'-nitro-N-nitrosoguanidine (ENNG), and the indirect-acting (i.e., requiring metabolic activation) dimethyl (DMN) and diethyl nitrosamines (DEN). ENU is the most widely studied and understood alkylating agent and as such has been instrumental in contributing to the knowledgebase in this field. Immunohistochemistry studies clearly indicate the presence of alkylated DNA following exposure to ENU in both somatic cells and spermatogonia (Kamino et al. 1995; Seiler et al. 1997; van Zeeland et al. 1990).

Cancer is a critical endpoint in human health risk assessment.   It is embedded in regulatory frameworks for human health protection in many countries (see OSHA 2023 for examples of US regulations and European Parliament 2022 for examples of regulations in Europe).

adjacent

Moderate

High non-adjacent

Moderate

High non-adjacent

Low

Moderate non-adjacent

Moderate

High non-adjacent

Low

Moderate

Moderate Mixed

Moderate

All life stages

Not Specified Not Specified

AOPWiki 2016-11-29T18:41:16

2023-09-25T16:26:51