Relationship: 24: Alkylation, DNA leads to Inadequate DNA repair

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

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.

Key Event Relationship Description

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.

Evidence Supporting this KER

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.

References

Basu, A.K. and J.M. Essigmann (1990), "Site-specific alkylated oligodeoxynucleotides: Probes for mutagenesis, DNA repair and the structure effects of DNA damage", Mutation Research, 233: 189-201.

Beranek, D.T. (1990), "Distribution of methyl and ethyl adducts following alkylation with monofunctional alkylating agents", Mutation Research , 231(1): 11-30.

Bronstein, S.M., J.E. Cochrane, T.R. Craft, J.A. Swenberg and T.R. Skopek (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., T.R. Skopek and J.A. Swenberg (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.

Drabløs, F., E. Feyzi, P.A. Aas, C.B. Vaagbø, B. Kavli, M.S. Bratlie, J. Peña-Diaz, M. Otterlei, G. Slupphaug and H.E. Krokan (2004), "Alkylation damage in DNA and RNA--repair mechanisms and medical significance", DNA Repair, 3(11): 1389-1407.

Duncan, T., S.C. Trewick, P. Koivisto, P.A. Bates, T. Lindahl and B. Sedgwick B (2002), "Reversal of DNA alkylation damage by two human dioxygenases", Proc. Natl. Acad. Sci. USA, 99(26): 16660-16665.

Fang, Q., S. Kanugula, J.L. Tubbs, T.A. Tainer and A.E. Pegg (2010), "Repair of O4-alkylthymine by O6-alkylguanine-DNA alkyltransferases", J. Biol. Chem. 12(285): 885-895.

Frei, J.V., D.H .Swenson, W. Warren, P.D. Lawley (1978), "Alkylation of deoxyribonucleic acid in vivo in various organs of C57BL mice by the carcinogens N-methyl-N-nitrosourea, N-ethyl-N-nitrosourea and ethyl methanesulphonate in relation to induction of thymic lymphoma. Some applications of high-pressure liquid chromatography", Biochem. J., 174(3): 1031-1044.

Kaina, B., M. Christmann, S. Naumann and W.P. Roos (2007), "MGMT: Key node in the battle against genotoxicity, carcinogenicity and apoptosis induced by alkylating agents", DNA Repair, 6: 1079–1099.

Latypov, V.F., J.L. Tubbs, A.J. Watson, A.S. Marriott, G. McGown, M. Thorncroft, O.J. Wilkinson, P. Senthong, A. Butt, A.S. Arvai, C.L. Millington, A.C. Povey, D.M. Williams, M.F. Santibanez-Koref, J.A. Tainer and G.P. Margison GP (2012), "Atl1 regulates choice between global genome and transcription-coupled repair of O(6)-alkylguanines", Mol. Cell, 47(1): 50-60.

Muller, L., E. Gocke, T. Lave and T. Pfister (2009), "Ethyl methanesulfonate toxicity in Viracept – a comprehensive assessment based on threshold data for genotoxicity", Toxicology Letters, 190: 317-329.

O’Brien, J.M., M. Walker, A. Sivathayalan, G.R. Douglas, C.L. Yauk and F. Marchetti (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.

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., 24(5): 618-639.

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.

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(1-2): 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(3): 205-211.

Shrivastav, N., D. Li and J.M. Essignmann (2010), "Chemical biology of mutagenesis and DNA repair: cellular response to DNA alkylation", Carcinogenesis, 31(1): 59-70.

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.

Wyatt, M.D., J.M. Allan, A.Y. Lau, T.E. Ellenberger, L.D. Samson (1999), "3-methyladenine DNA glycosylases: structure, function, and biological importance", Bioessays, 21(8): 668-676.

York S.J. and P. Modrich (2006), "Mismatch repair-dependent iterative excision at irreparable O6-methylguanine lesions in human nuclear extracts", J. Biol. Chem., 281(32): 22674-22683.

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

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

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

Key Event Relationship Description

Inadequate repair of DNA damage includes incorrect repair (i.e., incorrect base insertion), incomplete repair (i.e., accumulation of repair intermediates such as strand breaks, stalled replications forks, and/or abasic sites), and absent repair resulting in the retention of DNA damage.

It is well-established that DNA excision repair pathways require DNA strand breakage for removing the damaged sites; for example, base excision repair (BER) of oxidative lesions involves removal of oxidized bases by glycosylases followed by cleavage of the DNA strand 5’ from the abasic site. If the repair process is disrupted at this point, repair intermediates including single strand breaks (SSB) may persist in the DNA. A SSB can turn into a double strand break (DSB) if it occurs sufficiently close to another SSB on the opposite strand. SSBs can be converted into DSBs when helicase unwinds the DNA strands during replication. Furthermore, SSBs and abasic sites can act as replication blocks causing the replication fork to stall and collapse, giving rise to DSBs (Minko et al., 2016; Whitaker et al., 2017).

Evidence Supporting this KER

1. DNA strand breaks generated due to faulty attempted repair

Excision repair pathways require the induction of SSB as part of damage processing. Increases in DNA lesions may lead to the accumulation of intermediate SSB. Attempted excision repair of lesions on opposite strands can turn into DSBs if the two are in close proximity (Eccles et al., 2010). Generation of DSBs has been observed in both BER and nucleotide excision repair (NER) (Ma et al., 2009; Wakasugi et al., 2014).

Previous studies have demonstrated that an imbalance in one of the multiple steps of BER can lead to an accumulation of repair intermediates and failed repair. It is highly likely that a disproportionate increase in oxidative DNA lesions compared to the level of available BER glycosylases leads to an imbalance between lesions and the initiating step of BER (Brenerman et al., 2014). Accumulation of oxidative lesions, abasic sites, and SSBs generated from OGG1, NTH1, and APE1 activities would be observed as a result. Moreover, studies have reported accumulation of SSB due to OGG1- and NHT1-overexpression (Yang et al., 2004; Yoshikawa et al., 2015; Wang et al., 2018). BER repair intermediates have been observed to interfere with transcription as well (Kitsera et al., 2011). While overexpression may lead to imbalanced lyase activities that generate excessive SSB intermediates, deficiency of these enzymes is also known to cause an accumulation of oxidative lesions that could lead to strand breaks downstream. Hence, both the overexpression and deficiencies of repair enzymes can lead to strand breaks due to excessive activity or inadequate repair, respectively.

Similar to BER, NER involves coordinated dual incision of the DNA strand flanking bulky lesions, generating single-stranded DNA gaps as repair intermediates. If these intermediates persist due to incomplete repair or imbalance in downstream processing steps, they can contribute to the accumulation of strand breaks. For example, NER processing of UV-induced DNA damage can lead to the formation of DSBs, particularly through the persistence of single-stranded DNA gaps or secondary processing events (Wakasugi et al., 2014). In addition, structure-specific endonucleases involved in NER, such as XPF-ERCC1 and XPG, can introduce strand incisions that, when improperly coordinated or occurring at closely spaced lesions, may result in DSB formation (Riedl et al., 2003). Thus, dysregulation or saturation of NER can contribute to DNA strand break formation through the accumulation and improper resolution of repair intermediates.

2. DNA strand breaks generated due to replication stress caused by accumulated DNA lesions

Retention of DNA lesions (i.e., damaged bases and SSB) can interfere with the progression of the replication fork. Thymidine glycol is an example of an oxidative DNA lesion that acts as a replication block (Dolinnaya et al., 2013). Persistent replication fork stalling and/or dissociation of replication machinery are known to cause the replication fork to collapse, which generates highly toxic DSBs (Zeman and Cimprich, 2014; Alexander and Orr-Weaver, 2016). Fork stalling also increases the risk of two replication forks colliding with each other, generating DSBs.

Finally, the replication fork can collide with SSBs generated during BER, hindering the completion of repair and giving rise to DSBs (Ensminger et al., 2014).

In vitro studies with empirical evidence are shown below for select DNA repair pathways. These studies build in elements of essentiality (modulation of DNA repair), as well as dose and incidence concordance. The primary evidence is essentiality, where repair is genetically modulated in some way. Because multiple lines of evidence are considered within individual studies, we present the data by source of evidence (in vitro versus in vivo) rather than by type of empirical evidence (dose, incidence, or temporal concordance; essentiality) to avoid repetitive use of the same studies. This is a very data-rich area, and the examples presented here are intended to be illustrative rather than exhaustive, reflecting a broader and well-established body of evidence in which mechanistic understanding and essentiality provide strong overall support for this KER.

Inadequate repair of oxidative lesions

• Concentration concordance of strand breaks in repair-deficient and –proficient cells (insufficient repair) (Wu et al., 2008)
  • In a study using A549 human adenocarcinoma cells, DNA strand breaks in hOGG1-proficient and hOGG1-deficient cells were compared following exposure to increasing concentrations of bleomycin.
  • Strand breaks were measured as DNA migration length in alkaline comet assay after 3 hours of exposure to six increasing concentrations (0.05, 0.25, 0.5, 1, 5, and 10 mg/L).
  • Concentration-dependent increase in strand breaks was observed in both cell types; however, at all concentrations significantly more strand breaks (p<0.05) were present in the hOGG1-deficient cells than in the proficient cells, demonstrating insufficient repair of oxidative lesions leading to DNA strand breaks.
  • Thus, this evidence supports the essentiality of inadequate DNA repair as a modulator of the downstream KE.
• Incomplete OGG1-initiated base excision repair (BER) leads to DNA strand breaks (Wang et al., 2018):
  • In a study using mouse embryonic fibroblasts (MEF), Ogg1+/+ and Ogg1-/- cells were treated with increasing concentrations of H2O2 for varying durations
    Higher levels of 8-oxodG were detected in Ogg1-/- cells compared to Ogg1+/+ cells after treatment with 400 µM H2O2 at all time points (5, 15, 30, 60, and 90 min)
    • Demonstrates insufficient removal of 8-oxo-dG in OGG1-deficient cells
  • Significantly  more strand breaks, as indicated by the higher % of TUNEL-positive cells (p<0.001), were detected in Ogg1+/+ cells compared to Ogg1-/- cells after exposure to 400 µM H2O2 for 3 hours
    • Both cell types showed a very similar increase in DNA strand breaks at lower concentrations (50, 100, and 200
      µM) and there was no significant difference between Ogg1+/+ and Ogg1-/- cells at these concentrations – this suggests that up to a certain level of oxidative damage, OGG1-initiated BER does not exacerbate strand breaks but when oxidative stress is excessive (at 400µM in this study), OGG1-initiated BER is compromised and leads to increased strand breaks (incomplete repair)
  • Finally, DNA strand breaks in both cell types were measured using both alkaline and neutral comet assay after a 30- minute exposure to 400µM H2O2; while there was an increase in the olive tail moment (indicating DNA strand breaks) in both cell types compared to the control, the increase of strand breaks in Ogg1+/+ cells was significantly larger than in Ogg1-/- cells in both assays (p<0.001)

Inadequate repair of alkylated DNA

• Interference of N-methylpurine DNA glycosylase (MPG)-initiated BER by replication leading to strand breaks (Ensminger et al., 2014)
  • A549 human alveolar basal epithelial cells were exposed to increasing concentrations of methylmethane sulfonate (MMS) for 1 hour and replicating cells were labeled using a thymidine analogue, 5-ethynyl-2’-desoxyuridine (EdU).
  • In S-phase cells, MMS concentration-dependent increase in γH2AX foci was detected (70 foci/cell at the highest concentration). In contrast, γH2AX foci were not detected G1- and G2-phase cells until the highest concentration (15 foci/cell).
  • MPG-depleted cells in S-phase showed no significant increase in γH2AX foci, while the control cells showed significant MMS concentration-dependent increases.
  • These results suggest interference of MPG-initiated BER by replication, leading to DSBs, and that the depletion of MPG decreases the probability of strand breaks in S-phase (evidence of essentiality of ‘inadequate repair’ to KEdown). 

• Depletion of O6-alkylguanine-DNA alkyltransferase (AGT/MGMT) enhances the genotoxic effects of alkylating agents
  • Numerous studies have demonstrated that genetic depletion or pharmacological inhibition of AGT/MGMT increases cellular sensitivity to alkylating chemotherapeutic agents (e.g., enhanced cytotoxicity, mutagenicity, and chromosomal damage) (Dolan et al., 1990, 1991; reviewed by Rabik et al., 2006). Although many of these studies did not directly measure DNA strand breaks, they provide strong evidence that AGT/MGMT-mediated repair is essential for preventing the persistence and biological consequences of O6-alkylguanine lesions.
• MGMT depletion increases alkylation-induced DNA strand breaks (direct evidence of essentiality)
  • Roos et al. (2004) exposed human peripheral lymphocytes to methylating agents, N-methyl-N’-nitro-N-nitrosoguanidine (MNNG) or temozolomide, following pharmacological inhibition of MGMT using O6-benzylguanine. In proliferating lymphocytes, MGMT depletion significantly increased DNA DSBs, as indicated by the neutral comet assay and γH2AX phosphorylation. Similar effects were induced by temozolomide. These findings provide direct evidence linking impaired MGMT-mediated repair to increased DNA strand breaks.
  • Carlsson et al. (2025) exposed human HepG2-CYP2E1 cells to N-nitrosodimethylamine (NDMA), a methylating agent that generates O6-methylguanine after metabolic activation. NDMA induced a concentration-dependent increase in O6-methylguanine adducts 2 h post-exposure, with adduct levels persisting and further increasing at 24 h. NDMA also induced strand breaks, as indicated by the alkaline comet assay and increased γH2AX foci formation, and micronuclues formation. Inhibition of MGMT by O6-benzylguanine significantly enhanced these responses compared to NDMA treatment alone. In contrast, MGMT inhibition did not significantly affect DNA strand break formation induced by N-nitrosodiethanolamine (NDELA) or N-nitrosomethylaniline (NMA), consistent with differences in the types of DNA lesions generated by these compounds and indicating that the contribution of MGMT-mediated repair depends on lesion chemistry.
  • These findings provide direct evidence that inadequate MGMT-mediated repair of O6-methylguanine lesions promotes the accumulation of downstream DNA strand breaks and chromosomal damage.

Inadequate mismatch repair

• Incomplete/incorrect mismatch repair (MMR) leads to DNA strand breaks (Peterson-Roth et al., 2005):
  • MLH1 (MMR protein)-deficient and -proficient HCT116 human colon cancer cells were treated with 30µM K₂CrO₄ (DNA crosslinking, Cr adducts, protein-DNA crosslinking, DNA oxidation) for 3, 6, and 12 hours and γH2AX foci (biomarker of DNA DSB) were scored by fluorescence microscopy
  • At 6 and 12 hours, MLH1+ cells had higher percentage of γH2AX foci than MLH1- cells
  • The futile repair model of MMR suggests that strand breaks arise from MMR attempting repeatedly to repair the newly synthesized strand opposite adducts in S and G2 phases; approximately 80% of the γH2AX-positive MLH1+ cells were in G2 phase 12 hours after a 3-hour exposure to 20 µM Cr(VI), while the level was five times lower in MLH1- cells, suggesting that the MMR-induced DSB occurred following DNA synthesis; this supports the futile repair model and demonstrates inadequate repair

Inadequate Repair of DSBs

• Rydberg et al. [2005] exposed GM38 primary human dermal fibroblasts to increasing doses of linear electron transfer (LET) radiation of helium and iron ions (Rydberg et al., 2005).
  • The cells were allowed to recover for 16 hours following irradiation.
  • Unrepaired DSBs were measured after recovery using PFGE.
  • There was a dose-dependent increase in unrepaired DSBs due to both ion exposures.
  • Increase in persistent unrepaired DSBs with increasing dosage indicates exceeded repair capacity.
• DSB repair was also monitored by measuring γH2AX foci 0.05 - 24 hours after irradiation.
  • DSBs decreased over time and less than 1 foci per cell on average remained in MRC-5 cells 24hours after 0.02, 0.2 and 2 Gy exposures.
  • Repair was slower in 180BR cells, particularly for the 2 Gy exposure, where 20 foci per cell remained after 24 h. 
  • A follow-up study by the same group, found similar results for MRC-5 and 180BR cells exposed to 0.02 and 0.2 Gy of X-rays (Kühne et al., 2004). 
• Rothkamm and Löbrich (2003) exposed MRC-5 primary human lung fibroblasts (repair-proficient) and 180BR DNA ligase IV-deficient human fibroblasts to 10 and 80 Gy of X-rays (Rothkamm and Lobrich, 2003).
  • DNA ligase IV deficiency results in impaired NHEJ
  • DSB repair was monitored using PFGE by measuring the % of DSBs remaining after 0.25, 2, and 24 h following irradiation.
  • DSBs decreased over time and, eventually, less than 10% of the DSBs remained in MRC-5 cells after 24h following both 80 and 10 Gy exposures.
  • Repair was noticeably slower in 180BR cells, where the clearance of DSBs was hindered and approximately 40 and 20% of the DSBs remained at 24 hours following 80 and 10 Gy exposures, respectively.
  • The above demonstrates defective DNA repair leading to persistent DSBs.

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

References

Alexander, J., Orr-Weaver, T. (2016), Replication fork instability and the consequences of fork collisions from rereplication, Genes Dev, 30:2241-2252.

Carlsson, M. J., Herzog, N., Felske, C., Ackermann, G., Regier, A., Wittmann, S., Cereijo, R. F., Sturla, S. J., Küpper, J.-H. & Fahrer, J. (2025). The DNA Repair Protein MGMT Protects against the Genotoxicity of N‑Nitrosodimethylamine, but Not N‑Nitrosodiethanolamine and N‑Nitrosomethylaniline, in Human HepG2 Liver Cells with CYP2E1 Expression. Chemical Research in Toxicology, 38(6), 1134–1146. https://doi.org/10.1021/acs.chemrestox.5c00133

Dolan, M. E., Moschel, R. C. & Pegg, A. E. (1990). Depletion of mammalian O6-alkylguanine-DNA alkyltransferase activity by O6-benzylguanine provides a means to evaluate the role of this protein in protection against carcinogenic and therapeutic alkylating agents. Proceedings of the National Academy of Sciences, 87(14), 5368–5372. https://doi.org/10.1073/pnas.87.14.5368

Dolan, M. E., Mitchell, R. B., Mummert, C., Moschel, R. C. & Pegg, A. E. (1991). Effect of O6-benzylguanine analogues on sensitivity of human tumor cells to the cytotoxic effects of alkylating agents. Cancer Research, 51(13), 3367–3372.

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

Dahle, J., Brunborg, G., Svendsrud, D., Stokke, T., Kvam, E. (2008), Overexpression of human OGG1 in mammalian cells decreases ultraviolet A induced mutagenesis, Cancer Lett, 267:18-25.

Dolinnaya, N., Kubareva, E., Romanova, E., Trikin, R., Oretskaya, T. (2013), Thymidine glycol: the effect on DNA molecular structure and enzymatic processing, Biochimie, 95:134-147.

Eccles, L., Lomax, M., O’Neill, P. (2010), Hierarchy of lesion processing governs the repair, double-strand break formation and mutability of three-lesion clustered DNA damage, Nucleic Acids Res, 38:1123-1134.

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Relationship: 3801: Increase, DNA strand breaks leads to Apoptosis

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

This KER applies to eukaryotic cells exposed to genotoxic stressors that induce persistent or unrepaired DNA strand breaks. However, the applicability of this KER is context-dependent and influenced by cell type, developmental stage, DNA repair capacity, and DDR signaling status.

Key Event Relationship Description

DNA strand breaks activate a series of DNA damage response (DDR) pathways that together determine whether damaged cells go through cell cycle arrest, repair, or cell death. When the levels of DNA damage exceed the DNA repair capacity, DDR signaling shifts the cellular response toward elimination of damaged cells by various forms of cell death. Thus, DNA strand breaks are potent inducers of programmed cell death (i.e., apoptosis) when damage is irreparable or sustained.

Evidence Supporting this KER

The empirical support for this KER is strong based on a large body of literature connecting DNA strand breaks and the induction of apoptosis. Both SSBs and DSBs are frequently associated with increased apoptosis in a dose-concordant manner in somatic and germ cells, as well as animal models (Roos and Kaina, 2006). This concordance has been observed following exposures to many DNA-damaging agents; select examples in different tissues and models are provided below.

Evidence in vivo (exposure)

Saha et al. (2014) demonstrated clear temporal and dose concordance between DSB formation and apoptosis in the embryonic mouse brain. Mice were exposed in utero to acute doses of 10, 25, 50, 100 or 200 mGy X-rays, and embryo brains were examined 1 hour or 6 hour later. The number of 53BP1 foci per cell was quantified as a marker of DSB induction, while TUNEL staining (Terminal deoxynucleotidyl UTP Nick End Labeling) was used to monitor apoptosis. Significant increases in 53BP1 foci formation (at 1 hour) and TUNEL+ apoptotic cells (at 6 hours) were observed starting at 10 mGy, indicating that the two KEs occur within the same dose range and that DSBs occur earlier than apoptosis. At 6 hours post-irradiation, while the number of 53BP1 foci at lower doses remained relatively unchanged, it was reduced at50 mGy and returned to the control level at 100 mGy, indicating efficient repair. Despite this apparent resolution of DNA damage, apoptosis continued to increase in a dose-dependent manner, suggesting that even low and transient DSBs are sufficient to induce apoptosis in this sensitive tissue (Saha et al., 2014).

Acetaminophen (200 mg/kg) induces DNA fragmentation and apoptosis in hepatocytes after intraperitoneal administration in mice (Bajt et al., 2011). Liver cells collected from exposed mice showed the release of histone-associated DNA fragments, mitochondrial pro-apoptotic proteins (apoptosis-inducing factor AIF, Smac, endonuclease G, and cytochrome c) into the cytosol at 6 hours post-treatment, accompanied by increased TUNEL+ staining at both 6 and 24 hours. This study also demonstrates the essentiality of mitochondrial pathways through the use of AIF-deficient mice. When treated with acetaminophen, these mice showed a marked reduction in presence of DNA strand breaks (TUNEL assay), % DNA fragmentation, and the expression of apoptotic markers compared to wild-type counterparts. This indicates that AIF is a necessary mediator of DNA fragmentation and apoptosis (Bajt et al., 2011). However, detection of DNA fragments does not allow discrimination between upstream DNA damage and downstream apoptotic DNA fragmentation (see Uncertainties and Inconsistencies). As only a single dose and timepoint were assessed for most markers, the data primarily support co-occurrence.

Evidence in vivo (genetic manipulation)

Using Nestin-Cre mice, Rodrigues et al. (2013) showed that deletion of Nibrin (Nbn, a key sensor of DSBs) and Atm kinase in neural stem/progenitor cells led to DSB accumulation (γH2AX+ cells, p53 stabilization) and concordantly enhanced apoptosis (TUNEL+ cells and cleaved caspase-3 staining) in specific regions of the developing brain (cerebellum, ganglionic eminences) and eye (lens) on embryonic days E15.5 and E17.5. The incidence of γH2AX+ cells (~6,000 to 10,000 cells/mm²) was consistently higher than that of TUNEL+ cells (~250 to 800 cells/mm²) and caspase-3+ cells (~200 to 800 cells/mm²) in the embryonic brains (Figure 1). A similar pattern was observed in eye lens and retina, supporting incidence concordance. Greater impairment of DDR signaling (Nbn/Atm^Nes-Cre double deletion) induced more DSBs and higher apoptosis than Nbn single deletion alone (Nbn^Nes-Cre) (Figure 1). This establishes a graded response-response relationship between DSB formation and apoptosis (Rodrigues et al., 2013). However, in the retina region, apoptosis induced by Nbn deletion was prevented by simultaneous inactivation of ATM, despite the presence of DNA damage. These findings provide direct evidence of essentiality, showing that key signaling pathways downstream of DNA strand breaks are required for the induction of subsequent apoptotic cell death. The observed differences across tissue suggest that these responses are context-dependent and may be mediated by tissue-specific pathways (Rodrigues et al., 2013).

Loss of Apc (adenomatous polyposis coli) in mouse liver tissue led to an early increase in DSBs, as indicated by elevated γH2AX and Rad51 staining at day 4, accompanied by upregulation of DNA damage checkpoint proteins p53 and p21 (Méniel et al., 2015). This was followed by a subsequent increase in apoptotic cells at day 6. Quantitative immunohistochemistry analysis revealed that the incidence of γH2AX-positive and Rad51-positive cells was consistently higher than that of caspase-3-positive apoptotic cells. At day 4, γH2AX-positive cells increased from 0.9% in WT to 26.2% in Apc-deficient mice and 67.1% in Apc/p53 double knockout mice, while Rad51 increased from 2% in WT to 9.9% and 20.7% after genetic manipulation (Figure 2). At the same timepoint, caspase-3-positive cells remained low (~0.36-1.34% across genotypes). A similar pattern was observed at day 6. Additional deletion of p53 further increased markers of DNA damage, indicating that p53 plays an essential role in regulating DSB levels following Apc loss. Together, these findings support incidence concordance and temporal sequence in which DSBs precede activation of checkpoint signaling and subsequent induction of p53-dependent apoptosis.

Xiao et al. (2023) established a myocardial ischemia/reperfusion mouse model to study the role of a long non-coding RNA (cardiac ischemia reperfusion associated Ku70 interacting lncRNA, or CIRKIL) in coronary artery disease. Overexpression of CIRKIL in mouse hearts and 100 μmol/L H₂O₂-treated mouse cardiomyocytes induced parallel increases in DSBs (γH2AX expression), p53 activation, and apoptotic markers, including the ratio of pro-apoptotic Bax and anti-apoptotic Bcl-2 proteins (Bax/Bcl-2), caspase-3 activity, and TUNEL+ cells. Conversely, knockdown of CIRKIL, as well as single knockout or double knockout of CIRKIL and its partner Ku70, alleviated these myocardial injury and reduced markers of both DNA strand breaks and apoptosis. The consistent bidirectional changes in these endpoints by multiple genetic interventions supports the role of the CIRKIL/Ku70 pathway in linking DNA strand breaks to apoptotic responses, and provides indirect evidence of essentiality (Xiao et al., 2023).

Together, the in vivo evidence provides strong empirical support for temporal concordance between strand breaks and apoptosis. Concurrent changes in both KEs were often observed at the same dose level, supporting qualitative dose concordance. A few studies support incidence concordance. In addition, the data highlight that the progression from DNA strand breaks to apoptosis is influenced by tissue-specific sensitivity and mechanisms, which should be considered when evaluating the applicability of this KER.

Evidence in vitro

Roos et al. (2004) provided important mechanistic evidence linking DNA alkylation, impaired repair, DNA strand breaks, and apoptosis in human lymphocytes. O6-methylguanine (O6-MeG) is a key DNA lesion associated with apoptosis, and its removal is mediated by O6-MeG-DNA methyltransferase (MGMT). To prevent repair of O6-MeG, cells were pre-treated with O6-benzylguanine to inactivate MGMT, followed by treatment with O6-MeG-generating agents, N-methyl-N’-nitro-N-nitrosoguanidine (MNNG) or an anticancer drug temozolomide. In MGMT-depleted proliferating lymphocytes, O6-MeG lesions were processed by mismatch repair during DNA replication, leading to the formation of DSBs, as demonstrated by neutral comet assay and γH2AX expression analysis. MNNG (10 or 25 μM) induced DSBs within 3 hours and the level peaked at 24 hours post-exposure. Apoptosis (measured by sub-G1 DNA content analysis or Annexin V/PI staining) was only observed after 24 hours (and persisted up to 96 hours) following MNNG treatment (at multiple concentrations, up to 20 μM), occurring later than DSB formation. Similar effects were induced by temozolomide. Apoptosis was associated with p53 stabilization and upregulation of the Fas receptor, and was significantly reduced (~61%) by treatment with an anti-Fas neutralizing antibody. In contrast, non-proliferating cells showed minimal apoptosis at lower MNNG concentrations (≤ 10 μM), indicating that DNA replication and mismatch repair is required in converting O6-MeG to DSBs. Furthermore, ionizing radiation (5 Gy), which directly induces DSBs, triggered apoptosis in both proliferating and non-proliferating lymphocytes. Together, these data suggest that O6-MeG is converted during DNA replication into DSBs that trigger apoptosis via p53- and Fas-dependent pathways. In addition, MGMT inactivation provides direct evidence of essentiality, as impaired DNA repair enhances the persistence of DNA strand breaks and contributes to apoptosis.

He et al. (2018) studied the effects of loperamide (an antidiarrheal agent) on DNA damage and apoptosis in leukemia cell lines and human primary leukemia cells. Cells exposed to 5, 10, 20 μM loperamide for 24 hours exhibited clear evidence for DNA strand breaks, which were detected by the alkaline comet assay (identifying SSBs and alkali-labile sites) and increased expression of γH2AX. In the same concentration range, a dose-dependent increase in the percentage of apoptotic cells was quantified by Annexin V/7-aminoactinomycin D+ staining and flow cytometry. The induction of apoptosis was further confirmed by increased protein expression of two apoptotic markers, cleaved-caspase-3 and cleaved poly (ADP‑ribose) polymerase (PARP), and decreased expression of an anti‑apoptosis protein Mcl‑1 at 10 μM and 20 μM. Activation of the ATM‑CHK2 pathway following 5‑20 µM loperamide treatment supported the induction of a DDR pathway in leukemia cells. As an example, the effects of loperamide on strand break formation and apoptosis in Molt-4 acute lymphocytic leukemia cells are shown in Figure 3. Similar effects were observed in Thp1 acute myelocytic leukemia cells and primary human leukemia cells (He et al., 2018).

Similarly, exposure to quercetin (40-80 μM) and the positive control paclitaxel (0.45 μM) for 48 hours induced simultaneous strand breaks and apoptosis in MGMT-positive human glioblastoma T98G cells (Wang et al., 2023). DNA strand breaks were assessed using the alkaline and neutral comet assays, and γH2AX staining, capturing both SSBs and DSBs. Apoptosis was assessed by the TUNEL assay, Annexin V-FITC and PI double staining, the ratio of Bax/Bcl-2, and the expression of apoptosis-related proteins, including cleaved caspase-9, cleaved caspase-3, and cleaved PARP. Quercetin (starting at 40 μM) and paclitaxel induced concentration-dependent increases in tail moment (tail length × % tail DNA) in the comet assays and γH2AX expression, indicating DSB formation. At the same concentrations and timepoint, both compounds significantly increased apoptotic cells, accompanied by a higher Bax/Bcl-2 ratio and elevated expression of apoptosis-related markers. Notably, quercetin (≥ 40 μM) also downregulated MGMT expression and activity, suggesting reduced MGMT-mediated DNA repair (Wang et al., 2023). These data support dose and temporalconcordance between DNA strand breaks and apoptosis; the concurrent downregulation of MGMT supports a role of insufficient DNA repair in the accumulation of DNA strand breaks and subsequent apoptosis.

Yauk et al. (2016) studied micronucleus induction and apoptosis in human lymphoblastoid TK6 cells following 4-hour exposures to six genotoxic chemicals: 2-aminoanthracene (0.75-2 μg/mL), acetaminophen (1-10 mM), cyclophosphamide (1-20 μM), dibenz[a,h]anthracene (1-280 μg/mL), dimethylnitrosamine (1-10 mM), furan (2-5 mM) (Figure 4). Micronucleus frequency serves as a marker of chromosomal damage resulting from unrepaired DNA lesions (including DSBs), but it is not a direct measure of DNA strand breaks. Consistent dose-dependent increases in both micronucleus frequency and the number of apoptotic cells were observed 20 hours post-exposure across all tested chemicals (Figure 4). In general, significant increases in apoptosis occurred at slightly lower concentrations than those required to induce micronucleus formation, and fewer cells exhibited chromosome damage than apoptosis, indicating a lack of incidence and only partial dose concordance. These observations likely reflect differences in assay sensitivity and biological progression, as micronucleus formation is a late-stage cytogenetic outcome that requires progression through mitosis. Therefore, these differences do not contradict the overall relationship. As both endpoints were measured at the same time, these data support temporal concordance of this KER.

Liu et al. (2014) showed that inhibition of the PI3K/AKT pathway (a regulator of DDR) by BKM120 led to DNA damage and increased apoptosis in human and murine hepatocellular carcinoma cell lines (Huh7 and BNL). Cells pretreated with 1 μM BKM120 for 1 hour were irradiated (10 Gy for Huh7 cells and 5 Gy for BNL cells). Radiation induced DSB formation (assessed by γH2AX staining) within 30 minutes and the presence of the PI3K inhibitor led to prolonged foci retention (≥ 8 hours in Huh7 cells and ≥ 4 hours in BNL cells). In parallel, PI3K inhibition increased radiation-induced apoptosis, as indicated by elevated cleaved caspase-3 expression and increased Annexin V-positive cells at 24 hours. Co-treatment with rapamycin, an mTOR inhibitor downstream of PI3K, further enhanced both DSB persistence and apoptosis (Liu et al., 2014). These findings demonstrate strong temporal concordance, with DSBs occurring earlier than apoptotic responses; parallel increases were observed across experimental conditions. Although dose-response data are not available, modulation of DNA damage persistence through PI3K/AKT and downstream mTOR inhibition provides indirect essentiality support for the link between DNA strand breaks and apoptosis.

Additional empirical evidence from Chen et al. (2022) supports this KER in human nasopharyngeal carcinoma cells. Two cell lines, SUNE1 and HONE1, were transfected to overexpress ubiquitin-specific protease 44 (USP44). USP44 regulates ubiquitin signaling at DSB sites, thereby modulating DDR and repair processes. Following irradiation (6 Gy), both cell lines exhibited rapid induction of DNA strand breaks, as shown by significant increases in tail moment in the neutral comet assay and higher γH2AX foci numbers within 30 minutes to 4 hours post-exposure. Minimal or no DNA damage was observed at 24 hours, indicating efficient repair at later timepoints. In the same system, USP44 overexpression promoted radiation-induced apoptosis, demonstrated by Annexin V/PI staining at 24 hours in vitro and increased caspase-3 staining in subcutaneous xenograft tumors in mice (Chen et al., 2022). The alignment between early DSB formation and subsequent apoptotic responses supports strong temporal concordance; concurrent increases in both KEs following USP44 overexpression provide indirect essentiality evidence for this KER.

CDKN2AIP is a cell cycle-associated protein that has been implicated in the regulation of DNA damage repair, p53-dependent apoptosis, and spermatogenesis. An in vitro study by Cao et al. (2022) using Cdkn2aip-knockdown TM4 mouse testicular Sertoli cells exposed to γ-irradiation (5 Gy) showed impaired DSB repair, evidenced by persistent γH2AX and 53BP1 foci formation at 2h, 12h, and 24h, together with a significant increase in apoptosis measured by flow cytometry and TUNEL at 24h (Cao et al., 2022). This study provides evidence for temporal concordance and essentiality.

Overall, across multiple somatic cell types, radiation-induced, chemically or genetically induced DNA strand breaks are consistently associated with an increase in apoptosis. These studies demonstrate strong temporal concordance, with DNA strand breaks occurring early or concurrently with apoptotic responses. Evidence for dose concordance is supported in several studies but dose-response data are not always available. Evidence for incidence concordance is limited in vitro. The consistent direction of responses across experimental conditions provide strong empirical support for the relationship between these two KEs in somatic cells.

Evidence in germ cells

A study by Hamer et al., (2003) provided evidence linking DSBs and apoptotic responses in male germ cells. They first demonstrated the presence of basal γH2AX in testes of wild-type FvB/NAU mice. Following a single ionizing radiation (4 Gy), γH2AX foci were rapidly (within 3 hours) induced in spermatogonia, spermatocytes, and round spermatids. In spermatogonia, γH2AX and p53 colocalized in the nucleus and their interaction was enhanced following irradiation. These findings support co-occurrence of DSBs and p53-dependent apoptosis in spermatogonia.

Habas et al. reported the effects of known mutagens on testicular germ cells isolated from 10-12-week-old NMRI mice (National Medical Research Institute) or male Sprague-Dawley rats. Isolated testicular germ cells (spermatogonia, spermatocytes and spermatids) were exposed for 1 hour to one of seven genotoxic compounds at 50, 500, or 1000 μM, and were immediately subjected to alkaline comet or TUNEL analyses. The compounds included doxorubicin (Dox), which induces DNA strand breaks via topoisomerase II inhibition (Habas et al., 2017), and a panel of mutagens targeting germ cells at specific stages: alkylating agents N-ethyl-N-nitrosourea (ENU) and N-ethyl-N-nitrosourea (MNU), which target pre-meiotic spermatogonia; 6-mercaptopurine (6-MP) and 5-bromo-20-deoxy-uridine (5-BrdU), which produce the greatest responses in early meiotic spermatocytes; and methyl methanesulphonate (MMS) and ethyl methanesulphonate (EMS) that primarily target post-meiotic germ cells (Habas et al., 2016). Representative results (Figure 5) show highly consistent dose-dependent increases in both % tail DNA in the alkaline comet assay and the percentage of TUNEL+ apoptotic cells across male germ cells at all stages, with both effects being statistically significant at the same exposure concentrations (Habas et al., 2016, 2017). These data support dose and temporal concordance while extending the applicability of this KER to male germ cells.

Cao et al. (2022) demonstrated that loss of CDKN2AIP leads to a concurrent increase in DNA strand breaks and apoptosis in male germ cells. CDKN2AIP preferentially expresses in spermatocyte and spermatids and acts as a central regulator of DSBs repair. Analyses of Cdkn2aip-/- male C57BL/6 mice revealed a significant accumulation of DSBs (γH2AX analyses) in pachytene spermatocytes from postnatal day P35 testes, along with a higher incidence of TUNEL foci per cells in the seminiferous tubules (Cao et al., 2022). These data indicate a clear association between increased DSBs and male germ cell apoptosis, and provide evidence for the essential role of DNA damage repair in preventing DSB accumulation and subsequent apoptotic cell death.

Suh et al. (2006) examined oocytes from Balb/c mice exposed to γ-irradiation (0, 0.1, 0.45 Gy) on postnatal day 5. At 24 hours post-exposure, on average, oocytes exposed to 0.1 Gy had an average of three γH2AX-marked DSBs with minimal apoptosis; oocytes receiving 0.45 Gy had about ten DSBs and underwent apoptosis within 24 hours, as indicated by condensed chromatin and positive TUNEL staining (Suh et al., 2006). Consistent with early DNA damage induction, Stringer et al. (2020) reported dose- and time-dependent increases in nuclear localization of phosphorylated ATM in mouse oocytes as early as 30 minutes following irradiation (0.2, 0.45, 7 Gy), indicating activation of DNA damage responses. The presence of DSBs was evidenced by the formation of γH2AX foci at all doses. Apoptosis was indirectly monitored by follicle survival at 24 hours and oocyte retrieval rate. At the same timepoints and dose range, Rad51 (a marker of homologous recombination) localized to DNA damage sites in over 90% of oocytes in primordial follicles, whereas DNA-dependent protein kinase catalytic subunit (DNA-PKcs, a marker of non-homologous end joining) was mostly present in granulosa cells; this difference suggests cell type-specific utilization of repair pathways. Importantly, intervention studies in this system provide indirect evidence of essentiality. In Tap63 knockout mice (deficient in DNA damage-induced apoptotic signaling), primordial follicles were preserved despite the presence of DNA damage. Conversely, pharmacological inhibition of Rad51 increased persistence of unrepaired DNA damage and apoptosis, suggesting that efficient DNA repair mitigated the progression from DNA strand breaks to cell death.

Experimental evidence from isolated germ cells demonstrates that DNA strand breaks are closely associated with increased apoptosis across multiple stages of spermatogenesis and in oocytes. These findings support the applicability of this KER to germ cells and reinforce dose and temporal concordance.

The biological plausibility of this KER is strong and supported by extensive understanding of the DDR pathways (Jackson and Bartek, 2009). The mechanistic processes that link DNA strand breaks to apoptosis are well characterized and detailed in many excellent reviews including Kaina (2003), Roos and Kaina (2006), and Zio et al. (2012). Both p53-dependent (encoded by Tp53) and p53-independent mechanisms contribute to DNA strand break-induced apoptosis (Roos and Kaina, 2006). It is important to note that activation of p53 and other DDR pathways does not immediately trigger apoptosis. Rather, the cellular response depends on the extent and persistence of DNA damage. In many cases, initial DDR activation promotes cell cycle arrest and DNA repair (reviewed by Williams and Schumacher, 2016). When DNA damage is extensive, irreparable, or persistent, the DDR signaling cascade shifts to favour programmed cell death, in part through the action of p53 and other stress sensors (Maréchal and Zou, 2013). This conditional response reflects a protective mechanism that preserves genomic integrity and prevents the propagation of cells with damaged DNA.

Sensing of DNA strand breaks by serine/threonine kinases

The serine/threonine kinases, ATM (ataxia telangiectasia mutated) and ATR (ATM- and Rad3-Related), are key sensors and mediators of cellular responses to DNA strand breaks in cells (Maréchal and Zou, 2013).

ATM is predominantly activated by double strand breaks (DSBs). The MRN (Mre11- Rad50-Nbs1) complexrecognizes DSB ends and recruits ATM to the damage sites (Maréchal and Zou, 2013). In undamaged cells, ATM exists predominantly as inactive dimers/oligomers; in the presence of DSBs, ATM undergoes autophosphorylation anddissociates into active monomers that phosphorylate a series of proteins involved in cell cycle arrest, DNA repair, and apoptosis, such as CHK2, Brca1, H2AX, and p53 (Lee, 2005). Of these, phosphorylation of histone H2AX at serine 139 (γH2AX) is one of the earliest cellular responses to DSBs. γH2AX foci form rapidly (within minutes) at sites of damage and are widely recognized as a sensitive marker of DNA damage following chemical exposure in both somatic cells and germ cells (Hamer et al., 2003). In the testis, basal γH2AX signaling is observed across multiple stages of germ cells and is associated with p53-mediated apoptosis signaling in spermatogonia following stress (Hamer et al., 2003).

ATR kinase, on the other hand, responds to a broader spectrum of DNA damage. ATR is activated by replication protein A (RPA)-coated single-stranded DNA (ssDNA), which arises during replication stress or as intermediates in DNA repair (Maréchal and Zou, 2013). ATR activation requires its binding partner, ATRIP (ATR-interacting protein), which recognizes RPA-coated ssDNA and localizes ATR to the damage sites (Maréchal and Zou, 2013). Both single strand breaks (SSBs) and DSBs (when resected by nucleases) can generate ssDNA and thus activate ATR (Maréchal and Zou, 2013).

p53-dependent DNA strand break-triggered apoptosis

Both ATM and ATR kinases phosphorylate and activate p53 transcription factor, a master regulator of apoptosis.ATM primarily phosphorylates p53 at serine-15, whereas ATR targets both serine-15 and serine-37 (Tibbetts et al., 1999).These partially overlapping activities ensure p53 activation during genotoxic stress. Once p53 is activated by ATM/ATR, it promotes the transcription of genes that favour apoptosis over survival, particularly when DNA damage is persistent or irreparable. To accomplish this, p53 activates pro-apoptotic genes (e.g., Puma, Bax, Apaf-1, Noxa), while inactivating anti-apoptotic factors such as members in the Bcl-2 family. These pro-apoptotic factors increase the permeability of the mitochondrial membrane and signal the release of cytochrome c (Wawryk-Gawda et al., 1998). Cytochrome c then binds Apf-1 adaptor and procaspase-9, aggregating the proteins and activating caspase-9, which in turn activates caspase-3 to induce apoptosis (Elmore, 2007).

p53-independent DNA strand break-triggered apoptosis

Cells can also respond to DNA strand breaks and undergo apoptosis through several p53-independent mechanisms (Roos and Kaina, 2006). DNA strand breaks induce ATM/ATR-dependent phosphorylation of checkpoint proteins CHK1 or CHK2, which can trigger apoptosis independently of p53 by stimulating E2F1-dependent expression of pro-apoptotic factor p73 (Urist et al., 2004). A newly identified p53-independent apoptosis pathway is triggered by DNA strand breaks through ribosome stalling mediated by Schlafen 11 (SLFN11, a tRNase). Upon DNA damage, SLFN11 causes ribosome stalling and global translation inhibition. The stalled ribosomes are recognized by ZAKα, a ribosome-associated stress sensor, which then initiates a MAPK signaling cascade that directly activates the mitochondrial apoptosis machinery, independent of the canonical p53 pathway (Boon et al., 2024). Other factors may be indirectly activated by DNA strand breaks and contribute to p53-independent apoptosis, such as pathways involving Bcl-2 degradation, caspase-2 activation, and NF-κB dependent Fas ligand transcription (Roos and Kaina, 2006).

In the female germ line, p63, a homologue of p53, acts in a conserved mechanism to preserve genomic fidelity. Specifically, a p63 isoform TAp63 is essential in the process of DNA damage-induced oocyte death not involving p53 (Suh et al., 2006). TAp63 is constitutively expressed in oocytes during meiotic arrest. In response to DNA damage, TAp63 is phosphorylated and binds to p53 cognate DNA sites and results in transcriptional activation of apoptosis pathways in female germ line (Suh et al., 2006).

A primary uncertainty in this KER arises from the use of the TUNEL assay, which can detect both the primary DNA strand breaks and the secondary DNA fragmentation that occurs during late-stage apoptosis, necrosis, or severe oxidative stress. This overlap makes it difficult to distinguish whether the DNA damage is an initiating event or a downstream consequence, particularly when both KEs are measured at the same timepoint (Bajt et al., 2011). For stressors like acetaminophen that can induce extensive necrosis, interpreting the TUNEL signal as a specific marker of apoptosis may confound the causal interpretation of this KER. To reduce this uncertainty, we recommend studies using multiple timepoints to establish temporal concordance, and using direct DNA damage assays (e.g., comet assays, γH2AX staining) and apoptosis specific markers (e.g., caspase activation or Annexin V staining) to strengthen confidence in the conclusions.

This KER is context-specific. Factors like developmental stage, activity of the repair pathways, and cellular sensitivity can modulate this KER. During ovarian reserve formation, meiotic germ cells naturally accumulate high levels of programmed DSBs that are efficiently repaired. In female FVB/N mice, Zhou et al. (2021) reported increased levels of DSBs from embryonic day 13.5 to postnatal day 2, with 23.94% to 83.76% germ cells stained positive for γH2AX. Despite the high incidence of DSBs, no detectable germ cell apoptosis was observed based on caspase-3 and TUNEL staining. A similar phenomenon is observed in the male germ line. During spermatogenesis, programmed DSBs occur during meiosis and chromatin remodeling processes (e.g., histone-to-protamine replacement during spermiogenesis) (Talibova et al., 2022) . Together, these observations indicate that the apoptotic response is not determined by the presence of DSBs per se, but their persistence and repair failure are critical modifiers.

Moreover, some studies report apoptosis in the absence of detectable DNA strand breaks. For example, in mice exposed daily to furan for 28 days, no treatment-related induction of DNA strand breaks or DNA cross links was noted in liver cells, as measured by the alkaline comet assay and γH2AX foci formation at 24 hours after last administration (Cordelli et al., 2010). However, a significant increase in TUNEL+ apoptotic cells was observed at the highest dose. While this suggests that apoptosis can arise through alternative pathways, the absence of detectable DNA damage may reflect limitations in the sampling time and assay sensitivity, as DNA lesions measured by the comet assay can be transient and rapidly repaired.

References

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Relationship: 3802: Apoptosis leads to Decrease, Sperm count

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

This KER is primarily supported by in vivo mammalian studies, particularly in rodents. The KER is most applicable when apoptosis is measured in early spermatogenesis, as apoptosis in late-stage germ cells plausibly affects sperm quality rather than sperm quantity. The sperm counts should be assessed following a delay consistent with the time required for the affected cells to progress through spermatogenesis. While the underlying biological processes are conserved across eukaryotic species that produce sperm, quantitative relationships may vary depending on species-specific spermatogenic kinetics.

Key Event Relationship Description

Apoptosis in germ cells directly depletes the developing germ cell population, whereas apoptosis in Sertoli cells and Leydig cells can indirectly reduce sperm counts by disrupting structural support and endocrine signaling in the testis that is required to sustain spermatogenesis.

Evidence Supporting this KER

The biological plausibility of this KER is strong, based on extensive mechanistic understanding of spermatogenesis and the role of apoptosis in regulating testicular cell populations.

Spermatogenesis is a continuous and highly coordinated process of germ cell proliferation and differentiation, leading to the production of mature spermatozoa. Apoptosis (programmed cell death) is an important mechanism that removes damaged cells and maintains tissue homeostasis (Shaha et al., 2010). During spermatogenesis, physiologic apoptosis occurs in all testicular cells (Oldereid et al., 2001). Meanwhile, stress-induced apoptosis can be triggered by exogenous stressors, such as environmental toxicants, heat, and radiation, which can markedly increase apoptotic cell death in testis (Shaha et al., 2010).

Mechanistically, the primary link between apoptosis and reduced sperm counts is the loss of developing male germ cells, which decreases the number of cells available to complete spermatogenesis and produce mature spermatozoa. However, apoptosis of testicular Sertoli and Leydig cells can also contribute to reduced sperm counts through indirectmechanisms, such as disrupting structural support and hormonal regulation of spermatogenesis.

Apoptosis in male germ cells

In the early stage of spermatogenesis, spermatogonial stem cells (SSCs) maintain the male germline through self-renewal, while a subset of SSCs commits to differentiation to produce mature sperm (Nakagawa et al., 2010). As long-term germ cell reservoirs, SSCs have enhanced DNA repair capacity, which renders them less sensitive to damage or stress-induced apoptosis compared with their differentiating progeny (Meistrich et al., 1982b; Rübe et al., 2011). However, excessive apoptosis of SSCs at higher doses can compromise the ability of the testis to replenish the pool of developing germ cells and cause sustained or irreversible impairment of spermatogenesis (Meistrich et al., 2013).

Differentiating spermatogonia are particularly sensitive to stress-induced apoptosis because they undergo rapid mitotic divisions (Meistrich et al., 1982a, 1982b, 2013). Apoptosis of differentiating spermatogonia results in a progressive loss of the more mature germ cells in a process known as maturation depletion (Meistrich et al., 2013). Although later-stage germ cells continue to develop, minimal replacement occurs from progenitor cells. Consequently, the pool of developing germ cells is depleted over time, leading to reductions in sperm counts that become evident after a delay. This delay reflects the time required for developing germ cells to complete spermatogenesis before the effects of progenitor cell depletion become apparent in mature sperm output. As a result, the timing of the decline in sperm counts is generally consistent with the duration of spermatogenesis in the species of interest (Meistrich et al., 1992).

Primary spermatocytes undergo extensive meiotic reorganization and division to generate haploid spermatids. Disruption of meiotic progression induces checkpoint mediated apoptosis to eliminate defective cells, particularly at the pachytene stage (Roeder and Bailis, 2000; Li et al., 2025). Compared with pre-meiotic germ cells, apoptosis in spermatocytes is often detected during meiotic progression rather than as an immediate response following exposure (Meistrich et al., 1982b, 2013). The subsequent reduction in sperm counts is usually temporary when SSCs survive, as these remaining stem cells rapidly replenish spermatogenic cells and restore sperm counts in ~45 days in mice and ~12 weeks in humans (Meistrich et al., 2013). 

After meiosis, spermatids differentiate into mature spermatozoa during spermiogenesis. At this stage, male germ cells undergo chromatin and morphological remodeling when the sperm genome becomes highly condensed and transcriptionally silenced (Aitken et al., 2010). In parallel, mitochondria are relocalized, either becoming confined to the sperm midpiece or transferred into residual bodies for degradation and elimination (Varuzhanyan and Chan, 2020). These structural changes limit the execution of canonical caspase-dependent apoptotic pathways. Instead, post-meiotic germ cells undergo truncated or abortive apoptosis, which is mediated primarily by reactive oxygen species and lipid peroxidation; the formation of lipid adducts leads to a rapid loss of sperm motility within hours, reflecting immediate functional loss (Aitken and Baker, 2013). Notably, these cells remain viable, and this form of apoptosis does not typically result in immediate DNA fragmentation and will not be detected by the TUNEL assay (Aitken and Baker, 2013). Accordingly, compared with apoptosis in early-stage germ cells, apoptosis occurring at post-meiotic stages is less likely to cause substantial germ cell loss and reduction in sperm counts despite causing functional defects.

Apoptosis in other testicular cells

In addition to germ cells, apoptosis can also happen in other testicular cells and indirectly influence the progression of spermatogenesis. Sertoli cells provide essential structural support in the seminiferous epithelium and play multiple roles in maintaining testicular functions, including supporting germ cell development, facilitating germ cell movement across the epithelium, enabling spermiation (release of mature spermatids), and secreting regulatory factors (Murphy and Richburg, 2015). As one Sertoli cell only has a finite supportive capacity, the size of Sertoli cell population is a key determinant of the total germ cell numbers and, indirectly, Leydig cell numbers in the testis (Rebourcet et al., 2017). Under physiological conditions, spontaneous apoptosis is an important mechanism to maintain the Sertoli cell to germ cell ratio. Toxicant-induced Sertoli cell apoptosis has been reported to cause germ cell loss through loss of structural and metabolic support, ultimately leading to reduced sperm output (Murphy and Richburg, 2015). Among germ cell populations, the developing spermatocytes are the most sensitive to Sertoli cell-selective apoptosis, while spermatogonia are relatively resistant (Murphy and Richburg, 2015). Leydig cells are responsible for testosterone synthesis, which is essential for multiple key processes during spermatogenesis, including germ cell survival and progression (Smith and Walker, 2014). Hormonal deprivation significantly increases the incidence of apoptotic germ cells, particularly late-stage spermatocytes that are androgen-dependent (Hikim et al., 1997). Excessive apoptosis in Leydig cells is strongly linked to lower intratesticular testosterone levels, and therefore can impair germ cell survival, indirectly leading to reduced sperm counts.

Together, these well-established mechanisms provide strong biological support for a causal relationship between increased apoptosis in testicular cells and reduced sperm counts.

The empirical support for this KER is strong. Concordant increases in apoptosis in testicular cells and reductions in sperm counts have been consistently observed across multiple in vivo rodent studies, and numerous intervention studies provide evidence of essentiality by demonstrating preservation or recovery of sperm counts following attenuation of apoptotic signaling. Temporal concordance is strongly supported by the well-established kinetics of spermatogenesis, although it is often inferred rather than directly measured within individual studies. Quantitative dose-response information is limited because many studies employ single exposure or intervention conditions. In addition, apoptosis frequently co-occurs with oxidative stress and inflammatory responses, which may contribute to spermatogenic impairment and complicate attribution to apoptosis alone. Nevertheless, these limitations do not detract from the overall consistency, biological coherence, and weight of evidence supporting this KER.

A large number of studies demonstrated the use of chemicals with anti-apoptotic, anti-inflammatory, and anti-oxidant properties to attenuate toxicant-induced testicular injury. These studies consistently show that attenuation of apoptosis is associated with recovery of sperm counts, providing supportive, albeit indirect, evidence for the essentiality of apoptosis. Since this is a data-rich field and the biological plausibility is strong, select empirical studies are presented as examples.

In a study by Yaman et al (2018), 30 day-old male Wistar albino rats were exposed to a single intraperitoneal dose of 5 mg/kg cisplatin. Half of the rats were euthanized at 31 days of age (pre-pubertal) and observed for the presence of testicular cell apoptosis; the other half was euthanized at 90 days (fertile age) to evaluate epididymal sperm counts. In exposed pre-pubertal rats, 107 ± 5.9 apoptotic (TUNEL-positive) cells were observed in 100 seminiferous tubule cross-sections, compared with only 24.6 ± 6.1 cells in control rats (p<0.001). The TUNEL+ cells were predominantlyspermatogonia and spermatocytes. Consistent with the expected temporal sequent, the average sperm counts measuredlater in adulthood were significantly reduced by ~54% (175 ± 67.1 million/mL in treated rats compared to 379.2 ± 45.9 million/mL observed in the controls) (Figure 1). Moreover, co-treatment with L-carnitine (250 mg/kg, for three consecutive days) significantly decreased the numbers of TUNEL+ cells in the tubules and resulted in a near-complete restoration of sperm counts (Figure 1). This study provides strong evidence of temporal concordance and indirect evidence of essentiality, as modulation of apoptosis was associated with recovery of sperm production.

Udefa et al. (2020) reported preventive effects of a tigernut plant extract on lead acetate-induced testicular damage in rats. Male Wistar rats were injected intraperitoneally with 20 mg/kg lead acetate once daily, with or without oral co-treatment with the extract (500 or 1000 mg/kg) for three weeks. Lead exposure significantly increased apoptosis in the testes, evidenced by pro-apoptotic shift in the Bax/Bcl-2 ratio (~7-fold increase) and elevated cleaved caspase-3 levels (~2.3-fold increase), indicating activation of pro-apoptotic pathways. Concordant decreases in serum and testicular testosterone levels suggested toxicity to Leydig cells. These changes were accompanied by a ~32% reduction in epididymal sperm counts. Co-treatment with the plant extract reversed changes in apoptotic markers and restored sperm counts in a dose-dependent manner. A negative correlation (r= -0.813, p<0.05) between the Bax/Bcl-2 ratio and sperm counts was reported, based on a small number of treatment group-level data. Since the Bax/Bcl-2 ratio is an indirect marker of apoptotic signaling, this correlation provides only limited quantitative support for a direct response-response relationship between the two KEs.

Oyovwi et al. (2023) investigated whether quercetin, a bioflavanol known for anti-oxidant and anti-apoptotic properties, could mitigate testicular dysfunction as a side effect of the antiepileptic drug levetiracetam (LEV). Male Wistar rats (10-12 weeks old) were exposed to saline or LEV (300 mg/kg/day) by oral gavage for 56 days, a period that covers a full cycle of spermatogenesis in rats. A separate group of rats were co-treated with quercetin (20 mg/kg/day) during days 28-56, administered 30-minute after each LEV dose. LEV exposure altered apoptosis-related factors in testicular tissues, including decreased expression of anti-apoptotic protein Bcl-2, and increased pro-apoptotic signaling. Specifically, the cytochrome c level in mitochondrial supernatant was increased by ~60%, while expression of caspase-3 and p53, and sperm DNA fragmentation index (indirectly predicted by aniline blue staining) showed about 2 to 3-fold increases relative to controls (Figure 2). These changes were accompanied by lower testicular weights, a ~30% decrease in epididymal sperm counts, and histological evidence of germ cell loss at all stages, along with decreases in Sertoli cell and Leydig cell numbers. Co-treatment with quercetin reversed all changes in apoptotic markers to control levels and prevented the LEV-induced reductions in testicular weights and sperm counts. This study provides high-quality evidence supporting essentiality of apoptosis in male germ cells.

In a mouse study examining the effects of X-ray radiation on male infertility, selenium nanoparticles (SeNPs) and a probiotic Lactobacillus casei were shown to mitigate testicular damage through anti-apoptotic and anti-oxidant mechanisms (Ehghaghi et al., 2022). Sixty-four Syrian male mice were divided into eight groups (n=8 per group) and orally treated with vehicle (PBS), SeNPs (0.2 mg/kg/day), probiotic (1 × 10⁸ CPU), or their combination. Four groups received whole body X-ray radiation (2 Gray for 5 minutes) after each daily oral dosing for 30 days, and the other four groups were used as non-irradiated controls. Apoptosis of testicular cells was assessed using Annexin V and propidium iodide (PI) double staining and flow cytometry. In the absence of irradiation, none of the groups exhibited significant changes in testicular parameters (Figure 3). X-ray exposure caused a ~20-fold increase in late apoptotic testicular cells, alongside a ~65% decrease in spermatogonia cell numbers and > 50% decrease in testicular sperm counts (Figure 3). Treatment with either SeNPs or the probiotic reduced apoptotic signaling, as indicated by downregulation of caspase-3 and caspase-9 transcripts. These changes were accompanied by partial restoration of spermatogonia and sperm counts. The combined treatment restored both apoptotic markers and downstream endpoints to near-control levels.

In addition to the select studies presented above, multiple rodent studies have reported chemo-preventive effects of anti-apoptotic interventions and corresponding recovery of impaired spermatogenesis caused by a wide range of stressors, including high-fat diet (Abdulwahab et al., 2021), nanomaterials (reviewed by Sun et al., 2025), and toxicants such as aflatoxin B1 (Ijaz et al., 2023), tartrazine (Essawy et al., 2024), atrazine (reviewed by Abarikwu et al., 2023), suggesting a broad applicability of this KER. Mitigation of testicular cells apoptosis is consistently concordant with preservation of sperm counts.

Together, these findings demonstrate concordant changes in apoptosis and sperm counts across treatment groups and provide indirect evidence of essentiality, as attenuation of apoptosis (and other stress responses) was associated with recovery of sperm production. Although evidence for dose-response relationships is limited, the studies generally support dose and temporal alignment and a moderate empirical relationship for this KER.

A key uncertainty arises from the use of whole testis homogenates to measure apoptotic markers, which reflect mixed cell populations and do not distinguish responses of germ cells and somatic cells. Histological evaluation of seminiferous tubules supports germ cell loss as a primary driver of reduced sperm counts (Yaman et al., 2018; Udefa et al, 2020; Ehghaghi et al., 2022; Oyovwi et al., 2023; Ijaz et al., 2023). However, reductions in Sertoli cell and Leydig cell numbers can occur in parallel (Oyovwi et al., 2023). Therefore, the reduced sperm output is likely caused by combined effects on mixed populations of testicular cells and cumulative damages over the course of spermatogenesis, although the relative contribution of each cell type is not clear.

An apparently inconsistent study was noted. In a rat study, exposure to 10 mg/kg of sodium arsenite for 14 days induced a near 100% increase in the mRNA expression of Bax and caspase-3, and a ~40% decrease in Bcl-2 expression in testicular tissues, indicating activation of apoptotic signaling. Although histological analysis showed remarkable loss of spermatogenic cells, particularly spermatogonia, no changes were observed in testicular weights or epididymal sperm counts. Suppression of apoptotic signaling by carvacrol treatment did not alter sperm density across groups (Gur et al., 2023). The inconsistency likely reflects a limitation of the study design rather than a lack of biological linkage between apoptosis and sperm counts. Samples were collected immediately following the 14-day exposure period , which is insufficient to observe downstream changes in sperm output given the duration of spermatogenesis in rats.

Multiple biological pathways can influence sperm output and introduce variability and uncertainty in this KER.Inflammation and oxidative stress frequently co-occur with apoptosis, and therefore reductions in sperm output may reflect overlapping or combined effects of these stress-related responses. In many studies, attenuation of apoptosis is often accompanied by simultaneous suppression of oxidative stress and inflammatory markers (Udefa et al., 2020; Oyovwi et al, 2023). Given the extensive interplay among stress responsive pathways, it is difficult to separate their individual contributions. In addition, disruption of hormonal regulation in the hypothalamic-pituitary-gonadal axis can indirectly affect spermatogenesis and sperm production. Such interacting mechanisms may partly explain inconsistencies observed across studies and should be considered when interpreting this KER.

References

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Relationship: 3771: Alkylation, DNA leads to Decrease, Sperm count

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

This KER is supported primarily by mammalian data (rodents and humans) and is applicable to other eukaryotic species that produce sperm. DNA adducts can occur in any cell type, but this KER is specific to the male germline. Although reduced sperm count can only be measured after sexual maturity, DNA alkylation may occur during fetal, juvenile, or adult life stages. Exposure during early development can impair germ cell populations and lead to delayed reductions in sperm production that become evident after reproductive maturation.

Key Event Relationship Description

DNA alkylating agents can form covalent adducts in the DNA of any exposed cell types; however, actively proliferating germ cells, particularly spermatogonia, are especially susceptible to the cytotoxic consequences of alkylation damage. During DNA replication, alkyl adducts can interfere with replication fork progression and activate DNA damage response pathways. Persistent or unrepaired lesions can result in cell cycle arrest and apoptosis, leading to depletion of differentiating germ cell populations and, consequently, reduced sperm production.

Evidence Supporting this KER

The biological plausibility of this KER is strong. DNA alkylation refers to the covalent addition of alkyl groups (e.g., methyl, ethyl, propyl) to nucleotides at noncanonical positions in DNA, generating lesions that interfere with DNA replication, transcription, and chromosome integrity (Soll et al., 2017). These lesions can occur across all stages of spermatogenesis and trigger downstream signalling cascades that lead to apoptosis. Across species, actively proliferating cells have long been shown to be particularly susceptible to chemotherapeutic alkylating drugs due to ongoing DNA replication; this provides a conserved biological basis for alkylation-induced impairment of spermatogenesis (Drewinko et al., 1981; Meistrich, 1982a).

Unrepaired alkylation damage in male germ cells can block DNA replication and induce base mispairing and strand breaks, which activate DNA damage response pathways and lead to cell cycle arrest and selective elimination of damaged cells through programmed cell death (Kaina, 2003). This stringent quality control mechanism is important to preserve germline genomic integrity; however, it inevitably reduces the supply of cells progressing through spermatogenesis (Agarwal et al., 2020; Li et al., 2025). Therefore, DNA alkylation-induced apoptosis in germ cells can plausibly lead to reduced sperm counts.

The consequences of alkylation-induced apoptosis on sperm counts vary depending on the developmental stage at which the damage occurs. Alkylation of DNA in proliferating or differentiating germ cells at early (pre-meiotic) stages compromises the ability of the testis to replenish the pool of developing germ cells and can cause sustained or irreversible reductions in sperm counts (Meistrich et al., 2013).

Meiosis gives rise to spermatocytes, which have tightly regulated meiotic DNA damage checkpoints to coordinate repair of endogenous programmed DNA strand breaks (Hunter, 2015). Alkylation-induced DNA lesions in meiotic cells can increase the overall repair burden and interfere with the tightly regulated repair of programmed meiotic breaks. When lesions persist, checkpoint signaling promotes the elimination of defective meiotic cells (Roeder and Bailis, 2000; Li et al., 2025). However, this effect is typically temporary because surviving SSCs can repopulate the seminiferous epithelium over time (Meistrich, 1982a).

In contrast, post-meiotic germ cells in late spermatogenesis are largely DNA repair deficient; thus, DNA damage at this stage can only be repaired after fertilization through maternal repair mechanisms (Olsen et al., 2005; Newman et al., 2021). Alkylation of DNA in post-meiotic germ cells does not typically drive germ cell death (and thus reduced sperm counts) contributing instead to sperm DNA damage and reduced sperm quality.

In addition to germ cells, alkylation damage in testicular Sertoli and Leydig cells may also indirectly contribute to reduced sperm counts through disruption of structural support and hormonal regulation required for spermatogenesis. However, differentiating germ cells are considered particularly sensitive targets of alkylating agents due to ongoing DNA replication, and are likely the primary drivers of alkylation-induced impairment of sperm production.

In addition to impacts on adult spermatogenesis, clinical and experimental studies demonstrate that exposure to alkylating chemotherapeutic agents during prepubertal or adolescent ages can impair germ cell populations and lead to persistent reductions in sperm production detectable in adulthood (reviewed by Delessard et al., 2019). Alterations in Sertoli cells and Leydig cells have also been reported following chemotherapy exposure, albeit the effects are inconsistent across studies and do not yet allow definitive conclusions (Delessard et al., 2019). These observations in cancer survivors and experimental models support the relevance of early-life stages and the delayed manifestation of impaired spermatogenesis following chemotherapy.

Together, these well-established biological processes support strong plausibility for a causal relationship between DNA alkylation damage and reduced sperm counts. While alkylation damage can occur across all stages of spermatogenesis, effects on sperm counts are primarily driven by damage to proliferating and meiotic germ cells and have typically been observed following exposure in sexually mature individuals. However, exposures during early developmental stages may result in more persistent or severe impairments due to disruption of the germ cell pool, with effects becoming apparent later in adulthood.

Strong empirical evidence is available from experimental animal models and human clinical observations. Although direct measurement of both DNA adducts and sperm counts in the same study is limited, a substantial body of evidence demonstrates consistent patterns of temporal and dose concordance across studies examining well-characterized DNA alkylating agents. Human data further support the biological relevance of this relationship.

Altakroni et al. (2021) analyzed 105 semen samples from men undergoing assisted reproduction treatment (ART)and quantified the levels of N7-methyldeoxyguanosine (N7-MedG), a biomarker of exposure to methylating agents, in sperm genomic DNA using an immunoslotblot assay. A negative correlation was noted between N7-MedG concentrations and sperm concentration (Spearman's rank correlation coefficient= -0.32, p<0.001). Logistic regression analyses demonstrated that the elevated N7-MedG level was significantly associated with low sperm concentration (odds ratio [OR] = 6.46, 95% CI: 1.51-27.7); this association remained strong after adjustment for other DNA damage markers, including the percentage of sperm with high DNA damage and median % tail DNA measured by the neutral comet assay (adjusted OR = 6.42, 95% CI: 1.47-28.0) (Altakroni et al., 2021). This study provides evidence linking DNA alkylation to reduced sperm counts in humans.

It is well characterized that various DNA adducts rapidly form in mouse male germ cells within two hours after exposure to alkylating agents: N-ethyl-N-nitrosourea (ENU), ethyl methanesulfonate (EMS), and diethyl sulfate (DES). The frequency of adduct formation increases with dose, supporting dose-dependent activation of the upstream KE. Data for ENU exposure are shown as examples (Figure 1) (van Zeeland et al., 1990).

Accordingly, DNA adduct formation can be reasonably inferred following exposures to well-characterized DNA alkylating agents. Empirical evidence supporting this KER primally comes from studies in animals and humans. Below, we summarize human clinical observations and selected data on several alkylating medications as examples to illustrate consistent patterns of reduced sperm counts following exposure.

Evidence in human

Azoospermia (absence of sperm in semen, < 0.1 million cells/mL) and oligozoospermia (semen with a low sperm count, 0.1-10 million cells/mL) are well-recognized side effects of cancer treatment with alkylating agents in men of reproductive age (reviewed by Howell and Shalet, 2005; Okada and Fujisawa, 2018). Dramatic declines in ejaculated sperm counts can occur within 1-2 months after receiving the treatment, while azoospermia usually occur after 2 months (Meistrich et al., 2013). The duration and reversibility of treatment-induced azoospermia is largely dependent on the dose of the drugs administered (Okada and Fujisawa, 2018).

Clinical studies provide evidence linking treatment with alkylating agents to reduced sperm counts in humans. More than 50% of patients with malignant lymphomas developed azoospermia after receiving chlorambucil treatment at a cumulative dose of more than 400 mg (Richter et al., 1970; Cheviakoff et al., 1973). Similarly, in 35 patients treated with cyclophosphamide, sperm production in 22 patients declined to azoospermic levels, and 6 declined to oligozoospermic levels within a few months after treatment initiation (Meistrich et al., 1992). In this cohort, patients received less than 7.5 g/m² (median dose 4.1 g/m²) of cyclophosphamide restored sperm production to a normal range within 5 years, whereas only 10% of patients recovered when treated with higher doses (> 7.5 g/m²), likely reflecting SSC depletion due to cumulative cytotoxicity (Meistrich et al., 1992).

Consistent patterns have also been reported for chemotherapy regimens containing alkylating agents. For instance, MVPP (mustine, vinblastine, procarbazine, and prednisolone) and COPP (cyclophosphamide, vincristine, procarbazine, and prednisolone) caused azoospermia in more than 90% of adult patients (reviewed by Howell and Shalet, 2005). When the doses of alkylating agents were reduced in these regimens, gonadotoxicity was also reduced, suggesting the role of DNA alkylation in causing reduced sperm counts and a dose-dependent relationship (Okada and Fujisawa, 2018).

Long-term reproductive effects of alkylating drugs and irradiation during early life stages have been reported. Green et al. (2014) evaluated semen parameters in 214 adult male survivors of childhood cancer who had received alkylating chemotherapeutic drugs. Azoospermia was diagnosed in 53 (25%) of participants and oligospermia in 59 (28%); the mean cumulative alkylating agent dose was higher in both groups than in participants with normospermia. Moreover, alkylating agent exposure was negatively correlated with sperm concentrations (r=-0.37, p<0.0001); specifically, each 1000 mg/m² increase in cumulative dose increased the odds of azoospermia by 22% (Green et al., 2014).

Beaud et al. (2019) revealed correlations between sperm parameters and treatment characteristics in 13 childhood cancer survivors. The participants received various doses of Mustargen®, procarbazine, or cyclophosphamide around puberty. Despite the small sample size, a strong link was observed between lower sperm counts and the use of alkylating agents (Spearman's rank correlation coefficient r=-0.62, p≤0.05), but not the use of other two non-alkylating drug classes, vinca alkaloids (r=0.07, p>0.05) and anthracyclines (r=-0.02, p>0.05) (Beaud et al., 2019).

Evidence in animal models in vivo

Meistrich (1982a) examined the effects of triethylenethiophosphoramide (thio-TEPA) on testicular cell survival and sperm production in C₃H mice. Male mice were injected i.p. with eight doses of thio-TEPA (0-35 mg/kg), and the number of sperm heads was counted in testis homogenates. Thio-TEPA exerts its anticancer activity through the formation of alkyl DNA adducts (Musser et al., 1989; Grigorii et al., 1991). A clear dose-dependent decrease in sperm head numbers was observed 8 weeks post-treatment. At doses ≥ 10 mg/kg, sperm counts were reduced to less than 10% of the control level, while the highest dose (35 mg/kg) reduced the number to ~1% of control (Meistrich, 1982a). These findings demonstrate dose concordance, with increasing exposure to this known alkylating agents associated with progressively greater reductions in sperm counts, and these decreases occur subsequent to treatment with alkylating agents, consistent with temporal concordance.

The same research group further examined spermatogenesis in C₃H mice following single i.p. injections of 14 chemotherapeutic drugs, including 7 alkylating agents (Meistrich et al., 1982b). Eleven days post-treatment, histological analyses showed that spermatogonial cells were the most sensitive germ cell population to cell death, whereas the spermatocytes and spermatids were relatively resistant at this time point, consistent with known biology. Dose-dependent loss of differentiating spermatogonia was observed following treatment with lomustine (40 mg/kg), mustargen (3 mg/kg), carmustine (10 and 60 mg/kg), mitomycin C (1.5 and 5 mg/kg), chlorambucil (20 mg/kg), thio-TEPA (3 and 26 mg/kg), and procarbazine (120 and 800 mg/kg). Assessment of sperm production at later time points supports temporal concordance between alkylation-induced germ cell loss and downstream reductions in sperm counts. Testicular sperm counts in testis homogenates measured 29 days post-treatment confirmed moderate to strong killing effects of alkylating agents on differentiating spermatogonia; all drugs reduced sperm counts to less than 10% of control within the dose range tested. Sperm counts measured 56 days after treatment reflect stem cell survival and recovery capacity of sperm production. At this timepoint, lomustine and mustargen induced up to a 25% decrease in sperm head numbers, whereas carmustine, chlorambucil, mitomycin C, and procarbazine reduced sperm counts to 30-60% of control levels. Nearly no sperm were counted in testes treated with thio-TEPA at 26 mg/kg, indicating minimal stem cell survival and recovery. Thus, a large number of alkylating agents lead to spermatogonial cell death and decline in sperm counts.

Busulfan, a potent bifunctional alkylating agent, produced similar effects. Bucci and Meistrich (1987) reported clear dose-dependent decreases in testicular sperm counts in C₃H mice exposed to busulfan (6, 13, 20, 28, 40 mg/kg) via i.p. injection. The dose required to reduce sperm head numbers by 50% of controls was estimated at 10 mg/kg at 29 days and 8 mg/kg at 56 days post-treatment. At 44 weeks, testicular and epididymal sperm counts remained significantly decreased by more than 50% in mice treated with busulfan ≥ 20 mg/kg compared to controls, while sperm counts returned to normal ranges in the lower dose groups (Bucci and Meistrich, 1987). These data demonstrate delayed onset and persistence of reduced sperm counts following exposure to a potent alkylating agent, supporting strong dose and temporal concordance and limited recovery of SSCs at higher doses.

Similar findings have been reported in a non-human primate model, supporting the conservation of this KER across species. Hermann et al. (2009) reported the effects of a single intravenous exposure to busulfan (4, 8, 12 mg/kg) exposure on spermatogenesis in adult rhesus macaques. Semen samples were collected weekly and total sperm counts per ejaculate were monitored for up to 52 weeks. Exposure to all three doses of busulfan caused dose-dependent and progressive decreases in sperm counts, which declined to zero by 7 weeks in animals treated with 12 mg/kg busulfan, and by 10 weeks following treatment with 4 mg/kg and 8 mg/kg busulfan. Testis volumes were also decreased correspondingly. Animals treated with 4 mg/kg busulfan restored normal spermatogenesis by 24 weeks following the exposure, albeit the sperm count is still lower than those in the controls. In contrast, sperm counts remained undetectable 52 weeks following exposure to 8 mg/kg and 12 mg/kg busulfan, suggesting SSC depletion and a longe-term loss of fertility. This study provides strong evidence for temporal and dose concordance, as reductions in sperm counts occur after exposure with a delay consistent with spermatogenic progression, with higher doses leading to more rapid and sustained depletion of sperm.

Repeated administration of a potent alkylating agent, ENU (100 mg/kg/week for three weeks), in C57BL/6J mice caused a delayed but significant reduction in sperm counts (Yin et al., 2014). No significant change in sperm count was observed during the first two weeks following the initial exposure, confirming that lack of apoptotic response in post-meiotic germ cells; however, sperm counts declined markedly from weeks 3-8 and reached near-zero levels by week 8. Sperm counts then gradually recovered, reaching approximately 85% of control levels by week 12, indicating profound but reversible impairment of spermatogenesis following ENU exposure (Figure 2). This study provides indirect support for essentiality through a stop-start (stressor removal) design.

In another Balb/c mouse study examining germline chromatin damage and sperm counts, ENU was used as a positive control. A single i.p. injection of 75 mg/kg of ENU at 8 weeks of age resulted in a delayed ~43% decrease in cauda epididymal sperm counts at 18 weeks of age, relative to control, accompanied by increased sperm DNA fragmentation and elevated germline mutation frequencies (Swayne et al., 2012). These studies provide strong temporal evidence.

Prenatal exposure studies provide evidence that early life exposure to alkylating agents and ionizing radiation can cause long-term effects on spermatogenesis. Comish et al. (2014) exposed pregnant 129 mice to either whole-body irradiated (0.8 Gy) or cyclophosphamide (7.5 mg/kg, i.p.) on embryonic days E10.5 and E11.5, corresponding to a critical period of primordial germ cell development. Male offspring were evaluated for testicular development and sperm production. At 4 weeks of age, male offspring from both treatment groups presented significantly reduced testicular weights (~70% of control), indicative of significant germ cell loss. The average number of atrophic Sertoli cell-only seminiferous tubules increased from 1.7 per testis section in controls to 4.8-5.8 tubules per section in irradiated and cyclophosphamide-treated animals. At 8 weeks of age, testicular weight remained reduced to 78% of control values, while testicular head counts and epididymal sperm counts were reduced to 62%-70% of control levels. These findings demonstrate that alkylation damage during fetal life can result in long-lasting germ cell depletion and reductions in sperm production in adulthood.

Together, these findings demonstrate that alkylation-induced genomic damage in male germ cells is consistently associated with impaired spermatogenesis and reduced sperm output. Although direct measurement of DNA adduct formation and sperm counts within the same study is uncommon, parallel evidence from studies of well-characterized alkylating agents shows consistent, dose-dependent changes in both KEs, supporting dose concordance. Significant reductions in sperm output are typically observed weeks to months following exposure, reflecting the time required for species-specific spermatogenic transit and confirming temporal concordance. The extent, duration, and reversibility of these effects highly depend on exposure dose, survival of SSCs, and spermatogenic kinetics in the test species.

Several uncertainties remain in this KER. First, DNA alkyl adducts in germ cells and sperm output are rarely measured in the same study, and many studies rely on surrogate markers of DNA damage, such as DNA strand breaks, making it difficult to attribute downstream effects specifically to DNA alkylation.

Second, reduced sperm count can arise through multiple mechanisms. In addition to direct alkylation-induced DNA damage in germ cells, other defects such as spermiation failure (i.e., retention of mature spermatozoa in late stage tubules) (Meistrich et al., 1982b), Sertoli cell dysfunction (Cui et al. 2024), and hormonal disruption (Sriram et al., 2024) may co-occur with alkylation damage and contribute to reduced testicular or epididymal sperm output. Moreover, alkylating agents have been shown to induce epigenetic alterations in sperm, including changes in DNA methylation at specific fertility-related genes (Altakroni et al., 2021). These alterations are associated with sperm DNA damage and reduced sperm counts (Jenkins et al., 2016); however, reported correlations between DNA damage markers and methylation are gene-specific and inconsistent across studies (Altakroni et al., 2021). As such, the relative contribution of epigenetic dysregulation to reduced sperm counts in this context remains uncertain.

Although Altakroni et al. (2021) reported an association between N7-MedG levels and decreased sperm concentration in 105 patients, this result was not replicated in a smaller follow-up study (n=16) (Altakroni et al., 2025). The latter study was based on a limited number of samples with N7-MedG measurements, which may have reduced statistical power. No significant correlations between N7-MedG levels, sperm concentration, or neutral comet assay measures (% tail DNA), despite observing a significant association with reduced fertilization rates.

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