Fluoroquinolone mitochondrial and genomic DNA adduct

What could be possible consequences of covalent bond mitochondrial and genomic DNA adduct by Fluoroquinolones, our study was done by Fluoroquinolone Toxicity Study NFP. All 50+ participates who took Ciprofloxacin and Levofloxacin were positive. In vitro study showed DNA adduct after 3.61 min. We are FQAD victims not scientists. We would like to know what may be consequences of DNA adduct.

QPCR: a tool for analysis of mitochondrial and nuclear DNA damage in ecotoxicology

The quantitative PCR (QPCR) assay for DNA damage and repair has been used extensively in laboratory species. More recently, it has been adapted to ecological settings. The purpose of this article is to provide a detailed methodological guide that will facilitate its adaptation to additional species, highlight its potential for ecotoxicological and biomonitoring work, and critically review the strengths and limitations of this assay. Major strengths of the assay include very low (nanogram to picogram) amounts of input DNA direct comparison of damage and repair in the nuclear and mitochondrial genomes, and different parts of the nuclear genome detection of a wide range of types of DNA damage very good reproducibility and quantification applicability to properly preserved frozen samples simultaneous monitoring of relative mitochondrial genome copy number and easy adaptation to most species. Potential limitations include the limit of detection (

1 lesion per 10 5 bases) the inability to distinguish different types of DNA damage and the need to base quantification of damage on a control or reference sample. I suggest that the QPCR assay is particularly powerful for some ecotoxicological studies.


“Our results suggest that the m.2352T>C polymorphism has a strong clinical effect on the risk of fibromyalgia and possibly other chronic pain conditions.” the authors

They extracted the genomic DNA and then did deep sequencing and found that the FM patients stood out in another way: they and the vulvar vestibulitis patients were the only pain groups to have a significantly increased frequency of a specific genetic polymorphism i.e. small change in one of the mtDNA genes.

(Vulvar vestibulitis refers to a condition characterized by severe pain in the vagina during intercourse, when using tampons or even when sitting on something like a bicycle seat. The pain can also be constant.)

Tilburg hypothesized genetic problems with energy production – plus stress in FM – could result in more pain.

In both groups, but more so in FM, an alteration in a single nucleotide (single nucleotide polymorphism (SNP) (m.2352T>C (rs28358579)) showed up with increased frequency in the MT-RNR2 ribosomal gene in women who a particular form of the gene called the c (minor) allele. An allele refers to a variant form of a gene and minor alleles refer to the second most common form of a gene in the population.

This small genetic alteration was also more evident, but to much lesser degree, in the other CPCC’s.

This gene, remarkably, had been unassessed in pain or other conditions, yet the evidence indicated that a small alteration in the gene was contributing to an increased risk for fibromyalgia and vulvar vestibulitis and perhaps other chronic pain conditions. (The polymorphism also increased the risk of a woman having more than one complex chronic pain condition.)

(One reason the polymorphism hasn’t been assessed much is that it’s found on an allele (the c (minor) allele) that is only very rarely found in Caucasian women and most genetic analyses have been done on Caucasian women. It is not currently assessed in 23andMe.)

Then they tested their finding in another pain cohort – the Orofacial Pain: Prospective Evaluation and Risk Assessment (OPPERA) Study cohort – which contained 52 women with FM and over a 1,000 without it. The researchers must have been dancing a jig when they learned that their initial finding stood up: people with FM indeed had significantly higher rates of the polymorphism.

Their deep dive into the mtDNA appeared to have uncovered the first evidence of genetic weakness for coming down with a “complex persistent pain condition”.

The effect was strong in fact, it appeared to be unusually strong. The authors reported:

“The replicated genetic effect size of the C allele on the disease risk (OR 5.1 and 4.3 in discovery and replication cohort, respectively) is impressive and has little precedence within the field of common diseases.”

An OR (odds ratio) of 5.1, if I have it right, means that having that particular SNP on that allele increased a woman’s odds of having fibromyalgia being present five-fold. That appears to be a huge number for a single gene polymorphism.

While having FM was significantly associated with having this polymorphism, the alternative c allele that the SNP was found in, was more much prominent in African American women and almost never found in women who identified themselves as Caucasian (1%). (Indeed, the odds ratio boomed in African American women to 7.6). Because the C allele had a stronger effect in a mixed population than in the African American population, it may have strongly affected other racial groups as well.

The SNP is located in the mitochondria’s 16S rRNA gene in the ribosome. Since this SNP has rarely been tested for, it hasn’t been associated with any other diseases or conditions. It’s not clear what it’s doing, but it may be hampering the ribosome’s ability to translate mRNA into proteins.

Mitochondrial functioning impacted

Next, they determined whether the polymorphism was actually impacting mitochondrial function. Obtaining ten cell lines from women with and without the polymorphism, they assessed the mitochondrial functioning of each one. Cells carrying the minor allele on which the SNP was found showed no difference in mitochondrial functioning (mitochondrial membrane potential) when put in glucose media (glycolysis), but when put in galactose media, their mitochondria membrane potential declined – indicating those cells were producing less ATP.


DNA sample preparation

mtDNA was extracted from human isolated muscle mitochondria and from murine (NMRI strain) muscle and liver mitochondria essentially as described previously [6]. nDNA was extracted from nuclei isolated from NMRI mouse livers essentially as described [7]. The polymerase chain reaction (PCR)-amplified fragments PCR-I and PCR-II of human mtDNA were obtained using the primers 5′-TAGAAACCGTCTGAACTATC-3′ (forward) and 5′-CCACAGATTTCAGAGCATT-3′ (reverse) for PCR-I and 5′-CACATTACAGTCAAATCCCT-3′ (forward) and 5′-TTGTATTGATGAGATTAGTA-3′ (reverse) for PCR-II. The amplified DNA fragments correspond to the sequences from nucleotide (nt) 7171 to nt 7611 (PCR-I 421 bp) and nt 15761 to nt 16487 (PCR-II 727 bp) in the human mitochondrial genome (GenBank accession number X93334 ref. [8]). The PCR products were purified using the PCR Clean-Up kit (Boehringer Mannheim, Mannheim, Germany) and were reconstituted in phosphate-buffered saline (PBS) before intra-articular (i.a.) injection (5 μg per knee joint) in NMRI mice. The GpC–ODN, 5′-TCCATGAGCTTCCTGATGCT-3′, and oxoGpC–ODN, 5′-TCCATGAXCTTCCTGATGCT-3′, where X = 8-oxodG, were synthesized by SGSDNA (Stockholm, Sweden).

DNA injections in mice

mtDNA, nDNA, and ODN samples (20 μl vol containing 5 μg DNA or 10 nmol ODN) were injected i.a. in the knees of female, 6- to 8-week-old mice [BALB/c mice from ALAB, Stockholm, Sweden CB17 and severe combined immunodeficiency (SCID) mice from M&B, Bomholtvej, Denmark NMRI mice from B&K, Universal AB, Sollentuna, Sweden]. All animals were housed in the animal facility of the Department of Rheumatology and Inflammation Research, University of Göteborg (Sweden), under standard conditions. The i.a.-injected mice were killed after 3 or 14 days, and the joints were removed for histopathology or immunohistochemistry.

In vivo depletion of immune cells

Female NMRI mice were depleted of peripheral blood monocytes by treatment with etoposide (Bristol Myers Squibb AB, Bromma, Sweden), which was injected subcutaneously (s.c.) at a dose of 12.5 mg/kg daily, starting 2 days before DNA injection, and continued for the course of the experiment (3 days). This treatment selectively depletes monocytes, as previously demonstrated [9]. The control mice received s.c. injections of PBS. Female BALB/c mice (6–8 weeks old) were depleted of granulocytes by intraperitoneal (i.p.) pretreatment with 1 mg of the monoclonal antibody (mAb) RB6-8C5 [10] 2 h before the i.a. injections of mtDNA. Control mice were pretreated with an immunoglobulin G (IgG) rat anti-ovalbumin (anti-OVA) mAb.

Histopathology and immunohistochemistry of mouse joints

The histopathological and immunohistochemical examinations of mouse joints were performed as described previously [4]. The severity of arthritis in knee-joint sections was evaluated by a blinded observer and assessed on a scale of 0–3, where 0 = no signs of inflammation, 1 = mild inflammation characterized by hyperplasia of the synovial lining layer, and 2–3 = increasing levels of inflammation characterized by influx of inflammatory cells into the synovial tissue. For reference purposes, the joint shown in Fig. 1a rates a score of 2, and that seen in Figure 1b is scored as 0.

Antisense ODN to nuclear factor (NF)-κB

The expression of the p65 subunit of NF-κB was blocked in female NMRI mice by i.p. administration of 900 μg antisense ODN, 5′-GAAACAGATCGTCCATGGT-3′, or the mismatched control ODN, 5′-GAAACAGATCGTCTATGGT-3′, 2 days before i.a. injection of mtDNA. These phosphorothioate-modified ODNs were synthesized by CyberGene AB (Huddinge, Sweden). No arthritis was observed in histopathological sections of the joints of mice injected i.p. with the antisense or the mismatched ODN alone at a dose of 900 μg per mouse.

Proliferation and tumor necrosis factor α (TNF-α) measurements

Naïve, murine spleen cells from four individual NMRI mice were incubated at a concentration of 2 × 10 6 cells/ml in Iscove’s medium containing 10% fetal calf serum. The cells were cultured at 37°C in 5% CO2 in the presence of 10 μg/ml mtDNA, 10 μg/ml nDNA, or medium alone for 24 h (TNF-α assay) or for 68 h (proliferation assay). The 24-h culture supernatants were assayed for TNF-α production by enzyme-linked immunosorbent assay (ELISA R&D Systems, Minneapolis, MN). To study proliferative responses, the 68-h cultures were pulsed with 3 H-thymidine for 4 h, the cells were harvested onto filters, and 3 H-thymidine incorporation was measured in a β-counter.

Analysis of human synovial fluids

PCR analysis for the presence of free (extracellular) mtDNA was performed on SF samples aspirated from 54 patients presenting with RA at Sahlgrenska University Hospital (Göteborg, Sweden). The control SF samples (n=17) were collected at autopsy from control subjects (none of whom showed any signs of RA or osteoarthritis). The samples were centrifuged at 500 g for 10 min and then at 2000 g for 10 min, and the supernatants were boiled for 10 min. Following centrifugation in a microfuge (11,000 g for 15 min at 4°C), the supernatants were analyzed by PCR using the human mtDNA-specific ODN primers, 5′-GCTCTCCATGCATTTGGTAT-3′ (forward primer) and 5′-TTGTATTGATGAGATTAGTA-3′ (reverse primer). The reaction mixtures contained 4 μM each forward and reverse primers in a total volume of 25 μl, which consisted of 2.5 μl-extracted SF in 10 mM Tris-HCl, pH 8.3, 50 mM KCl, 4.0 mM MgCl2, 0.2 mM deoxy-unspecified nucleoside 5′-triphosphates (deoxy-adenosine 5′-triphosphate, -cytidine 5′-triphosphate, -guanosine 5′-triphosphate, and -thymidine 5′-triphosphate), and 1.25 U Taq DNA polymerase (AmpliTaq Gold, Perkin-Elmer, Branchburg, NJ). All SF samples were subjected to 45 cycles of PCR amplification, each consisting of a 12-min hot start at 94°C, 30 s denaturation at 94°C, 30 s annealing at 60°C, and extension at 72°C for 1 min. Samples that showed the 453-bp PCR product on silver-stained sodium dodecyl sulfate-polyacrylamide gel electrophoresis gels were considered to be positive for the presence of mtDNA.


We used benzo[a]pyrene (B[a]P) to investigate the relationship between induced mtDNA damage and mutation. B[a]P is an established mutagen that has been shown to induce 40- to 90-fold more lesions in mtDNA than in nDNA (9,19). B[a]P is present in a wide range of combustion products, including tobacco smoke, coal tar and vehicular exhaust (13), and requires metabolic activation by cytochrome P450 isozymes, followed by epoxide hydrolase, to form mutagenic metabolites (e.g. benzo(a)pyrene-7,8-diol-9,10-epoxide, BPDE), which form bulky helix-distorting lesions by covalently modifying DNA.

To maximize the likelihood of mtDNA damage, we implemented a 28-day sub-chronic dosing regimen, which included three concentrations of B[a]P (25, 50 or 75 mg/kg body weight/day). As B[a]P requires metabolic conversion to a DNA-reactive substance (i.e. BPDE), we elected to study mtDNA mutagenesis in the liver, a well-known site of this activation (42�). Additionally, to assess the impact of mutagen treatment in a highly proliferative tissue, bone marrow was also included in our study. Both tissues have been shown to be exceptionally sensitive to DNA-damaging agents in nuclear and mitochondrial DNA, including B[a]P-induced damage. Moreover, previous work has shown that B[a]P significantly increases mutations in nuclear DNA, at similar doses, and in the same tissues tested as in the present study (34). However, the potential effect of B[a]P exposure on the induction of mtDNA mutations in vivo has not been examined.

To this end, we extracted DNA from liver and bone marrow cells to explore the possible effect of B[a]P treatment on mtDNA mutagenesis using the dRMC assay that builds upon the RMC methodology, and 3D assay (30,31). The dRMC and 3D assays (Figure ​ (Figure1) 1 ) have been used to quantify point mutations and deletion mutations in both humans and mice previously (30,45,46)(Supplementary Figure S1).

Illustrated overview of the 3D and dRMC assays for the quantification of mitochondrial mutations. (1) Whole cell DNA is extracted. (2) mtDNA is incubated with TaqI restriction endonuclease, which recognizes 5′-TCGA-3′ sites. mtDNA that are wild-type at TaqI sites (WT, blue), will be cleaved, whereas mtDNA with a mutation in the mutation target site (red) will be resistant to cleavage. A control region devoid of TaqI site(s) (purple) is used to quantify total mtDNA copies interrogated. (3) Digested DNA is added to a PCR mastermix with site-specific primers which flank the mutational target and Taqman probes, and then partitioned into thousands of 1 nl droplets in an oil immersion. The control region and mtDNA with mutations in the target site act as substrates for amplification, whereas mtDNA which are WT at the mutational target are not. (4) Droplets are thermal cycled to amplify target DNA as well as release the Taqman probe fluorophore from its quencher through Taq polymerase's inherent exonuclease activity. The ongoing rounds of amplification displace and cleave more probe, accumulating fluorescence. (5) Post-amplification, droplets are detected and their fluorescence is quantified. Mutation frequency is calculated by dividing the mutant concentration by the concentration of the control region.

Effect of B[a]P exposure on the frequency of mtDNA point mutation

Whole-cell DNA was extracted from frozen bone marrow and liver for mtDNA mutation analysis from B[a]P-exposed and control mice after 28 days of daily treatment and 3 post-exposure rest days. In bone marrow mtDNA isolated from B[a]P-treated mice, the mutation burdens ordered by increasing daily dose of B[a]P, were 3.8 ± 1.1, 4.2 ± 1.2, 3.8 ± 1.4 × 10 𢄦 bp and 2.6 ± 0.5, 2.4 ± 0.6, 2.0 ± 1.0 × 10 𢄦 bp (Figure ​ (Figure2A), 2A ), for the 12S rRNA region and ND5 site, respectively. In untreated mice, bone marrow mtDNA mutation frequencies were 3.5 ± 0.7 × 10 𢄦 bp and 1.7 ± 0.2 × 10 𢄦 bp, at the 12S and ND5 sites, respectively. No significant increases, or dose-dependent changes (ANOVA, multiple-comparisons corrected t-test), were observed between control (Figure ​ (Figure2A) 2A ) and treatment groups.

B[a]P treatment does not increase the frequency of mitochondrial point mutations. Mice were treated daily with B[a]P or vehicle for 28 consecutive days and tissues collected three days later. After DNA extraction, mutation frequency per bp (± s.e.m.) was determined via dRMC within the 12S rRNA and ND5 genes in mouse mtDNA. B[a]P did not induce mutations in (A) bone marrow (P = 0.66, 12S rRNA locus P = 0.21, ND5 locus one-way ANOVA) or (B) liver isolates of mice treated (P = 0.98, 12S rRNA locus P = 0.98, ND5 locus one-way ANOVA).

mtDNA isolates from the liver of B[a]P-treated mice displayed mutation frequencies, ordered by increasing dose, of 2.3 ± 0.5, 2.4 ± 0.1, 2.5 ± 0.2 × 10 𢄦 bp and 2.0 ± 0.8, 2.1 ± 0.9, and 2.0 ± 0.7 × 10 𢄦 bp for 12S rRNA and ND5 sites (Figure ​ (Figure2B), 2B ), respectively. The mutation frequency of untreated liver mtDNA at the 12S rRNA and ND5 sites were 2.6 ± 1.0 × 10 𢄦 bp and 2.2 ± 1.00 × 10 𢄦 bp. As with bone marrow, liver mtDNA mutation frequency was unaffected by B[a]P exposure. Thus, in both tissues, B[a]P exposure did not affect the frequency of point mutations in mtDNA.

Incidence of mtDNA deletions following mutagen exposure

The bulky adducts induced by B[a]P may underlie the lack of conversion into mtDNA point mutations, as the strand-distorting lesion produced by B[a]P strongly inhibits mitochondrial replication and thus lesion bypass (47). Polymerase stalling, however, has been hypothesized to cause deletion mutations (47,48). The observed lack of point mutation induction (Figure ​ (Figure2) 2 ) in mtDNA may be predicated upon blocked replication instead of error-prone polymerase bypass or DNA repair processes. Thus, we speculated that this would lead to polymerase stalling at the sites of damaged bases and, potentially, induce the formation of large mtDNA deletions.

To examine this possibility, we employed the 3D assay (31) to quantify mtDNA deletions following exposure to B[a]P. 3D can evaluate the presence of deletions in the mitochondrial genome, such as the 𠆌ommon deletion’: a 3.8 kb region in mouse mtDNA that shows preferential excision and end-joining due to sequence microhomology (49,50). The frequency of mtDNA deletions in our control mice was 1.1 ± 0.4 and 111.4 ± 31.0 deletions per 10 7 genomes for bone marrow and liver, respectively. These values are complementary to those found in previous studies for similarly aged mice, where liver showed the highest frequency of deletions (49). Deletion frequencies in treated mice, ordered by increasing doses of B[a]P were: 1.1 ± 0.50, 0.9 ± 0.40 and 1.1 ± 0.2 copies per 10 7 genomes in bone marrow (Figure ​ (Figure3A) 3A ) and, 71.4 ± 15.5, 47.6 ± 14.3 and 84.1 ± 33.2 deletions per 10 7 genomes, in liver (Figure ​ (Figure3B). 3B ). As with point mutations, B[a]P exposure did not significantly change the frequency of deletions at any dose or in either tissue (ANOVA, multiple-comparisons adjusted t-test).

B[a]P treatment does not induce mitochondrial deletions. Following 28 days of treatment with B[a]P no significant induction of deletion mutation frequency, (± s.e.m.) per mitochondrial genome was determined via 3D in (A) bone marrow (P = 0.94 one-way ANOVA) and (B) liver mtDNA(P = 0.37 one-way ANOVA).

B[a]P adducts in mtDNA and nDNA

Although the induction of mtDNA damage induced by B[a]P is extensively described, the unexpected lack of mutation induction in mtDNA following B[a]P exposure prompted us to address the possibility that damage was not induced in our test animals. To quantify the potential induction of B[a]P induced damage, we extracted DNA from bone marrow and liver tissues 24 h post-treatment with an acute dose of 75 mg B[a]P/kg body weight. As adducts formed by B[a]P inhibit polymerase extension, we quantified their presence via long-range quantitative PCR (13,51). This sensitive assay quantifies lesions that inhibit polymerase extension, and is not specific to one species of DNA adduct or lesion. B[a]P induced 0.29 ± 0.10 lesions per 10 kb (Figure ​ (Figure4A, 4A , P < 0.05, one-tailed Welch's t-test) and 0.26 ± 0.09 lesions per 10 kb (Figure ​ (Figure4B, 4B , P < 0.05, one-tailed Welch's t-test) in bone marrow and liver mtDNA, respectively. DNA samples were also processed for nDNA lesions using quantitative PCR directed to portions of the lacZ transgene. B[a]P induced 1.27 ± 0.40 lesions per 10 kb (Supplementary Figure S2A, P < 0.01, one-tailed Welch's t-test) and 0.66 ± 0.11 lesions per 10 kb (Supplementary Figure S2B, P < 0.05, one-tailed Welch's t-test) in bone marrow and liver nDNA, respectively. Thus, the lack of induced point and deletion mutations in the mitochondrial genome following 28 days of daily B[a]P exposures cannot be explained by the absence of damage induction. A single exposure of 75 mg B[a]P/kg body weight introduced 29 lesions per 10 6 bp in mtDNA with the potential to stall or inhibit polymerase extension. Yet despite the abundance of B[a]P-induced DNA lesions, no significant induction of mutation is observed in the mitochondrial genome following 28 days of daily B[a]P exposures.

B[a]P treatment induces mtDNA adducts. The presence of lesions (± s.e.m.) in mouse bone marrow and liver mtDNA was enumerated by quantitative PCR. Mice were treated with a single, acute dose of B[a]P at either 0 or 75 mg/kg body weight. DNA was extracted from bone marrow and liver tissues 24 h after treatment. B[a]P induces significant adduct burden in each tissue mtDNA (* indicates P < 0.05 one-tailed Welch's t-test). (A) Lesions formed in bone marrow. (B) Lesions formed in liver.

Nuclear B[a]P-induced mutagenesis

Mutation and damage burdens in mtDNA are typically described with comparisons to nDNA. Therefore, we sought to place the observed mitochondrial resistance to mutation in the context of the nuclear genome. We had selected B[a]P as our test mutagen, as previous reports had demonstrated preferential B[a]P adduct formation in mtDNA compared to nDNA (9). As such, we had hypothesized the mitochondrial genome would be more sensitive to B[a]P-induced mutation than the nuclear genome. In our evaluation of induced nDNA mutation and damage, we utilized the Muta TM Mouse transgenic rodent, which harbors a stably integrated lacZ transgene incorporated into a recoverable lambda phage shuttle vector. The shuttle vector can readily be recovered by packaging in phage particles that are subsequently used to infect galactose-sensitive bacteria (52,53). In the presence of P-gal, only those phages that receive a mutant lacZ can form plaques, allowing quantification of the mutant frequency in the nDNA (33,54). The mutant frequency in untreated animals was 4.3 ± 0.9 × 10 𢄥 in bone marrow, and 6.3 ± 0.6 × 10 𢄥 in liver. Contrary to our observations in the mitochondrial genome, B[a]P exposure resulted in a dose-dependent increase in nuclear mutant frequencies in both tissues, with 203 ± 35.8, 344 ± 75.0 and 679 ± 59.8 mutants × 10 𢄥 in bone marrow and 26 ± 4.8, 96 ± 14.5 and 219.0 ± 59.8 mutants × 10 𢄥 in liver samples (Figure ​ (Figure5A, 5A , P ≤ 0.0001 in bone marrow Figure ​ Figure5B, 5B , P ≤ 0.001 in liver, chi-square test). Additionally, we were able to confirm increased mutant frequencies using a nuclear version of the dRMC that quantifies mutations within the lacZ transgene. Mutation frequencies in untreated animals were 2.9 ± 1.7 × 10 𢄦 bp in bone marrow and 1.6 ± 0.4 × 10 𢄦 bp in liver tissues, whereas mutation frequency in mice exposed to 75 mg/kg body weight/day B[a]P was 35.2 ± 14.4 × 10 𢄦 bp in bone marrow and 29.5 ± 8.3 × 10 𢄦 bp in liver (Supplementary Figure S3, P ≤ 0.05 for bone marrow and liver, one-tailed Welch's-adjusted t-test). These results show a clear differential response between mtDNA and nDNA to B[a]P-induced mutagenesis.

B[a]P treatment results in a dose-dependent increases in the frequency of nuclear DNA transgene (lacZ) mutants. After 28 days of daily treatment with B[a]P, DNA was extracted from mouse tissues 3 days post-exposure. Mutant frequency (± s.e.m.) in mouse nuclear DNA displayed significant, dose-dependent increases (**P < 0.01 ***P < 0.001 ****P < 0.0001 Welch's-adjusted t-test). (A) Nuclear mutant frequency induced in bone marrow isolates (χ 2 = 4898, P ≤ 0.0001 Fisher's Exact, P ≤ 0.0001). (B) Nuclear mutant frequency induced in liver nuclear DNA (χ 2 = 897.2, P ≤ 0.0001 Fisher's Exact, P ≤ 0.0001).

Evaluation of ENU as a mitochondrial DNA mutagen

To explore whether the resistance of mtDNA to mutagenesis is unique to chemicals that induce bulky adducts, we investigated the potential of N-ethyl-N-nitrosourea (ENU) to induce mtDNA mutation. ENU is an alkylating agent that acts by transferring its ethyl group to oxygen or nitrogen radicals in nucleic acids (55). This primarily induces base mis-pairing and misincorporation by replicative polymerases without substantial stalling, which we hypothesized would encourage polymerase bypass errors rather than inhibit replication (56). Similar to our protocol for B[a]P exposure, we employed a 28 day, sub-chronic dosing regimen of 5 mg/kg body weight/day of ENU. As with the B[a]P-treated cohort, we evaluated mitochondrial point mutations and large deletions with dRMC and 3D, and quantified nDNA mutagenesis. In bone marrow, control and ENU-treated mouse mtDNA, point mutation frequencies were: 4.6 ± 1.5 × 10 𢄦 bp versus 4.5 ± 0.5 for the 12S rRNA locus and, 1.6 ± 0.3 × 10 𢄦 bp versus 1.9 ± 0.4 × 10 𢄦 bp for ND5 site (Figure ​ (Figure6A). 6A ). In liver, these frequencies were: 7.9 ± 4.8 × 10 𢄦 bp versus 14.8 ± 4.8 for the 12S rRNA site and 3.2 ± 0.4 × 10 𢄦 bp, and 4.4 ± 1.19 × 10 𢄦 for the ND5 site (Figure ​ (Figure6B). 6B ). Thus, mirroring our B[a]P results, the mtDNA point mutation frequency was not significantly altered at either mtDNA target and in either tissue after ENU exposure (bone marrow: P = 0.97, 12S rRNA locus P = 0.90, ND5 locus and in liver: P = 0.29, 12S rRNA locus P = 0.09, ND5 locus, Welch's unpaired t-test). 3D quantification of mtDNA 𠆌ommon’ deletions revealed 1.6 ± 0.3 and 1.5 ± 0.3 deletions per 10 7 genomes in untreated and treated bone marrow, respectively (Figure ​ (Figure7A). 7A ). In liver, these frequencies were 12.5 ± 3.8 deletions per 10 7 genomes and 20.4 ± 5.5 (Figure ​ (Figure7B). 7B ). In summary, regardless of the tissue of origin, ENU did not induce mtDNA point or deletion mutations.

Subchronic ENU treatment does not increase the frequency of mitochondrial point mutations. Mice were treated daily with vehicle or 5 mg/kg ENU for 28 consecutive days. Three days following treatment, DNA was extracted from bone marrow and liver. Mutation frequency per bp mtDNA (± s.e.m.) was determined via dRMC at TaqI restriction sites within the 12S rRNA and ND5 genes in mouse mitochondrial DNA. (A) Bone marrow mutation frequency (P = 0.97, 12S rRNA locus P = 0.90, ND5 locus Welch's unpaired t-test). (B) Liver mutation frequency (P = 0.29, 12S rRNA locus P = 0.09, ND5 locus Welch's unpaired t-test).

Subchronic ENU treatment does not induce deletions in mouse bone marrow and liver mtDNA. Deletion frequency per mitochondrial genome (± s.e.m.) was determined via 3D. (A) Bone marrow deletion frequency in mice treated with ENU (P = 0.82, two-tailed Welch's unpaired t-test). (B) Liver deletion frequency in mice treated with ENU (P = 0.09, two-tailed Welch's unpaired t-test).

Nuclear ENU-induced mutagenesis

Similar to our observations with B[a]P, quantification of nuclear mutants following ENU exposure showed, as expected, that ENU significantly induced mutant frequencies in both bone marrow and liver tissues. Specifically, we observed lacZ mutant frequencies of 5.0 ± 1.2 × 10 𢄥 in the untreated mice bone marrow, whereas 155.0 ± 11.1 mutants × 10 𢄥 were recovered from the ENU-treated mice. In liver, we quantified 3.0 ± 0.7 mutants × 10 𢄥 in the untreated cohort, and 27.0 ± 3.3 mutants × 10 𢄥 from their ENU-exposed counterparts (Supplementary Figure S4). The lacZ dRMC assay performed on these samples confirmed mutation induction by ENU in the nuclear genome. In these mice, mutant frequency of untreated animals was 1.1 ± 0.8 × 10 𢄦 bp in bone marrow and 1.6 ± 0.7 × 10 𢄦 bp in liver tissues, whereas mutation frequency in mice exposed to 5 mg/kg body weight/day ENU was 15.5 ± 4.6 × 10 𢄦 bp in bone marrow and 10.2 ± 4.67 × 10 𢄦 bp in liver (Supplementary Figure S5, P ≤ 0.05 for bone marrow and liver, one-tailed Welch's-adjusted t-test). As with the results of B[a]P exposure, these findings show a clear difference in the sensitivity of mtDNA and nDNA to ENU-induced mutagenesis. Intriguingly, although mtDNA copies per nuclear genome were unaffected by either B[a]P or ENU exposure in bone marrow (B[a]P, P = 0.37 ENU, P = 0.40, both Welch's-adjusted t-test), recovered liver tissue posted increased mtDNA copy number in B[a]P-exposed mice (ANOVA, P = 0.06 0 versus 75 mg/kg body weight/day P = 0.03, Welch's-adjusted t-test) and decreased mtDNA copy number in ENU-exposed mice (P = 0.03, Welch's-adjusted t-test), suggesting tissue- and compound-specific responses to mutagens that do not produce changes in the relative abundance of mtDNA copies (Supplementary Figure S6).

Mitochondrial dysfunction, mitophagy, and DNA damage

Mitochondria generate most of the cellular ATP through OXPHOS and the ETC. In addition, they supply cells with cofactors such as NAD + , NADH, and other intermediate metabolites that are required for cellular function. As each cell harbors multiple mitochondria, and mitochondrial damage accumulates over time, the average quality of the pool of mitochondria in each cell declines with age. This is consistent with the fact that several age-related diseases are characterized by loss of energy homeostasis and mitochondrial dysfunction [21] . The health of the mitochondrial pool is maintained by fission and fusion of individual mitochondria and autophagic removal of damaged mitochondria by mitophagy [93-95] . High cellular energy demand stimulates mitochondrial fusion and formation of mitochondrial networks [96, 97] . Fusion of mitochondria promotes mixing of their content and complementation of gene products, which in turn maintain respiratory capacity under stressful conditions. Fusion of mitochondria promotes maintenance of the mitochondrial membrane potential (MMP), increases ATP production, decreases mitophagy, and prevents cell death [94, 96-98] .

Failure to restore NAD + bioavailability, in response to DDR, negatively affects the cellular redox balance, which exacerbates oxidative and metabolic stress (Fig. 3) [99, 100] . DNA damage stimulates mitochondrial fusion [101] , mitochondrial OXPHOS activity, and cellular metabolism to supply cells with ATP and NAD + in order to restore cellular homeostasis [102, 103] . In cells that sustain high levels of DNA damage, persistent activation of DDR creates a high-energy demand, and a cascade of events lead to mitochondrial dysfunction and metabolic stress. Consistent with this, human diseases characterized by defects in DNA repair, including Cockayne Syndrome (CS) [75, 104] , xeroderma pigmentosum (XP) [105] , and ataxia telangiectasia (A-T) [106, 107] , are characterized by loss of energy homeostasis, and defective mitophagy. We propose that the pathways involved with the activation of PARP1 and the ensuing depletion of cellular NAD + and ATP contribute to the premature aging features in these diseases, as described above (Fig. 3).

Signaling pathways that link DNA damage to mitophagy

Damaged mitochondria are targeted for degradation by two pathways, one that is ubiquitin-dependent and one that is ubiquitin-independent. In ubiquitin-dependent mitophagy, damaged mitochondria are identified based on membrane potential. PTEN-induced putative kinase 1 (PINK1) is a depolarization sensor localized to the mitochondrial outer membrane (MOM), that is activated by autophosphorylation [108-110] in a membrane potential-dependent manner. Activated PINK1 phosphorylates parkin [111] and ubiquitin [112] , which ultimately recruits autophagy receptors [113] leading to degradation of the targeted organelle. In the ubiquitin-independent pathway autophagy receptors bind directly to the mitochondria and tether it to the nascent autophagosome [114-119] .

Several proteins link DNA damage signaling to mitophagy (Fig. 4). First, SIRT1 deacetylates and inactivates p53. This p53 was recently shown to repress the promoter activity, protein, and mRNA levels of PINK1, which downregulates mitophagy [120] . SIRT1 also stabilizes the MMP, most likely through regulation of PGC1α and mitochondrial uncoupling protein 2 (UCP2). In XPA cells, PINK1 is destabilized due to a high MMP, which causes defective mitophagy. Upregulation of UCP2 through activation of the NAD + –SIRT1–PGC1α axis restores mitophagy in these cells [105] . Yet another way SIRT1 promotes mitophagy is through its cooperation with AMP-activated protein kinase (AMPK). Upon activation, AMPK promotes several pathways that maintain cellular energy homeostasis, such as mitophagy, through phosphorylation of the pro-mitophagy factor UNC51-like kinase 1 (ULK1) [121] and the mTORC1 inhibitor tuberous sclerosis complex 2 (TSC2) [122] . AMPK is activated by increased levels of AMP, ADP, and also by SIRT1 through deacetylation of LKB1. AMPK activates SIRT1 by increasing the NAD + /NADH ratio [123] thus likely creating a positive activation feedback cycle [124] . The central DDR regulator, ATM, also activates AMPK, which inhibits mTORC1, and also activates NEMO-JNK, thereby promoting autophagy/mitophagy. Nevertheless, the mechanisms by which DNA damage modulates the efficiency of mitophagy are poorly understood. The extent and persistence of the DNA damage might determine the balance between inhibition and initiation of these processes. Therefore, this is an important area for future research.

Materials and methods

DNA samples

DNA was extracted from 10 HNSCC cell lines from 9 patients. Normal DNA was isolated from fibroblasts from the same patients. MtDNA sequences were compared in the normal and tumour cell line DNA to determine which alterations identified in the tumour cell lines were polymorphisms and which were mutations.

Five cell lines (UM-SCC-3, UM-SCC-10B, UM-SCC-11B, UM-SCC-27 and UM-SCC-68) were from recurrent or metastatic sites, and 5 (UM-SCC-11A, UM-SCC-16, UM-SCC-62, UM-SCC-82 and UM-SCC-95) were from primary tumours. UM-SCC-11A was cultured from a biopsy of a primary laryngeal cancer and UM-SCC-11B was derived from the laryngectomy specimen of the same patient after chemotherapy.

The tumour specimen site and the stage of disease, at the time the tissue was taken for culture, are shown in Table ​ TableI I .

Table I

Cell lineSpecimen SiteStage
UM–SCC𠄳Lymph nodeII
UM–SCC�Lymph nodeIII
UM–SCC�Larynx after cx.IV
UM–SCC�Lymph nodeII
UM–SCC�Tonsillar fossaIII
UM–SCC�Lymph nodeII

PCR amplification and sequencing

The following oligonucleotide primers were used for ND4 gene PCR amplification and sequencing:

fragment 1 (product size 812bp) from 10688bp to 11500bp,L 5’-TGGGCCAGCCCTACTAGTCT-𠆓 and R 5’-GTCAGGGGGTTGAGAATGAG-𠆓

fragment 2 (product size 712bp) from 11295bp to 12076bp, L 5’-TCACTCTCACTGCCCAAGAA-𠆓 and R 5’-GGAGAATGGGGGATAGGTGT-𠆓.

PCR reactions were carried out with 200-500 ng genomic DNA as template, PCR buffer (10x) 10 μl, Mg buffer (25 mM) 6 μl, dNTP (10 mM) 2 μl, primer (10 μM) 2 μl, Taq (5 μl) 0.5 μl (Promega, Wisconsin, Usa) and water for a total volume of 100 μl.

PCR conditions were: 1 cycle at 94 ଌ for 5 min followed by 30 cycles at 94 ଌ for 1 min, 60 ଌ 1 min, 72 ଌ 1 min, and final extention of 1 cycle at 72 ଌ for 7 min.

To confirm the correct size of the PCR products, 10 μl of amplified DNA was analyzed in 1.5% agarose gel.

The PCR products were purified using the Qiaquick PCR purification kit (Qiagen, Valencia, CA, Usa). The DNA concentrations were measured with a spectometer and adjusted to 10 ng/μl. The primer concentration used for sequencing was 3.2 pmol/μl.

DNA sequencing was carried out using an Applied Biosystems DNA sequencer model 377.

Mitochondrial Permeability Transition Pore

The permeability transition represents a sudden increase of the inner mitochondrial membrane permeability to solutes with a molecular weight lower than 1.5 kDa. Short-term openings of the permeability transition pore represent physiological adjustments to regulate Ca 2+ and ROS homeostasis, providing mitochondria with a fast mechanism to release Ca 2+ when this reaches a harmful concentration inside the mitochondrial matrix. Conversely, long-term opening of the pore is linked to mitochondrial dysfunction because its occurrence leads to mitochondrial depolarization, cessation of ATP synthesis, Ca 2+ release, pyridine nucleotide depletion, and inhibition of respiration. In vitro, long-term opening of the pore lead to matrix swelling, which, in turn, causes the mobilization of cytochrome c, the outer mitochondrial membrane rupture, and eventually the release of proapoptotic proteins such as cytochrome c. The precise molecular composition and identity of the mitochondrial permeability transition pore are still controversial. Candidate components of the pore are the adenine nucleotide translocase (ANT), the voltage dependent anion channel (VDAC), the phosphate carrier (PiC), and components of the ATP synthase (Bernardi and Di Lisa, 2015).

It has been suggested that taxane-induced and platinum-induced mitochondrial injury might be associated to a direct interaction with the permeability transition pore. Though there is a large body of evidence that therapeutic and toxic effects of cisplatin on cells is primarily a consequence of inter-strand and intra-strand covalent adduct formation with nucleic acids, it has been shown that platinum can also crosslink with proteins. Experiments in intact cells suggest that cisplatin, perhaps conjugated to glutathione, accumulates in mitochondria, rapidly impairs the oxygen consumption, and induces oxidative stress (Dzamitika et al., 2006 Garrido et al., 2008). Depending on the source, oxidative stress not only can originate from mitochondria but also from other organelles, especially endoplasmic reticulum (ER) and peroxisomes (Kaludercic et al., 2014). Co-localization experiments in Human Kidney-2 (HK-2) cells demonstrated that cisplatin-induced ROS is of mitochondrial origin (Choi et al., 2015). It has been shown that cisplatin can form crosslinks with the VDAC, facilitating mitochondrial membrane permeabilization, release of cytochrome c, and apoptosis (Yang et al., 2006). Short exposure to paclitaxel produces marked loss of renal tubules epithelial lining and damage of the brush border membranes with signs of both oncotic necrosis and apoptosis (Rabah, 2010). Paclitaxel induces an abrupt fall of the mitochondrial membrane potential and a loss of mitochondrial Ca 2+ . Because cyclosporin A, a de-sensitizer of the mitochondrial permeability transition pore, blocked paclitaxel-induced loss of mitochondrial Ca 2+ , the authors concluded that paclitaxel induced the opening of the mitochondrial permeability transition pore (Kidd et al., 2002).

The opening of the transition pore has also been associated with nephrotoxicity induced by some natural products very popular in traditional medicine. Xanthium strumarium, used in traditional Chinese medicine to treat nasal and sinus congestion, has been linked to several cases of poisoning with renal proximal tubular necrosis features which upon X. strumarium ingestion have been reported in the literature (Turgut et al., 2005). The seeds of this plant are enriched in carboxyatractyloside, a well-characterized inhibitor of the ANT. The binding of carboxyatractyloside to the ANT triggers the opening of the mitochondrial transition pore (Obatomi et al., 1998 Klingenberg, 2008). In 1992, a high prevalence of kidney disease in female patients ingesting slimming pills raised attention to the nephrotoxicity of aristolochic acids. Ever since, aristolochic acids are considered a group of toxins that can cause end-stage renal failure associated with urothelial carcinomas (Han et al., 2019). Aristolochic acid is transported into the proximal tubular cells by the OATs (Bakhiya et al., 2009 Xue et al., 2011). Studies in HK-2 cells showed that aristolochic acid exposure caused ATP depletion, mitochondrial membrane depolarization, cytochrome c release, and increase of caspase 3 activity. These toxic effects were attenuated by cyclosporin A, a known �sensitizer” of the opening of the pore (Bernardi, 1996). Experiments in isolated mitochondria showed that aristolochic acid inhibited the activity of the mitochondrial ANT (Qi et al., 2007).


In the present study, we showed that workers with exposure to PAHs above 3 μmol 1-pyrenol/mol creatinine had significantly higher mtDNAcn in PBLs compared with controls. Workers also exhibited higher levels of genetic and chromosomal alterations in anti-BPDE–DNA adducts, micronuclei, telomere length, and epigenetic changes (i.e., p53 hypomethylation) as previously reported (6, 31). Increased levels of mtDNAcn in PBLs have been associated with a future risk of lung cancer (22). Our findings extend those observations by showing that increased mtDNAcn may also occur in healthy individuals who are occupationally exposed to considerably higher levels of carcinogenic PAH(BP) than Biological Exposure Index proposed by Jongeneelen (34), i.e., 1-pyrenol of 2.28 μmoles/mol creatinine. At this value, corresponding to the post-shift excretion value at an environmental exposure equal to the airborne threshold limit value of coal tar pitch volatiles (i.e., 0.2 mg/m 3 of “benzene soluble matter”, ACGIH (35), coke-oven workers have been shown to be at a 30% increased risk of lung cancer (34). Alterations in mtDNA have long been suggested to contribute to the development of lung cancer (for a review see ref. 36). However, whether mtDNAcn has a direct role in lung carcinogenesis is still under investigation. Increased mtDNAcn and a concurrent decline in mitochondrial function of salivary cells have been shown in response to tobacco smoke (18, 19), as well as in normal adjacent lung tissues of patients with cancer (17). Individuals with higher blood mtDNAcn at baseline have higher risk of developing lung cancer (22, 37). In addition, mtDNAcn alterations are associated with impaired apoptosis and subsequent increased cellular proliferation (38), as well as with nDNA mutations due to aberrant mtDNA insertion into the nuclear genome (39). The findings of the present study are suggestive of potential roles of mtDNAcn in PAH-induced carcinogenesis. However, whether mtDNAcn changes contribute to determining increased risks of malignancies in PAH-exposed individuals remains to be determined. Our findings on high PAH-exposed workers indicate that lymphocyte mtDNAcn may represent a novel marker specifically associated with levels of PAH exposure above the risk threshold. Increased PBL mtDNAcn has been associated with exposures derived from combustion processes such as benzene (40, 41) and particulate matter (32). Our results may indicate a potential role of PAHs, also a product of combustion, in those associations.

In our study, we found that mtDNAcn was correlated with genotoxic anti-BPDE–DNA adduct formation that, in addition to offering an assessment of the dose of carcinogens close to the molecular targets, represents a measure of cumulative exposure to carcinogenic PAHs due to the longer life span of PBL DNA compared with urine metabolites. Because adducts detection was carried out DNA aliquots derived from the same DNA samples where mtDNAcn were determined, we cannot exclude that part of anti-BPDE–DNA adducts may have mtDNA origin. Therefore, at least part of the correlation observed between anti-BPDE–DNA adduct level and mtDNAcn might depend on direct adduct formation to the mtDNA, rather than on interconnections between the nuclear and mitochondrial compartment. However, mtDNA is a small molecule of approximately 15,000 base pairs. Even considering a potential, albeit high, number of 10,000 mtDNA copies in one single cell, the total number of mtDNA base pairs in a cell would amount to just approximately 150 million base pairs, i.e., only 5% of the approximately 3 billion bp in the human nuclear genome. Nonetheless, the lipophilic character of BP and its metabolites, coupled with the very high ratio of lipid/DNA in mitochondria may facilitate the access of anti-BPDE to the mtDNA. Also, anti-BPDE has 40 to 90-fold higher affinity for mtDNA than for nDNA (9–11). Compared with nDNA, mtDNA has diminished protective histones and DNA repair capacity, and is therefore particularly susceptible to DNA damage. Consequently, the contribution of mtDNA-bound anti-BPDE could be a relevant portion of the total cellular burden of DNA adducts.

Mitochondria have been shown to compensate for damage and dysfunction by replicating their mtDNA and increasing mtDNAcn (16). A rise in mtDNA content has been directly associated with DNA damage (42, 43) and reduced respiratory chain function, secondary to oxidative damage (17, 44). In our study, anti-BPDE-related mtDNAcn changes were detected at high PAH-exposure doses. Although PAHs are still considered the primary genotoxic carcinogens produced by coal combustion emissions (for a comprehensive review see ref. 3 and references therein), the presence in coke emissions of toxic metals, or alternatively of reactive oxygen species produced by PAH (12) or metal (45) metabolism, might have contributed, along with BP exposure, in determining the mtDNAcn alteration observed in our study. Also, alterations of mitochondrial lipids and proteins produced by PAHs and/or anti-BPDE may have operated as concurrent events contributing to the increased mtDNAcn. However, we cannot exclude that different socio-economic statuses might have contributed, along with PAH exposure (for which we have supplied a measure of internal dose and biologically effective dose, i.e., the specific promutagenic anti-BPDE–DNA), in the increase of mtDNAcn that we observed in PBLs of coke-oven workers.

Interrelationships between the mtDNA and nDNA is however suggested in our study by the finding that individuals with lower mtDNAcn, even after adjusting for age, also had lower telomere length. This finding specifically suggests a link between nuclear telomere attrition, a marker of biologic aging, and mitochondrial alterations. This observation is in line with previous findings by Sahin and colleagues. (46) that showed a potential unifying mechanism connecting the nucleus and mitochondria in cellular aging. In that work, progressive nuclear telomere shortening, mediated by the activation of a p53-dependent pathway, was found to determine a reduction of mitochondrial function and mtDNAcn (46).

The present study has several strengths. The enrollment of the study participants was carefully designed to minimize potential confounding and increase the capability to reveal PAH effects by selecting non–current smoking males, all living in the same residential area. The selection of non–current smokers minimized the probability that the observed associations were dependent on factors other than the occupational PAH exposure. We also evaluated several other potential sources of PAH exposure, including dietary PAHs, indoor PAH exposure, and environmental PAH exposure, which showed no differences between coke-oven workers and controls. Our study had reliable measurements of PAH(BP) internal and target doses. Also, we measured in the study participants, biomarkers of genetic instability and methylation that allowed for characterizing the intercorrelation between mtDNAcn and nDNA alterations. Finally, the results of this study appear to be biologically plausible and the direction of the effects is consistent with the available literature data on mtDNAcn mechanisms.

We also recognize limitations to our study. This is a small-sized study and its results need to be confirmed in a larger independent investigation. Its cross-sectional design does not allow for investigating the temporal relationship of PAH exposure with mtDNAcn, as well as of the biomarkers of damage, genetic instability, and DNA methylation. The absence of air monitoring, as well as of repeated biologic sampling, are also limitations of the study exposure assessment strategy. However, PAH exposure was assessed using biomarkers of internal dose (urinary 1-pyrenol) and target dose (anti-BPDE–DNA adduct), which may more appropriately represent the effective exposure dose. To limit confounding, we matched coke-oven workers and controls for their individual characteristics, including age, gender, and ethnicity. In addition, we adjusted the analysis contrasting high-exposed workers, as well those based on continuous exposure or biomarker variables, for age. Because of the limited number of study subjects, it is possible that the associations observed were due to confounding or chance. The small sample size might have also caused false negative findings. For instance, we did not find any difference in mtDNAcn between low-exposed coke-oven workers and controls. Future studies with augmented sample size are warranted to better characterize the effects of PAH exposure on mtDNAcn at low doses.

In conclusion, coke-oven workers exposed to high levels of PAHs exhibited significantly higher PBL mtDNAcn, as well as genetic alterations in nDNA (i.e. anti-BPDE–DNA adduct, micronuclei, shorter telomere length, p53 hypomethylation). Individuals with shorter telomere length showed lower mtDNAcn, thus linking PAH exposure and mitochondrial dysfunction with cellular aging. These features were found in PBLs of individuals chronically exposed to PAHs. As previous investigations have shown that increased mtDNAcn is predictive of future risk of lung cancer, the results of the present study are highly suggestive that mtDNAcn may serve as a biomarker of cancer risk due to PAH exposure. Although these results imply a role of mtDNAcn in PAH carcinogenesis, whether mtDNAcn mediates the risk of lung cancer, determined by PAH exposure, should be determined in future mechanistic investigations.


Non-ionising radiations, electromagnetic fields (EMF) such as radiofrequency (RF), or power frequency radiation have become very common in everyday life. All of these exist as low frequency radiation which can come from wireless cellular devices or through electrical appliances which induce extremely low frequency radiation (ELF). Exposure to these radioactive frequencies has shown negative affects on the fertility of men by impacting the DNA of the sperm and deteriorating the testes [2] as well as an increased risk of tumor formation in salivary glands. [3] [4] The International Agency for Research on Cancer considers RF electromagnetic fields to be possibly carcinogenic to humans, however the evidence is limited. [5]

Radiation and medical imaging Edit

Advances in medical imaging has resulted in increased exposure of humans to low doses of ionizing radiation. Radiation exposure in pediatrics has been shown to have a greater impact as children's cells are still developing. [2] The radiation obtained from medical imaging techniques is only harmful if consistently targeted multiple times in a short space of time. Safety measures have been introduced in order to limit the exposure of harmful ionizing radiation such as the usage of protective material during the use of these imaging tools. A lower dosage is also used in order to fully rid the possibility of a harmful effect from the medical imaging tools. The National Council on Radiation Protection and Measurements along with many other scientific committees have ruled in favor of continued use of medical imaging as the reward far outweighs the minimal risk obtained from these imaging techniques. If the safety protocols are not followed there is a potential increase in the risk of developing cancer. This is primarily due to the decreased methylation of cell cycle genes, such as those relating to apoptosis and DNA repair. The ionizing radiation from these techniques can cause many other detrimental effects in cells including changes in gene expression and halting the cell cycle. However, these results are extremely unlikely if the proper protocols are followed. [1] [4]

Target theory concerns the models of how radiation kills biological cells and is based around two main postulates:

  1. "Radiation is considered to be a sequence of random projectiles
  2. the components of the cell are considered as the targets bombarded by these projectiles" [6]

Several models have been based around the above two points. From the various proposed models three main conclusions were found:

  1. Physical hits obey a Poisson distribution
  2. Failure of radioactive particles to attack sensitive areas of cells allow for survival of the cell
  3. Cell death is an exponential function of the dose of radiation received as the number of hits received is directly proportional to the radiation dose all hits are considered lethal [7]

Radiation exposure through ionizing radiation (IR) affects a variety of processes inside of an exposed cell. IR can cause changes in gene expression, disruption of cell cycle arrest, and apoptotic cell death. The extent of how radiation effects cells depends on the type of cell and the dosage of the radiation. Some irradiated cancer cells have been shown to exhibit DNA methylation patterns due to epigenetic mechanisms in the cell. In medicine, medical diagnostic methods such as CT scans and radiation therapy expose the individual to ionizing radiation. Irradiated cells can also induce genomic instability in neighboring un-radiated cells via the bystander effect. Radiation exposure could also occur via many other channels than just ionizing radiation.

The basic ballistic models Edit

The single-target single-hit model Edit

In this model a single hit on a target is sufficient to kill a cell [7] The equation used for this model is as follows:

Where k represents a hit on the cell and m represents the mass of the cell.

The n-target single-hit model Edit

In this model the cell has a number of targets n. A single hit on one target is not sufficient to kill the cell but does disable the target. An accumulation of successful hits on various targets leads to cell death. [7] The equation used for this model is as follows:

Where n represents number of the targets in the cell.

The linear quadratic model Edit

The equation used for this model is as follows: [7]

where αD represents a hit made by a one particle track and βD represents a hit made by a two particle track and S(D) represents the probability of survival of the cell.

The three lambda model Edit

This model showed the accuracy of survival description for higher or repeated doses. [7]

The equation used for this model is as follows:

The linear-quadratic-cubic model Edit

The equation used for this model is as follows: [7]

Sublesions hypothesis models Edit

The repair-misrepair model Edit

This model shows the mean number of lesions before any repair activations in a cell. [7]

The equation used for this model is as follows:

where Uo represents the yield of initially induced lesions, with λ being the linear self-repair coefficient, and T equaling time

The lethal-potentially lethal model Edit

This equation explores the hypothesis of a lesion becoming fatal within a given of time if it is not repair by repair enzymes. [7]

The equation used for this model is as follows:

T is the radiation duration and tr is the available repair time.

The saturable repair model Edit

This model illustrates the efficiency of the repair system decreasing as the dosage of radiation increases. This is due to the repair kinetics becoming increasingly saturated with the increase in radiation dosage. [7]

The equation used for this model is as follows:

n(t) is the number of unrepaired lesions, c(t) is the number of repair molecules or enzymes, k is the proportionality coefficient, and T is the time available for repair.

Radiation hormesis Edit

Hormesis is the hypothesis that low levels of disrupting stimulus can cause beneficial adaptations in an organism. [8] The ionizing radiation stimulates repair proteins that are usually not active. Cells use this new stimuli to adapt to the stressors they are being exposed to. [9]

Radiation-Induced Bystander Effect (RIBE) Edit

In biology, the bystander effect is described as changes to nearby non-targeted cells in response to changes in an initially targeted cell by some disrupting agent. [10] In the case of Radiation-Induced Bystander Effect, the stress on the cell is caused by ionizing radiation.

The bystander effect can be broken down into two categories, long range bystander effect and short range bystander effect. In long range bystander effect, the effects of stress are seen further away from the initially targeted cell. In short range bystander, the effects of stress are seen in cells adjacent to the target cell. [10]

Both low linear energy transfer and high linear energy transfer photons have been shown to produce RIBE. Low linear energy transfer photons were reported to cause increases in mutagenesis and a reduction in the survival of cells in clonogenic assays. X-rays and gamma rays were reported to cause increases in DNA double strand break, methylation, and apoptosis. [10] Further studies are needed to reach a conclusive explanation of any epigenetic impact of the bystander effect.

Formation of ROS Edit

Ionizing radiation produces fast moving particles which have the ability to damage DNA, and produce highly reactive free radicals known as reactive oxygen species (ROS). The production of ROS in cells radiated by LDIR (Low-Dose Ionizing Radiation) occur in two ways, by the radiolysis of water molecules or the promotion of nitric oxide synthesis (NOS) activity. The resulting nitric oxide formation reacts with superoxide radicals. This generates peroxynitrite which is toxic to biomolecules. Cellular ROS is also produced with the help of a mechanism involving nicotinamide adenosine dinucleotide phosphate (NADPH) oxidase. NADPH oxidase helps with the formation of ROS by generating a superoxide anion by transferring electrons from cytosolic NADPH across the cell membrane to the extracellular molecular oxygen. This process increases the potential for leakage of electrons and free radicals from the mitochondria. The exposure to the LDIR induces electron release from the mitochondria resulting in more electrons contributing to the superoxide formation in the cells.

The production of ROS in high quantity in cells results in the degradation of biomolecules such as proteins, DNA, and RNA. In one such instance the ROS are known to create double stranded and single stranded breaks in the DNA. This causes the DNA repair mechanisms to try to adapt to the increase in DNA strand breaks. Heritable changes to the DNA sequence have been seen although the DNA nucleotide sequence seems the same after the exposure with LDIR. [11]

Activation of NOS Edit

The formation of ROS is coupled with the formation of nitric oxide synthase activity (NOS). NO reacts with O2 − generating peroxynitrite. The increase in the NOS activity causes the production of peroxynitrite (ONOO-). Peroxynitrite is a strong oxidant radical and it reacts with a wide array of biomolecules such as DNA bases, proteins and lipids. Peroxynitrite affects biomolecules function and structure and therefore effectively destabilizes the cell. [11]

Mechanism of oxidative stress and epigenetic gene regulation Edit

Ionizing radiation causes the cell to generate increased ROS and the increase of this species damages biological macromolecules. In order to compensate for this increased radical species, cells adapt to IR induced oxidative effects by modifying the mechanisms of epigenetic gene regulation. There are 4 epigenetic modifications that can take place:

  1. formation of protein adducts inhibiting epigenetic regulation
  2. alteration of genomic DNA methylation status histone interactions affecting chromatin compaction
  3. modulation of signaling pathways that control transcription factor expression

ROS-mediated protein adduct formation Edit

ROS generated by ionizing radiation chemically modify histones which can cause a change in transcription. Oxidation of cellular lipid components result in an electrophilic molecule formation. The electrophilic molecule binds to the lysine residues of histones causing a ketoamide adduct formation. The ketoamide adduct formation blocks the lysine residues of histones from binding to acetylation proteins thus decreasing gene transcription. [11]

ROS-mediated DNA methylation changes Edit

DNA hypermethylation is seen in the genome with DNA breaks at a gene-specific basis, such as promoters of regulatory genes, but the global methylation of the genome shows a hypomethylation pattern during the period of reactive oxygen species stress. [12]

DNA damage induced by reactive oxygen species results in increased gene methylation and ultimately gene silencing. Reactive oxygen species modify the mechanism of epigenetic methylation by inducing DNA breaks which are later repaired and then methylated by DNMTs. DNA damage response genes, such as GADD45A, recruit nuclear proteins Np95 to direct histone methyltransferase's towards the damaged DNA site. The breaks in DNA caused by the ionizing radiation then recruit the DNMTs in order to repair and further methylate the repair site.

Genome wide hypomethylation occurs due to reactive oxygen species hydroxylating methylcytosines to 5-hydroxymethylcytosine (5hmC). [13] The production of 5hmC serves as an epigenetic marker for DNA damage which is recognizable by DNA repair enzymes. The DNA repair enzymes attracted by the marker convert 5hmC to an unmethylated cytosine base resulting in the hypomethylation of the genome. [14]

Another mechanism that induces hypomethylation is the depletion of S-adenosyl methionine synthetase (SAM). The prevalence of super oxide species causes the oxidization of reduced glutathione (GSH) to GSSG. Due to this, synthesis of the cosubstrate SAM is stopped. SAM is an essential cosubtrate for the normal functioning of DNMTs and histone methyltrasnferase proteins.

ROS-mediated post-translation modification Edit

Double stranded DNA breaks caused by exposure to ionizing radiation are known to alter chromatin structure. Double stranded breaks are primarily repaired by poly ADP (PAR) polymerases which accumulate at the site of the break leading to activation of the chromatin remodeling protein ALC1. ALC1 causes the nucleosome to relax resulting in the epigenetic up-regulation of genes. A similar mechanism involves the ataxia telangiectasia mutated (ATM) serine/threonine kinase which is an enzyme involved in the repair of double stranded breaks caused by ionizing radiation. ATM phosphorylates KAP1 which causes the heterochromatin to relax, allowing increased transcription to occur.

The DNA mismatch repair gene (MSH2) promoter has shown a hypermethylation pattern when exposed to ionizing radiation. Reactive oxygen species induce the oxidization of deoxyguanosine into 8-hydroxydeoxyguanosine (8-OHdG) causing a change in chromatin structure. Gene promoters that contain 8-OHdG deactivate the chromatin by inducing trimethyl-H3K27 in the genome. Other enzymes such as transglutaminases (TGs) control chromatin remodeling through proteins such as sirtuin1 (SIRT1). TGs cause transcriptional repression during reactive oxygen species stress by binding to the chromatin and inhibiting the sirtuin 1 histone deacetylase from performing its function. [11]

ROS-mediated loss of epigenetic imprinting Edit

Epigenetic imprinting is lost during reactive oxygen species stress. This type of oxidative stress causes a loss of NF- κB signaling. Enhancer blocking element CCCTC-binding factor (CTCF) binds to the imprint control region of insulin-like growth factor 2 (IGF2) preventing the enhancers from allowing the transcription of the gene. The NF- κB proteins interact with IκB inhibitory proteins, but during oxidative stress IκB proteins are degraded in the cell. The loss of IκB proteins for NF- κB proteins to bind to results in NF- κB proteins entering the nucleus to bind to specific response elements to counter the oxidative stress. The binding of NF- κB and corepressor HDAC1 to response elements such as the CCCTC-binding factor causes a decrease in expression of the enhancer blocking element. This decrease in expression hinders the binding to the IGF2 imprint control region therefore causing the loss of imprinting and biallelic IGF2 expression. [11]

After the initial exposure to ionizing radiation, cellular changes are prevalent in the unexposed offspring of irradiated cells for many cell divisions. One way this non-Mendelian mode of inheritance can be explained is through epigenetic mechanisms. [11]

Ionizing radiation and DNA methylation Edit

Genomic instability via hypomethylation of LINE1 Edit

Ionizing radiation exposure affects patterns of DNA methylation. Breast cancer cells treated with fractionated doses of ionizing radiation showed DNA hypomethylation at the various gene loci dose fractionation refers to breaking down one dose of radiation into separate, smaller doses. [15] Hypomethylation of these genes correlated with decreased expression of various DNMTs and methyl CpG binding proteins. LINE1 transposable elements have been identified as targets for ionizing radiation. The hypomethylation of LINE1 elements results in activation of the elements and thus an increase in LINE1 protein levels. Increased transcription of LINE1 transposable elements results in greater mobilization of the LINE1 loci and therefore increases genomic instability. [11]

Ionizing radiation and histone modification Edit

Irradiated cells can be linked to a variety of histone modifications. Ionizing radiation in breast cancer cell inhibits H4 lysine tri-methylation. Mouse models exposed to high levels of X-ray irradiation exhibited a decrease in both the tri-methylation of H4-Lys20 and the compaction of the chromatin. With the loss of tri-methylation of H4-Lys20, DNA hypomethylation increased resulting in DNA damage and increased genomic instability. [11]

Loss of methylation via repair mechanisms Edit

Breaks in DNA due to ionizing radiation can be repaired. New DNA synthesis by DNA polymerases is one of the ways radiation induced DNA damage can be repaired. However, DNA polymerases do not insert methylated bases which leads to a decrease in methylation of the newly synthesized strand. Reactive oxygen species also inhibit DNMT activity which would normally add the missing methyl groups. This increases the chance that the demethylated state of DNA will eventually become permanent. [16]

Epigenetic affects on a developing brain Edit

Chronic exposure to these types of radiation can have an effect on children from as early as when they are fetuses. There have been multiple cases reported of hindrance in the development of the brain, behavioral changes such as anxiety, and the disruption of proper learning and language processing. An Increase in the cases of ADHD behavior and autism behavior has been shown to be directly correlated with the exposure of EMF waves. The World Health Organization has classified RFR as a possible carcinogen for its epigenetic effects on DNA expression. The exposure to EMF waves on a consistent 24hr basis has shown to lower the activity of miRNA in the brain affecting developmental and neuronal activity. This epigenetic change causes the silencing of necessary genes along with the change in expression of other genes integral for the normal development of the brain. [2]

MGMT- and LINE1- specific DNA methylation Edit

DNA methylation influences tissue responses to ionizing radiation. Modulation of methylation in the gene MGMT or in transposable elements such as LINE1 could be used to alter tissue responses to ionizing radiation and potentially opening new areas for cancer treatment.

MGMT serves as a prognostic marker in glioblastoma. Hypermethylation of MGMT is associated with the regression of tumors. Hypermethylation of MGMT silences its transcription inhibiting alkylating agents in tumor killing cells. Studies have shown patients who received radiotherapy, but no chemotherapy after tumor extraction, had an improved response to radiotherapy due to the methylation of the MGMT promoter.

Almost all human cancers include hypomethylation of LINE1 elements. Various studies depict that the hypomethylation of LINE1 correlates with a decrease in survival after both chemotherapy and radiotheraphy.

Treatment by DNMT inhibitors Edit

DMNT inhibitors are being explored in the treatment of malignant tumors. Recent in-vitro studies show that DNMT inhibitors can increase the effects of other anti-cancer drugs. Knowledge of in-vivo effect of DNMT inhibitors are still being investigated. The long term effects of the use of DNMT inhibitors are still unknown. [16]