Browse Subject Areas

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

Diet-Induced Obesity Modulates Epigenetic Responses to Ionizing Radiation in Mice

Diet-Induced Obesity Modulates Epigenetic Responses to Ionizing Radiation in Mice

  • Guillaume Vares, 
  • Bing Wang, 
  • Hiroko Ishii-Ohba, 
  • Mitsuru Nenoi, 
  • Tetsuo Nakajima


Both exposure to ionizing radiation and obesity have been associated with various pathologies including cancer. There is a crucial need in better understanding the interactions between ionizing radiation effects (especially at low doses) and other risk factors, such as obesity. In order to evaluate radiation responses in obese animals, C3H and C57BL/6J mice fed a control normal fat or a high fat (HF) diet were exposed to fractionated doses of X-rays (0.75 Gy ×4). Bone marrow micronucleus assays did not suggest a modulation of radiation-induced genotoxicity by HF diet. Using MSP, we observed that the promoters of p16 and Dapk genes were methylated in the livers of C57BL/6J mice fed a HF diet (irradiated and non-irradiated); Mgmt promoter was methylated in irradiated and/or HF diet-fed mice. In addition, methylation PCR arrays identified Ep300 and Socs1 (whose promoters exhibited higher methylation levels in non-irradiated HF diet-fed mice) as potential targets for further studies. We then compared microRNA regulations after radiation exposure in the livers of C57BL/6J mice fed a normal or an HF diet, using microRNA arrays. Interestingly, radiation-triggered microRNA regulations observed in normal mice were not observed in obese mice. miR-466e was upregulated in non-irradiated obese mice. In vitro free fatty acid (palmitic acid, oleic acid) administration sensitized AML12 mouse liver cells to ionizing radiation, but the inhibition of miR-466e counteracted this radio-sensitization, suggesting that the modulation of radiation responses by diet-induced obesity might involve miR-466e expression. All together, our results suggested the existence of dietary effects on radiation responses (especially epigenetic regulations) in mice, possibly in relationship with obesity-induced chronic oxidative stress.


Over the last century, ample evidence has been accumulating of a strong link between carcinogenesis and exposure to ionizing radiation [1]. The deleterious effects of ionizing radiation are considered to result mainly from the direct induction of DNA damage in target cells, but also from non-targeted effects (such as radiation-induced genomic instability and bystander effect) [2]. Ionizing radiation can cause chromosomal aberrations, including acentric fragments, which upon cell replication are excluded from the main nuclei and form micronuclei (MN) [3]. The monitoring of MN in erythrocytes of the bone marrow is an effective tool to assess the genotoxic effects of ionizing radiation even at low levels. Radiation-induced genomic instability is a well-documented phenomenon, which can be observed even in the progeny of irradiated animals, suggesting the involvement of epigenetic mechanisms [4], such as the modulation of genome methylation or the regulation of micro-RNA expression. Recent data suggest that even low dose radiation exposures could also result in epigenetic modifications [5].

A major question for radiation biology is to understand how other risk factors could influence the biological effects of ionizing radiation, especially in terms of cancer risk [6], or reversely how ionizing radiation exposures could potentialize other risk factors. Obesity and associated disorders are increasingly becoming a global health challenge. Obesity is a major risk factor for various metabolic syndromes (such as insulin resistance, type 2 diabetes and nonalcoholic fatty liver disease) [7][9] and for initiation of cancer at several organ sites, including breast, endometrium and kidney [10]. Weight gain and obesity was also related to mortality from liver cancer, pancreatic cancer, non-Hodgkin lymphoma and myeloma [11]. Nonalcoholic fatty liver disease can evolve into nonalcoholic steatohepatitis, which can lead to primary hepatocellular carcinoma [12]. There is compelling evidence that oxidative stress and inflammation are major mechanisms involved in metabolic disorders associated with obesity [13].

The susceptibility to develop diet-induced obesity (DIO) and metabolic syndromes strongly depends on the genetic background, with heritability estimates ranging from 40% to 70% in human populations [14], [15]. Furthermore, a number of studies have shown that various mouse strains and their genetic variants show different metabolic phenotypes [16]. For example, the liver response to a sustained high-fat diet was characterized by the activation of peroxisomal β-oxidation in C57BL/6J mice and by lipogenesis in 129Sc mice [17]. While C57BL/6J mice exhibited severe obesity and diabetes after being fed a high-fat diet, C3H mice fed a similar diet showed normal glucose tolerance and no hepatic steatosis even though they exhibited a significant weight increase [18]. The C57BL/6J mouse strain has become a popular experimental model for evaluating the molecular mechanisms and consequences of DIO, because they develop insulin resistance, hyperglycemia and obesity in a way that closely matches the development of human metabolic syndrome [19]. In comparison, other strains, such as C3H/He, 129/Sv and A/J mice are more resistant to diabetes and obesity [20].

Emerging evidence suggests that dietary effects [21], as well as carcinogenesis [22], involve epigenetic mechanisms, such as DNA methylation changes or miRNA regulations. On the one hand, aberrant gene silencing through promoter hypermethylation has been shown to be a crucial event during early neoplastic progression [23]. High fat diet-induced obesity and metabolic syndromes can also modulate the methylation pattern of various gene promoters in human and animal models, such as leptin [24], peroxisome proliferator activated receptor γ (PPARγ) [25] or microsomal triglyceride transfer protein (Mttp) [26]. On the other hand, there is now ample (and still growing) data describing cancer-associated miRNA regulations, implying the crucial role of miRNAs in maintaining cellular homeostasis [27].

While radiation-induced carcinogenesis was significantly reduced in C3H mice after calorie restriction [28], there is a scarcity of data concerning the modulation of radiation responses by high fat diets or obesity. In a mouse model of malignant glioma, high fat low-carbohydrate ketogenic diet significantly enhanced the effectiveness of radiation therapy [29]. Obesity also increased non-ionizing UV radiation-induced oxidative stress [30].

In this study, we compared the short-term biological responses to ionizing radiation in C57BL/6J and C3H mice fed a control normal fat diet or a high fat (HF) diet. First, we evaluated whether DIO could influence radiation-induced genotoxicity. Then, because liver is one of the main target sites for metabolic syndrome, we measured immediate epigenetic regulations in relationship with diet and ionizing radiation (gene repression through promoter hypermethylation, miRNA regulations).

Materials and Methods


14 weeks-old C57BL/6J DIO mice and 5 weeks-old C3H/HeJ mice were purchased from Charles River Laboratories (Yokohama, Japan) and SLC, Inc. (Hamamatsu, Japan), respectively. C57BL/6J DIO mice are a model of pre-diabetic type 2 diabetes with elevated blood glucose and impaired glucose tolerance. C3H mice are HF diet-induced obesity resistant. Mice were maintained for 3 (C57BL/6J) or 12 weeks (C3H) in a conventional animal facility under a 12-h light/12-h dark photoperiod (lights on from 7∶00 a.m. to 7∶00 p.m.). Animals were housed in autoclaved cages with sterilized wood chips and allowed free access to 10% fat (D12450B, Research Diets Inc., New Brunswick, NJ, USA) or 60% fat laboratory chow (D12492, Research Diets Inc.) and acidified water (pH = 3.0±0.2). The 60% fat chow will be referred to as “high fat diet” (HF diet). All experimental protocols involving mice were reviewed and approved by The Institutional Animal Care and Use Committee of the National Institute of Radiological Sciences (NIRS) and were performed in accordance with the NIRS Guidelines for the Care and Use of Laboratory Animals.

Cell culture and free fatty acid treatment

AML12 mouse liver cells (ATCC, CRL-2254) were grown in DMEM/F12 medium (Gibco, Gaithersburg, MD, USA) supplemented with 0.005 mg/ml insulin, 0.005 mg/ml transferrin, 5 ng/ml selenium, and 40 ng/ml dexamethasone (ITS Media Supplement, Sigma-Aldrich, St Louis, MO, USA) and 10% fetal bovine serum (FBS, Nichirei Biosciences, Tokyo, Japan). Oleic acid and palmitic acid (Sigma-Aldrich) were conjugated to bovine serum albumin at a molar ratio of 6∶1. Cells were treated with a mixture of oleic and palmitic acid for 24 hours before irradiation.


Mice were exposed to four times daily doses of 0.75 Gy X-rays (0.25 Gy/min) generated by a Pantak 320S machine (Shimadzu, Japan) operated at 200 kVp and 20 mA, using a 0.50-mm Al +0.50-mm Cu filter. An exposure rate meter (AE-1321M, Applied Engineering Inc., Japan) was used for the dosimetry. Mice were sacrificed one day after the final irradiation.

AML12 cells were irradiated in serum-free medium 72 hours after plating, using an X-ray generator (ISOVOLT Titan-320, General Electric, Fairfield, CT, USA) at a dose-rate of 0.9 Gy/min.

Bone marrow micronucleus test

Diet- and radiation-induced genotoxicity was assessed using the standard bone marrow MN test according to the method published by Hayashi et al. [31]. Bone marrow smears were prepared from both femurs. Micronuclei were counted in immature polychromatic erythrocytes (PCEs) and in mature normochromatic erythrocytes (NCEs). The frequencies of micronuclei in PCEs and NCEs are referred to as MNPCEs and MNNCEs, respectively. The ratio of PCEs to mature NCEs (P/N ratio) is an indicator of the relative proliferation rate in the erythroid lineage, and its decrease is considered to be an indicator of mutagen-induced cytotoxicity [32]. At least 15000 cells (PCE+NCE) per mouse were counted and the data for each experimental point were obtained from at least 5 mice.

Flow cytometry

To identify cells of the erythroid lineage, bone marrow cells were incubated immediately after collection, with FITC-conjugated TER-119 antibody (557915, BD Pharmingen, San Diego, CA, USA) for 20 minutes at 4°C then washed, fixed with paraformaldehyde/methanol [33] and resuspended in cold PBS with 3% FBS, 50 µg/mL propidium iodide and 1 µg/mL RNase. The cell cycle repartition of erythroid cells was then analyzed on a FACSCalibur flow cytometer (Becton Dickinson, San Jose, CA, USA) with the CellQuest software (Becton Dickinson). Cell cycle analysis was performed using ModFit LT software (Verity Software House, Topsham, ME, USA).

Methylation PCR array

For methylation analysis of mouse liver tissues, the EpiTect Methyl qPCR profiling service (Filgen, Nagoya, Japan) was used. Genomic DNA was extracted from C57BL/6J mouse livers using the QIAmp DNA Mini Kit (Qiagen, Chatsworth, CA). The methylation profiling of 22 liver cancer-related genes (Ep300, Cdkn1b, Cdkn2a, Fhit, Pycard, Tnfrsf10d, Cdh1, Dlc1, Opcml, Reln, Ccnd2, Cdkn1a, Rassf1, Wt1, Socs1, Msh2, Msh3, Gstp1, Reln, E2f1, Runx3, Sfrp2) was measured using EpiTect Methyl qPCR arrays according to the manufacturer’s instructions (Qiagen). This system does not require bisulfite conversion of DNA and relies on methylation-sensitive restriction enzymes. Briefly, 1 µg DNA was digested overnight at 37°C with the Methyl-Profiler DNA Methylation Enzyme Kit (Qiagen) containing methylation-sensitive and methylation-specific restriction enzymes, which digest unmethylated and methylated DNA, respectively. Then enzymatic activity was inactivated at 65°C for 20 min. The remaining DNA in each individual enzyme reaction was marked with SYBR Green then quantified using a 7500 real-time PCR system (Applied Biosystems, Foster City, CA, USA) using primers specific to one of the 22 liver cancer-related genes. The relative proportions of unmethylated and methylated DNA were then measured relative to the total DNA for each gene (measured when no restriction enzymes were added).

Methylation-specific PCR

Genomic DNA was extracted from C57BL/6J mouse livers using the NucleoSpin TriPrep kit (Macherey-Nagel, Düren, Germany) according to the manufacturer’s instructions. 500 ng DNA was denatured using 0.3M NaOH (42°C, 30 min) then bisulfite modification was carried out by adding 1020 µL 40.5% sodium bisulfite, 60 µL 10 mM hydroquinone and 10 µL H2O, overlaying the mix with mineral oil and incubating at 55°C for 16 hours. The modified DNA was then purified using the Wizard DNA cleanup system (Promega) and resuspended in 100 µL TE. The purified DNA samples were then denatured (with NaOH) and precipitated with ethanol and stored at –20°C until use. The promoter methylation status of p16, Mgmt and Dapk genes was determined by methylation-specific PCR using primer sets described by Sharpless et al., Yamada et al. and Mittag et al, respectively [34][36].

Microarray analysis of miRNA levels

Total RNA was extracted from the livers of C57BL/6J mice (n = 3 per experimental condition) using TRIzol reagent (Invitrogen, Carlsbad, CA, USA). The profiling of miRNA expression levels was performed using the miRCURY LNA miRNA profiling service (Filgen). The quality of total RNA was checked using a Bioanalyzer 2100 (Agilent, Santa Clara, CA, USA), then RNA was labeled with Hy3 and hybridized to miRCURY LNA microRNA arrays 7th generation (Exiqon, Vedbaek, Denmark). Slides were scanned using a laser scanner (Molecular Devices, Sunnyvale, CA, USA) and the resulting images were analyzed using Array-Pro v.4.5 (Media Cybernetics, Bethesda, MD, USA). Following background correction, quantile normalization was applied, so that the distribution remains the same across arrays [37]. Unsupervised hierarchical clustering and heatmap vizualisation of the samples for murine probes were performed using the CIMminer online software ( Differentially expressed miRNA were identified by using the t-test within Significance Analysis of Microarrays (SAM) and median normalization [38]. The Diana miRPath tool v.2.1 [39] coupled with microT-CDS database was used to determine KEGG pathway [40] enrichment within the putative targets for the differentially expressed miRNAs. All array data have been submitted to the Gene Expression Omnibus (GEO) under the accession number GSE47956.

Real-time PCR

For the validation of methylated genes expression, we used primers for Socs1, Ep300, p16, Mgmt and Dapk (QuantiTect Primer Assay, Qiagen); for the evaluation of miR-466e-5p gene targets expression, we used primers for Zfp704, Cplx2 and Cnot7 [41], [42]; for the validation of miRNA microarrays, we used primers for miR-466e-5p, miR-21-3p and miR-185-3p (MystiCq microRNA qPCR Assay Primer, Sigma-Aldrich). Mouse liver RNA was reverse-transcribed using the RT2 first strand kit (SABiosciences, Frederick, MD, USA) for gene expression experiments and MystiCq microRNA cDNA Synthesis Mix (Sigma-Aldrich) for miRNA expression experiments. Then quantitative real-time PCR reactions were performed in triplicate with an Applied Biosystems 7300 Real-Time PCR system (Life Technologies), using the RT2 SYBR Green PCR Master Mix (SABiosciences) for gene expression experiments and the MystiCq microRNA SYBR Green qPCR ReadyMix for miRNA expression experiments, according to the manufacturers’ instructions.

Measurement of ROS levels

Intracellular levels of reactive oxygen species (ROS) in AML12 cells were measured using 5-(and-6)-chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H2DCFDA, Molecular Probes, Eugene, OR, USA). Cells were plated in 12-well plates and loaded with pre-warmed PBS containing a mixture of oleic and palmitic acid for 12 hours, then 10 µM CM-H2DCFDA was added and cells were incubated for 40 minutes. Fluorescence intensities were measured using a SpectraMax M5 microplate reader (Molecular Devices, Sunnyvale, CA, USA) (excitation at 493 nm, emission at 520 nm). Unstained cells were used as negative control.

Clonogenic assay

miR-466e inhibitor (Sigma-Aldrich) was transfected 48 hours before irradiation into AML12 cells using Lipofectamine RNAiMax reagent (Invitrogen), according to the manufacturer’s instructions (lipofectamine alone was added to control cells). Free fatty acids were added 24 hours before irradiation. After irradiation, cells treated or not with fatty acids were seeded at defined densities and incubated for 10–14 days then stained. Colonies with >50 cells were scored and surviving fractions were determined after correcting for the plating efficiency. Survival curve data were fitted to the linear-quadratic model [43] and are presented as the mean of at least three independent experiments.


Diet-induced obesity does not modulate radiation-induced genotoxicity in the bone marrow

Pre-diabetic obese C57BL/6J mice fed a high fat (HF) diet for several weeks kept gaining weight in our laboratory (Figure S1). C3H mice also accumulated additional weight when fed a HF diet, but did not exhibit metabolic syndromes. In order to evaluate whether DIO could influence radiation-induced genotoxicity in C57BL/6J and C3H mice, we measured bone marrow micronuclei frequencies after irradiation (Figure 1). Exposure to ionizing radiation resulted in increased MNPCE incidence regardless of diet in both mouse strains. No significant difference in MNPCE frequencies was observed between obese and non- obese animals, suggesting that DIO did not modulate radiation-induced genotoxicity.

Figure 1. Bone marrow micronucleus test.

A and C: Frequency of micronuclei in the bone marrow cells of C57BL/6J (A) and C3H (C) mice. B and D: ratio of polychromatic erythrocytes to normochromatic erythrocytes (P/N ratio) in the bone marrow of C57BL/6J (B) and C3H (D) mice. *p<0.05 compared to non-irradiated control (for irradiated mice fed a normal diet) or obese mice (for irradiated mice fed a HF diet).

A significant decrease in P/N ratios was observed after irradiation in non-obese C57BL/6J and C3H mice (Figure 1B, D), indicative of cytotoxic damage or radiation-induced inhibition of proliferation. However, while exposure to ionizing radiation did also result in lower P/N ratio in obese C3H mice, C57BL/6J did not exhibit a significant radiation-induced decrease in P/N ratio. In order to determine whether the modulation of P/N ratio involved any modulation of erythroid cell proliferation, we studied the cell cycle repartition of nucleated erythroid progenitor and precursor cells in the bone marrow (Figure 2). TER-119+ cells are of the erythroid lineage and represent approximately 20% of the total cell number in the bone marrow. No significant difference in cell proliferation was observed in C57BL/6J mice after irradiation or depending on diet. In C3H mice, erythroid progenitors and precursor cells exhibited a lower entry into S/G2/M after irradiation or HF diet (p<0.01). The lower percentage of S/G2/M cells in irradiated animals suggested that the observed decreased P/N ratio resulted from an inhibition of cell proliferation of erythroid progenitors and precursors in response to radiation. However, surprisingly, C3H mice fed a HF diet did not show decreased numbers of erythroid cells in S/G2/M after irradiation, which may suggest that in obese mice, the cytotoxic effects of radiation affect mainly PCEs but not erythroid progenitors.

Figure 2. Cell cycle repartition of bone marrow TER-119+ nucleated cells.

TER-119 was used as a lineage marker for erythroid cells. Analysis was performed in C57BL/6J (A) and C3H (B) mice using ModFit LT software. Results are the average of three independent experiments. Error bars represent standard deviation. *p<0.05 compared to control.

Diet-induced obesity is associated with immediate modulation of promoter methylation in the liver

Even though DIO did not apparently modulate radiation-induced genotoxicity, we wondered whether immediate effects at the molecular or cellular level could be observed, which could ultimately influence radiation sensitivity and overall radiation risks in obese mice. Because liver is a major target organ for the metabolic syndromes resulting from DIO (including fatty liver disease), we then focused our investigations on effects in the liver.

Promoter-specific hypermethylation is a frequent event in liver carcinogenesis [44]and is dependent on diet [45]. For this reason, we measured the methylation status of 27 gene promoters in the livers of C57BL/6J mice, using methylation PCR arrays and MSP (Figure 3). Methylation array provided us with methylation levels in the promoters of 24 liver cancer-related genes (Figure 3A). The percentages of highly-methylated DNA (as a fraction of total input DNA) never exceeded 10% for most of the genes, except E1A binding protein p300 (Ep300) and suppressor of cytokine signaling 1 (Socs1). It exceeded 20% for Ep300 and Socs1 genes in non-irradiated animals fed a HF diet, but it dropped to 7.8% and 12.4%, respectively, after irradiation. Our results suggested that Ep300 expression might be inhibited in obese mice through promoter methylation, but not after irradiation. The results were less clear for Socs1, whose methylation levels were already elevated in control animals.

Figure 3. Gene promoter methylation in C57BL/6J mouse livers.

A: Heatmap representing the promoter methylation status for 24 liver cancer-related genes in the livers of non-irradiated and irradiated C57BL/6J mice fed a normal or HF diet. B: Representative gels showing the methylation-specific PCR analysis of the promoter methylation status for p16, Mgmt and Dapk genes.

Previous investigations have described in various human and animal models the epigenetic regulation of promoter methylation for the tumor suppressors p16 and Dapk, as well as for the O6-methylguanine-DNA methyltransferase Mgmt [46][49]. For this reason, we also measured by MSP the methylation status of these three genes in our model (Figure 3B). While no methylation was observed in control animals, HF diet triggered the methylation of p16, Mgmt and Dapk gene promoters (both in irradiated and in non-irradiated animals). Mgmt promoter was also methylated after irradiation in mice fed a normal diet. Interestingly, expression levels of Ep300, Socs1, p16, Mgmt and Dapk in obese and irradiated mice were significantly lower than in control mice (Figure 3C).

Overall our results suggest that the expression levels of several anti-tumour genes might be modulated by rapid promoter methylation in obese mice, with potential consequences in terms of radiation responses or carcinogenic risk (if sustained over the long-term).

Diet-induced obesity influences micro-RNA regulations in the liver after irradiation

In addition to the regulation of gene expression by promoter methylation, diet-related epigenetic processes can also involve miRNAs. miRNA expression profiles in C57BL/6J mouse livers were analyzed by using locked nucleic acid (LNA) [50] miRNA arrays. Both HF diet and irradiation resulted in few consistent miRNA regulations among the sampled animals (three animals per experimental group) (Figure S2). Therefore SAM analysis was performed to compare miRNA expression levels between each experimental group (Table 1). SAM analysis provided a limited number of miRNAs that were consistently deregulated in every animal of each experimental group. Most of the observed modulations were upregulations. When compared to control animals (normal diet, non-irradiated), only one miRNA, miR-466e-5p, was upregulated in obese mice (no miRNA was downregulated). The putative targets genes of miR-466e-5p were identified by miRPath (using microT-CDS database) and the Biosynthesis of fatty acids KEGG pathway was identified by pathway enrichment analysis (p<0.01). After irradiation, 33 miRNAs were upregulated and 1 was downregulated (Table 1). 39 KEGG pathways (p<0.01) were found to be altered by miRPath (Table 2), including Pathways in cancer, TGF-β signaling, MAPK signaling, Focal adhesion, Apoptosis and Wnt signaling pathway, which are all known functions associated with radiation exposure [51][53]. On the contrary, in irradiated obese mice, only one miRNA (miR-3961) was upregulated compared to control and no miRNA was modulated compared to non-irradiated obese mice: the above-mentioned radiation-induced miRNA regulations were not observed in obese mice. We verified by real-time quantitative PCR the expression levels of miR-466e-3p, miR-185-3p and miR-21-3p; both miR-466-e-3p and miR-21-3p were upregulated after high-fat diet and irradiation, respectively (Table S1). To evaluate the biological significance of miR-466e-3p upregulation in obese mice, we measured by real-time quantitative PCR the expression levels of three target genes: Zfp704, Cnot7 and Cplx2 (Figure 4). These three genes were downregulated in mice fed a HF diet.

Figure 4. Expression levels of miR-466e-3p target genes.

Expression levels of Zfp704, Cnot7 and Cplx2, relative to control, were measured by quantitative real-time PCR. Results are the average of three independent experiments. Error bars represent standard deviation.

Table 1. miRNA regulations in the livers of non-irradiated and irradiated C57BL/6J mice fed a normal or HF diet, determined using Significance Analysis of Microarrays (SAM).

Table 2. Altered KEGG pathways (p<0.01) associated with the putative target genes of the miRNAs regulated by ionizing radiation exposure in mice fed a normal diet.

Free fatty acids generate oxidative stress and sensitize liver cells to ionizing radiation

In order to mimic in vitro the effects of obesity on radiation responses in the mouse liver, we treated AML12 murine liver cells with free fatty acids (FFAs: palmitic acid and oleic acid) for 12 to 24 hours. Twelve hours after FFA administration, ROS levels increased significantly, reminiscing of chronic oxidative stress in C57BL6 DIO mouse livers; the addition of miR-466e inhibitor did not influence ROS levels (Figure 5). The reproductive clonogenic viability of AML12 cells 24 hours after FFA treatment was determined using the clonogenic assay (Figure 6). The dose that gave 5% mean clonogenic survival (DL95) was lower after FFA treatment (FFA: 5.27 Gy, Control: 6.12 Gy), indicating that FFAs sensitized cells to ionizing radiation. However, FFA-triggered sensitization was counteracted by the administration of miR-466e inhibitor, suggesting that miR-466e expression might influence radiation-sensitivity.

Figure 5. ROS levels in AML12 cells.

ROS levels were measured after treatment with free fatty acids (FFAs) oleic acid and palmitic acid and with miR-466e inhibitor. Results are the average of three independent experiments. Error bars represent standard deviation. Asterisks denote significant differences (t-test, *p<0.01).

Figure 6. Dose-response curves for clonogenic survival of AML12 cells.

AML12 cells were treated for 24 hours with free fatty acids (FFAs: oleic acid, palmitic acid) alone (circles, dotted line) or both with miR-466e inhibitor and FFAs (triangles, continuous line) then exposed to various doses of ionizing radiation. Lipofectamine alone was added to control cells (squares, continuous line). Lines represented fitted curves according to linear quadratic regression. The red dotted line indicates 5% clonogenic survival (DL95). Results are the average of at least three independent experiments. Error bars represent standard deviation. Statistical significance of the difference between dose-response curves was performed using one-way Analysis of Variance (one-way ANOVA) with Bonferroni correction for pairwise group comparisons. *p<0.01 compared to control.


Given the importance of diet in the development of obesity-associated metabolic syndromes, a significant amount of interest has been given to the molecular analysis of metabolic disruptions in obesity. In particular, there is growing evidence showing that diet can strongly influence epigenetic processes [21]. For this reason, we wondered whether metabolic disturbances associated with obesity were susceptible to influence radiation sensitivity.

In order to understand the combinational effects of DIO and ionizing radiation in mice, we exposed C57BL/6J and C3H mice to fractionated doses of X-rays. DIO mice exhibit excessive body weight, body fat accumulation, prediabetes type 2 and metabolic syndrome. C3H mice were also studied because they are comparatively resistant to metabolic syndrome and atherosclerosis [54]. High 60% fat (HF) chow provided an energy density of 5.24 kcal/g (compared to 3.85 kcal/g for the control 10% fat chow) and mice fed a HF diet exhibited an accumulation of body fat and became obese, while mice fed a control diet did not.

There are ample amounts of evidence describing increased bone marrow MN frequency after radiation exposure in mice [55][59]. On the contrary, HF diet did not modulate bone marrow MN frequency both in non-irradiated and irradiated mice, in contradiction with human data describing a correlation between peripheral blood MN frequency and pathologies associated with metabolic syndrome and obesity [60]. The P/N ratio was higher in C3H than in C57BL/6J control mice, suggesting that these mice have a higher rate of erythropoiesis. Whereas no significant proliferation changes were observed in C57BL/6J mice, both exposure to radiation and HF diet resulted in lower erythroid precursors cell proliferation in C3H mice. In obese animals, it was suggested that bone marrow adipogenesis may impair erythropoiesis, resulting in anemia [61]. Rapidly cycling early erythroid progenitors are highly radiosensitive and encounter cell cycle damage checkpoints and apoptosis in response to radiation, compared to more mature erythroid cells [62]. Interestingly, this inhibition of proliferation was not observed in obese C3H mice after irradiation, suggesting that the progenitor cells might not respond to radiation exposure the same way in obese mice than in control mice, or that in response to both HF diet and radiation exposure, erythropoiesis might be stimulated. Additional investigations are necessary to understand the underlying mechanisms.

Obesity and obesity-related complications play a significant role in the pathogenesis of liver diseases and liver cancer [63], [64]; for this reason, we focused our analysis on the interplay between radiation-responses and DIO in the mouse liver. Because DNA methylation is known to play a crucial role in carcinogenesis by regulating the expression of tumor suppressor or DNA repair genes [65], we measured the methylation profiles of several genes associated with liver cancer. While short-term modulation of promoter methylation was previously observed in C57BL/6J mice exposed to acute or chronic X-ray irradiation [66], only the Mgmt gene presented a methylated form after irradiation in animals fed a normal diet. Mgmt plays a major role in protecting cells from the genotoxic and carcinogenic effects of methylating mutagens [67]. Increased methylation of the Mgmt gene promoter might result in lower Mgmt gene expression levels, in contradiction with previous observations showing that genotoxic stress (including ionizing radiation) can induce Mgmt expression in rat liver cells [68], [69]. However, MSP does only provide qualitative results at best and additional investigations are required to measure Mgmt gene expression levels in our model. Our results also suggested that DIO was associated with increased hypermethylation of Ep300, p16 and Dapk gene promoters. Inactivation of tumor suppressor p16 by aberrant methylation is frequently observed in carcinomas and precancerous lesions of various organs [48], [49]. Similarly, aberrant methylation of the apoptosis-related Dapk promoter is extremely common in cancers [34], [46]. These tumor suppressor genes exhibited promoter hypermethylation only in obese animals, suggesting potential effects in terms of radiation response and carcinogenesis. Expression levels of p16, Mgmt, Dapk, Ep300 and Socs1 were systematically lower in obese and irradiated animals, suggesting that promoter methylation contributes to gene repression in these animals.

In a similar model of DIO in C57BL/6J mice, microarray analysis revealed that the expression of 97 genes was significantly modulated in the livers of mice fed a HF diet, compared to mice fed a normal-fat diet [70]. These genes were mainly involved in metabolism, stress defense mechanisms (such as protection against oxidative damage) and inflammatory processes. Other microarray analyses revealed that a number of genes are deregulated in response to ionizing radiation [71], [72], including at very low dose-rates [73], [74]. Because gene expression can be modulated by miRNAs, we studied diet- and radiation-associated miRNA regulations in C57BL/6J mouse livers.

Recent studies have described obesity-related miRNA regulations in mouse and human models. These miRNAs were involved in various processes such as adipocyte differentiation, metabolic integration, insulin resistance and appetite regulation [75]. HF diet-associated miRNA up- (miR-22, miR-342-3p, miR-142-3p and others) and down-regulations (miR-200b, miR-200c, miR-204 and others) were observed in the adipose tissues of C57BL/6J mice [76]. In the liver, there is convincing evidence that miR-122 is involved in cholesterol and lipid metabolism, and the silencing of miR-122 has resulted in decreased cholesterol levels in mice and monkeys [77][79]. We did not observe any significant modulation of miR-122 in the livers of obese and/or irradiated mice. However, using SAM analysis, we identified a new HF diet-associated miRNA upregulation (miR-466e); additionally, the modulation of miR-709 expression was under the fold-change cut-off value of 1.5 but was potentially associated with HF diet as suggested by SAM analysis. miR-709 was recently described as a tumor-suppressor miRNA targeting Myc, Akt and Ras, which participates in the regulation of apoptosis through the miR-15a/miR-16-1 pathway [80], [81]. HF diet resulted in the down-regulation of three putative miR-466e target genes (Zfp704, Cnot7, Cplx2), and it is highly likely that a number of genes are regulated in response to miR-466e up-regulation. A functional analysis of the putative target genes of miR-709 and miR-466e suggested that these miRNAs might be involved in lipid metabolism. miR-466e gene is included in the Chromosome 2 miRNA cluster (C2MC, or Sfmbt2 cluster), a mouse-specific large miRNA cluster located in intron 10 of the Sfmbt2 gene. C2MC contains 71 closely related miRNA genes modulated in response to cellular stress (nutrition deprivation, hyperglycemia, hypertonic stress, oxidative stress, xenobiotic stress, aging…) and involved in cell proliferation and apoptosis, cell fate decision and immune response [82][85].

For this reason, we hypothesized that miR-466e might also be involved in stress response in the mouse liver, and we evaluated in vitro the effects of free fatty acid (FFA) administration and the potential role of miR-466e in the response of AML12 mouse liver cells to radiation. Although miR-466e inhibition did not modulate FFA-induced ROS levels, it counteracted the radiosensitization of AML12 cells by FFAs. Interestingly, it was recently shown that oxidative stress triggered acetylation in the miR-466h-5p promoter region, leading to the activation of this miRNA closely related to miR-466e [82]. Further investigations are needed to understand whether miR-466e is regulated through similar mechanisms, and to decipher the biological function of miR-466e.

We observed that a number of miRNAs were upregulated after irradiation in mice fed a normal diet. A number of other studies have recently identified miRNAs associated with radiation responses [86], [87]. In particular, miR-21, which targets many genes in the apoptotic pathway, was consistently seen to be upregulated after exposure to ionizing radiation in various models [88][91]; miR-21 was also upregulated in radiation-induced thymic lymphoma in BALB/c mice [92]. Many of the deregulated miRNAs in our study are not yet fully characterized and additional investigations will likely provide more insights into their potential involvement in radiation responses. Many of the molecular pathways identified by the miRPath tool in irradiated mice fed a normal diet were known to be associated with radiation responses and/or cancer, such as the TGF-β [93], MAPK [94], ErbB [95] and Wnt [53] signaling pathways, apoptosis, etc. Interestingly, these radiation-induced miRNA regulations were not observed in irradiated obese mice. This could not be explained by a previous up-regulation associated with HF diet, since only one miRNA was deregulated in obese mice.

We therefore suggest that the molecular pathways normally involved in radiation responses are disrupted in obese mice. The question remains whether those might influence long-term effects, such as radiation-induced carcinogenesis. It is highly possible that because obese animals experience chronic oxidative stress (as reported by others and suggested by our immunohistochemistry results) [96], they are further sensitized to ionizing radiation risks. 8-OHdGs in CpG sequences were shown to inhibit the methylation of neighboring cytosine residues, resulting in DNA hypomethylation [97]. However, obesity was associated in our model with increased promoter methylation for several genes.

A substantial body of evidence shows that oxidative stress is a major contributor to the deleterious effects of obesity [98]. The expansion of abdominal fat results in elevated levels of free fatty acids (FAAs) [99], [100], which stimulate reactive oxygen species (ROS) production [98] and induce hepatic insulin resistance as well as increased glucose production. Furthermore, decreased antioxidant capacity levels and increased DNA damage were described in relationship with obesity-induced chronic oxidative stress [96]. The disruption of genetic and epigenetic mechanisms by obesity-induced chronic oxidative stress might result in a deficient response to ionizing radiation. For example, it was shown that the deregulation of protein kinase C (PKC) signaling, an early responder to ionizing radiation [101], is a crucial event in obesity-related metabolic syndrome and oxidative stress [102], [103]. Irradiation in obese animals might thus result in increased cancer risk. Furthermore, miRNA regulations (such as the upregulation of miR-466e) are likely to play a significant role in regulating radiation response.

Overall, it is necessary to better understand the crosstalk between obesity-related metabolic syndrome and radiation responses, both at low and high dose ranges, in order to better evaluate the risks and effects of ionizing radiation resulting from natural or medical exposures, or to develop specific radiation-therapy regimens adapted to each patient characteristics.

Supporting Information

Figure S1.

Time-course of weight gain in mice. C57BL/6J (left) and C3H (right) mice were fed a normal (black symbols) and a HF diet (colored symbols).


Figure S2.

Heatmap of miRNA expression in the livers of obese and/or irradiated C57BL/6J DIO mice. Unsupervised hierarchical clustering of 1157 mouse miRNAs (100% of known mouse miRNAs) was performed. C: control (mice No. 116, 117, 118); HF: HF diet (mice No. 106, 107, 108); R: irradiated (mice No. 111, 112, 113); HFR: HF diet, irradiated (mice No. 101, 102, 103).


Table S1.

Real-time PCR verification of miRNA expression. Expression ratios (compared to control) of miR-466e-3p, miR-185-3p and miR-21-3p were measured by RT-PCR. Green values indicate miRNAs identified as upregulated in the SAM microarray analysis.



We thank Ms Hiromi Arai, Taeko Iwai and Kyoko Sakuma for their technical assistance.

Author Contributions

Conceived and designed the experiments: GV TN MN. Performed the experiments: GV TN MN BW HI. Analyzed the data: GV TN. Contributed reagents/materials/analysis tools: GV TN MN BW HI. Wrote the paper: GV.


  1. 1. Little JB (2000) Ionizing Radiation. In: Holland-Frei Cancer Medicine. Hamilton (ON): BC Decker. pp. Chapter 14.
  2. 2. Wright EG (2010) Manifestations and mechanisms of non-targeted effects of ionizing radiation. Mutat Res 687: 28–33.
  3. 3. Heddle JA, Carrano AV (1977) The DNA content of micronuclei induced in mouse bone marrow by gamma-irradiation: evidence that micronuclei arise from acentric chromosomal fragments. Mutat Res 44: 63–69.
  4. 4. Ilnytskyy Y, Kovalchuk O (2011) Non-targeted radiation effects-an epigenetic connection. Mutat Res 714: 113–125.
  5. 5. Bernal AJ, Dolinoy DC, Huang D, Skaar DA, Weinhouse C, Jirtle RL (2013) Adaptive radiation-induced epigenetic alterations mitigated by antioxidants. FASEB J 27: 665–671.
  6. 6. Wakeford R (2012) Radiation effects: Modulating factors and risk assessment – an overview. Ann ICRP 41: 98–107.
  7. 7. Könner AC, Brüning JC (2012) Selective insulin and leptin resistance in metabolic disorders. Cell Metab 16: 144–152.
  8. 8. Marchesini G, Moscatiello S, Domizio SD, Forlani G (2008) Obesity-Associated Liver Disease. Journal of Clinical Endocrinology & Metabolism 93: s74–s80.
  9. 9. Centers for Disease Control and Prevention (CDC) (2004) Prevalence of overweight and obesity among adults with diagnosed diabetes–United States, 1988–1994 and 1999–2002. MMWR Morb Mortal Wkly Rep 53: 1066–1068.
  10. 10. Bianchini F, Kaaks R, Vainio H (2002) Overweight, obesity, and cancer risk. Lancet Oncol 3: 565–574.
  11. 11. Calle EE, Rodriguez C, Walker-Thurmond K, Thun MJ (2003) Overweight, obesity, and mortality from cancer in a prospectively studied cohort of U.S. adults. N Engl J Med 348: 1625–1638.
  12. 12. Torres DM, Harrison SA (2008) Diagnosis and therapy of nonalcoholic steatohepatitis. Gastroenterology 134: 1682–1698.
  13. 13. Fernández-Sánchez A, Madrigal-Santillán E, Bautista M, Esquivel-Soto J, Morales-González A, et al. (2011) Inflammation, oxidative stress, and obesity. Int J Mol Sci 12: 3117–3132.
  14. 14. Lu Y, Loos RJ (2013) Obesity genomics: assessing the transferability of susceptibility loci across diverse populations. Genome Med 5: 55.
  15. 15. Elks CE, den Hoed M, Zhao JH, Sharp SJ, Wareham NJ, et al. (2012) Variability in the heritability of body mass index: a systematic review and meta-regression. Front Endocrinol (Lausanne) 3: 29.
  16. 16. Nishikawa S, Yasoshima A, Doi K, Nakayama H, Uetsuka K (2007) Involvement of sex, strain and age factors in high fat diet-induced obesity in C57BL/6J and BALB/cA mice. Exp Anim 56: 263–272.
  17. 17. Sabidó E, Wu Y, Bautista L, Porstmann T, Chang C-Y, et al. (2013) Targeted proteomics reveals strain-specific changes in the mouse insulin and central metabolic pathways after a sustained high-fat diet. Molecular Systems Biology 9.
  18. 18. Lee YJ, Ko EH, Kim JE, Kim E, Lee H, et al. (2012) Nuclear receptor PPARγ-regulated monoacylglycerol O-acyltransferase 1 (MGAT1) expression is responsible for the lipid accumulation in diet-induced hepatic steatosis. Proc Natl Acad Sci U S A 109: 13656–13661.
  19. 19. Collins S, Martin TL, Surwit RS, Robidoux J (2004) Genetic vulnerability to diet-induced obesity in the C57BL/6J mouse: physiological and molecular characteristics. Physiol Behav 81: 243–248.
  20. 20. Almind K, Kahn CR (2004) Genetic Determinants of Energy Expenditure and Insulin Resistance in Diet-Induced Obesity in Mice. Diabetes 53: 3274–3285.
  21. 21. Supic G, Jagodic M, Magic Z (2013) Epigenetics: a new link between nutrition and cancer. Nutr Cancer 65: 781–792.
  22. 22. Choi JD, Lee J-S (2013) Interplay between Epigenetics and Genetics in Cancer. Genomics Inform 11: 164–173.
  23. 23. Jones PA, Baylin SB (2007) The epigenomics of cancer. Cell 128: 683–692.
  24. 24. Milagro FI, Campión J, García-Díaz DF, Goyenechea E, Paternain L, et al. (2009) High fat diet-induced obesity modifies the methylation pattern of leptin promoter in rats. J Physiol Biochem 65: 1–9.
  25. 25. Fujiki K, Kano F, Shiota K, Murata M (2009) Expression of the peroxisome proliferator activated receptor γ gene is repressed by DNA methylation in visceral adipose tissue of mouse models of diabetes. BMC Biology 7: 38.
  26. 26. Chang X, Yan H, Fei J, Jiang M, Zhu H, et al. (2010) Berberine reduces methylation of the MTTP promoter and alleviates fatty liver induced by a high-fat diet in rats. J Lipid Res 51: 2504–2515.
  27. 27. Palanichamy JK, Rao DS (2014) miRNA dysregulation in cancer: towards a mechanistic understanding. Front Genet 5: 54.
  28. 28. Yoshida K, Inoue T, Nojima K, Hirabayashi Y, Sado T (1997) Calorie restriction reduces the incidence of myeloid leukemia induced by a single whole-body radiation in C3H/He mice. Proc Natl Acad Sci U S A 94: 2615–2619.
  29. 29. Abdelwahab MG, Fenton KE, Preul MC, Rho JM, Lynch A, et al. (2012) The ketogenic diet is an effective adjuvant to radiation therapy for the treatment of malignant glioma. PLoS One 7: e36197.
  30. 30. Katiyar SK, Meeran SM (2007) Obesity increases the risk of UV radiation-induced oxidative stress and activation of MAPK and NF-kappaB signaling. Free Radical Biology and Medicine 42: 299–310.
  31. 31. Hayashi M, Sofuni T, Ishidate M (1984) Kinetics of micronucleus formation in relation to chromosomal aberrations in mouse bone marrow. Mutat Res 127: 129–137.
  32. 32. Suzuki Y, Nagae Y, Li J, Sakaba H, Mozawa K, et al. (1989) The micronucleus test and erythropoiesis. Effects of erythropoietin and a mutagen on the ratio of polychromatic to normochromatic erythrocytes (P/N ratio). Mutagenesis 4: 420–424.
  33. 33. Pollice AA, Philip Mccoy J, Shackney SE, Smith CA, Agarwal J, et al. (1992) Sequential paraformaldehyde and methanol fixation for simultaneous flow cytometric analysis of DNA, cell surface proteins, and intracellular proteins. Cytometry 13: 432–444.
  34. 34. Mittag F, Kuester D, Vieth M, Peters B, Stolte B, et al. (2006) DAPK promotor methylation is an early event in colorectal carcinogenesis. Cancer Lett 240: 69–75.
  35. 35. Sharpless NE, Bardeesy N, Lee KH, Carrasco D, Castrillon DH, et al. (2001) Loss of p16Ink4a with retention of p19Arf predisposes mice to tumorigenesis. Nature 413: 86–91.
  36. 36. Yamada H, Vijayachandra K, Penner C, Glick A (2001) Increased sensitivity of transforming growth factor (TGF) beta 1 null cells to alkylating agents reveals a novel link between TGFbeta signaling and O(6)-methylguanine methyltransferase promoter hypermethylation. J Biol Chem 276: 19052–19058.
  37. 37. Bolstad BM, Irizarry RA, Astrand M, Speed TP (2003) A comparison of normalization methods for high density oligonucleotide array data based on variance and bias. Bioinformatics 19: 185–193.
  38. 38. Tusher VG, Tibshirani R, Chu G (2001) Significance analysis of microarrays applied to the ionizing radiation response. Proceedings of the National Academy of Sciences 98: 5116–5121.
  39. 39. Papadopoulos GL, Alexiou P, Maragkakis M, Reczko M, Hatzigeorgiou AG (2009) DIANA-mirPath: Integrating human and mouse microRNAs in pathways. Bioinformatics 25: 1991–1993.
  40. 40. Kanehisa M, Goto S (2000) KEGG: kyoto encyclopedia of genes and genomes. Nucleic Acids Res 28: 27–30.
  41. 41. Ron-Harel N, Segev Y, Lewitus GM, Cardon M, Ziv Y, et al. (2008) Age-dependent spatial memory loss can be partially restored by immune activation. Rejuvenation Res 11: 903–913.
  42. 42. Gupta RK, Arany Z, Seale P, Mepani RJ, Ye L, et al. (2010) Transcriptional control of preadipocyte determination by Zfp423. Nature 464: 619–623.
  43. 43. Brenner DJ, Hlatky LR, Hahnfeldt PJ, Huang Y, Sachs RK (1998) The linear-quadratic model and most other common radiobiological models result in similar predictions of time-dose relationships. Radiat Res 150: 83–91.
  44. 44. Raggi C, Invernizzi P (2013) Methylation and liver cancer. Clin Res Hepatol Gastroenterol 37: 564–571.
  45. 45. Schwenk RW, Jonas W, Ernst SB, Kammel A, Jähnert M, et al. (2013) Diet-dependent alterations of hepatic Scd1 expression are accompanied by differences in promoter methylation. Horm Metab Res 45: 786–794.
  46. 46. Gozuacik D, Kimchi A (2006) DAPk Protein Family and Cancer. Autophagy 2: 74–79.
  47. 47. Christmann M, Verbeek B, Roos WP, Kaina B (2011) O(6)-Methylguanine-DNA methyltransferase (MGMT) in normal tissues and tumors: enzyme activity, promoter methylation and immunohistochemistry. Biochim Biophys Acta 1816: 179–190.
  48. 48. Nuovo GJ, Plaia TW, Belinsky SA, Baylin SB, Herman JG (1999) In situ detection of the hypermethylation-induced inactivation of the p16 gene as an early event in oncogenesis. Proc Natl Acad Sci U S A 96: 12754–12759.
  49. 49. Zang J-J, Xie F, Xu J-F, Qin Y-Y, Shen R-X, et al. (2011) P16 gene hypermethylation and hepatocellular carcinoma: a systematic review and meta-analysis. World J Gastroenterol 17: 3043–3048.
  50. 50. Kauppinen S, Vester B, Wengel J (2006) Locked nucleic acid: high-affinity targeting of complementary RNA for RNomics. Handb Exp Pharmacol : 405–422.
  51. 51. Barcellos-Hoff MH (2005) Integrative radiation carcinogenesis: interactions between cell and tissue responses to DNA damage. Semin Cancer Biol 15: 138–148.
  52. 52. Dent P, Yacoub A, Contessa J, Caron R, Amorino G, et al. (2003) Stress and radiation-induced activation of multiple intracellular signaling pathways. Radiat Res 159: 283–300.
  53. 53. Su W, Chen Y, Zeng W, Liu W, Sun H (2012) Involvement of Wnt signaling in the injury of murine mesenchymal stem cells exposed to X-radiation. Int J Radiat Biol 88: 635–641.
  54. 54. Wang SS, Shi W, Wang X, Velky L, Greenlee S, et al. (2007) Mapping, Genetic Isolation, and Characterization of Genetic Loci That Determine Resistance to Atherosclerosis in C3H Mice. Arteriosclerosis, Thrombosis, and Vascular Biology 27: 2671–2676.
  55. 55. Deimling LI, Machado FLS, Welker AG, Peres LM, Santos-Mello R (2009) Micronucleus induction in mouse polychromatic erythrocytes by an X-ray contrast agent containing iodine. Mutat Res 672: 65–68.
  56. 56. Sudheer Kumar M, Unnikrishnan MK, Uma Devi P (2003) Effect of 5-aminosalicylic acid on radiation-induced micronuclei in mouse bone marrow. Mutat Res 527: 7–14.
  57. 57. Singh SP, Singh S, Jain V (1990) Effects of 5-bromo-2-deoxyuridine and 2-deoxy-D-glucose on radiation-induced micronuclei in mouse bone marrow. Int J Radiat Biol 58: 791–797.
  58. 58. Jagetia GC, Aruna R (1999) Teniposide (VM-26) treatment enhances the radiation-induced micronuclei in the bone marrow of mouse. Mutat Res 425: 87–98.
  59. 59. Xu W, Shen X, Yang F, Han Y, Li R, et al. (2012) Protective effect of polysaccharides isolated from Tremella fuciformis against radiation-induced damage in mice. J Radiat Res 53: 353–360.
  60. 60. Andreassi MG, Barale R, Iozzo P, Picano E (2011) The association of micronucleus frequency with obesity, diabetes and cardiovascular disease. Mutagenesis 26: 77–83.
  61. 61. Payne MWC, Uhthoff HK, Trudel G (2007) Anemia of immobility: caused by adipocyte accumulation in bone marrow. Med Hypotheses 69: 778–786.
  62. 62. Peslak SA, Wenger J, Bemis JC, Kingsley PD, Frame JM, et al. (2011) Sublethal radiation injury uncovers a functional transition during erythroid maturation. Exp Hematol 39: 434–445.
  63. 63. Wree A, Kahraman A, Gerken G, Canbay A (2011) Obesity affects the liver - the link between adipocytes and hepatocytes. Digestion 83: 124–133.
  64. 64. Wellen KE, Hotamisligil GS (2005) Inflammation, stress, and diabetes. J Clin Invest 115: 1111–1119.
  65. 65. Futscher BW (2013) Epigenetic changes during cell transformation. Adv Exp Med Biol 754: 179–194.
  66. 66. Kovalchuk O, Burke P, Besplug J, Slovack M, Filkowski J, et al. (2004) Methylation changes in muscle and liver tissues of male and female mice exposed to acute and chronic low-dose X-ray-irradiation. Mutat Res 548: 75–84.
  67. 67. Christmann M, Kaina B (2012) O(6)-methylguanine-DNA methyltransferase (MGMT): impact on cancer risk in response to tobacco smoke. Mutat Res 736: 64–74.
  68. 68. Grombacher T, Kaina B (1995) Constitutive expression and inducibility of O6-methylguanine-DNA methyltransferase and N-methylpurine-DNA glycosylase in rat liver cells exhibiting different status of differentiation. Biochim Biophys Acta 1270: 63–72.
  69. 69. Fritz G, Tano K, Mitra S, Kaina B (1991) Inducibility of the DNA repair gene encoding O6-methylguanine-DNA methyltransferase in mammalian cells by DNA-damaging treatments. Mol Cell Biol 11: 4660–4668.
  70. 70. Kim S, Sohn I, Ahn J-I, Lee K-H, Lee YS, et al. (2004) Hepatic gene expression profiles in a long-term high-fat diet-induced obesity mouse model. Gene 340: 99–109.
  71. 71. Roudkenar MH, Li L, Baba T, Kuwahara Y, Nakagawa H, et al. (2008) Gene Expression Profiles in Mouse Liver Cells after Exposure to Different Types of Radiation. J Radiat Res 49: 29–40.
  72. 72. Amundson SA, Bittner M, Meltzer P, Trent J, Fornace AJ (2001) Induction of gene expression as a monitor of exposure to ionizing radiation. Radiat Res 156: 657–661.
  73. 73. Vares G, Uehara Y, Ono T, Nakajima T, Wang B, et al. (2011) Transcription factor-recognition sequences potentially involved in modulation of gene expression after exposure to low-dose-rate γ-rays in the mouse liver. J Radiat Res 52: 249–256.
  74. 74. Uehara Y, Ito Y, Taki K, Nenoi M, Ichinohe K, et al. (2010) Gene Expression Profiles in Mouse Liver after Long-Term Low-Dose-Rate Irradiation with Gamma Rays. Radiat Res.
  75. 75. Heneghan HM, Miller N, Kerin MJ (2010) Role of microRNAs in obesity and the metabolic syndrome. Obes Rev 11: 354–361.
  76. 76. Chartoumpekis DV, Zaravinos A, Ziros PG, Iskrenova RP, Psyrogiannis AI, et al. (2012) Differential expression of microRNAs in adipose tissue after long-term high-fat diet-induced obesity in mice. PLoS One 7: e34872.
  77. 77. Girard M, Jacquemin E, Munnich A, Lyonnet S, Henrion-Caude A (2008) miR-122, a paradigm for the role of microRNAs in the liver. J Hepatol 48: 648–656.
  78. 78. Krützfeldt J, Rajewsky N, Braich R, Rajeev KG, Tuschl T, et al. (2005) Silencing of microRNAs in vivo with 'antagomirs'. Nature 438: 685–689.
  79. 79. Esau C, Davis S, Murray SF, Yu XX, Pandey SK, et al. (2006) miR-122 regulation of lipid metabolism revealed by in vivo antisense targeting. Cell Metab 3: 87–98.
  80. 80. Tang R, Li L, Zhu D, Hou D, Cao T, et al. (2012) Mouse miRNA-709 directly regulates miRNA-15a/16-1 biogenesis at the posttranscriptional level in the nucleus: evidence for a microRNA hierarchy system. Cell Res 22: 504–515.
  81. 81. Li X, Sanda T, Look AT, Novina CD, von Boehmer H (2011) Repression of tumor suppressor miR-451 is essential for NOTCH1-induced oncogenesis in T-ALL. J Exp Med 208: 663–675.
  82. 82. Druz A, Son Y-J, Betenbaugh M, Shiloach J (2013) Stable inhibition of mmu-miR-466h-5p improves apoptosis resistance and protein production in CHO cells. Metab Eng 16: 87–94.
  83. 83. Druz A, Betenbaugh M, Shiloach J (2012) Glucose depletion activates mmu-miR-466h-5p expression through oxidative stress and inhibition of histone deacetylation. Nucleic Acids Res 40: 7291–7302.
  84. 84. Luo Y, Liu Y, Liu M, Wei J, Zhang Y, et al. (2014) Sfmbt2 10th intron-hosted miR-466(a/e)-3p are important epigenetic regulators of Nfat5 signaling, osmoregulation and urine concentration in mice. Biochim Biophys Acta 1839: 97–106.
  85. 85. Zheng GXY, Ravi A, Gould GM, Burge CB, Sharp PA (2011) Genome-wide impact of a recently expanded microRNA cluster in mouse. Proc Natl Acad Sci U S A 108: 15804–15809.
  86. 86. Metheetrairut C, Slack FJ (2013) MicroRNAs in the ionizing radiation response and in radiotherapy. Current Opinion in Genetics & Development 23: 12–19.
  87. 87. Ding N, Wu X, He J, Chang L, Hu W, et al. (2011) Detection of Novel Human MiRNAs Responding to X-ray Irradiation. J Radiat Res 52: 425–432.
  88. 88. Shi Y, Zhang X, Tang X, Wang P, Wang H, et al. (2012) MiR-21 is continually elevated long-term in the brain after exposure to ionizing radiation. Radiat Res 177: 124–128.
  89. 89. Vincenti S, Brillante N, Lanza V, Bozzoni I, Presutti C, et al. (2011) HUVEC respond to radiation by inducing the expression of pro-angiogenic microRNAs. Radiat Res 175: 535–546.
  90. 90. Wagner-Ecker M, Schwager C, Wirkner U, Abdollahi A, Huber PE (2010) MicroRNA expression after ionizing radiation in human endothelial cells. Radiation Oncology 5: 25.
  91. 91. Simone NLAS, Ly BPA, Saleh DA, Savage ADA, DeGraff JEA, et al. (2009) Ionizing Radiation-Induced Oxidative Stress Alters miRNA Expression. PLoS One 4: e6377.
  92. 92. Liu C, Li B, Cheng Y, Lin J, Hao J, et al. (2011) MiR-21 plays an important role in radiation induced carcinogenesis in BALB/c mice by directly targeting the tumor suppressor gene Big-h3. Int J Biol Sci 7: 347–363.
  93. 93. An YS, Kim M-R, Lee S-S, Lee Y-S, Chung E, et al. (2013) TGF-β signaling plays an important role in resisting γ-irradiation. Exp Cell Res 319: 466–473.
  94. 94. Dent P, Yacoub A, Fisher PB, Hagan MP, Grant S (2003) MAPK pathways in radiation responses. Oncogene 22: 5885–5896.
  95. 95. Runkle EA, Zhang H, Cai Z, Zhu Z, Karger BL, et al. (2012) Reversion of the ErbB malignant phenotype and the DNA damage response. Exp Mol Pathol 93: 324–333.
  96. 96. Demirbag R, Yilmaz R, Gur M, Celik H, Guzel S, et al. (2006) DNA damage in metabolic syndrome and its association with antioxidative and oxidative measurements. Int J Clin Pract 60: 1187–1193.
  97. 97. Weitzman SA, Turk PW, Milkowski DH, Kozlowski K (1994) Free radical adducts induce alterations in DNA cytosine methylation. Proc Natl Acad Sci U S A 91: 1261–1264.
  98. 98. Vincent HK, Innes KE, Vincent KR (2007) Oxidative stress and potential interventions to reduce oxidative stress in overweight and obesity. Diabetes Obes Metab 9: 813–839.
  99. 99. Baldeweg SE, Golay A, Natali A, Balkau B, Del Prato S, et al. (2000) Insulin resistance, lipid and fatty acid concentrations in 867 healthy Europeans. European Group for the Study of Insulin Resistance (EGIR). Eur J Clin Invest 30: 45–52.
  100. 100. Laws A, Hoen HM, Selby JV, Saad MF, Haffner SM, et al. (1997) Differences in Insulin Suppression of Free Fatty Acid Levels by Gender and Glucose Tolerance Status. Arteriosclerosis, Thrombosis, and Vascular Biology 17: 64–71.
  101. 101. Choi EK, Rhee YH, Park HJ, Ahn SD, Shin KH, et al. (2001) Effect of protein kinase C inhibitor (PKCI) on radiation sensitivity and c-fos transcription. Int J Radiat Oncol Biol Phys 49: 397–405.
  102. 102. De Marchi E, Baldassari F, Bononi A, Wieckowski MR, Pinton P (2013) Oxidative Stress in Cardiovascular Diseases and Obesity: Role of p66Shc and Protein Kinase C. Oxid Med Cell Longev. 2013: 564961.
  103. 103. Nakajima T (2008) Positive and negative regulation of radiation-induced apoptosis by protein kinase C. J Radiat Res (Tokyo). 49: 1–8.