Oxidative Medicine and Cellular Longevity

Oxidative Medicine and Cellular Longevity / 2019 / Article
Special Issue

Oxidative Stress and Tissue Repair: Mechanism, Biomarkers, and Therapeutics

View this Special Issue

Review Article | Open Access

Volume 2019 |Article ID 3010342 | https://doi.org/10.1155/2019/3010342

Jinlong Wei, Bin Wang, Huanhuan Wang, Lingbin Meng, Qin Zhao, Xinyu Li, Ying Xin, Xin Jiang, "Radiation-Induced Normal Tissue Damage: Oxidative Stress and Epigenetic Mechanisms", Oxidative Medicine and Cellular Longevity, vol. 2019, Article ID 3010342, 11 pages, 2019. https://doi.org/10.1155/2019/3010342

Radiation-Induced Normal Tissue Damage: Oxidative Stress and Epigenetic Mechanisms

Guest Editor: Reggiani Vilela Gonçalves
Received07 Jul 2019
Revised23 Oct 2019
Accepted24 Oct 2019
Published12 Nov 2019

Abstract

Radiotherapy (RT) is currently one of the leading treatments for various cancers; however, it may cause damage to healthy tissue, with both short-term and long-term side effects. Severe radiation-induced normal tissue damage (RINTD) frequently has a significant influence on the progress of RT and the survival and prognosis of patients. The redox system has been shown to play an important role in the early and late effects of RINTD. Reactive oxygen species (ROS) and reactive nitrogen species (RNS) are the main sources of RINTD. The free radicals produced by irradiation can upregulate several enzymes including nicotinamide adenine dinucleotide phosphate oxidase (NADPH oxidase), lipoxygenases (LOXs), nitric oxide synthase (NOS), and cyclooxygenases (COXs). These enzymes are expressed in distinct ways in various cells, tissues, and organs and participate in the RINTD process through different regulatory mechanisms. In recent years, several studies have demonstrated that epigenetic modulators play an important role in the RINTD process. Epigenetic modifications primarily contain noncoding RNA regulation, histone modifications, and DNA methylation. In this article, we will review the role of oxidative stress and epigenetic mechanisms in radiation damage, and explore possible prophylactic and therapeutic strategies for RINTD.

1. Introduction

Cancer is one of the most challenging diseases in modern times. In 2015, China reported about 4.2 million new cancer cases and 2.8 million cancer-related deaths [1]. Radiotherapy (RT) is currently one of the leading therapeutic approaches for several cancers; however, it carries the potential to cause injury to normal tissue, with both short-term and long-term side effects. In recent years, studies have shown that the oxidation/reduction (redox) system was associated with several types of damage after radiation exposure [2]. In addition, the redox system is related to epigenetic regulation and can regulate the expression of microRNAs (miRNAs) and other molecules, thus playing a role in sustained oxidative damage after radiation [3].

Cells and tissues are composed of about 80% or more water, and most of the radiation damage occurs due to the radiolysis of water, which induces the production of reactive oxygen species (ROS) and reactive nitrogen species (RNS) [4]. ROS and RNS are the main sources of radiation-induced normal tissue damage (RINTD). The generation of ROS induces molecular changes and causes oxidative damage to proteins, lipids, and DNA. It can also activate signal transduction pathways and early-response transcription factors [5]. The redox system plays an important role in acute radiation damage and is responsible for some radiation-induced early and late effects including inflammation, out-of-field effects, fibrosis, bystander effects, and others [69].

In recent years, several studies have demonstrated that epigenetic modulators play an important role in normal tissue damage, after redox-induced ionizing radiation. Epigenetic modifications are made up of the heritable changes in the expression of the gene that do not influence the sequence of the DNA. In mammals, epigenetic modifications primarily contain noncoding RNA regulation, histone modifications (methylation, phosphorylation, and acetylation), and DNA methylation. Epigenetic changes can be reversible and can easily respond to natural bioactive dietary compounds [10]. Afanas’ev et al. has reported that free radicals such as NO and ROS can regulate and control the epigenetic processes [11]. In addition, the regulation of some miRNAs may decrease or increase the oxidative damage [11].

In regard to the damage caused by RT, treatment strategies are still lacking. Here, we review the role of oxidative stress and epigenetic mechanisms in radiation damage to explore possible therapeutic strategies for RINTD.

2. Oxidative Stress

Oxidative stress is involved in the development of many diseases including RINTD. The redox system plays an important role in the early and late effects of RINTD [12]. When cells are exposed to radiation, they immediately form free radicals with a half-life of nanoseconds. The redox system begins producing free radicals a few hours after exposure, with the potential to last for years [13, 14]. The free radicals produced by ionizing radiation can upregulate several enzymes, including nicotinamide adenine dinucleotide phosphate oxidase (NADPH oxidase), lipoxygenases (LOXs), nitric oxide synthase (NOS), and cyclooxygenases (COXs). Their effects on mitochondrial function are distinct. These enzymes are expressed in specific ways in various cells, tissues, and organs (Figure 1).

2.1. NADPH Oxidases

NADPH oxidase (NOX) is thought to be a membrane-bound oxidoreductase. It can transfer electrons from NADPH to the oxygen molecules. In addition, some subtypes of these enzymes have been found in cells [12]. NADPH oxidase enzymes such as DUOX1, DUOX2, and NOX1-5 are the most crucial subtypes. They participate in the process of respiratory chain rupture after radiation [15]. These enzymes have the ability to transfer electrons across the plasma membrane and produce superoxide and other downstream ROS. However, the tissue distribution and activation mechanisms of the individual members of the NOX family are undoubtedly different [16]. In addition, many inflammatory cytokines and chemokines such as TGF-β, TNF-α, IL-1, and IFN-γ are involved in the NOX system activation [17]. NADPH oxidase enzymes play a key role in acute and chronic oxidative stress in bystander and directly irradiated cells [18]. Also, it has been shown that the expression of NOX2 and NOX4 can be upregulated in nontargeted tissues [19].

NOX1 is the first homolog of NOX2 described (then called gp91phox) [20, 21]. NOX1 can be expressed in a variety of cells including endothelial cells in the uterus, prostate, and placenta, as well as osteoclasts. It can also be expressed in some malignant tumors including colon cancer and melanoma [2224]. Choi et al. reported that the NOX1-specific inhibitor can limit radiation-induced collagen deposition and fibroblastic changes in the endothelial cells, thereby alleviating pulmonary fibrosis induced by radiation [25]. In addition, the production of ROS was significantly decreased after inhibition of NOX1.

NOX2 is considered the prototype of the NADPH oxidase. A report by Kim et al. confirmed that NOX2 was involved in radiation-induced salivary gland damage. After 56 days of exposure to 18 Gy radiation, the expression of the NOX2 gene in the salivary glands of rats was significantly increased. In addition, the apoptotic genes such as caspase-9 and MAPKs, including p-38 and JNK, participate in NOX2 signaling cascades [26]. Experiments conducted by Narayanan et al. demonstrated that irradiation of human lung fibroblasts generated O2•- and H2O2 with alpha particles. The plasma membrane-bound NOX2 is responsible for the production of O2•- and H2O2 [27]. Datta et al. confirmed that long-range mitochondrial dysfunction and the increased NADPH oxidase, including NOX1 and NOX2 activity, are the main factors for radiation-induced continuous oxidative stress in the intestinal epithelial cells [28].

NOX3 was first described in the year 2000 based on its similarity to the sequences of other NOX subtypes [29], although the function of the protein was first studied in 2004 [30, 31]. At present, the study of NOX3 in radiation damage is limited. Shin et al. confirmed that the expression of NOX3 was upregulated after the irradiation of the oral mucosa of rats. The increased expression of NOX3 is thought to be related to the necrotic inflammatory exudate and oral mucosa ulcers [32].

NOX4 was initially thought to be an NADPH oxidase homolog, highly expressed in the kidney [33, 34]. Pazhanisamy et al. found that systemic irradiation in mice can selectively induce sustained oxidative stress in hematopoietic stem cells (HSCs), at least in part, by the upregulation of NOX4. The increased production of ROS by NOX in HSCs can mediate radiation-induced hematopoietic genomic instability [35]. The experimental results of Wang et al. showed that systemic irradiation selectively induces chronic oxidative stress in HSCs, at least in part by the upregulation of NOX4 expression, thereby giving rise to the induction of HSC senescence and residual bone marrow damage [36]. In addition, the TGF-β-NOX4 pathway may be responsible for the continuous production of ROS/NO and the subsequent genomic instability after bone marrow irradiation [37].

NOX5, found in the lymphoid and testis, contains an N-terminal extension with three EF hands, and it can produce superoxide dismutase and conduct H+ ions when intracellular free Ca2+ increases [3840]. NOX5 may participate in Ca2+-activated, redox-dependent processes of spermatozoa and lymphocytes including cytokine secretion, cell proliferation, and sperm-oocyte fusion [41]. Weyemi et al. showed that the two members of NADPH oxidase, NOX4 and NOX5, participated in the process of radiation-induced DNA damage in human primary fibroblasts.

Currently, there is a small amount of evidence supporting the role of DUOX1/DUOX2 in chronic oxidative stress. Both DUOX1 and DUOX2 are highly expressed in the thyroid gland [42, 43]. Furthermore, DUOX1 can be found in the prostate and airway epithelia [4447]. DUOX2 has been described in the salivary gland, airway epithelia, prostate, rectal mucosa, duodenum, cecum, and colon [4449]. Ameziane-El-Hassani et al. demonstrated that radiation-induced DUOX1-dependent H2O2 production by NADPH oxidase was delayed in a dose-dependent manner for several days. In addition, p38 MAPK, which was activated after irradiation, can increase DUOX1 through the expression of IL-13, giving rise to sustained DNA damage and growth stagnation [50]. Wu and Doroshow showed that IL-4/IL-13 can induce the expression of DUOX2/DUOXA2 and the production of ROS in human colon and pancreatic cancer cells, which in turn may be related to the occurrence of inflammatory gastrointestinal malignancies [51]. There are some studies that have shown the upregulation of DUOX1 and DUOX2 in the lung and heart. Some radioprotectors and antioxidants such as melatonin, metformin, and selenium have been shown to reduce the expression of these genes following exposure to ionizing radiation, relieving the heart damage and lung damage caused by radiation [5256]. However, the relationship between DUOX2 expression and radiation-induced carcinogenesis has not been established and demands further verification.

2.2. COX-2

Cyclooxygenase-2 (COX-2), an isoform of cyclooxygenase, is responsible for the time-dependent and localized production of prostaglandins (PGs) at inflammatory sites [57], including tissues exposed to ionizing radiation. COX-2 plays a crucial role in the inflammatory response that converts arachidonic acid released by membrane phospholipids into PGs. In addition, ROS production is a standard secondary byproduct of arachidonic acid metabolism in the synthesis of PGE2 [58]. Several studies have shown that increased COX-2 expression is related to radiation toxicity after the irradiation of organs, such as the lungs, heart, kidneys, intestines, colon, and the brain [59, 60]. Other studies have reported that COX-2 is involved in the pathogenesis of vascular damage, atherosclerosis, and fibrosis induced by ionizing radiation [61]. Cheki et al. demonstrated that celecoxib, an inhibitor of COX-2, can decrease dermal inflammation, MCP-1 mRNA expression, and radiation-induced skin reactions [62]. In addition, several studies have investigated the role of nonsteroidal anti-inflammatory drugs (NSAIDS) as an inhibitor of COX on radiation damage in the lung and joints [6365]. Clinically approved inhibitors are represented by NSAIDs like aspirin or ibuprofen and by selective COX-2 inhibitors such as celecoxib [60].

2.3. LOXs

LOXs are enzymes that dioxygenate unsaturated fatty acids, which can initiate lipoperoxidation of the membrane, synthesize signaling molecules, or induce cell structural and metabolic changes [66]. Currently, the role of LOX in radiation damage has been reported. Matyshevskaia et al. demonstrated that activated LOX is involved in the production of ROS after exposure and plays an important role in the process of radiation-induced DNA fragmentation in lymphocytes [67]. Another experimental study showed that LOX was activated immediately after exposure to thymocytes. High LOX activity was observed in cells within an hour after irradiation. In addition, radiation-induced generation of lipid peroxides may be a factor in LOX activation [68]. In another study, Halle et al. showed that chronic adventitial inflammation, vasa vasorum expansion, and 5-LOX upregulation are involved in radiation-induced arterial damage in cancer survivors. In previously irradiated arterial segments, the expression of 5-LOX was increased in exogenous macrophages surrounding vascular dilatation [69].

2.4. Nitric Oxide

Under conditions of stress, including inflammation, inducible nitric oxide synthase (iNOS) was thought to be the primary source of nitric oxide (NO) and played an important role in carcinogenesis and the oxidative stress process. NO is generated by macrophages under the stimulation of inflammation through the iNOS enzyme, and it can interact with the mitochondria-derived superoxide to further produce peroxynitrite [70]. iNOS enzymes play a key role in the radiation damage via posttranslational regulation of the BER pathway of DNA repair. The main effect of NO is nitroacetylation of 8-xoguanine glycosylase (Ogg1). Ogg1 inhibition by NO can result in increased accumulation of oxidative DNA lesions [71, 72]. Malaviya et al. noted that iNOS is involved in radiation-induced lung damage. In addition, there are complex interactions between oxidative and nitrosative stress, as well as inflammatory pathways that mediate lung damage after radiation [73]. In another study, the inhibitors of iNOS such as aminoguanidine and N-nitro-L-arginine methyl ester have been shown to reduce radiation-induced lung damage [74, 75]. In a study by Ohta el al., increased levels of NO were directly related to the radiation dose, and NO levels increased in the first few hours after receiving the radiation [76].

The role of NO in the radiation-induced bystander effect has been explored. The peculiarity of NO as a redox signaling molecule is partly due to its hydrophobic properties and relative stability [77]. The hydrophobicity of NO allows it to diffuse through the cytoplasm and plasma membrane, allowing this kind of signaling molecule to readily diffuse from irradiated cells to bystander cells without the involvement of gap junction intercellular communication. NO generated in the irradiated tissues can mediate cellular regulation through posttranslational modification of many regulatory proteins [78]. Ghosh et al. have shown that activated iNOS in irradiated cells are crucial to the bystander response. In addition, lipopolysaccharide-induced iNOS activity and the production of NO after irradiation increased bystander cell DNA damage [79, 80]. Han et al. showed that within 30 min of low-dose alpha-particle irradiation, NO played an important role in the process of DNA double-strand breaks in bystander cells [81].

2.5. Oxidative Stress and Inflammation

Inflammatory responses are thought to play an important role in redox activation. Normal cells that are directly exposed to irradiation or ROS will give rise to nuclear and mitochondrial DNA damages, which can lead to cell death via processes such as mitotic catastrophe, apoptosis, and necrosis [82]. Necrosis can trigger the release of inflammatory cytokines such as IL-1, IL-4, IL-13, and other inflammatory mediators, while apoptosis may cause the release of anti-inflammatory cytokines including TGF-β and IL-10 [83, 84]. ROS are the main cause of RINTD. The continuous formation of ROS after IR exposure can be the source of radiosensitivity of the T lymphocytes and other cells [85]. Moreover, ROS can activate the NF-κB signaling pathway along with proinflammatory cytokines. NF-κB plays a key role in chronic inflammatory diseases after RT [86]. Inflammatory cytokines and growth factors can give rise to a variety of signaling cascades, such as NADPH oxidase, COX-2, and iNOS [87]. It has also been reported that COX-2 is an important gene which can mediate the subsequent inflammatory responses [88]. Mitochondria is thought to be an energy and free radical reservoir. In normal conditions, antioxidant defense systems neutralize superoxide and form free radicals and protect cells from oxidative damage resulting from mitochondrial activity [89]. However, mitochondrial dysfunction and apoptosis can be induced by ROS, pro-IL-1β, iNOS, and inflammatory responses. Also, studies have shown that the ROS-derived NOX system participated in mitochondrial dysfunction and in the subsequent production of ROS in this organelle [90]. Next, the damaged mitochondria will release ROS and activate the nucleotide-binding domain and the leucine-rich-repeat-containing family pyrin 3 (NLRP3) inflammasome pathway [91]. The activation of the NLRP3 inflammasome is the platform of caspase-1 activation. Finally, it will lead to the secretion of proinflammatory cytokines IL-18 and IL-1β [92]. Recent experiments and studies have shown that the upregulation of the NLRP3 inflammasome has a big impact on radiation damage [9396]. Chronic inflammatory processes can exist for ages after irradiation, and the immune system does not suppress them. This is related to chronic oxidative damage giving rise to the genomic instability and impaired normal tissue function [97].

2.6. Oxidative Stress and Cellular Senescence

Cellular senescence can be induced by ionizing radiation. Radiation-induced senescence is mainly one of the mechanisms in radiation-induced pathological changes. Radiation-induced cellular senescence is thought to be caused by p53 activation, which is associated with a radiation-induced double-strand DNA break [98]. However, the exact mechanism of inducing cellular senescence is still unclear, but the involvement of ROS has been widely reported [99, 100]. The study of Kobashigawa et al. suggested that the delayed increase of intracellular oxidative stress levels plays a key role in the process of radiation-induced cellular senescence by p53 accumulation [101]. Sakai et al. showed that NOX4 can mediate the production of ROS in radiation-induced senescent cells and lead to normal tissue damage after irradiation through recruitment of inflammatory cells and intensification of tissue inflammation [102].

3. Epigenetic Mechanisms

3.1. Epigenetics and Cancer

Cancer is commonly thought to be caused by genetic alterations including deletions, insertions, mutations, recombination, copy number gains, single-nucleotide polymorphisms, and genomic instability [103, 104]. The latest evidence suggests that cancer may occur without changes in the nucleotide sequence, by means of alleged epigenetic alterations. Combinational crosstalk between epigenetic alterations and genetics has been known to play a role in the development, progression, and recurrence of cancer [105]. Miousse et al. reported that epigenetic alterations are among the driving forces of irradiation-induced carcinogenesis, by observing long interspersed nucleotide element 1 DNA methylation changes in the mouse hematopoietic system after irradiation [106].

Epigenetic dysregulation including increased activity of histone deacetyltransferases (HDACs), DNA methyltransferases (DNMTs), and changes in the noncoding RNA expression, can give rise to changes in gene transcription and expression, which regulate cell cycle, cell differentiation and proliferation, and apoptosis [107, 108]. Yi et al. showed that the combined action of DNMT and HDAC inhibitors could stagnate the cell cycle at the G2/M phase and suppress the growth of endometrial cancer by upregulating E-cadherin and downregulating Bcl-2 [107]. Choi et al. noted that DNMTs including DNMT3A, DNMT3B, and DNMT1 are overexpressed in the hepatocellular carcinoma compared with noncancerous liver samples [109]. One such study has demonstrated that HDAC5 can promote glioma cell proliferation by upregulating the expression of Notch 1 at both the mRNA and the protein level [110]. In addition, HDAC5 can also promote human HCC cell proliferation by upregulating the expression of Six1 [111]. Epigenetic regulation, as a molecular target for cancer prevention and therapy, has aroused wide interest. For example, some studies showed that (-)-epigallocatechin-3-gallate, a main component of green tea, could possibly bind with the DNMTs, reducing the methylation activity of cancer cells through epigenetic mechanisms, and thus leading to cancer prevention or treatment [112, 113]. At present, there is increasing evidence that targeting epigenetic modifications is an effective cancer prevention strategy.

3.2. Epigenetics and RINTD

In recent years, the relationship between epigenetic mechanisms and radiation damage has been studied extensively [114116]. Currently, epigenetic mechanisms such as DNA methylation and miRNA and histone modifications are reported to be associated with radiation damage. These mechanisms are summarized in Table 1.


Epigenetic mechanismsIrradiation organEpigenetic functionsTarget genes/proteinsDamage effectsReference

DNA methylationBrainIncreased expression of DNMT1 and 3aIncreased expression of TET1 and TET3 proteinsRadiation-induced cognitive dysfunctionAcharya et al. [116]
ThymusDecreased expression of DNMT1, 3a, and 3bDecrease in the levels of methyl-binding proteins MeCP2 and MBD2Increased the risk of radiation-inducedleukemia and thymic lymphomaPogribny et al. [119]
Human breast cancer cells (MDA-MB-231)Decreased DNMT1 expressionDownregulation of RB1 expressionDNA damage and apoptosisAntwih et al. [118]
BrainDecreased expression of DNMT1, 3a, and 3bDecrease in the levels of methyl-binding protein MeCP2Bystander effect in the spleenKoturbash et al. [120]

Histone methylationIntestineIncreased expression of histone H3 methylationRadiation-induced intestinal damageHerberg et al. [124]

Histone acetylationSkinInhibition of HDAC activityRadiation-induced skin damage and carcinogenesisZhang et al. [125]

Regulation of miRNAsHematopoietic systemUpregulation of miR-30a-3p, miR-30c-5p, etc.Radiation-induced hematopoietic damageAcharya et al. [129]
LungUpregulation of miR-19a-3p, miR-144-5p, and miR-144-3pRadiation-induced lung injuryGao et al. [135]
SpleenIncreased expression of miR-34aUpregulation of gene p53Radiation-induced spleen damageGhosh et al. [137]
Hematopoietic and osteoblast cellsIncreased expression of miR-30cSuppression of gene REDD1Radiation-induced hematopoietic and osteoblast cell damageLi et al. [131]

3.2.1. DNA Methylation

DNA methylation is a crucial means of epigenetic modification, which primarily occurs in the CPG islands of the gene promoter regions. Multiple DNMT functions are required to establish and maintain DNA methylation patterns [117]. Therapeutic radiation can give rise to biological responses to confront the subsequent DNA damage and genomic stress, to avoid cell death. Antwih et al. showed that the DNA methylation response to radiation is parallel to the classical biological responses to radiation. The differential methylation level of DNA repair, cell cycle, and apoptosis pathways varied with different radiation doses [118]. Fractionated low-dose radiation exposure has been reported to cause the accumulation of DNA damage and profound alterations in DNA methylation in the murine thymus. This could be the source of the risk of radiation-induced leukemia and thymic lymphoma [119].

Acharya et al. showed that neuroepigenetic mechanisms played an important role in affecting the functional and structural changes in the brain and in cognition after irradiation. In irradiated mice with cognitive impairment, 5-hydroxymethylcytosine and 5-methylcytosine were detected in the region of the hippocampus consistent with increased levels of Ten Eleven Translocation- (TET-) 1, TET3, and DNMT3a. DNMT3a and TET enzymes including TET1 and TET3 are related to addiction behavior and memory formation. In addition, they found an obvious improvement in the epigenetic effects of irradiation by inhibiting methylation using 5-iodotubercidin [116]. Koturbash et al. demonstrated the role of epigenetic effects in maintaining the long-term, persistent bystander effect in the spleen, in vivo. After localized cranial irradiation for 24 h and 7 months, the levels of methyltransferases DNMT3a, DNMT3b, and DNMT1 and methyl-binding protein MeCP2 in the spleen tissue were significantly decreased [120].

The above results indicate that radiation can cause changes in DNA methylation to modify and regulate the expression of related genes and proteins, thus causing the corresponding tissue and organ damage. Future research is essential to confirm the role of DNA methylation in radiation-induced normal tissue damage. In addition, DNA methylation can be used as a target to prevent and treat radiation damage in the future.

3.2.2. Histone Modifications

Histone modification is rarely studied in radiation-induced normal tissue damage. Most reports have focused on the regulatory role of histone modification in radiation approaches for killing tumor cells [121123]. Histone modifications include methylation, phosphorylation, and acetylation. Herberg et al. showed that mismatch repair-deficiency leads to genome-wide changes in histone H3 methylation profiles prior to tumorigenesis. Analogous changes constitute a lasting epigenetic feature of radiation-induced DNA damage [124]. Zhang et al. showed that solar-simulated ultraviolet radiation can influence both histone acetyltransferase and HDAC activities causing decreased histone acetylation. This could be the main cause for radiation-induced skin DNA damage [125]. Further research is needed to verify the role of histone modifications in radiation damage.

3.2.3. Regulation of miRNAs

MiRNAs combined with mRNAs can lead to posttranscriptional degradation or repression [126]. It is well known that the role of miRNAs in ROS production and oxidative stress is to increase the superoxide level by suppressing antioxidant enzymes. A good example is the upregulation of miR-21 in both targeted and bystander cells. MiR-21, which is triggered by TGF-β, can inhibit SOD2 gene expression, giving rise to a decrease in the activity of SOD2 and damage to irradiated and bystander cells by superoxide [3, 127]. In addition, the SOD activity and glutathione level were inhibited which have been revealed in nontargeted lung tissues [128]. Many studies have shown a link between miRNA regulation and RINTD. miRNAs can play an important role in the early evaluation of radiation-induced hematopoietic damage, as functional dosimeters of radiation [129, 130]. Li et al. also demonstrated that miR-30c plays a key role in radiation-induced hematopoietic and osteoblast cell damage, possibly by regulating the expression of the gene REDD1 [131].

Another study by Li et al. reported that the isomer of vitamin E, delta-tocotrienol, can inhibit radiation-induced miR-30 and protect human and mice CD34+ cells from radiation damage by inhibiting IL-1β-induced NF-κB/miR-30 signaling [132].

Radiation-induced lung damage includes chronic fibrosis and acute pneumonia [133]. miRNAs have been reported in many diseases including those with lung involvement [134]. Gao et al. showed that miR-19a-3p, miR-144-5p, and miR-144-3p are upregulated in rats 2 weeks after thorax irradiation [135]. Recently, Xie et al. also studied the response of lung miRNA expression to radiation-induced lung damage in rats [136]. MiRNAs may serve as biomarkers for early stages of radiation-induced lung damage.

Radiation-induced spleen damage has also been reported in recent years. Ghosh et al. reported that whole-body radiation exposure resulted in higher expression of miRNAs in the spleen tissue on day 4 and on day 250. In addition, the vitamin E analog gamma-tocotrienol can modulate radiation-induced miRNA expression in the mouse spleen, preventing radiation damage to the spleen [137].

Collectively, miRNAs can serve as promising candidates for radiation biodosimetry. In addition, the prevention and treatment of RINTD through miRNA regulation may have a promising future.

4. Conclusions

In summary, oxidative stress responses and epigenetic mechanisms play important roles in RINTD. The redox system and various oxidases upregulated by free radicals and generated by radiation, including NADPH oxidase, LOXs, NOS, and COXs, participated in RINTD through different regulatory mechanisms. ROS and NOS produced by inflammatory cells and mitochondria are involved in oxidative damage to bystander cells and untargeted tissues. In addition, a variety of inflammatory factors including the NLRP3 inflammasome play an important role in radiation-induced oxidative stress damage. Epigenetic mechanisms such as DNA methylation, regulation of miRNAs, and histone modifications have been extensively studied in recent years in relation to RINTD. New progress has been made in the field of radiation damage treatment through the regulation of epigenetic mechanisms. With a further understanding of oxidative stress and epigenetic regulatory mechanisms, we hope to better explore the preventive and therapeutic strategies in RINTD in the future.

Conflicts of Interest

The authors report no conflicts of interest in this work.

Authors’ Contributions

Jinlong Wei and Bin Wang contributed equally to this work.

Acknowledgments

We would like to thank Editage (http://www.editage.cn) for English language editing. This work was supported in part by grants from the National Natural Science Foundation of China (grant no. 81570344 to Ying Xin), the Norman Bethune Program of Jilin University (grant no. 2015225 to Ying Xin and no. 2015203 to Xin Jiang), and the Science and Technology Foundation of Jilin Province (grant no. 20180414039GH to Ying Xin and no. 20190201200JC to Xin Jiang).

References

  1. W. Chen, R. Zheng, P. D. Baade et al., “Cancer statistics in China, 2015,” CA: A Cancer Journal for Clinicians, vol. 66, no. 2, pp. 115–132, 2016. View at: Publisher Site | Google Scholar
  2. D. R. Spitz, E. I. Azzam, J. J. Li, and D. Gius, “Metabolic oxidation/reduction reactions and cellular responses to ionizing radiation: a unifying concept in stress response biology,” Cancer Metastasis Reviews, vol. 23, no. 3-4, pp. 311–322, 2004. View at: Publisher Site | Google Scholar
  3. Y. Jiang, X. Chen, W. Tian, X. Yin, J. Wang, and H. Yang, “The role of TGF-beta1-miR-21-ROS pathway in bystander responses induced by irradiated non-small-cell lung cancer cells,” British Journal of Cancer, vol. 111, no. 4, pp. 772–780, 2014. View at: Publisher Site | Google Scholar
  4. E. Vorotnikova, R. A. Rosenthal, M. Tries, S. R. Doctrow, and S. J. Braunhut, “Novel synthetic SOD/catalase mimetics can mitigate capillary endothelial cell apoptosis caused by ionizing radiation,” Radiation Research, vol. 173, no. 6, pp. 748–759, 2010. View at: Publisher Site | Google Scholar
  5. P. Dent, A. Yacoub, P. B. Fisher, M. P. Hagan, and S. Grant, “MAPK pathways in radiation responses,” Oncogene, vol. 22, no. 37, pp. 5885–5896, 2003. View at: Publisher Site | Google Scholar
  6. M. Najafi, A. Shirazi, E. Motevaseli et al., “The melatonin immunomodulatory actions in radiotherapy,” Biophysical Reviews, vol. 9, no. 2, pp. 139–148, 2017. View at: Publisher Site | Google Scholar
  7. R. Yahyapour, D. Shabeeb, M. Cheki et al., “Radiation protection and mitigation by natural antioxidants and flavonoids: implications to radiotherapy and radiation disasters,” Current Molecular Pharmacology, vol. 11, no. 4, pp. 285–304, 2018. View at: Publisher Site | Google Scholar
  8. R. Yahyapour, E. Motevaseli, A. Rezaeyan et al., “Mechanisms of radiation bystander and non-targeted effects: implications to radiation carcinogenesis and radiotherapy,” Current Radiopharmaceuticals, vol. 11, no. 1, pp. 34–45, 2018. View at: Publisher Site | Google Scholar
  9. R. Yahyapour, A. Salajegheh, A. Safari et al., “Radiation-induced non-targeted effect and carcinogenesis; implications in clinical radiotherapy,” Journal of Biomedical Physics & Engineering, vol. 8, no. 4, pp. 435–446, 2018. View at: Google Scholar
  10. S. M. Meeran, A. Ahmed, and T. O. Tollefsbol, “Epigenetic targets of bioactive dietary components for cancer prevention and therapy,” Clinical Epigenetics, vol. 1, no. 3-4, pp. 101–116, 2010. View at: Publisher Site | Google Scholar
  11. I. Afanas’ev, “Mechanisms of superoxide signaling in epigenetic processes: relation to aging and cancer,” Aging and Disease, vol. 6, no. 3, pp. 216–227, 2015. View at: Publisher Site | Google Scholar
  12. R. Yahyapour, E. Motevaseli, A. Rezaeyan et al., “Reduction-oxidation (redox) system in radiation-induced normal tissue injury: molecular mechanisms and implications in radiation therapeutics,” Clinical and Translational Oncology, vol. 20, no. 8, pp. 975–988, 2018. View at: Publisher Site | Google Scholar
  13. F. Sieber, S. A. Muir, E. P. Cohen et al., “High-dose selenium for the mitigation of radiation injury: a pilot study in a rat model,” Radiation Research, vol. 171, no. 3, pp. 368–373, 2009. View at: Publisher Site | Google Scholar
  14. W. Zhao and M. E. Robbins, “Inflammation and chronic oxidative stress in radiation-induced late normal tissue injury: therapeutic implications,” Current Medicinal Chemistry, vol. 16, no. 2, pp. 130–143, 2009. View at: Publisher Site | Google Scholar
  15. G. R. Drummond, S. Selemidis, K. K. Griendling, and C. G. Sobey, “Combating oxidative stress in vascular disease: NADPH oxidases as therapeutic targets,” Nature Reviews Drug Discovery, vol. 10, no. 6, pp. 453–471, 2011. View at: Publisher Site | Google Scholar
  16. K. Bedard and K. H. Krause, “The NOX family of ROS-generating NADPH oxidases: physiology and pathophysiology,” Physiological Reviews, vol. 87, no. 1, pp. 245–313, 2007. View at: Publisher Site | Google Scholar
  17. A. Panday, M. K. Sahoo, D. Osorio, and S. Batra, “NADPH oxidases: an overview from structure to innate immunity-associated pathologies,” Cellular & Molecular Immunology, vol. 12, no. 1, pp. 5–23, 2015. View at: Publisher Site | Google Scholar
  18. K. Mortezaee, N. H. Goradel, P. Amini et al., “NADPH oxidase as a target for modulation of radiation response; implications to carcinogenesis and radiotherapy,” Current Molecular Pharmacology, vol. 12, no. 1, pp. 50–60, 2019. View at: Publisher Site | Google Scholar
  19. M. Najafi, A. Shirazi, E. Motevaseli et al., “Melatonin modulates regulation of NOX2 and NOX4 following irradiation in the lung,” Current Clinical Pharmacology, vol. 14, 2019. View at: Publisher Site | Google Scholar
  20. B. Bánfi, A. Maturana, S. Jaconi et al., “A mammalian H+ channel generated through alternative splicing of the NADPH oxidase homolog NOH-1,” Science, vol. 287, no. 5450, pp. 138–142, 2000. View at: Publisher Site | Google Scholar
  21. Y. A. Suh, R. S. Arnold, B. Lassegue et al., “Cell transformation by the superoxide-generating oxidase Mox1,” Nature, vol. 401, no. 6748, pp. 79–82, 1999. View at: Publisher Site | Google Scholar
  22. Z. Sun and F. Liu, “Association of Nox1 and vinculin with colon cancer progression,” Cancer Investigation, vol. 31, no. 4, pp. 273–278, 2013. View at: Publisher Site | Google Scholar
  23. F. Liu, A. M. Gomez Garcia, and F. L. Meyskens Jr., “NADPH oxidase 1 overexpression enhances invasion via matrix metalloproteinase-2 and epithelial-mesenchymal transition in melanoma cells,” The Journal of Investigative Dermatology, vol. 132, no. 8, pp. 2033–2041, 2012. View at: Publisher Site | Google Scholar
  24. X. J. Fu, Y. B. Peng, Y. P. Hu, Y. Z. Shi, M. Yao, and X. Zhang, “NADPH oxidase 1 and its derived reactive oxygen species mediated tissue injury and repair,” Oxidative Medicine and Cellular Longevity, vol. 2014, Article ID 282854, 10 pages, 2014. View at: Publisher Site | Google Scholar
  25. S. H. Choi, M. Kim, H. J. Lee, E. H. Kim, C. H. Kim, and Y. J. Lee, “Effects of NOX1 on fibroblastic changes of endothelial cells in radiation‑induced pulmonary fibrosis,” Molecular Medicine Reports, vol. 13, no. 5, pp. 4135–4142, 2016. View at: Publisher Site | Google Scholar
  26. J. H. Kim, K. M. Kim, M. H. Jung et al., “Protective effects of alpha lipoic acid on radiation-induced salivary gland injury in rats,” Oncotarget, vol. 7, no. 20, pp. 29143–29153, 2016. View at: Publisher Site | Google Scholar
  27. P. K. Narayanan, E. H. Goodwin, and B. E. Lehnert, “α particles initiate biological production of superoxide anions and hydrogen peroxide in human cells,” Cancer Research, vol. 57, no. 18, pp. 3963–3971, 1997. View at: Google Scholar
  28. K. Datta, S. Suman, B. V. S. Kallakury, and A. J. Fornace, “Exposure to heavy ion radiation induces persistent oxidative stress in mouse intestine,” PLoS One, vol. 7, no. 8, article e42224, 2012. View at: Publisher Site | Google Scholar
  29. H. Kikuchi, M. Hikage, H. Miyashita, and M. Fukumoto, “NADPH oxidase subunit, gp91(phox) homologue, preferentially expressed in human colon epithelial cells,” Gene, vol. 254, no. 1-2, pp. 237–243, 2000. View at: Publisher Site | Google Scholar
  30. B. Bánfi, B. Malgrange, J. Knisz, K. Steger, M. Dubois-Dauphin, and K. H. Krause, “NOX3, a superoxide-generating NADPH oxidase of the inner ear,” The Journal of Biological Chemistry, vol. 279, no. 44, pp. 46065–46072, 2004. View at: Publisher Site | Google Scholar
  31. R. Paffenholz, R. A. Bergstrom, F. Pasutto et al., “Vestibular defects in head-tilt mice result from mutations in Nox3, encoding an NADPH oxidase,” Genes & Development, vol. 18, no. 5, pp. 486–491, 2004. View at: Publisher Site | Google Scholar
  32. Y. S. Shin, H. A. Shin, S. U. Kang et al., “Effect of epicatechin against radiation-induced oral mucositis: in vitro and in vivo study,” PLoS One, vol. 8, no. 7, article e69151, 2013. View at: Publisher Site | Google Scholar
  33. M. Geiszt, J. B. Kopp, P. Várnai, and T. L. Leto, “Identification of renox, an NAD(P)H oxidase in kidney,” Proceedings of the National Academy of Sciences of the United States of America, vol. 97, no. 14, pp. 8010–8014, 2000. View at: Publisher Site | Google Scholar
  34. A. Shiose, J. Kuroda, K. Tsuruya et al., “A novel superoxide-producing NAD(P)H oxidase in kidney,” The Journal of Biological Chemistry, vol. 276, no. 2, pp. 1417–1423, 2001. View at: Publisher Site | Google Scholar
  35. S. K. Pazhanisamy, H. Li, Y. Wang, I. Batinic-Haberle, and D. Zhou, “NADPH oxidase inhibition attenuates total body irradiation-induced haematopoietic genomic instability,” Mutagenesis, vol. 26, no. 3, pp. 431–435, 2011. View at: Publisher Site | Google Scholar
  36. Y. Wang, L. Liu, S. K. Pazhanisamy, H. Li, A. Meng, and D. Zhou, “Total body irradiation causes residual bone marrow injury by induction of persistent oxidative stress in murine hematopoietic stem cells,” Free Radical Biology & Medicine, vol. 48, no. 2, pp. 348–356, 2010. View at: Publisher Site | Google Scholar
  37. H. Zhang, Y. A. Wang, A. Meng et al., “Inhibiting TGFbeta1 has a protective effect on mouse bone marrow suppression following ionizing radiation exposure in vitro,” Journal of Radiation Research, vol. 54, no. 4, pp. 630–636, 2013. View at: Publisher Site | Google Scholar
  38. B. Bánfi, F. Tirone, I. Durussel et al., “Mechanism of Ca2+ activation of the NADPH oxidase 5 (NOX5),” The Journal of Biological Chemistry, vol. 279, no. 18, pp. 18583–18591, 2004. View at: Publisher Site | Google Scholar
  39. G. Cheng, Z. Cao, X. Xu, E. van Meir, and J. D. Lambeth, “Homologs of gp91phox: cloning and tissue expression of Nox3, Nox4, and Nox5,” Gene, vol. 269, no. 1-2, pp. 131–140, 2001. View at: Publisher Site | Google Scholar
  40. N. Salles, I. Szanto, F. Herrmann et al., “Expression of mRNA for ROS-generating NADPH oxidases in the aging stomach,” Experimental Gerontology, vol. 40, no. 4, pp. 353–357, 2005. View at: Publisher Site | Google Scholar
  41. B. Bánfi, G. Molnár, A. Maturana et al., “A Ca2+-activated NADPH oxidase in testis, spleen, and lymph nodes,” The Journal of Biological Chemistry, vol. 276, no. 40, pp. 37594–37601, 2001. View at: Publisher Site | Google Scholar
  42. X. De Deken, D. Wang, M. C. Many et al., “Cloning of two human thyroid cDNAs encoding new members of the NADPH oxidase family,” The Journal of Biological Chemistry, vol. 275, no. 30, pp. 23227–23233, 2000. View at: Publisher Site | Google Scholar
  43. C. Dupuy, R. Ohayon, A. Valent, M. S. Noël-Hudson, D. Dème, and A. Virion, “Purification of a novel flavoprotein involved in the thyroid NADPH oxidase. Cloning of the porcine and human cdnas,” The Journal of Biological Chemistry, vol. 274, no. 52, pp. 37265–37269, 1999. View at: Publisher Site | Google Scholar
  44. R. Forteza, M. Salathe, F. Miot, R. Forteza, and G. E. Conner, “Regulated hydrogen peroxide production by Duox in human airway epithelial cells,” American Journal of Respiratory Cell and Molecular Biology, vol. 32, no. 5, pp. 462–469, 2005. View at: Publisher Site | Google Scholar
  45. C. Schwarzer, T. E. Machen, B. Illek, and H. Fischer, “NADPH oxidase-dependent acid production in airway epithelial cells,” The Journal of Biological Chemistry, vol. 279, no. 35, pp. 36454–36461, 2004. View at: Publisher Site | Google Scholar
  46. M. Geiszt, J. Witta, J. Baffi, K. Lekstrom, and T. L. Leto, “Dual oxidases represent novel hydrogen peroxide sources supporting mucosal surface host defense,” The FASEB Journal, vol. 17, no. 11, pp. 1502–1504, 2003. View at: Publisher Site | Google Scholar
  47. D. Wang, X. de Deken, M. Milenkovic et al., “Identification of a novel partner of duox: EFP1, a thioredoxin-related protein,” The Journal of Biological Chemistry, vol. 280, no. 4, pp. 3096–3103, 2005. View at: Publisher Site | Google Scholar
  48. C. Dupuy, M. Pomerance, R. Ohayon et al., “Thyroid oxidase (THOX2) gene expression in the rat thyroid cell line FRTL-5,” Biochemical and Biophysical Research Communications, vol. 277, no. 2, pp. 287–292, 2000. View at: Publisher Site | Google Scholar
  49. R. A. El Hassani, N. Benfares, B. Caillou et al., “Dual oxidase2 is expressed all along the digestive tract,” American Journal of Physiology-Gastrointestinal and Liver Physiology, vol. 288, no. 5, pp. G933–G942, 2005. View at: Publisher Site | Google Scholar
  50. R. Ameziane-El-Hassani, M. Talbot, M. C. de Souza Dos Santos et al., “NADPH oxidase DUOX1 promotes long-term persistence of oxidative stress after an exposure to irradiation,” Proceedings of the National Academy of Sciences of the United States of America, vol. 112, no. 16, pp. 5051–5056, 2015. View at: Publisher Site | Google Scholar
  51. Y. Wu and J. H. Doroshow, “Abstract 5358: IL-4/IL-13 induce Duox2/DuoxA2 expression and reactive oxygen production in human pancreatic and colon cancer cells,” Cancer Research, vol. 74, Supplement 19, pp. 5358–5358, 2014. View at: Publisher Site | Google Scholar
  52. R. Azmoonfar, P. Amini, H. Saffar et al., “Metformin protects against radiation-induced pneumonitis and fibrosis and attenuates upregulation of dual oxidase genes expression,” Advanced Pharmaceutical Bulletin, vol. 8, no. 4, pp. 697–704, 2018. View at: Publisher Site | Google Scholar
  53. P. Amini, S. Kolivand, H. Saffar et al., “Protective effect of selenium-L-methionine on radiation-induced acute pneumonitis and lung fibrosis in rat,” Current Clinical Pharmacology, vol. 14, no. 2, pp. 157–164, 2019. View at: Publisher Site | Google Scholar
  54. S. Kolivand, P. Amini, H. Saffar et al., “Selenium-L-methionine modulates radiation injury and Duox1 and Duox2 upregulation in rat’s heart tissues,” Journal of Cardiovascular and Thoracic Research, vol. 11, no. 2, pp. 121–126, 2019. View at: Publisher Site | Google Scholar
  55. A. Aliasgharzadeh, B. Farhood, P. Amini et al., “Melatonin attenuates upregulation of Duox1 and Duox2 and protects against lung injury following chest irradiation in rats,” Cell Journal, vol. 21, no. 3, pp. 236–242, 2019. View at: Publisher Site | Google Scholar
  56. B. Farhood, A. Aliasgharzadeh, P. Amini et al., “Radiation-induced dual oxidase upregulation in rat heart tissues: protective effect of melatonin,” Medicina, vol. 55, no. 7, p. 317, 2019. View at: Publisher Site | Google Scholar
  57. J. Y. Fu, J. L. Masferrer, K. Seibert, A. Raz, and P. Needleman, “The induction and suppression of prostaglandin H2 synthase (cyclooxygenase) in human monocytes,” The Journal of Biological Chemistry, vol. 265, no. 28, pp. 16737–16740, 1990. View at: Google Scholar
  58. J. Song, Y. Wei, Q. Chen, and D. Xing, “Cyclooxygenase 2-mediated apoptotic and inflammatory responses in photodynamic therapy treated breast adenocarcinoma cells and xenografts,” Journal of Photochemistry and Photobiology B, vol. 134, pp. 27–36, 2014. View at: Publisher Site | Google Scholar
  59. D. Wang and R. N. Dubois, “The role of COX-2 in intestinal inflammation and colorectal cancer,” Oncogene, vol. 29, no. 6, pp. 781–788, 2010. View at: Publisher Site | Google Scholar
  60. M. Laube, T. Kniess, and J. Pietzsch, “Development of antioxidant COX-2 inhibitors as radioprotective agents for radiation therapy—a hypothesis-driven review,” Antioxidants, vol. 5, no. 2, p. 14, 2016. View at: Publisher Site | Google Scholar
  61. A. Rezaeyan, G. H. Haddadi, M. Hosseinzadeh, M. Moradi, and M. Najafi, “Radioprotective effects of hesperidin on oxidative damages and histopathological changes induced by X-irradiation in rats heart tissue,” Journal of Medical Physics, vol. 41, no. 3, pp. 182–191, 2016. View at: Publisher Site | Google Scholar
  62. M. Cheki, R. Yahyapour, B. Farhood et al., “COX-2 in radiotherapy: a potential target for radioprotection and radiosensitization,” Current Molecular Pharmacology, vol. 11, no. 3, pp. 173–183, 2018. View at: Publisher Site | Google Scholar
  63. T. A. Zykova, F. Zhu, X. Zhai et al., “Resveratrol directly targets COX-2 to inhibit carcinogenesis,” Molecular Carcinogenesis, vol. 47, no. 10, pp. 797–805, 2008. View at: Publisher Site | Google Scholar
  64. P. Rao and E. E. Knaus, “Evolution of nonsteroidal anti-inflammatory drugs (NSAIDs): cyclooxygenase (COX) inhibition and beyond,” Journal of Pharmacy & Pharmaceutical Sciences, vol. 11, no. 2, pp. 81s–110s, 2008. View at: Publisher Site | Google Scholar
  65. D. Citrin, A. P. Cotrim, F. Hyodo, B. J. Baum, M. C. Krishna, and J. B. Mitchell, “Radioprotectors and mitigators of radiation-induced normal tissue injury,” The Oncologist, vol. 15, no. 4, pp. 360–371, 2010. View at: Publisher Site | Google Scholar
  66. M. Maccarrone, “Lipoxygenases, apoptosis, and the role of antioxidants,” in Photoprotection, Photoinhibition, Gene Regulation, and Environment, B. Demmig-Adams, W. W. Adams, and A. K. Mattoo, Eds., vol. 21 of Advances in Photosynthesis and Respiration, pp. 321–332, Springer, Dordrecht, 2006. View at: Publisher Site | Google Scholar
  67. O. P. Matyshevskaia, V. N. Pastukh, and V. A. Solodushko, “Inhibition of lipoxygenase activity reduces radiation-induced DNA fragmentation in lymphocytes,” Radiatsionnaia Biologiia, Radioecologiia, vol. 39, no. 2-3, pp. 282–286, 1999. View at: Google Scholar
  68. O. E. Grichenko, A. C. Pushin, V. V. Shaposhnikova, M. K. H. Levitman, and N. Korystov Iu, “Analysis of 15-lipoxygenase activity in irradiated thymocytes,” Izvestiia Akademii Nauk. Seriia Biologicheskaia, vol. 5, pp. 517–521, 2004. View at: Google Scholar
  69. M. Halle, T. Christersdottir, and M. Back, “Chronic adventitial inflammation, vasa vasorum expansion, and 5-lipoxygenase up-regulation in irradiated arteries from cancer survivors,” The FASEB Journal, vol. 30, no. 11, pp. 3845–3852, 2016. View at: Publisher Site | Google Scholar
  70. F. Aktan, “iNOS-mediated nitric oxide production and its regulation,” Life Sciences, vol. 75, no. 6, pp. 639–653, 2004. View at: Publisher Site | Google Scholar
  71. M. Najafi, E. Motevaseli, A. Shirazi et al., “Mechanisms of inflammatory responses to radiation and normal tissues toxicity: clinical implications,” International Journal of Radiation Biology, vol. 94, no. 4, pp. 335–356, 2018. View at: Publisher Site | Google Scholar
  72. M. Jaiswal, N. LaRusso, R. A. Shapiro, T. R. Billiar, and G. J. Gores, “Nitric oxide-mediated inhibition of DNA repair potentiates oxidative DNA damage in cholangiocytes,” Gastroenterology, vol. 120, no. 1, pp. 190–199, 2001. View at: Publisher Site | Google Scholar
  73. R. Malaviya, A. J. Gow, M. Francis, E. V. Abramova, J. D. Laskin, and D. L. Laskin, “Radiation-induced lung injury and inflammation in mice: role of inducible nitric oxide synthase and surfactant protein D,” Toxicological Sciences, vol. 144, no. 1, pp. 27–38, 2015. View at: Publisher Site | Google Scholar
  74. Y. Nozaki, Y. Hasegawa, A. Takeuchi et al., “Nitric oxide as an inflammatory mediator of radiation pneumonitis in rats,” The American Journal of Physiology, vol. 272, 4 Part 1, pp. L651–L658, 1997. View at: Publisher Site | Google Scholar
  75. C. Tsuji, S. Shioya, Y. Hirota et al., “Increased production of nitrotyrosine in lung tissue of rats with radiation-induced acute lung injury,” American Journal of Physiology-Lung Cellular and Molecular Physiology, vol. 278, no. 4, pp. L719–L725, 2000. View at: Publisher Site | Google Scholar
  76. S. Ohta, S. Matsuda, M. Gunji, and A. Kamogawa, “The role of nitric oxide in radiation damage,” Biological & Pharmaceutical Bulletin, vol. 30, no. 6, pp. 1102–1107, 2007. View at: Publisher Site | Google Scholar
  77. J. S. Beckman and W. H. Koppenol, “Nitric oxide, superoxide, and peroxynitrite: the good, the bad, and ugly,” American Journal of Physiology-Cell Physiology, vol. 271, no. 5, pp. C1424–C1437, 1996. View at: Publisher Site | Google Scholar
  78. V. A. Yakovlev, “Role of nitric oxide in the radiation-induced bystander effect,” Redox Biology, vol. 6, pp. 396–400, 2015. View at: Publisher Site | Google Scholar
  79. S. Ghosh, D. K. Maurya, and M. Krishna, “Role of iNOS in bystander signaling between macrophages and lymphoma cells,” International Journal of Radiation Oncology, Biology, Physics, vol. 72, no. 5, pp. 1567–1574, 2008. View at: Publisher Site | Google Scholar
  80. V. A. Yakovlev, “Nitric oxide-dependent downregulation of BRCA1 expression promotes genetic instability,” Cancer Research, vol. 73, no. 2, pp. 706–715, 2013. View at: Publisher Site | Google Scholar
  81. W. Han, L. Wu, S. Chen et al., “Constitutive nitric oxide acting as a possible intercellular signaling molecule in the initiation of radiation-induced DNA double strand breaks in non-irradiated bystander cells,” Oncogene, vol. 26, no. 16, pp. 2330–2339, 2007. View at: Publisher Site | Google Scholar
  82. J. Pugin, “How tissue injury alarms the immune system and causes a systemic inflammatory response syndrome,” Annals of Intensive Care, vol. 2, no. 1, p. 27, 2012. View at: Publisher Site | Google Scholar
  83. B. Frey, M. Rückert, L. Deloch et al., “Immunomodulation by ionizing radiation-impact for design of radio-immunotherapies and for treatment of inflammatory diseases,” Immunological Reviews, vol. 280, no. 1, pp. 231–248, 2017. View at: Publisher Site | Google Scholar
  84. Y. Shen, X. Jiang, L. Meng, C. Xia, L. Zhang, and Y. Xin, “Transplantation of bone marrow mesenchymal stem cells prevents radiation- induced artery injury by suppressing oxidative stress and inflammation,” Oxidative Medicine and Cellular Longevity, vol. 2018, Article ID 5942916, 13 pages, 2018. View at: Publisher Site | Google Scholar
  85. Y. Ogawa, T. Kobayashi, A. Nishioka et al., “Radiation-induced reactive oxygen species formation prior to oxidative DNA damage in human peripheral T cells,” International Journal of Molecular Medicine, vol. 11, no. 2, pp. 149–152, 2003. View at: Google Scholar
  86. K. Mortezaee, M. Najafi, B. Farhood, A. Ahmadi, D. Shabeeb, and A. E. Musa, “NF-κB targeting for overcoming tumor resistance and normal tissues toxicity,” Journal of Cellular Physiology, vol. 234, no. 10, pp. 17187–17204, 2019. View at: Publisher Site | Google Scholar
  87. V. Bours, G. Bonizzi, M. Bentires-Alj et al., “NF-κB activation in response to toxical and therapeutical agents: role in inflammation and cancer treatment,” Toxicology, vol. 153, no. 1-3, pp. 27–38, 2000. View at: Publisher Site | Google Scholar
  88. S. M. Meeran, S. Akhtar, and S. K. Katiyar, “Inhibition of UVB-induced skin tumor development by drinking green tea polyphenols is mediated through DNA repair and subsequent inhibition of inflammation,” The Journal of Investigative Dermatology, vol. 129, no. 5, pp. 1258–1270, 2009. View at: Publisher Site | Google Scholar
  89. M. A. Packer and M. P. Murphy, “Peroxynitrite formed by simultaneous nitric oxide and superoxide generation causes cyclosporin-A-sensitive mitochondrial calcium efflux and depolarisation,” European Journal of Biochemistry, vol. 234, no. 1, pp. 231–239, 1995. View at: Publisher Site | Google Scholar
  90. R. A. Kowluru and M. Mishra, “Oxidative stress, mitochondrial damage and diabetic retinopathy,” Biochimica et Biophysica Acta (BBA) - Molecular Basis of Disease, vol. 1852, no. 11, pp. 2474–2483, 2015. View at: Publisher Site | Google Scholar
  91. F. Ortiz, D. Acuña-Castroviejo, C. Doerrier et al., “Melatonin blunts the mitochondrial/NLRP3 connection and protects against radiation-induced oral mucositis,” Journal of Pineal Research, vol. 58, no. 1, pp. 34–49, 2015. View at: Publisher Site | Google Scholar
  92. J. C. Leemans, S. L. Cassel, and F. S. Sutterwala, “Sensing damage by the NLRP3 inflammasome,” Immunological Reviews, vol. 243, no. 1, pp. 152–162, 2011. View at: Publisher Site | Google Scholar
  93. R. Han, D. Wu, S. Deng, T. Liu, T. Zhang, and Y. Xu, “NLRP3 inflammasome induces pyroptosis in lung tissues of radiation-induced lung injury in mice,” Xi Bao Yu Fen Zi Mian Yi Xue Za Zhi, vol. 33, no. 9, pp. 1206–1211, 2017. View at: Google Scholar
  94. D. Shin, G. Lee, S. H. Sohn et al., “Regulatory T cells contribute to the inhibition of radiation-induced acute lung inflammation via bee venom phospholipase A2 in mice,” Toxins, vol. 8, no. 5, p. 131, 2016. View at: Publisher Site | Google Scholar
  95. S. H. Sohn, J. M. Lee, S. Park et al., “The inflammasome accelerates radiation-induced lung inflammation and fibrosis in mice,” Environmental Toxicology and Pharmacology, vol. 39, no. 2, pp. 917–926, 2015. View at: Publisher Site | Google Scholar
  96. Y. G. Liu, J. K. Chen, Z. T. Zhang et al., “NLRP3 inflammasome activation mediates radiation-induced pyroptosis in bone marrow-derived macrophages,” Cell Death & Disease, vol. 8, no. 2, article e2579, 2017. View at: Publisher Site | Google Scholar
  97. R. Yahyapour, P. Amini, S. Rezapoor et al., “Targeting of inflammation for radiation protection and mitigation,” Current Molecular Pharmacology, vol. 11, no. 3, pp. 203–210, 2018. View at: Publisher Site | Google Scholar
  98. K. Suzuki, I. Mori, Y. Nakayama, M. Miyakoda, S. Kodama, and M. Watanabe, “Radiation-induced senescence-like growth arrest requires TP53 function but not telomere shortening,” Radiation Research, vol. 155, no. 1, pp. 248–253, 2001. View at: Publisher Site | Google Scholar
  99. N. J. Linford, S. E. Schriner, and P. S. Rabinovitch, “Oxidative damage and aging: spotlight on mitochondria,” Cancer Research, vol. 66, no. 5, pp. 2497–2499, 2006. View at: Publisher Site | Google Scholar
  100. S. E. Schriner, N. J. Linford, G. M. Martin et al., “Extension of murine life span by overexpression of catalase targeted to mitochondria,” Science, vol. 308, no. 5730, pp. 1909–1911, 2005. View at: Publisher Site | Google Scholar
  101. S. Kobashigawa, G. Kashino, H. Mori, and M. Watanabe, “Relief of delayed oxidative stress by ascorbic acid can suppress radiation-induced cellular senescence in mammalian fibroblast cells,” Mechanisms of Ageing and Development, vol. 146-148, pp. 65–71, 2015. View at: Publisher Site | Google Scholar
  102. Y. Sakai, T. Yamamori, Y. Yoshikawa et al., “NADPH oxidase 4 mediates ROS production in radiation-induced senescent cells and promotes migration of inflammatory cells,” Free Radical Research, vol. 52, no. 1, pp. 92–102, 2018. View at: Publisher Site | Google Scholar
  103. J. S. You and P. A. Jones, “Cancer genetics and epigenetics: two sides of the same coin?” Cancer Cell, vol. 22, no. 1, pp. 9–20, 2012. View at: Publisher Site | Google Scholar
  104. M. Verma, D. Seminara, F. J. Arena, C. John, K. Iwamoto, and V. Hartmuller, “Genetic and epigenetic biomarkers in cancer: improving diagnosis, risk assessment, and disease stratification,” Molecular Diagnosis & Therapy, vol. 10, no. 1, pp. 1–15, 2006. View at: Publisher Site | Google Scholar
  105. M. Verma, “Cancer control and prevention: nutrition and epigenetics,” Current Opinion in Clinical Nutrition and Metabolic Care, vol. 16, no. 4, pp. 376–384, 2013. View at: Publisher Site | Google Scholar
  106. I. R. Miousse, J. Chang, L. Shao et al., “Inter-strain differences in LINE-1 DNA methylation in the mouse hematopoietic system in response to exposure to ionizing radiation,” International Journal of Molecular Sciences, vol. 18, no. 7, p. 1430, 2017. View at: Publisher Site | Google Scholar
  107. T. Z. Yi, J. Li, X. Han et al., “DNMT inhibitors and HDAC inhibitors regulate E-cadherin and Bcl-2 expression in endometrial carcinoma in vitro and in vivo,” Chemotherapy, vol. 58, no. 1, pp. 19–29, 2012. View at: Publisher Site | Google Scholar
  108. N. Reichert, M. A. Choukrallah, and P. Matthias, “Multiple roles of class I HDACs in proliferation, differentiation, and development,” Cellular and Molecular Life Sciences, vol. 69, no. 13, pp. 2173–2187, 2012. View at: Publisher Site | Google Scholar
  109. M. S. Choi, Y. H. Shim, J. Y. Hwa et al., “Expression of DNA methyltransferases in multistep hepatocarcinogenesis,” Human Pathology, vol. 34, no. 1, pp. 11–17, 2003. View at: Publisher Site | Google Scholar
  110. Q. Liu, J.-M. Zheng, J.-K. Chen et al., “Histone deacetylase 5 promotes the proliferation of glioma cells by upregulation of Notch 1,” Molecular Medicine Reports, vol. 10, no. 4, pp. 2045–2050, 2014. View at: Publisher Site | Google Scholar
  111. G. W. Feng, L. D. Dong, W. J. Shang et al., “HDAC5 promotes cell proliferation in human hepatocellular carcinoma by up-regulating Six1 expression,” European Review for Medical and Pharmacological Sciences, vol. 18, no. 6, pp. 811–816, 2014. View at: Google Scholar
  112. M. Z. Fang, Y. Wang, N. Ai et al., “Tea polyphenol (–)-epigallocatechin-3-gallate inhibits DNA methyltransferase and reactivates methylation-silenced genes in cancer cell lines,” Cancer Research, vol. 63, no. 22, pp. 7563–7570, 2003. View at: Google Scholar
  113. A. S. Tsao, D. Liu, J. Martin et al., “Phase II randomized, placebo-controlled trial of green tea extract in patients with high-risk oral premalignant lesions,” Cancer Prevention Research, vol. 2, no. 11, pp. 931–941, 2009. View at: Publisher Site | Google Scholar
  114. C. Weigel, P. Schmezer, C. Plass, and O. Popanda, “Epigenetics in radiation-induced fibrosis,” Oncogene, vol. 34, no. 17, pp. 2145–2155, 2015. View at: Publisher Site | Google Scholar
  115. C. Weigel, M. R. Veldwijk, C. C. Oakes et al., “Epigenetic regulation of diacylglycerol kinase alpha promotes radiation- induced fibrosis,” Nature Communications, vol. 7, no. 1, article 10893, 2016. View at: Publisher Site | Google Scholar
  116. M. M. Acharya, A. A. D. Baddour, T. Kawashita et al., “Epigenetic determinants of space radiation-induced cognitive dysfunction,” Scientific Reports, vol. 7, no. 1, article 42885, 2017. View at: Publisher Site | Google Scholar
  117. H. Denis, M. N. Ndlovu, and F. Fuks, “Regulation of mammalian DNA methyltransferases: a route to new mechanisms,” EMBO Reports, vol. 12, no. 7, pp. 647–656, 2011. View at: Publisher Site | Google Scholar
  118. D. A. Antwih, K. M. Gabbara, W. D. Lancaster, D. M. Ruden, and S. P. Zielske, “Radiation-induced epigenetic DNA methylation modification of radiation-response pathways,” Epigenetics, vol. 8, no. 8, pp. 839–848, 2013. View at: Publisher Site | Google Scholar
  119. I. Pogribny, I. Koturbash, V. Tryndyak et al., “Fractionated low-dose radiation exposure leads to accumulation of DNA damage and profound alterations in DNA and histone methylation in the murine thymus,” Molecular Cancer Research, vol. 3, no. 10, pp. 553–561, 2005. View at: Publisher Site | Google Scholar
  120. I. Koturbash, A. Boyko, R. Rodriguez-Juarez et al., “Role of epigenetic effectors in maintenance of the long-term persistent bystander effect in spleen in vivo,” Carcinogenesis, vol. 28, no. 8, pp. 1831–1838, 2007. View at: Publisher Site | Google Scholar
  121. H. D. Halicka, X. Huang, F. Traganos, M. A. King, W. Dai, and Z. Darzynkiewicz, “Histone H2AX phosphorylation after cell irradiation with UV-B: relationship to cell cycle phase and induction of apoptosis,” Cell Cycle, vol. 4, no. 2, pp. 339–345, 2005. View at: Google Scholar
  122. B. Maroschik, A. Gürtler, A. Krämer et al., “Radiation-induced alterations of histone post-translational modification levels in lymphoblastoid cell lines,” Radiation Oncology, vol. 9, no. 1, p. 15, 2014. View at: Publisher Site | Google Scholar
  123. S. Matsuda, K. Furuya, M. Ikura, T. Matsuda, and T. Ikura, “Absolute quantification of acetylation and phosphorylation of the histone variant H2AX upon ionizing radiation reveals distinct cellular responses in two cancer cell lines,” Radiation and Environmental Biophysics, vol. 54, no. 4, pp. 403–411, 2015. View at: Publisher Site | Google Scholar
  124. M. Herberg, S. Siebert, M. Quaas et al., “Loss of Msh2 and a single-radiation hit induce common, genome-wide, and persistent epigenetic changes in the intestine,” Clinical Epigenetics, vol. 11, no. 1, p. 65, 2019. View at: Publisher Site | Google Scholar
  125. X. Zhang, T. Kluz, L. Gesumaria, M. S. Matsui, M. Costa, and H. Sun, “Solar simulated ultraviolet radiation induces global histone hypoacetylation in human keratinocytes,” PLoS One, vol. 11, no. 2, article e0150175, 2016. View at: Publisher Site | Google Scholar
  126. D. P. Bartel, “MicroRNAs: target recognition and regulatory functions,” Cell, vol. 136, no. 2, pp. 215–233, 2009. View at: Publisher Site | Google Scholar
  127. B. Farhood, N. H. Goradel, K. Mortezaee et al., “Intercellular communications-redox interactions in radiation toxicity; potential targets for radiation mitigation,” Journal of Cell Communication and Signaling, vol. 13, no. 1, pp. 3–16, 2019. View at: Publisher Site | Google Scholar
  128. M. Najafi, R. Fardid, M. A. Takhshid, M. A. Mosleh-Shirazi, A. H. Rezaeyan, and A. Salajegheh, “Radiation-induced oxidative stress at out-of-field lung tissues after pelvis irradiation in rats,” Cell Journal, vol. 18, no. 3, pp. 340–345, 2016. View at: Google Scholar
  129. S. S. Acharya, W. Fendler, J. Watson et al., “Serum microRNAs are early indicators of survival after radiation-induced hematopoietic injury,” Science Translational Medicine, vol. 7, no. 287, article 287ra69, 2015. View at: Publisher Site | Google Scholar
  130. M. Port, F. Herodin, M. Valente et al., “MicroRNA expression for early prediction of late occurring hematologic acute radiation syndrome in baboons,” PLoS One, vol. 11, no. 11, article e0165307, 2016. View at: Publisher Site | Google Scholar
  131. X. H. Li, C. T. Ha, D. Fu, and M. Xiao, “Micro-RNA30c negatively regulates REDD1 expression in human hematopoietic and osteoblast cells after gamma-irradiation,” PLoS One, vol. 7, no. 11, article e48700, 2012. View at: Publisher Site | Google Scholar
  132. X. H. Li, C. T. Ha, D. Fu, M. R. Landauer, S. P. Ghosh, and M. Xiao, “Delta-Tocotrienol suppresses radiation-induced microRNA-30 and protects mice and human CD34+ cells from radiation injury,” PLoS One, vol. 10, no. 3, article e0122258, 2015. View at: Publisher Site | Google Scholar
  133. M. E. Robbins, J. K. Brunso-Bechtold, A. M. Peiffer, C. I. Tsien, J. E. Bailey, and L. B. Marks, “Imaging radiation-induced normal tissue injury,” Radiation Research, vol. 177, no. 4, pp. 449–466, 2012. View at: Publisher Site | Google Scholar
  134. Y. Huang, Q. Hu, Z. Deng, Y. Hang, J. Wang, and K. Wang, “MicroRNAs in body fluids as biomarkers for non-small cell lung cancer: a systematic review,” Technology in Cancer Research & Treatment, vol. 13, no. 3, pp. 277–287, 2014. View at: Publisher Site | Google Scholar
  135. F. Gao, P. Liu, J. Narayanan et al., “Changes in miRNA in the lung and whole blood after whole thorax irradiation in rats,” Scientific Reports, vol. 7, no. 1, article 44132, 2017. View at: Publisher Site | Google Scholar
  136. L. Xie, J. Zhou, S. Zhang et al., “Integrating microRNA and mRNA expression profiles in response to radiation-induced injury in rat lung,” Radiation Oncology, vol. 9, no. 1, p. 111, 2014. View at: Publisher Site | Google Scholar
  137. S. P. Ghosh, R. Pathak, P. Kumar et al., “gamma-Tocotrienol modulates radiation-induced microRNA expression in mouse spleen,” Radiation Research, vol. 185, no. 5, pp. 485–495, 2016. View at: Publisher Site | Google Scholar

Copyright © 2019 Jinlong Wei et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


More related articles

 PDF Download Citation Citation
 Download other formatsMore
 Order printed copiesOrder
Views2804
Downloads1006
Citations

Related articles

Article of the Year Award: Outstanding research contributions of 2020, as selected by our Chief Editors. Read the winning articles.