In huge DNA molecules in the cell, nucleus genes are present. These genes are responsible for the development and function of the cell and the whole organism, because they code proteins. Due to this fact, unrepaired or misrepaired DNA lesions, which can lead to gene mutations, can promote changes in the structure of the encoded protein or lead to the decrease or complete loss of its expression. The types of DNA lesions occurring were already discussed in Sect. 3.3. Based on the current experimental and theoretical evidence, the most repair-resistant lesions are not the single ones but a combination of them in a short DNA segment of 10–20 bp called clustered damage. Clustered DNA lesions are considered the signature of ionizing radiations especially for particle radiation [45]. Various studies suggest that the probability for a break or other DNA lesion to be incorrectly processed and amended is fairly low when damage is spatially separated but increases drastically when multiple breaks and/or non-break lesions coincide. For an analytical description of DNA repair pathways, the reader can refer to Sect. 3.4. As was already mentioned in Sect. 3.3, the DNA molecule consists of nucleotides (deoxyribose + phosphate group + base), which can be for simplicity named based on the four bases [adenine (A), cytosine (C), guanine (G), thymine (T)]. Thus, the DNA alphabet is a very easy one; it only consists of four letters. These four letters are then combined to give rise to groups of three, which define the amino acids that are then the new alphabet for the translation to proteins. For more details on DNA-to-RNA transcription and RNA-to-protein translation, see for example [62]. Even if the cells have a very sophisticated DNA damage response and repair system, it may happen that not all the damage is removed. A mutation is when a permanent change in the DNA sequence occurs. Mutations can be divided into somatic or germline mutation in terms of what kind of cell is affected. A germline mutation occurs in a sperm or in an egg and can be passed to offspring. Somatic mutations occur in cells of the body and cannot be passed to next generations. Mutations can also be grouped as point or chromosomal mutations. Point mutations are when a single nucleotide is replaced with another single nucleotide, or deleted, or inserted in a place that it should not be. Point mutations do not always have significant consequences on the encoded protein. For example, as is shown in Table 3.4, the mutation can be silent. This means that even if there is a change in the original DNA sequence, the final product of the transcription will be the same, because there are several combinations of the DNA alphabet that lead to the same amino acid. In other cases, the mutation can lead to the change of the final amino acid (missense mutation) or to the creation of a stop codon (nonsense mutation), which then affects the final protein.Table 3.4
Point mutations and their consequences

Point mutations

No mutation

Silent

Missense

Nonsense

DNA

TTC

TTT

TCC

ATC

mRNA

AAG

AAA

AGG

UAG

Amino acid

Lysine (Lys)

Lysine (Lys)

Arginine (Arg)

Stop


Mitotic cell death, also called mitotic catastrophe (MC), is the process when a cell dies during or right after mitosis [63]. It can be triggered by DNA damage and its mis- and unrepair and therefore through radiation. MC can be both a caspase-dependent, regulated and caspase-independent, unregulated pathway of cell death. Some characteristic morphologies can be found in Fig. 3.14a.

8 micrographs. a. A set of 5 micrographs of cells with alpha-tub and D A P I depict control, P D T, T x, and N c. b. A set of 3 micrographs depicts non-senescent and senescent cells.

Fig. 3.14
Morphologies of mitotic catastrophe (a) and senescence (b). (a) Fluorescence image of cancer cells undergoing mitosis. The DNA is labeled with DAPI and mitotic spindles using α-tubulin staining. The cells exhibiting mitotic catastrophe are treated with photodynamic therapy (PDT), Taxol (Tx), or nocodazole (Nc). The control shows normal mitotic spindles. The treated cells show various types of altered spindles and mitosis. Scale bar: 10 μm. Reproduced with permission (CCBY) from Mascaraque et al. [64]. (b) Phase-contrast images of Chang cells. Senescence was induced using 1 mM of deferoxamine. (Reproduced with permission (CCBY) from Kwon et al. [65])


8 micrographs. a. A set of 5 micrographs of cells with alpha-tub and D A P I depict control, P D T, T x, and N c. b. A set of 3 micrographs depicts non-senescent and senescent cells.


Senescence in biology refers to a process by which a cell ages and permanently and irreversibly stops dividing but does not die [63]. The number of senescent cells increases with age, but senescence also plays an important role during development as well as during wound healing and can be triggered by radiation. In culture, senescent cells exhibit a different morphology compared to non-senescent cells, called “fried egg” appearance (see Fig. 3.14b). It was shown that among other features, the radiation dose plays a major role in the induction of either senescence or apoptosis and necrosis. In some cell lines, senescence is the major response to low doses of radiation, whereas higher doses lead to apoptosis or necrosis. In IR-treated tissue, enhanced senescence may lead to pathogenic onsets, such as loss of organ function.


Recent advances in the field indicate that a further consequence of DNA damage misrepair or unrepair can be the release of cytoplasmic DNA that can also trigger immune responses. In general, it is widely accepted that immune signaling can be activated by the presence of DNA in unusual locations, such as the cytoplasm or the endosomes, as DNA is normally located in the nucleus of eukaryotic cells. Emerging evidence indicates a cross talk between DNA repair machinery and the immune system, and more specifically it has been discovered that DDR factors like DNA repair proteins can enhance innate immune signaling [66]. Defects in DDR and proper processing of DNA damage can therefore trigger a multitude of cellular phenotypes, including autoinflammatory disease, cellular senescence, and cancer. Genotoxic agents such as radiations or high oxidative stress can act as the primary instigators for immune signaling activation through the release of a wide range of biological and chemical factors often referred to as “danger signals” or damage-associated molecular patterns (DAMPs) [67] (Box 3.10).


Genomic instability (GI) collectively refers to a status of increased DNA changes, chromosomal rearrangements, and enhanced tendency for genetic alterations occurring during cell division.

Unrepaired or misrepaired DNA lesions can lead to chromosomal mutations, which can lead to cell death or loss of genetic material, thus promoting GI.

Mitotic cell death is the process of a cell dying in relation to mitosis and can be triggered by radiation-induced damages.

Senescence is the status of irreversible cell cycle arrest, which occurs naturally during aging but can be triggered by radiation, which can lead to pathological onsets.

Cytoplasmic DNA and DNA repair defects can trigger immune response.


Genomic instability (GI) collectively refers to a status of increased DNA changes, chromosomal rearrangements, and enhanced tendency for genetic alterations occurring during cell division.


Unrepaired or misrepaired DNA lesions can lead to chromosomal mutations, which can lead to cell death or loss of genetic material, thus promoting GI.


Mitotic cell death is the process of a cell dying in relation to mitosis and can be triggered by radiation-induced damages.


Senescence is the status of irreversible cell cycle arrest, which occurs naturally during aging but can be triggered by radiation, which can lead to pathological onsets.


Cytogenetic techniques can be used to analyze chromosomal aberrations in metaphase and morphological abnormalities of DNA content in interphase nuclei. The applicability of these aberrations in the fields of biological dosimetry, clinical cytogenetics, and environmental monitoring is based on a large number of radiobiological and DNA-repair theories.


As described before, when cells are exposed to a variety of genotoxic agents (chemical/physical/radiation/DNA-damaging agents), they cause defects in DNA, chromosomes, and other cellular components. Radiation induces extensive DNA damage such as DSBs that, if misrepaired or unrepaired, ordinarily result in asymmetrical chromosome rearrangements and exchanges, which may lead to formation of small chromatinic bodies also known as micronuclei (MN) (see Fig. 3.15). MN are tiny extranuclear bodies that contain damaged chromosome fragments and/or whole chromosomes that were not incorporated into the nucleus after cell division and are surrounded by a membrane. As a variety of genotoxic agents may damage DNA and the mitotic machinery by multiple mechanisms, leading to MN formation, MN are not IR specific.

A chart lists several genotoxic agents and describes genotoxic damages that result in micronuclei, nuclear bud, and nucleoplasmic bridge with their respective symptoms.

Fig. 3.15
Mechanisms by which genotoxic agents cause micronuclei and other nuclear anomalies. Micronuclei (MN) can originate from lagging acentric chromosomes or chromatid fragments or whole chromosomes at anaphase in mitosis. Nuclear bud (NBUD) formation represents the process of extrusion of the amplified/surplus DNA, DNA repair-recombinational protein complexes, and possibly excess chromosomes from aneuploidic cells. Nucleoplasmic bridges (NPBs) originate from dicentric chromosomes. This arises because the centromeres of dicentric chromosomes are often pulled in opposite directions and defective separation of sister chromatids occurs during anaphase leading to bridge formation, which can be observed as an NPB in telophase


A chart lists several genotoxic agents and describes genotoxic damages that result in micronuclei, nuclear bud, and nucleoplasmic bridge with their respective symptoms.


It is now well established that MN are formed from acentric chromatid fragments caused by misrepaired or unrepaired DNA breaks or lagging acentric chromosomes due to mitotic spindle failure at an anaphase. Additionally, the formation of DNA DSBs and MN is sometimes the result of simultaneous excision repair of damages (e.g., 8-oxo-deoxyguanosine) and inappropriate bases’ (e.g., uracil) incorporation in proximity on opposite complementary DNA strands.


A whole chromosome lagging behind (chromosome mal-segregation) during anaphase also results in MN formation. Mal-segregation usually happens due to absence or inappropriate attachment of spindle microtubules to chromosome kinetochore. However, the potential mechanisms behind the formation of MN are hypomethylation repeat sequences in centromeric and pericentromeric DNA, defects in kinetochore proteins or assembly, dysfunctional spindle, defective anaphase checkpoint genes, and malfunctioning in cell cycle control system. Sometimes, mis-segregation events occur when the centromeres of the dicentric chromosomes are pulled towards opposite poles of cells with sufficient forces to detach the chromosome from spindle during anaphase, thus resulting in micronucleus formation from whole chromosome loss.


Furthermore, multiple extrachromosomal acentric double minutes (DMs), cytogenetic hallmarks of genomic amplification, can aggregate after DNA damage and generate cytoplasmic MN that are subsequently eliminated from the cell.


Other nuclear anomalies such as nucleoplasmic bridges (NPBs) and nuclear buds (NBUDs) (see Fig. 3.15) are sensitive and reliable biomarkers for early genotoxic instability and chromosomal breakages and rearrangements. NPBs originate as an aftereffect of misrepair of DNA strand breaks or failure of complete chromatid separation to opposite poles of the cell during anaphase. It can also originate from telomere end-to-end fusion mechanism, a fundamental indication of and a marker for loss of telomere function, which is caused by (a) excessively short telomeres, (b) dysfunctional telomeres due to loss of telomere-binding proteins without telomere erosion, (c) inappropriate assembly of telomere-capping protein structure, (d) defects in recombinational repair proteins, or (e) lack of telomeres. Another distinctive nuclear anomaly, NBUDs, is one of the precursors of MN and is associated with chromosomal instability events. Most NBUDs originate from interstitial or terminal acentric fragments and represent the expulsion of undesirable amplified extrachromosomal DNA content, which localizes to specific sites at the periphery of the nucleus and is eventually eliminated via nuclear budding during the S phase of cell cycle. It is also plausible that NBUDs might occur after elimination of DNA repair-protein complexes in the cytoplasm (Box 3.11).


Micronuclei are small extranuclear bodies surrounded by a membrane that contain damaged chromosome fragments or even whole chromosomes. The genetic information encoded in the MN DNA will get lost and lead to large genomic consequences.

Chromosome segregation errors and/or fragment loss at anaphase (“inter-cell bridges”) and exclusion of acentric fragments from daughter nuclei lead to formation of MN in the cytoplasm.

Micronuclei occur outside the main cellular nucleus and are prone to rupturing, which leads to changes in DNA that can drive cancer development.

Extensive DNA damage may cause dicentric/concatenated ring chromosomes and acentric chromatid/chromosome fragments, which can result in the formation of a nucleoplasmic bridge (NPB) at anaphase and micronuclei, respectively.

Nuclear buds (NBUDs) are the result of elimination of amplified extrachromosomal DNA, which adheres to the nucleus by a thin nucleoplasmic connection, and are observed as double minute-type micronucleus bodies.


Micronuclei are small extranuclear bodies surrounded by a membrane that contain damaged chromosome fragments or even whole chromosomes. The genetic information encoded in the MN DNA will get lost and lead to large genomic consequences.


Chromosome segregation errors and/or fragment loss at anaphase (“inter-cell bridges”) and exclusion of acentric fragments from daughter nuclei lead to formation of MN in the cytoplasm.


Micronuclei occur outside the main cellular nucleus and are prone to rupturing, which leads to changes in DNA that can drive cancer development.


Extensive DNA damage may cause dicentric/concatenated ring chromosomes and acentric chromatid/chromosome fragments, which can result in the formation of a nucleoplasmic bridge (NPB) at anaphase and micronuclei, respectively.


Nuclear buds (NBUDs) are the result of elimination of amplified extrachromosomal DNA, which adheres to the nucleus by a thin nucleoplasmic connection, and are observed as double minute-type micronucleus bodies.


Micronucleus assays are frequently used to assess genotoxicity and cytotoxicity of different chemical and physical factors, including IR-induced DNA damage. The cytokinesis-block micronucleus assay can measure MN, NPBs, and NBUDs. A diverse range of reliable micronucleus tests (Fig. 3.16) are executed with different cell types, eventually reflecting chromosomal aberrations, ongoing DNA injury, initial stage in the development of genomic instability, and tumorigenesis. In the widely used cytokinesis-blocked MN assay, MN are scored in once-divided binucleated cells, where cytokinesis is blocked with addition of cytochalasin B, an inhibitor of microfilament ring assembly necessary for the completion of cytokinesis. In order to get statistically solid results, a huge amount of cells need to be scored. Therefore, automatic analysis of MN boosts the reliability of the assays. Concomitantly, it increases the statistical validity after analyzing a large number of cells in one go. Additionally, the existing automatic/semiautomatic micronucleus scoring by microscopic systems, by flow cytometry and imaging flow cytometry, gives high accuracy and sensitivity and leads to rapid analysis (Box 3.12).

An illustration lists the different factors of the cytokinesis-blocked micronucleus cytome assay, the mammalian erythrocyte micronucleus assay, the buccal micronucleus cytome assay, and the micronucleus assay in other cell types.

Fig. 3.16
Depending on the cell type, different micronucleus assays can be employed to assess and determine the genotoxicity and cytotoxicity of different chemical and physical factors. Applications of each assay are outlined in their respective boxes. The most popular CBMN assay can be applied to cultured human lymphocytes or cell lines to measure MN and other chromosomal instability biomarkers such as NPBs and NBUD. The mammalian erythrocyte micronucleus assay is performed on immature erythrocytes from bone marrow to determine cytogenetic damage after radiation exposure. The buccal micronucleus cytome assay is done in rapidly dividing buccal epithelial exfoliated cells (oral cavity) to analyze MN and other cytogenetic biomarkers (source of DNA damage, cytotoxicity, etc.). Occasionally, MN assay is performed on nasal mucosa cells or urine-derived cells for detection of chromosomal damage caused by environmental and lifestyle factors, occupational exposures, prognosis of cancer, and certain diseases. Although the objective and method of performance are similar to CBMN or bone marrow MN assays, these tests have not gained much popularity so far


An illustration lists the different factors of the cytokinesis-blocked micronucleus cytome assay, the mammalian erythrocyte micronucleus assay, the buccal micronucleus cytome assay, and the micronucleus assay in other cell types.


Micronucleus assays are used to assess genotoxicity and cytotoxicity of radiation.

Depending on cell type, different MN assays are used.

Automated analysis of MN boosts the reliability and statistical validity.


Micronucleus assays are used to assess genotoxicity and cytotoxicity of radiation.


Chromosomal mutations, also called chromosomal aberrations (CA), are observed at the first mitosis after irradiation and are those that incorporate chromosomal changes, such as deletions, inversions, insertions, substitutions, duplications, or translocations of parts of chromosomes. For better understanding, some types of mutations are shown in Fig. 3.17.

A schematic diagram depicts 5 different kinds of mutations in chromosomes. These are duplication, inversion with flip, deletion, translocation with the exchange of chromosomes, and insertion.

Fig. 3.17
Types of chromosomal mutations. Nonlethal aberrations are observed at the first mitosis after irradiation. Duplication: one or more copies of a DNA segment/a region of a chromosome are formed. Inversion: A segment of a chromosome breaks off and reinserts in reverse orientation within the same chromosome. Deletion: A part of a chromosome/one or more nucleotides from a segment of DNA are missing or deleted. Translocation: It involves two chromosomes in which a piece of one chromosome breaks off and rejoins to another chromosome. Insertion: A segment of one chromosome is removed and inserted to another chromosome or the same chromosome


A schematic diagram depicts 5 different kinds of mutations in chromosomes. These are duplication, inversion with flip, deletion, translocation with the exchange of chromosomes, and insertion.


The mutations shown can also lead to other aberrations. Three which should be mentioned are dicentric and ring chromosomes as well as acentric fragments as shown in Fig. 3.18. A dicentric chromosome is created when two chromosomes with two centromeres are fused. In metaphase, they are visible as one chromosome with two centromeres. This aberration will most likely die during mitosis. Acentric fragments are either fragments of a single chromosome or fused parts of different chromosomes containing no centromere. A ring chromosome is a chromosome which has two breaks on the opposing ends and is fused to form a ring. Both aberrations cannot be pulled into a daughter cell and most likely will, together with the encoded genetic information, be lost during mitosis [68]. According to the severity of the chromosomal aberration, the cell will more likely die; in some cases, it can get transformed to a cancer cell or, in case of germ line cell or a cell in early embryogenesis, several genetic disorders can occur [69]. For a more detailed view on this, refer to Chaps. 2 and 7.

A set of chromosomes at the metaphase stage and exposed to gamma radiation depicts abnormal dicentric, fragmented, and ring chromosomes.

Fig. 3.18
Human metaphase cell irradiated with 5 Gy gamma rays. Two dicentric chromosomes, three acentric fragments, and a ring chromosome could be found. From https://www.qst.go.jp/site/nirs-english/1369.html (accessed 05/2022)


A set of chromosomes at the metaphase stage and exposed to gamma radiation depicts abnormal dicentric, fragmented, and ring chromosomes.


The frequency of radiation-induced CAs rises with increasing radiation dose to the cells. Different types of CAs depend on the phase of cell cycle at which the nucleus is exposed to irradiation. Chromosome-type aberrations (Table 3.5) occur when pre-synthetic phase (G1) is exposed to irradiation, while chromatid-type aberrations (Table 3.6) appear if irradiation occurs during post-synthetic phase (G2). In chromosome-type aberrations, more than one break is unable to rejoin at the correct ends that often results in abnormal chromosomes. There is much hidden damage present, some of which is transmitted to future cell generations. In contrast, radiation can induce chromatid aberrations during late S and G2 phases, when sister chromatids are being duplicated and the DNA DSBs may result in chromatid breaks (deletions), interchanges, or triradials. Mostly, sister chromatids or non-sister chromatids of homologous chromosomes are affected by all the breaks and rejoins. The chromosomal aberrations serve as a biological dosimeter—an indicator of radiation exposure. Furthermore, radiation-induced CAs delineate an early marker of late effects, including cell killing and transformation.Table 3.5
Chromosome-type aberrations

Dicentrics

When G1 phase is exposed to irradiation, it causes chromatid breaks in two different chromosomes, which rejoin during S phase and can be seen as dicentric at M phase. Two centromeres in one chromosome appear in dicentrics via breakage-fusion-bridge cycle. These are relatively easy to detect and the main aberration used for biodosimetry

Chromosomal gap

Random achromatic lesions can occur at both the chromatids of a metaphase chromosome, which can be visible as non-stained/lightly stained thinner region. The width of this region is less than the width of chromatid arm

Acentric chromosomal fragments

When single or double breaks occur in the same chromosome arm, either at the end of a chromosome or between centromere and telomere region, it will produce terminal or interstitial acentric fragments, respectively. These acentric chromosomal fragments (without centromere) are lost during anaphase. These are generally associated with dicentric chromosomes

Ring chromosome (centric ring/acentric ring)

Usually, they result from two terminal breaks in both chromosome arms (chromatids), followed by fusion of the broken ends together to form a circular (centric ring) chromosome, leading to the loss of genetic material. Alternatively, the subtelomeric sequences or telomere-telomere fusion with no deletion also results in complete acentric ring chromosomes

Terminal and interstitial deletion (excess acentrics)

A terminal deletion is the loss of the end of a chromosome (telomere), leaving longer acentric fragment than the width of the chromatid. Interstitial deletion occurs when two breaks are induced in interstitial region and the terminal part rejoins the main body of the chromosome, generating double minutes as acentric fragments

Reciprocal translocation

Reciprocal (complete or two-way) translocations involve non-acrocentric chromosomes, and it occurs when two different (nonhomologous) chromosomes have exchanged segments with each other

Marker chromosome

Marker chromosomes are often referred to as mysterious supernumerary piece of chromosomal material. In addition to normal chromosomes, these are small additional structurally abnormal metacentric/centric chromosome fragments whose genetic origin is unknown; however, it can be determined by FISH analysis using specific probes
Table 3.6
Chromatid-type aberrations

Chromatid gaps (achromatic lesions)

Chromatid gap is a non-staining or very lightly stained region (achromatic lesion) of a single chromatid in which there is a minimal misalignment of the chromatid. The width of this region is less than the width of chromatid arm

Isochromatid deletions

The double breaks (often called isochromatid breaks) at the same position on both chromatids are an apparent exception to the definition of chromatid aberrations. They may be induced upon irradiation in the S and G2 phases of the cell cycle
Isochromatid deletions with complete and incomplete sister union (SU): The side-by-side ends of isochromatid breaks usually undergo a cross union to produce U-shaped fragments
Isochromatid deletion without unions (NU: nonunions): Occasionally, the sister union does not occur and such sister nonunions may be in either the proximal (centric) or the distal (acentric) fragments. They are cited as NUp (nonunion proximal) and NUd (nonunion distal), respectively

Terminal and interstitial deletion

Loss of terminal end of one of the chromatids of a chromosome

Symmetric interchanges

Symmetrical chromatid exchanges are equivalents of chromosome-type reciprocal translocation. Exchanges that yield a balanced interchange of genetic material between two identical sister chromatids (i.e., SCE) with no loss of genetic material and no mechanical problems at mitosis

Asymmetric interchanges

Inter-arm interchanges and asymmetrical chromatid exchanges are equivalents of chromosome-type dicentrics. The segments of chromatids are differently joined up, yielding an acentric and dicentric chromatid

Intra-chromatid exchanges/intra-arm interchanges

Chromatid exchanges may occur between non-sister chromatids of paired homologous chromosomes or between sister chromatids of a homologous chromosome. These exchanges may result in symmetrical or asymmetrical interchanged forms such as intra-chromatid exchange with centric ring, inter-chromatid exchange with dicentric, pericentric inversion, and duplication/deletion

Triradials

A three-armed configuration occurs when there is an interaction between one chromosome with an isochromatid deletion and a second having a chromatid deletion


A series of methods and techniques (Fig. 3.19) have been developing to assess stable or unstable type of CAs in order to evaluate the potential of a test compound (chemical/mutagen/radiation exposure). Human peripheral blood lymphocytes offer unique possibilities to study somatic cell division (in vitro) and thus have been utilized for detection of CAs (Box 3.13).

An illustration describes standard cytogenetic techniques such as chromosome banding, karyotyping, and flow cytometry on the left and 4 advanced cytogenetic techniques on the right, which are based on I S H, C G H, P C R, and southern blot.

Fig. 3.19
Techniques to assess constitutional or acquired chromosomal abnormalities using standard banding techniques (left) or advanced molecular cytogenetic techniques (right). Standard cytogenetic techniques are traditionally performed by karyotyping of stained metaphase chromosomes or by flow cytometry. Chromosome banding is used to produce alternating light and dark regions, also referred to as “cytogenetic bands,” along a chromosome with the use of special stains (abbreviations are listed below). Chromosome banding patterns are essential in pairing and ordering all the chromosomes, known as karyotyping. Flow cytometry-based procedures have been developed to assess numerical (ploidy) and structural (telomere length) chromosomal aberrations in mitotic cells largely based on DNA content. To overcome the limitations of the banding analysis, advanced cytogenetic techniques are introduced. In techniques based on ISH, fluorescently labeled “painting” probes are used to localize nucleic acid sequences. FISH identifies chromosomal rearrangements and mapping-specific genes on individual mitotic chromosomes. GISH determines the origin of genomes or chromatins in hybrids. RISH reveals cellular patterns of mRNA expression in cells. CGH-based techniques provide an overview of chromosome ploidy level (gain and loss) throughout the whole genome. CGH with the use of microarrays—aCGH—detects aneuploidies, deletions, duplications, and amplifications based on DNA content. Southern blotting and PCR-based molecular cytogenetic techniques have good potential to detect chromosomal abnormalities from trace amounts of specific regions of DNA/RNA. G-banding Giemsa banding, Q-banding quinacrine fluorescence banding, R-banding reverse banding, C-banding centromere banding, ISH in situ hybridization, FISH fluorescence in situ hybridization, GISH genomic in situ hybridization, RISH RNA in situ hybridization, CGH comparative genomic hybridization, aCGH array comparative genomic hybridization, QF-PCR quantitative fluorescence polymerase chain reaction, qPCR quantitative polymerase chain reaction, MAPH multiplex amplifiable probe hybridization, MLPA multiplex ligation-dependent probe amplification


An illustration describes standard cytogenetic techniques such as chromosome banding, karyotyping, and flow cytometry on the left and 4 advanced cytogenetic techniques on the right, which are based on I S H, C G H, P C R, and southern blot.


Radiation-induced breakage and improper rejoining in pre-replication (G1) chromosomes may lead to chromosome-type aberrations.

Radiation-induced breakage and inappropriate rejoining in post-replication (late S or G2) chromosomes may lead to chromatid-type aberrations.

Since the radiation-induced aberrations in G0 lymphocytes are of the chromosome type, all paired acentric fragments are to be classified as chromosome-type terminal deletions and not isochromatid deletions.

Unstable aberrations like dicentrics, rings, and anaphase bridges are lethal to cells and not passed on to the progeny. Small deletions and stable symmetric translocations are nonlethal and are passed on to the progeny; thus, they may have genetic consequences.


Radiation-induced breakage and improper rejoining in pre-replication (G1) chromosomes may lead to chromosome-type aberrations.


Radiation-induced breakage and inappropriate rejoining in post-replication (late S or G2) chromosomes may lead to chromatid-type aberrations.


Since the radiation-induced aberrations in G0 lymphocytes are of the chromosome type, all paired acentric fragments are to be classified as chromosome-type terminal deletions and not isochromatid deletions.


Unstable aberrations like dicentrics, rings, and anaphase bridges are lethal to cells and not passed on to the progeny. Small deletions and stable symmetric translocations are nonlethal and are passed on to the progeny; thus, they may have genetic consequences.


From the mentioned chromosomal mutations, the translocations are especially dangerous as, in contrast to many other types of chromosomal aberrations, they can be tolerated by the cells. They usually neither cause loss of genetic material nor mitotic cell death and are thus transmitted to the next cell generations. At the same time, translocations are highly oncogenic or affect cell physiology in other ways. Translocations may be simple; reciprocal; i.e., if chromatin fragments are exchanged between two chromosomes; or even complex [70]. Translocations mostly arise due to erroneous DNA end joining by classical NHEJ or mutagenic alternative repair pathways. Although homologous recombination is generally considered a highly precise repair mechanism, recombination between repetitive sequences especially in heterochromatin may also lead to chromatin exchanges [48]. In addition, HR can trigger chromosomal translocations when its intermediates are resolved by crossover between allelic or nonhomologous chromosomes [70]. Although translocations are not associated with extensive losses of the genetic material, they can generate fusion genes (and proteins) with aberrant, often oncogenic, functions. An example could be the reciprocal translocation t(9;22)(q34;q11) between genes BCR and ABL [71], which is responsible for the development of the well-known chronic myeloid leukemia (for terminology and categorization of translocation types, the reader is referred to specialized books on medical genetics or cytogenetics, e.g., Griffiths et al. [70]). In addition to formation of fusion genes, translocations may activate proto-oncogenes by repositioning them along or between the DNA molecules into a close proximity of a strong promoter of some other gene. If the reading frame of the translocated gene is shifted, its function may be lost. However, the gene activity can be changed also epigenetically, if a gene is moved into an incorrect chromatin environment. This is often a cause of the tumor suppressor silencing, after a tumor suppressor is translocated close to a genetically inactive heterochromatin domain. In the context of radiobiology, it is important to emphasize that cell exposures to different radiation types lead to different types of translocations. Cells irradiated with photonic radiation with low LET mostly contain interchromosomal translocations where one chromatin fragment is translocated to another chromosome or two fragments are reciprocally exchanged between two chromosomes. These lesions are usually simple, but the proportion of complex translocations increases with the radiation dose. Cells exposed to a particle high-LET radiation, on the other hand, mostly suffer from complex chromosomal translocations arising as the consequence of extensive chromatin fragmentation by highly localized energy deposition along the particle tracks [72]. For the same reason, high-LET radiation preferential generates intrachromosomal translocations affecting a single chromosome at multiple sites. To explain this phenomenon, it should be emphasized that chromosomes in the interphase cells occur in the form of chromosomal territories with only a limited extent of mutual intermingling along their borders, as explained in Sect. 3.5. Hence, the areas of chromosome territory borders where translocations between the neighboring chromosomes can be formed represent only a small proportion of the nuclear volume along the radiation particle track [53, 54]. With increasing doses and more particles transversing a single nucleus, however, extensive rearrangements of the genome affecting high numbers of chromosomes can be detected (Box 3.14).


Chromosomal translocations are the consequence of illegitimate rejoining of DNA double-strand breaks generated by radiation.

Chromosomal translocations pose a risk of formation of a fusion gene/protein with oncogenic functions; even single translocation may be a sufficient genetic defect to initiate leukemia.

While low-LET radiation generates mostly simple translocations, exposure to high-LET radiation leads to complex genotype rearrangements.

Due to the character of energy deposition, low-LET radiation produces predominantly interchromosomal translocations; higher occurrence of intrachromosomal translocations is then a sign of a high-LET exposure.


Chromosomal translocations are the consequence of illegitimate rejoining of DNA double-strand breaks generated by radiation.


Chromosomal translocations pose a risk of formation of a fusion gene/protein with oncogenic functions; even single translocation may be a sufficient genetic defect to initiate leukemia.


While low-LET radiation generates mostly simple translocations, exposure to high-LET radiation leads to complex genotype rearrangements.


Due to the character of energy deposition, low-LET radiation produces predominantly interchromosomal translocations; higher occurrence of intrachromosomal translocations is then a sign of a high-LET exposure.


Chromosome condensation, the landmark event at the onset of prophase, is the dramatic reorganization of the isolated patches of long thin chromatin strands at the nuclear periphery into compact short chromosomes that can be visualized at metaphase during mitosis or meiosis in eukaryotic cells. Maturation-promoting factor (also called mitosis-promoting factor or M phase-promoting factor, abbreviated MPF), the p34cdc2/cyclin B complex, serves as a master cell cycle regulator for the M-phase transition and chromatin condensation by phosphorylated condensins (Fig. 3.20). MPF activity mainly depends on the cellular concentration of cyclin B, which usually oscillates through cell cycle. During cell division, chromatin condenses and individualizes to discrete chromosomes, which are further segregated by mitotic spindle fibers. Once divided, chromatin decondenses to re-establish its interphase structure component facilitating DNA replication and protein-making processes.

A graphical representation of the relative concentration of cyclin and M P F expression during different phases of the cell cycle, namely, G 1, S, G 2, and M. It points to the M phase and S phase thresholds.

Fig. 3.20
The presence and action of MPF protein in the cell control premature chromosome condensation induction. Cyclin B oscillates through the cell cycle being undetectable during interphase, very low in G1, gradually increasing from S, reaching maximum in G2, and decreasing abruptly at G2/M transition. This corresponds to the MPF activity during cell cycle. MPF maturation/mitosis-promoting factor


A graphical representation of the relative concentration of cyclin and M P F expression during different phases of the cell cycle, namely, G 1, S, G 2, and M. It points to the M phase and S phase thresholds.


Chromosome condensation may also occur prematurely in interphase test cells when they are fused to mitotic cells or chemically using specific phosphatase inhibitors. The most common approach is the use of Chinese hamster ovary (CHO) cells as mitotic inducer cells. Following cell fusion, the MPF present in a mitotic cell interacts with the interphase nucleus causing dissolution of its nuclear membrane and premature chromosome condensation of interphase chromosomes. This phenomenon is known as premature chromosome condensation (PCC). The morphology of prematurely condensed chromosomes (PCCs) depends on the stage of the interphase cell in the cell cycle (i.e., G0, G1, S, and G2) (Fig. 3.21). PCCs in G0-phase cells exhibit single chromatids, highly condensed and distinct. During the G1 phase, G1-PCCs are despiralized single chromatid chromosomes, while chromosomes condensed during the S phase (S-PCCs) have a “pulverized” appearance because of less condensed chromatin at the sites of replication [73]. Condensation during the G2 phase (G2-PCCs) yields distinct elongated double-chromatid chromosomes. Consequently, cell fusion-mediated or chemical induction of PCCs has been proven a powerful cytogenetic tool in radiobiology to study the conversion of radiation-induced DNA lesions into chromosomal aberrations at various cell cycle stages since it enables visualization and quantification of radiation-induced numerical and structural chromosomal alterations directly in interphase cells.

Four sets of chromosomes depict different types of premature chromosome condensations during the G 0, G 1, S, and G 2 phases of the cell cycle. Centromeres are largely absent in the S and G 2 phases.

Fig. 3.21
Premature chromosome condensations (PCCs) at various stages of the cell cycle: darkly stained metaphase chromosomes belong to mitotic CHO cells, whereas the lighter stained to the interphase CHO cells. (a) G0-PCCs, (b) G1-PCCs, (c) S-PCCs (reproduced with permission (CCBY) from Pantelias et al. [73]), (d) G2-PCCs. CHO Chinese hamster ovary


Four sets of chromosomes depict different types of premature chromosome condensations during the G 0, G 1, S, and G 2 phases of the cell cycle. Centromeres are largely absent in the S and G 2 phases.


PCC can be induced either by fusion of human lymphocytes with mitotic cells (fusion-mediated PCC) or with the use of specific chemicals (chemical-induced PCC).


In the case of fusion-mediated PCC, the condensation was at first achieved with the use of fusogenic viruses (such as Sendai virus or its equivalent). However, an important disadvantage of this method is that the fusion efficiency depends on various notable factors [74]. These difficulties were overcome by using cell-fusing chemical agents (e.g., polyethylene glycol—PEG). PEG overcomes these difficulties and can be widely used for radiation cytogenetic studies.


Chemical-induced PCC exploits specific inhibitors for serine/threonine protein phosphatase, which can activate endogenous intracellular MPF, which is much simpler and easier than fusion-induced PCC. Chemicals that can be used for the achievement of drug-induced PCC are okadaic acid, calyculin A, 2-aminopurine, staurosporine, wortmannin, and sodium vanadate. A limitation of this method is that no PCC can be induced in G0 resting-phase cells (Box 3.15).


The appearance of a prematurely condensed interphase chromosome depends on the stage of cell cycle.

PCC can be done in two main ways either by the fusion of human lymphocytes with mitotic cells (fusion-mediated PCC) or by the use of chemicals (chemical-induced PCC).

G1-PCC displays very long single chromatids; PCC in an early, middle, and late S-phase cell shows crushed and pulverized appearance of both single and sister chromatids; G2-PCC demonstrates still long separated sister chromatids with no clearly visible centromere.

The dephosphorylated active form of MPF, a p34cdc2/cyclin B complex, promotes chromosome condensation in meiotic and mitotic cells.

Upon inhibition of protein phosphatase enzymes, cdc25 and cyclin B/cdc2 complex is activated which promotes condensation of chromosomes prematurely.


The appearance of a prematurely condensed interphase chromosome depends on the stage of cell cycle.


PCC can be done in two main ways either by the fusion of human lymphocytes with mitotic cells (fusion-mediated PCC) or by the use of chemicals (chemical-induced PCC).


G1-PCC displays very long single chromatids; PCC in an early, middle, and late S-phase cell shows crushed and pulverized appearance of both single and sister chromatids; G2-PCC demonstrates still long separated sister chromatids with no clearly visible centromere.


The dephosphorylated active form of MPF, a p34cdc2/cyclin B complex, promotes chromosome condensation in meiotic and mitotic cells.


Upon inhibition of protein phosphatase enzymes, cdc25 and cyclin B/cdc2 complex is activated which promotes condensation of chromosomes prematurely.


Because of its unique properties, PCC is used for visualizing and scoring chromosomal damage induced by radiation or other clastogenic agents, measuring the induction yield and repair kinetics of chromosome damage in cells at various cell cycle stages immediately after irradiation. It can also be used for the study of condensation dynamics and conformational changes that occur during the cell cycle. The data obtained using the PCC assay can correlate radiation-induced DNA damage and CAs observable at metaphase [75].


Mitotic cell fusion-induced PCC in human lymphocytes (G0-PCC) allows early detection of cytogenetic damage in interphase, the stage of human lymphocytes in peripheral blood, and is the most suitable technique especially for biodosimetry applications in radiation emergency accidents as well as for triage biodosimetry [76]. A later ring PCC (rPCC) assay is an alternative biodosimetry method to the “gold standard” cytogenetic approach (dicentric analysis in metaphase) for high-dose exposure to radiation and can be applied in a simulated mass casualty accident either after chemical induction of PCC [77] or by means of cell fusion providing a much faster assessment of dose [78].


During the last decade, it has been reported that high-LET radiation induces chromothripsis-like complex chromosomal alterations, resembling the phenomenon of chromothripsis appearing in tumors [79]. The term chromothripsis arises from the Greek dialect (chromo for chromosome and thripsis for shattering into pieces), and it was initially described in 2011 by Stephens et al. [80]. Rather than a progressive accumulation of sequential alterations induced in the genome, chromothripsis is a process where chromosome segments undergo tremendous but localized shattering and random rearrangements in a single catastrophic event. Inaccurate rejoining of the induced chromosome fragments results in a new genomic arrangement and the formation of complex chromosomal aberrations that may trigger carcinogenesis (Fig. 3.22).


The mechanisms responsible for chromothripsis are still under debate. However, studies have shown several situations that could be catastrophic for the cell and result in chromothripsis. One possible mechanism proposed is that DNA damage such as DSBs and chromosomal aberrations may cause aberrant mitosis and formation of MN including one or more chromosomes that may undergo localized shattering and chromothripsis. Chromosome shuttering and chromothripsis may emerge in MN when the main nucleus enters mitosis while DNA is still being replicated within micronuclei. Additionally, PCC induces a mechanical stress in the asynchronous micronucleated cells leading to chromosome shattering [73]. Random genomic rearrangements in micronuclei can then be integrated into the cell’s genome, triggering amplification of oncogenes and cancer development [81]. Other additional mechanisms have also been proposed, such as dicentric chromosome formation, telomere erosion, and abortive apoptosis [82].


Regarding radiation-induced chromothripsis-like chromosomal alterations, it was tested recently whether clustered DNA lesions and chromatin decompaction induced by high-LET irradiation can subsequently evolve in localized chromosome shattering in chromosome domains along the particle tracks. This is a critical risk for chromothripsis to occur, and the results obtained provided experimental evidence that high-LET particle radiation is effective in inducing chromothripsis-like aberrations, which can be used as a fingerprint of high-LET exposure [83]. These discoveries are valuable in the fields of radiation oncology and space radiation protection, since chromothripsis-like aberrations can be responsible for adverse effects and increase the hazard for secondary induced cancer.

A flow diagram depicts a chromosome with a centromere in the middle and telomeres leading to chromosomal shattering and abnormal re-ligation events, or random genomic rearrangements. In the end, the centromere shifts to the left.

Fig. 3.22
Schematic illustration of chromothripsis. It is a phenomenon where one single catastrophic event leads to a massive and localized shattering of one or few chromosomes. Shattered chromosome fragments are not properly rejoined resulting in a new genome configuration and a large number of complicated chromosomal aberrations


A flow diagram depicts a chromosome with a centromere in the middle and telomeres leading to chromosomal shattering and abnormal re-ligation events, or random genomic rearrangements. In the end, the centromere shifts to the left.


This chapter is dedicated to the importance of ionizing radiation-induced foci (IRIF) (Fig. 3.23) in DNA damage measurements. Traditional biomarkers of radiation exposures are chromosomal aberrations and micronuclei. In contrast to quantification of these biomarkers, which emerge due to repair errors in some cells only, IRIF of certain proteins and posttranslational modifications are formed in all cells on all DSB damages, almost immediately after irradiation. Hence, these IRIF can be considered specific biomarkers of DSB lesions [84]. This allows easier and faster victim triage. Moreover, naturally occurring amplification of the DSB damage signal, associated with extensive focal accumulation of γH2AX and numerous repair proteins at DSB sites (for detailed description on DNA repair, see Sect. 3.4), offers the unprecedented sensitivity of radiation dose estimation via the pure counting of IRIF on immunofluorescence microscopy images [84]. The radiation dose absorbed by the cells can be estimated by simple counting of such IRIF or, more automatically, by measuring the integrated intensity of the IRIF signal for high numbers of individual cells by flow cytometry [85]. Under the optimal conditions, especially the time range around 30 min after irradiation, the reported minimal detectable values lie in the range of mGy [86].

Three micrographs with two fluorescent dyes depict the presence of 53 B P 1, gamma H 2 A X, and 53 B P 1 plus gamma H 2 A X, respectively, at 5-micrometer scales.

Fig. 3.23
Radiation-induced DNA damage foci. 53BP1 (left, cyan) and γΗ2ΑΧ (middle, magenta) foci in HeLa cells irradiated with 1.2 Gy alpha particles and spatially fixed at 60 min postirradiation. Colocalization of γΗ2ΑΧ and 53BP1 foci is shown (right). Yellow line indicates the cell nucleus


Three micrographs with two fluorescent dyes depict the presence of 53 B P 1, gamma H 2 A X, and 53 B P 1 plus gamma H 2 A X, respectively, at 5-micrometer scales.


Furthermore, DNA damage induction and repair processes can be studied in individual cells using the IRIF assay. In practice, this is important in situations where individual cells can be differentially affected by irradiation, such as in the cases of a partial-body exposure. The ability to study individual cells is critically important also for radiobiological research as individual cells, even if irradiated homogeneously, appear in different phases of the cell cycle, belong to specific (cancer) cell clones, may be to a various extent affected by the bystander effect, etc.


On the other hand, the biochemical nature of IRIF means that their formation potentially depends on various factors, which may introduce some variability to DSB quantification. It remains a subject of discussion whether all DSBs necessarily require IRIF formation for successful repair. Additionally, some foci may persist at DSB sites even after the break rejoining. A real obstacle could follow from the fact that IRIF occur, to some extent, in nonirradiated cells. However, recent results have proved that the spontaneously forming foci differ in size and topology from the radiation-induced ones. So, staining patterns corresponding, for instance, to replication-stressed or apoptotic cells can be distinguished from IRIF related to DNA repair [87]. Importantly, this phenomenon is more prominent only in cancer cells, which are not relevant for biodosimetry. In any case, “the second γH2AX assay intercomparison exercise” carried out in the framework of the European biodosimetry network (RENEB) confirmed a high fidelity of irradiated victims’ triage (dose categorization, rather than dosimetry) based on IRIF detection of the postradiation modification of histone variant H2AX, called γH2AX [84].


γH2AX is formed by the phosphorylation of histone H2AX at ser139 [57]. This process is mediated by ATM, ATR, and DNA-PK kinases, appears in minutes after DNA breakage, and spreads over ~2 Mbps of DSB-surrounding chromatin. Due to this extent of chromatin modification, γH2AX can be microscopically visualized as compact IRIF at DSB sites of 400–600 nm size as described in Sect. 3.5.


The number of γH2AX foci at a particular time postirradiation corresponds to a dynamic equilibrium between the IRIF formation and disassembly as shown in Fig. 3.24. This is the reason why the maximum γH2AX numbers per cell are detected with a short delay after irradiation and the numbers of counted γH2AX are slightly lower compared to physically detected DNA breaks (PFGE, comet assay) [49].

A line graph with error plots depicts the number of foci versus time after irradiation for 1 and 2 gray units. Both lines exhibit a downward trend. A set of 6 microscopic images compares the foci at one hour and two hours of 1, 1, and 3 G y. 2 G y depicts a large cluster of foci.

Fig. 3.24
DNA repair kinetics. (a) Formation and disassembly of γH2AX foci in human cancer cells irradiated with 1 Gy or 2 Gy X-rays. (b) Representative microscopic images for γH2AX foci 1 h and 2 h after X-ray irradiation. (Reproduced with permission (CCBY) from Mariotti et al. [88])


A line graph with error plots depicts the number of foci versus time after irradiation for 1 and 2 gray units. Both lines exhibit a downward trend. A set of 6 microscopic images compares the foci at one hour and two hours of 1, 1, and 3 G y. 2 G y depicts a large cluster of foci.


For most cell types, the peak number of γH2AX is detected in the time window between 30 min and 1 h postirradiation on average, and some shift to later postirradiation times may appear in cancer cells as they often suffer from DSB repair defects. If the integrated γH2AX signal is measured by flow cytometry, the maximal values are measured later than with focus counting, at about 1 h postirradiation, as the size of γH2AX foci grows longer than their number [49]. After reaching the peak value, the number of γH2AX foci rapidly reduces (Fig. 3.24) and, at 24 h postirradiation, only few DSBs that are repaired only with difficulty persist in cells irradiated with medium doses (in order of Gy) of low-LET radiation. However, a substantial proportion of DSBs may still be detected at this late period of time or even after several days postirradiation in cells exposed to high-LET radiation or high doses of low-LET radiation. From the perspective of biodosimetry, this means that the highest precision of the absorbed dose estimation can be achieved in a few-hour window immediately after irradiation. This requirement can be fulfilled during planned medical care, where, in addition, the monitoring of γH2AX foci formation and disassembly (DSB repair kinetics) may be used to identify patients hypersensitive to radiotherapy or radioresistant tumors. However, in the case of unpredicted accidents with mass screening, IRIF-based biodosimetry must rely on the persistent foci due to the necessary reaction time. This requires suitable mathematical models for the absorbed dose estimation and restricts the method applicability to the acute photon dose range of ~0.5 to ~8.5 Gy and days after exposure (i.e., 1 day after 1 Gy and 14 days after 8.5 Gy) [89]. For military countermeasures, it should also be kept in mind that some chemical warfare agents, such as mustard gas, also generate γH2AX foci. Furthermore, background levels may vary due to non-irradiation-induced IRIF, which are also counted and vary individually, so this assay is best suited for triage rather for accurate dosimetry.


In addition to the analysis of γH2AX IRIF numbers, the spatial distribution of γH2AX foci can be determined by microscopy. This is an important advantage of microscopy over flow cytometry as low-LET and high-LET exposures can be distinguished according to nuclear topology of γH2AX foci [90] as described in Sect. 3.5. On the other hand, flow cytometry offers more room for automation than microscopy and can analyze much higher cell numbers, making it the more suitable method for routine biodosimetry in most circumstances (Box 3.16).


γH2AX IRIF form as the histone H2AX is phosphorylated after DSB induction.

γH2AX IRIF formation starts a few minutes after irradiation and peaks at 30 min–1 h postirradiation.

Especially after high-dose irradiation or irradiation with high-LET particles, persistent γH2AX IRIF are left after repair.

γH2AX IRIF can be used for triage-level biodosimetry by counting foci either in the first hours or persistent foci in microscopic images.


γH2AX IRIF form as the histone H2AX is phosphorylated after DSB induction.


γH2AX IRIF formation starts a few minutes after irradiation and peaks at 30 min–1 h postirradiation.


Especially after high-dose irradiation or irradiation with high-LET particles, persistent γH2AX IRIF are left after repair.


γH2AX IRIF can be used for triage-level biodosimetry by counting foci either in the first hours or persistent foci in microscopic images.


γH2AX attracts numerous proteins with specific signaling and/or repair functions to DSB sites. These proteins, in turn, form IRIF with protein-specific time occurrence and extent of colocalization with γH2AX. Hence, IRIF formed by numerous repair proteins can be used to quantify DSBs and estimate the absorbed dose in the same way as it was described above for γH2AX. Alternatively, repair protein and γH2AX foci can be detected simultaneously to enhance the fidelity of DSB evaluation. Furthermore, the protein composition and structure of IRIF protein complexes (e.g., their specific persistent homology at the nanoscale), and differences of these parameters in specific chromatin domains and after exposure to different types of ionizing radiation, help to understand the mechanisms of DNA repair.


Some proteins like 53BP1 form IRIF morphologically comparable to γH2AX foci. Others, which are required in only a few copies (Ku70 and Ku80), are too tiny and can be visualized only with electron microscopy or super-resolution optical microscopy [91]. Other proteins [such as MRE11, NBS1, or ATM (Fig. 3.25)] create small, but large enough, IRIF to be recognized by standard immunofluorescence microscopy. However, these proteins are, in addition to their IRIF location, also dispersed over the cell nucleus. As IRIF and free aggregates of these proteins may be similar in size, and cannot be discriminated by antibody staining, it is often difficult to reliably distinguish these IRIF from the background [52]. Depending on the function of a particular protein in the repair process, IRIF appear immediately (e.g., MRE11, NBS1, 53BP1) or only later after irradiation (BRCA1, BRCA2, RAD51, etc.). This timing may correspond with repair pathway specificity of a given protein. Some proteins, such as 53BP1 [57], are involved in the regulation of both major DSB repair pathways (NHEJ and HR), while other proteins are selective either for NHEJ or for HR (BRCA1, BRCA2, RAD51).

Two sets of micrographs with 2 B N of X-L-F and 2 hours of 1 gray unit depict, at the top, D A P I, RAD 51, p A T M, and merge and at the bottom, D A P I, RAD 51, gamma H 2 A X, and merge.

Fig. 3.25
DNA repair protein markers forming small foci. 2BN hTert (XLF-deficient) human fibroblasts were analyzed 2 h post-IR with 1 Gy. Cells were stained against DAPI, pATM, and RAD51, or DAPI, γH2AX, and RAD51. RAD51 is present in a subset of pATM and γH2AX foci. Reproduced with permission (CCBY) from Geuting et al. [92]. DAPI 4′,6-diamidino-2-phenylindole used for staining nuclei, XLF XRCC4-like factor


Two sets of micrographs with 2 B N of X-L-F and 2 hours of 1 gray unit depict, at the top, D A P I, RAD 51, p A T M, and merge and at the bottom, D A P I, RAD 51, gamma H 2 A X, and merge.


IRIF of repair pathway nonselective proteins, such as 53BP1, occur in all cells and colocalize with most γH2AX foci [57]. 53BP1 is thus a good DSB marker for biodosimetry, in addition to γH2AX. Moreover, 53BP1 foci have similar size and shape as γH2AX foci so that 53BP1 and γH2AX foci extensively colocalize (Fig. 3.23). This fact improves DSB detection in cells where both types of foci are labeled simultaneously. Co-labeling of γH2AX and 53BP1 foci may be especially useful when cells were exposed to low radiation doses generating only few DSBs or if cancer cells with a strong background signal are analyzed. A significant improvement of DSB number estimation due to γH2AX and 53BP1 co-detection is experienced also in cells exposed to high-LET radiation, where DSBs are extensively clustered and can be thus discriminated only with limitation. However, super-resolution microscopy methods, such as single-molecule localization microscopy (SMLM) or STED microscopy, are necessary for more precise analysis of IRIF foci or even their internal composition and arrangement [48, 57].


It should be noted that not all γH2AX foci necessarily colocalize with 53BP1 (or other repair proteins) at early time periods postirradiation. This includes also the period of 30 min postirradiation when the maximum γH2AX focus numbers are detected. On the other hand, at late time periods after irradiation, 53BP1 foci may persist in cells without being accompanied by γH2AX. These non-colocalizing 53BP1 foci probably label and protect incompletely repaired chromatin [93].


Moreover, as IRIF form also at sites of single-stranded DNA breaks (SSB) or oxidative base damages, co-labeling of γH2AX with suitable markers of these lesions (e.g., XRCC1 or OGG1) [94] can provide information on the complexity of individual DNA damage sites. This information may be correlated to various factors, such as the LET of the incidental radiation or chromatin density and genetic activity at DSB sites [16]. Table 3.7 shows a summary of the IRIF markers mentioned in this section and their occurrence (Box 3.17).Table 3.7
DNA repair proteins and occurrence

Protein/IRIF

Occurrence

γH2AX
53BP1
NBS1
MRE11
Ku70/80
RAD51
Brca1
Brca2

All DSB
All DSB
Part of MRN complex
Part of MRN complex
All DSB
Predominantly HR
Transition between NHEJ and HR
Predominantly HR


Repair protein IRIF, depending on the protein’s role throughout repair, can also be used for biodosimetry.

Repair protein IRIF can be used to understand repair mechanisms and pathways of individual DSB sites.

IRIF can be used to understand the effect of radiation of different LET.


Repair protein IRIF, depending on the protein’s role throughout repair, can also be used for biodosimetry.


Repair protein IRIF can be used to understand repair mechanisms and pathways of individual DSB sites.


IRIF can be used to understand the effect of radiation of different LET.


Exposure to IR induces oxidative damage to cellular molecules such as proteins, lipids, and DNA as a result of oxidative stress (OS), a consequence of the indirect effects of IR (see Chap. 2 and Sect. 3.2), as shown in Fig. 3.26. OS refers to a state of imbalance between oxidants and antioxidants, in favor of oxidants, due to either antioxidant depletion or oxidant accumulation. Oxidants include reactive oxygen (ROS) and nitrogen (RNS) species that comprise free radicals, which are characterized by oneself or more unpaired electrons in the outer shell, and non-radical reactive species. A list of radicals and non-radicals can be found in Table 3.8. Some of these species, e.g., superoxide and hydroxyl radical, are short-lived due to their high reactivity towards other molecules, while others, like hydrogen peroxide, are more stable. Among the ROS, the hydroxyl radical is particularly toxic and involved in the mediation of IR-induced lesions to cell biomolecules. By analogy to OS, nitrosative stress is mentioned when referring to RNS.

A flow diagram depicts the following steps. Plasma membrane damage, mitochondrion disruption with apoptosis, lipid peroxidation, nuclear and D N A damage, and oxidative stress that leads to protein denaturation.

Fig. 3.26
Possible ROS-mediated oxidative stress. Upon exposure to IR, oxidative stress can induce collateral damage, such as lipid peroxidation, protein denaturation, nuclear and DNA damage, mitochondrial damage, and apoptotic death by releasing cytochrome c. Oxidative stress owing to excess ROS generation induces overexpression of antioxidant enzymes in an attempt to control ROS levels. At high levels of oxidative stress, antioxidant defenses are overwhelmed, which leads to inflammatory and cytotoxic responses. (Reproduced with permission from Sanvicens and Marco [95]). NP nanoparticles, ROS reactive oxygen species
Table 3.8
List of free radicals and non-radicals

Free radicals

Non-radicals

Reactive oxygen species (ROS)

Superoxide °O2−

Hydrogen peroxide H2O2

Hydroxyl °OH

Singlet oxygen 1O2

Peroxyl °ROO

Ozone O3

Lipid peroxyl LO°O

Hypochlorous acid HOCl

Lipid peroxide LOOH

Reactive nitrogen species (RNS)

Nitric oxide °NO

Nitrous acid HNO2

Nitrogen dioxide °NO2

Peroxynitrite ONOO−

Dinitrogen trioxide N2O3


A flow diagram depicts the following steps. Plasma membrane damage, mitochondrion disruption with apoptosis, lipid peroxidation, nuclear and D N A damage, and oxidative stress that leads to protein denaturation.


Oxidants are produced from exogenous, such as air pollutants, xenobiotics, and IR, and endogenous sources as normal cellular metabolism by-products. Examples are the mitochondrial electron transport chain (ETC), nicotinamide adenine dinucleotide phosphate (NADPH) oxidase, xanthine oxidase, and peroxidases. Low to moderate ROS levels are crucial in physiological function of cell to avoid oxidative stress involved in aging and several neurodegenerative diseases, diabetes, cancer, atherosclerosis, etc. ROS are also signaling molecules involved in the IR non-targeted effects (see Chap. 2).


OS occurs in pathologic conditions, when the cellular antioxidant defenses are overwhelmed by free radicals and oxidants. Their great oxidative ability leads to oxidative damages to cellular biomolecules (DNA, proteins, and lipids) resulting in multiple damage affecting cell membrane, cellular signaling, and genome integrity. The accepted radiation biology paradigm considered DNA for a long time as the critical IR target and the primary cause for the harmful effects of IR, due to its content of genetic information, with nucleic acid damage being extensively characterized, without consideration that damaged lipids and proteins may also have detrimental effects on cellular function.


Further targets of radiation-generated ROS are lipids, major constituents of the cell membrane, because of their molecular structure containing abundant reactive double bonds [96]. Upon ROS reaction with polyunsaturated fatty acids (PUFA), chain reactions occur, leading to lipid peroxidation (LP) and generation of toxic decomposition products such as malondialdehyde (MDA), 4-hydroxy-2-nonenal (4-HNE), and isoprostanes (IsoPs), which are quantifiable markers of LP reactions. Biological LP consequences include changes in the permeability and fluidity of the membrane lipid bilayer, ion gradient disruption across membrane, and alterations in membrane-associated protein activity [96].


Potential oxidative damage to proteins is multiple, cysteine, methionine, and tyrosine residues. Chemical modifications include oxidation, carbonylation, and nitration and lead to posttranslational modifications inducing conformational changes affecting protein structure and function, i.e., loss of enzyme activity.


While the physical and chemical reactions initiated by radiation occur in less than a millisecond, the resulting biological effects may take hours, days, months, or years to be expressed and may differ among individuals due to varying intrinsic radiosensitivity. In particular, since the oxidative damage extent depends on the antioxidant availability, increased expression of antioxidant defense systems has been linked to decreased radiosensitivity [97].


OS also has a central role within the inflammatory process. ROS such as superoxide can rapidly combine with NO to form other RNS, such as peroxynitrite, and is 3–4 times faster than the dismutation of superoxide by the SOD. The RNS, in turn, induces nitrosative stress, which adds to the pro-inflammatory burden of ROS. Injured cells release chemoattractant molecules, and NO increases vascular permeability and vasodilation that trigger local inflammation. Neutrophils are the first inflammatory cells to arrive at the site of injury, and the increased expression of intercellular adhesion molecule 1 (ICAM-1) and platelet endothelial cell adhesion molecule 1 (PECAM-1) on disrupted endothelial surfaces contributes to neutrophil extravasation. When leukocytes come into contact with collagen fragments and fibronectin, they release pro-inflammatory cytokines like tumor necrosis factor alpha (TNF-α), IL-1, and IL-6 that increase ROS production and lead to even greater local inflammation that can perpetuate inducing chronic radiation injury, which in some cases develop into fibrosis [98] (Box 3.18).


Oxidative stress is characterized by an imbalance between prooxidant molecules and antioxidants.

Oxidative stress participates in the oxidative damage of cellular components.

Antioxidants play a key role in stopping the oxidative chain reactions by scavenging the free radical intermediates.

Excessive generation of ROS, that provokes mitochondrial DNA mutations, impairs the mitochondrial respiratory chain and modifies membrane permeability and mitochondria-associated defense systems.

Several biomarkers of oxidative stress exist and comprise direct ROS measurement, indirect measure of oxidative stress by quantifying oxidation products, and measure of antioxidant defenses.


Oxidative stress is characterized by an imbalance between prooxidant molecules and antioxidants.


Antioxidants play a key role in stopping the oxidative chain reactions by scavenging the free radical intermediates.


Excessive generation of ROS, that provokes mitochondrial DNA mutations, impairs the mitochondrial respiratory chain and modifies membrane permeability and mitochondria-associated defense systems.


Several biomarkers of oxidative stress exist and comprise direct ROS measurement, indirect measure of oxidative stress by quantifying oxidation products, and measure of antioxidant defenses.


In order to cope with ROS and RNS, living organisms have evolved essential antioxidant defense mechanisms (Fig. 3.27). These are classified as enzymatic and nonenzymatic systems or as high-molecular-weight and low-molecular-weight compounds. The first line of antioxidant defenses includes the highly abundant glutathione (GSH), catalase, glutathione peroxidase (GPx), and superoxide dismutase (SOD). GSH acts directly as an oxidant scavenger or indirectly as a cofactor of several enzymes such as the GPx. SOD exists in three isoforms using different metals as cofactors: SOD1, which is predominantly cytoplasmic; SOD2, which is mitochondrial; and SOD3, which is extracellular. SOD1 and SOD3 contain copper (Cu) and zinc (Zn), whereas SOD2 has manganese (Mn) in its active site. They catalyze the dismutation of °O2– to H2O2 afterwards converted to water by catalase, GPx, or peroxiredoxin (Prx). GPx transforms reduced GSH to its oxidized form (GSSG). GSH pool regenerates by de novo synthesis and glutathione reductase using NADPH as a reducing equivalent. GPx is also involved in hydroperoxide detoxification. Prx is involved in hydroperoxides and peroxynitrite detoxification, using thioredoxin (Trx) as a source of reducing equivalents. The most reactive and highly toxic °OH is produced from H2O2 in the presence of reduced transition metal, a reaction known as the Fenton reaction. Apart from GSH, nonenzymatic antioxidants include endogenous compounds which are produced in organism (uric acid, lipoic acid, l-arginine …) and exogenous compounds which are supplemented through the diet, i.e., carotenoids, ascorbic acid (vitamin C), vitamin E and derivatives (tocopherols and tocotrienols), polyphenols (curcumin, resveratrol, quercetin …), and others.

A flow diagram depicts the action of G S H reductase on G S H and G S S G, G S H peroxidase on R O O H, S O D 1 and 2 on O 2, catalase on H 2 O 2, p r x red on R O O H, t r x R 1 and 2 on t r x red and t r x o x.

Fig. 3.27
Antioxidant defense mechanisms


A flow diagram depicts the action of G S H reductase on G S H and G S S G, G S H peroxidase on R O O H, S O D 1 and 2 on O 2, catalase on H 2 O 2, p r x red on R O O H, t r x R 1 and 2 on t r x red and t r x o x.


Glutathione is the major low-molecular-weight thiol in mammals. It plays a key role in cell resistance against oxidative and nitrosative damage by providing reducing equivalents to enzymes involved in the metabolism of ROS, by eliminating potentially toxic oxidation products, and by reducing oxidized or nitrosated protein thiols. In its reduced form (GSH), glutathione is the principal intracellular antioxidant. The conversion of the oxidized form (GSSG) into GSH is done by glutathione reductase (GR) in the presence of NADPH, which is generated by glucose-6-phosphate dehydrogenase in the pentose phosphate pathway (Fig. 3.27). Hence, any damages to these enzymes can compromise GSH functions. The processes of glutathione synthesis, transport, utilization, and metabolism are tightly controlled to maintain intracellular glutathione homeostasis and redox balance. Glutathione is exclusively synthesized in the cytosol and about 85% of it remains there, mainly in the reduced form. The ratio of GSH:GSSG in the cytosol is conservatively estimated at about 10,000:1–50,000:1, and the concentration of the cytosolic GSH is as high as 10 mM, while GSSG in the cytosol is as low as nanomolar concentration [99]. Directly and indirectly, GSH effectively scavenges free radicals and other reactive species (e.g., hydroxyl radical, lipid peroxyl radical, peroxynitrite, and H2O2) through enzymatic reactions, such as those catalyzed by GPxs, glutathione-S-transferases (GST), formaldehyde dehydrogenase, maleylacetoacetate isomerase, and glyoxalase I (Fig. 3.27). GSH also helps to recover other important antioxidants as vitamin C.


OS was shown to promote the activation of redox-sensitive transcription factors such as the nuclear factor erythroid 2-related factor 2 (NRF2) and the nuclear factor kappa B (NF-κB). The NRF2 transcription factor plays a central role in the maintenance of cellular redox homeostasis via the coordinated transcriptional upregulation of numerous antioxidant proteins (Fig. 3.28). These include more than 500 genes that are crucial to metabolize electrophilic attack and protect against OS and inflammatory damage. Kelch-like ECH-associated protein 1 (KEAP1) is a key cytoplasmic repressor of NRF2. KEAP1 interaction with NRF2 leads to NRF2 proteasomal degradation. In the presence of OS or inducers, key “sensor” cysteine thiol groups on KEAP1 are modified, disrupting the degradation process and allowing NRF2 to directly translocate into the nucleus. NRF2 then upregulates the expression of enzymes involved in the synthesis and recycling of GSH, such as the catalytic and modulator subunits of glutamate–cysteine ligase (GCLC and GCLM), GR, GPx, SOD, and several GST. Moreover, several proteins within the redoxin family, such as Trx, TrxRs, Prxs, and sulfiredoxins, are also upregulated by NRF2 [100] as shown in Fig. 3.29. NRF2 stimulates the mitochondrial biogenesis program through activation of nuclear respiratory factor 1 and indirectly prevents/attenuates inflammation, because NRF2 activation results in the expression of previously mentioned antioxidant enzymes, which detoxify ROS, and in turn this reduces the expression of NLRP3 inflammasome and NFKβ (the main regulator of pro-inflammatory response). Moreover, NRF2 upregulates heme oxygenase activity (HO-1) and increases CO production, which in turn reduces NFKβ activity. In response to this, pro-inflammatory cytokine (IL6 and TNFa) production is reduced, and at the same time the production of anti-inflammatory cytokines (such as IL10) increases. As a consequence of these changes, NRF2 facilitates cells to survive oxidative stress and the inflammatory response that aggravates their cytotoxic effects (Fig. 3.29).

 A flowchart depicts that the reduced R O S levels lead to anti-inflammatory effects due to the decreased production of pro-inflammatory factors and increased production of anti-inflammatory cytokines.

Fig. 3.28
NRF2 protection against oxidative stress and excessive inflammatory responses involved in IR injury. NRF2 induces antioxidant response genes, like SOD, CAT, GPX, and GST that enhance ROS elimination. In addition, expression of enzymes such as GR and GS increases GSH cellular content and antioxidant capacity of the cell. Reduction in ROS levels decreases the expression of NFKβ, the main contributor to the inflammatory response. Moreover, NRF2 enhances the expression of HO-1 and its activity in the production of CO that reduces NFKβ activity, pro-inflammatory cytokine secretion (IL-6, TNFα, and IL-1β), and pro-inflammatory enzyme activity (COX-2 and iNOS). ARE antioxidant-responsive element, NRF2 NF-E2-related factor 2, SOD superoxide dismutase, CAT catalase, GPx glutathione peroxidase, GST glutathione S-transferase, GS glutathione synthetase, GR glutathione reductase, GSH glutathione, ROS reactive oxygen species, NFKβ nuclear factor kappa β, IL-6 and 10 interleukin 6 and 10, IL-1β interleukin 1 beta, TNFα tumor necrosis factor alpha, COX-2 cyclooxygenase 2, iNOS inducible nitric oxide synthase, HO-1 heme oxygenase 1

A flow diagram depicts that radiation generates R O S causing oxidative stress that weakens the mitochondrial membrane potential and leads to cytochrome c release and apoptosis.

Fig. 3.29
Mitochondria as the key player in radiation-induced oxidative stress-mediated apoptosis. Various stimuli like radiation or improper functioning of the oxidative phosphorylation induce oxidative stress via ROS production. This causes the mitochondria to dysfunction and subsequently leads to cell death by apoptosis. NAD+ nicotinamide adenine dinucleotide, NADH nicotinamide adenine dinucleotide hydrogen, H+ hydrogen, FAD flavin adenine dinucleotide, FADH2 flavin adenine dinucleotide hydrogen, ATP adenosine triphosphate, ADP adenosine diphosphate, Mn-SOD manganese superoxide dismutase, GPx glutathione peroxidase, H2O2 hydrogen peroxide, CuZn-SOD copper zinc superoxide dismutase, ROS reactive oxygen species, Bcl-2 B-cell lymphoma 2, Bax Bcl2-associated X, APAF1 apoptotic protease-activating factor 1


A flowchart depicts that the reduced R O S levels lead to anti-inflammatory effects due to the decreased production of pro-inflammatory factors and increased production of anti-inflammatory cytokines.


A flow diagram depicts that radiation generates R O S causing oxidative stress that weakens the mitochondrial membrane potential and leads to cytochrome c release and apoptosis.


Mts are double-membrane multifunctional organelles associated with biosynthesis, metabolism, cell survival, signaling of ROS, etc. In the late 1960s, it was found that radiation could significantly modify the structural form of mts and also the mitochondrial DNA. Human mtDNA is a 16,569 base pair (bp) double-strand circular DNA molecule containing 37 genes, encoding 13 polypeptides for the mt electron transport chain, 2 ribosomal RNA, and 22 transfer RNA for mt protein synthesis. Somatic cells have an average of 100–500 mts with 1–15 mtDNA molecules per mitochondrion.


Although nuclear DNA (nDNA) is the main IR target, mts are constantly removing excess ROS created during energy production and mtDNA is much more vulnerable to IR effects than nDNA. mtDNA is generally repaired less efficiently than nDNA [101], although it uses the same repair mechanisms such as BER, MMR, and HR but not NER and classical NHEJ. Furthermore, the histones for better exposure protection are lacking. Together, this leads to a mutation rate which is 10–1000 times higher than nDNA [102]. Both direct IR exposure and irradiated cell-conditioned medium induce mtDNA damage and alter directed protein synthesis. As a consequence, IR exposure can cause the loss of mt membrane potential, leading to mt undergoing either fission, division of one mitochondrion, or fusion, combination of several mitochondria, autophagy (mitophagy), apoptosis, modification in the mtDNA copy number per cell (mtDNAcn), and cause DNA damage and mutations, like point mutations or deletions. A common deletion mutation of 4977 base pair deletion in mtDNA genes coding for subunits of the mitochondrial ATPase, NADH dehydrogenase complex I, and cytochrome c oxidase is known as a marker for oxidative damage [101].


Changes in mtDNAcn or mutations in mtDNA both caused by high intra-mtROS control mt-dependent methylation potential of nDNA by decreasing methyltransferase activity and thus causing global DNA hypomethylation or changes in the expression of specific genes [103]. Global DNA methylation levels depend on human mtDNA variants and are also tissue specific and, therefore, may be connected with the differences in susceptibility to the pathogenic processes resulting from IR exposure and OS in different tissues [103].


OS also appears to target the mitochondrial DNA polymerase-γ activity required for replication and repair of mtDNA, thereby reducing the overall repair capacity. Therefore, subsequent to radiation exposure, mtDNA might be damaged, with an ensuing decrease in respiratory chain activity and decrease of mitochondrial function, giving rise to an increased ROS production. Moreover, mutations in mtDNA could lead to an increase in accessibility of reduced components of the ETCs to O2, which may result in an increase in prooxidant formation. The functional disablement can be weighed by the limitations of the complexes I and III of the mitochondria, reduction of succinate-induced respiratory competence, augmented ROS levels, and increased mitochondrial protein oxidation. The net consequence is persistent metabolic OS that continues to cause de novo oxidative damage to critical biological structures. Such mitochondrial dysfunction can lead to stress signals, which lead to reduced electron transport chain (ETC), and oxidative phosphorylation can cause imbalance in the mitochondrial ROS production, decrease in the mitochondrial membrane potential, and lesser cellular ATP or energy. Although mts are the main producer of ROS, mts themselves can be susceptible to the pathological outcomes once targeted by ROS. By triggering the mitochondrial stress and downstream signaling, the increased levels of free radicals linked to the mtDNA oxidative damage lead to apoptosis.


One of the crucial steps in the process of apoptosis is the permeability transition pore opening (mPTP), followed by drop in the mitochondrial membrane potential. Opening of the pore increases the permeability of the mitochondrial membrane to molecules, leading to mitochondrial swelling and necrosis. NO produced at the basal level (e.g., 5 μM) could S-nitrosylate cyclophilin D (CypD), a critical mPTP regulatory component. This prevents the association of CypD with mPTP that is required for opening the pore and confers a protection to the cell under a stress. On the other hand, NO produced at a high concentration (e.g., 500 μM) could produce peroxynitrite in the presence of large amounts of ROS. Peroxynitrite could oxidize mPTP leading to its opening, which would lead to the opening of mPTP, loss of ATP production, and necrosis. The damaged mitochondria generated excessive ROS like hydrogen peroxide and superoxide anion, which provokes the mitochondrion-driven ROS propagation. ROS themselves accelerate the production of mitochondrial ROS. This process is also called as ROS-instigated ROS release (RIRR) by initiating as inter-mitochondria signaling network [104] (Fig. 3.28). Oxidative insult by radiation to the mt alters the mitochondrial membrane potential and causes the leakage of cytochrome c from the inner membrane compartment, which elicits a sequence of signal transduction progression, the outcome of which is apoptotic cell death. Once the mitochondria are severely stressed, the pro-apoptotic factors like Bax create pores on the mitochondrial membrane, which lets the release of cytochrome c in the cell cytoplasm. It interacts with Apaf-1 to form a complex called apoptosome (Apaf-1, cytochrome c, and ATP). Caspase-9 then gets activated and commences the action of other caspases like caspase-3, -6, and -7. These lead to DNA fragmentation and cell degradation, thereby pushing the cells towards apoptosis. This kind of cell death is known as mitochondrial mediated cell death or intrinsic pathway of apoptosis (Fig. 3.29). However, in this case, apoptosis plays a role in abashing cells that induce excessive ROS.


Biomarkers of OS can be classified as molecules that are modified by interactions with ROS or molecules of the antioxidant system that change in response to increased OS. ROS levels can also be monitored using fluorescent probes of commercial kits, which specifically detect intracellular ROS such as H2O2, NO, or °O2−. However, assays that monitor ROS levels are unlikely to be useful for biomonitoring purposes due to the short half-life of ROS and the fact that the response is not specific to radiation exposure.


S-Glutathionylation is the posttranslational modification of protein cysteine residues by the addition of glutathione. This modification can prevent proteolysis caused by the excessive oxidation of protein cysteine residues under oxidative or nitrosative stress conditions. Measuring S-glutathionylation of the proteins as biomarkers (Fig. 3.30) is hampered by difficulty in accessing the tissue in which these modifications occur. Nevertheless, S-glutathionylation of hemoglobin has been proposed as a biomarker of OS strengthened by finding that it occurs in the circulating erythrocytes in parallel with S-glutathionylation of molecules in the vasculature or myocardium [105].

A flow diagram lists the oxidative products of proteins, polyunsaturated fatty acids, and deoxyribonucleic acid that are created by R N S or R O S.

Fig. 3.30
Main oxidative products of DNA, lipids, and proteins. Oxidative products (listed in gray boxes) are formed depending on the free radicals (RNS/ROS) and the biomolecule target (amino acids, proteins, phospholipids, nucleic acids). These products can be used as oxidative stress biomarkers. RNS reactive nitrogen species, ROS reactive oxygen species


A flow diagram lists the oxidative products of proteins, polyunsaturated fatty acids, and deoxyribonucleic acid that are created by R N S or R O S.


The participation of GSH in antioxidant reactions, either chemically or enzymatically via GPx, results in its own oxidation to GSSG. Decrease in intracellular GSH/GSSG ratio is one of the most used biomarkers of OS. In these conditions, GSSG is preferentially secreted out of the cell, and therefore, blood levels of GSH and GSSG may reflect changes in glutathione status in other less accessible tissues. 6 h after a single dose of irradiation (equivalent to 5 Gy), GSH/GSSG ratio decreases in blood. The decrease in GSH/GSSG is mainly due to an increase in the concentration of GSSG, because GSH levels do not change significantly [106].


Antioxidants protect the body from the harmful effects of free radical damage. Thus, measurement of antioxidant levels in target tissues or biofluids has been widely used to assess the extent of oxidant exposure and, in turn, OS. TAC is the measure of the free radical amount scavenged by a test solution, being used to evaluate the antioxidant capacity of biological samples (tissues or biofluids). The TAC system involves enzymes (SOD, CAT, GPxs, and other enzymes), endogenous antioxidants, and dietary antioxidants (mentioned before), which are generally decreased when OS increases. TAC can be easily measured in cells, tissue lysates, and biological fluids by commercial colorimetric kits and represents a global approach (integrated parameter considered as the cumulative effect of all antioxidants of the biological sample) if no specific antioxidant molecule is to be investigated. One of the critical points is that the results obtained with different methods are not always comparable, depending on the different technologies used for their assessment. Moreover, as mentioned by Dr. Sies (who coined the concept of oxidative stress): “neither the term ‘total’ nor the term ‘capacity’ are applicable to the in vivo assays using an arbitrarily selected oxidant generator assaying a sample removed from its biological context, which is characterized by enzymatic maintenance of steady state” [107]. For that reason, we agree with him “that investigators should measure individually parameters associated with oxidative stress (GSH, urate, ascorbate, tocopherol, etc.) and antioxidant enzymes activities (in tissues samples and lymphocytes (in the case of blood samples) if their want to have an idea of the exposure of the entire organism to oxidative stress” [108].


The “comet assay” and newer techniques [e.g., gas chromatography, high-pressure liquid chromatography (HPLC), immunoassays] can distinguish gross DNA damage produced by IR and damage from oxidation (for a detailed description, see Chap. 7). For low doses of radiation, the total number of induced DNA alterations is probably small when compared with the total number of equivalent alterations from endogenous sources. At DNA level, guanine is the most susceptible base to OS, and its oxidation at the C8 of the imidazole ring of deoxyguanosine generates 7,8-dihydro-8-oxo-2′deoxyguanosine (8-oxodG), which is the most predominant and stable DNA oxidative lesion in the genome (Fig. 3.30). A failure to repair oxidized bases creates a risk of mutation during DNA replication. For example, 8-oxodG mispairs with deoxyadenosine (dA) rather than deoxycytosine (dC) resulting in a C-A point mutation, thus increasing the risk of carcinogenesis. Besides the impact of confounding factors like age, sex, and smoking habits, with the help of correction factors, 8-oxodG levels are good and sensitive biological indicators of OS, which can be quantified in serum or urine samples, using HPLC coupled with mass spectrometry [109]. 8-OxodG can be removed by NER or BER with the action of 8-oxodG DNA glycosylase 1 (OGG1), a base excision DNA repair enzyme that cleaves the N-glycosidic bond between the base and the deoxyribose, generating an apurinic/apyrimidinic site (AP) and triggering the BER mechanism. DNA strand breaks and AP sites are effective substrates to activate DNA damage sensor PARP1. Overactivation of PARP1 is associated with apoptosis-inducing factor (AIF)-mediated and caspase-independent cell death. OGG1 seems to guard genome integrity through lesion repair or cell death depending on the magnitude of guanine oxidation. OGG1 may also be measured as an OS marker.


As previously mentioned, lipid peroxidation products include MDA, 4-HNE, or IsoPs and can be used as oxidative stress biomarkers (Fig. 3.30). The latter are prostaglandin-like molecules formed by the nonenzymatic peroxidation of arachidonic acid (AA). MDA may be formed as a result of enzymatic and free radical peroxidation of PUFAs containing at least three double bonds and is also formed during prostaglandin synthesis. MDA can also react with DNA bases to form deoxyguanosine, deoxyadenosine, and deoxycytidine adducts, and these DNA-MDA adducts have mutagenic effects. Phospholipids containing linoleic acid and AA are considered the main source for 4-HNE production. Many different analytical methods are available for the measurement of MDA, 4-HNE, or IsoPs in biological samples and are reviewed by Tsikas [96].


It has been estimated that proteins scavenge a majority (50–75%) of generated reactive species. To function as biomarkers, protein oxidation products must be stable, accumulate in detectable concentrations, and correlate with OS exposition. Protein carbonylation is an irreversible protein modification, associated with alterations in functional and structural integrity of proteins, contributing to cellular dysfunction and tissue damages. Due to relatively early formation during OS, higher stability in comparison to other oxidation products, and simple analysis methods, protein carbonyls are one of the most OS biomarkers. Protein carbonyls can be easily quantified in plasma, serum, tissue samples, and also saliva by enzyme-linked immunosorbent assay (ELISA) [110].


The reaction between °NO and °O2− forms peroxynitrite, which can nitrate tyrosine residues in proteins. This process is in competition with the enzymatic dismutation of °O2− and the diffusion of °NO across cells and tissues. Peroxynitrite-mediated damage has been implicated in a wide range of disease pathologies, and 3-nitrotyrosine (3-NT) and nitrated proteins have been established as a footprint of nitro/oxidative biomarker of progression and severity in conditions. The measurement of 3-NT can be performed in plasma, serum, as well as tissue samples by special mass spectrometry. Commercially available ELISAs are usually used in clinical studies due to standardization and easy sample preparation. In turn, several limitations have been highlighted in the literature, such as low sensitivity and minor specificity. This and other protein oxidation biomarkers in human diseases are extensively reviewed by Kehm et al. [110]. The advancement of proteomics will allow us to assess changes in proteins (including the assessment of carbonylated, S-glutathionylation, S-nitrated, and/or N-nitrated derivatives) that serve as biomarkers of exposure to IR. An overview of the oxidation products of DNA, lipids, and proteins formed can be found in Fig. 3.30.


The cell cycle is a fundamental process through which the cell grows and accurately duplicates the genetic material before it divides to give rise to two daughter cells. The cell cycle is divided into two phases: interphase in which the cell spends most of its time, followed by mitosis during which the cell divides into two daughter cells. The interphase has three distinct phases. The first phase is the G1 phase in which the cell grows and prepares itself for DNA synthesis. Second is the S phase, when the cell actually duplicate its DNA. The third phase is the G2 phase, where it prepares itself for mitosis. The duration of G1 varies considerably from cell to cell, while S, G2, and mitosis show less variation. Quiescence is a reversible state of a cell in which it does not divide but retains the ability to reenter cell cycle. This state is also called G0 phase.


The transition from one cell cycle phase to another is controlled by a variety of proteins, cyclins, and cyclin-dependent kinases. If the system identifies any inaccuracies, the transition from one phase to the next will be delayed and the cells arrested in the so-called cell cycle checkpoints [111]. Cells, which enter mitosis with unrepaired DNA damage, will most likely fail to divide properly resulting in cell death. In order to provide time for DNA damage repair or, if repair is not the best solution for inducing cell death, e.g. apoptosis, before DNA synthesis (S phase) and in particular mitosis is initiated, radiation induces arrest in checkpoints at the end of the G1 and G2 phases. Since the process that kills the cells after radiation damage is related to cell division, cells in G0 or cells which are differentiated or in senescence and have lost the ability to proliferate are very resistant to radiation [112].


Cell division is a highly regulated progression allowing cells to divide and to generate daughter cells. The regulation is necessary for the recognition and restoration of genetic injury along with the prevention of uninhibited cell division. It is regulated by cyclins and cyclin-dependent kinases (CDKs). CDKs are serine or threonine kinases, which unite with a separate subunit of functional cyclins, which presents domains essential for enzymatic activity. CDKs are known to have a crucial function not only in cell division but also in amending the transcription responses. Hence, the deregulation of CDKs is a characteristic of cancers and utilized for anticancer therapy purposes. On the other hand, cyclins establish the activity of CDKs as their levels keep changing during the cell cycle. Depending on their participation and function during the cell cycle, cyclins are divided into four categories: G1 cyclins, i.e., D cyclins; G1/S cyclin, i.e., cyclin E; S-phase cyclins, i.e., cyclins E and A; and M-phase cyclins, i.e., B cyclins. Researchers have discovered around 20 CDK-associated proteins, which makes the cell cycle a complex process that involves the combination of CDKs (Cdk1, Cdk2, Cdk3, Cdk5, Cdk4, Cdk6, Cdk7, Cdk8, etc.) and cyclins (A1, A2, B1, B2, B3, C, D1, D2, D3, E1, E2, F, etc.) in distinct phases of the cell cycle endowing extra governance to the cell cycle apparatus (Table 3.9 and Fig. 3.31). Cyclins impart the specificity for substrates and normal cell cycle regulation, which includes the subunit binding, localization, activation/deactivation, etc. to the Cdk/cyclin complexes [113].Table 3.9
Cyclins, CDKs, and their function throughout cell cycle

Cell cycle phase

Cyclins

CDKs

Functions

G1

Cyclin D

CDK 4, CDK6

Can act in response to external cues, e.g., growth factors and/or mitogens

G1/S

Cyclins E

CDK2

Control the centrosome duplication

S

Cyclins A and E

CDK2

The main targets are helicases and polymerases

M

Cyclins B

CDK1

Control G2/M checkpoint. The cyclins are produced in S phase but are inactive until the synthesis is entirely completed. Phosphorylate several downstream targets

A cyclic diagram of the cell cycle depicts the action of C D K 4 by 6 and cyclin D in the G 1 phase, C D K 2 and cyclin A in the S phase, C D K 1 and cyclin A in the G 2 phase, and C D K 1 and cyclin B in the M phase.

Fig. 3.31
Overview of cell cycle: functions of different phases, cyclins and CDKS, and CDIs


A cyclic diagram of the cell cycle depicts the action of C D K 4 by 6 and cyclin D in the G 1 phase, C D K 2 and cyclin A in the S phase, C D K 1 and cyclin A in the G 2 phase, and C D K 1 and cyclin B in the M phase.