CDKs have a very limited activity without the presence of a cyclin. To be an active kinase, it should be bound to its cyclin partner and its activity can be further altered by phosphorylation and association of additional proteins like p27. Every CDK/cyclin complex possesses a distinct function that is restricted to a specific cell cycle phase (Table 3.9). Cdk4 and/or Cdk6 are activated by D-type cyclins in the beginning of the G1 phase, and it commences phosphorylation of the retinoblastoma protein (Rb) family (Rb, p107, and p130). This releases the E2F transcription factor and causes the activation and transcription of the E2F-responsive genes that are necessary for the cell cycle progression. The cyclin A and E types are the early E2F-responsive genes. During the later G1 phase, cyclin E binds to Cdk2 to activate it and executes the phosphorylation of Rb (pocket proteins), provoking the further activation of the E2F intervened transcription. This assists in the crossing over of the cell cycle checkpoints at the periphery of the G1/S phase, and to S-phase commencement. Cdk2 unites with cyclin A and aids the progression of the S phase. During the inception of the S phase, A-type cyclins are synthesized, which phosphorylates proteins associated with DNA replication. Going further, at the time of G2/M transition, the activity of Cdk1/cyclin A is necessary for the induction of the prophase. Lastly, Cdk1/cyclin B complexes dynamically contribute to the completion of the mitosis process. Cdk1 activity fluctuates throughout cell cycle succession and is proficient of governing varied cell cycle adaptations (G1/S, S, and G2/M phases) by connecting with diverse cell cycle phase-associated specific cyclins, and several processes like action of CDK-activating kinase (CAK) and inhibitory phosphorylation on CDK. Regulating the cyclin levels and action of CDK inhibitors during the cell cycle assures that CDKs are active in the precise stage of the cell cycle. Cells exploit many processes such as transcriptional control of cyclin genes and breakdown of cyclins; the transcriptional control of the cyclin subunits is one way that ensures appropriate temporal expression of the cyclins and degradation of cyclins, to confine cyclins to the proper cell cycle phase and to keep them at the accurate concentration [114]. Ubiquitin-mediated protein degradation is one of the most crucial regulatory controls that confine the cyclins to the proper cell cycle phase. However, SCF (Skp1, Cullin, and F-box proteins) and APC/C (anaphase-promoting complex or cydosome) are two ubiquitin proteins involved in the degradation of cyclins. During the G1-S-phase transition, SCF controls degrading G1 cyclins (cyclin D), while APC/C degrades the cyclins of the S phase and mitosis, thus advancing the exit from mitosis. To control the CDK activity, the regulation of cyclin levels is not the only mechanism. Other mechanisms like activation and inhibition of phosphorylation actions on the CDK subunit and existence of inhibitors are critical in controlling cyclin-CDK activity [114].


CDK inhibitors are a family of proteins that can bind directly to the cyclin-CDK complex and hinder its activity. In the transition of the G1-S phases, these proteins play a very crucial role. CKIs implicated in controlling the S phase and mitotic CKIs are indispensable to avoid early commencement of the S- and M-phase CDKs. However, in human cancers, genes coding these CKIs are often mutated leading to aberrant cell cycle regulation. During normal or extreme conditions (DNA damage, telomere dysfunction, and stress), the functions and activities of the CDK/cyclin complexes are governed and controlled by two families of CKIs. The INK4 family comprises the p16INK4a, p15INK4b, p18INK4c, and p19INK4d which can specifically bind to Cdk4 and Cdk6 and hinder the activity of the D-type cyclin. The other Cip/Kip family (p21Cip1/Waf1/Sdi1, p27Kip1, p57Kip2) obstructs Cdk2/cyclin E, Cdk2/cyclin A, Cdk1/cyclin A, as well as Cdk1/cyclin B activity. The p21 protein hinders the formation of cyclin/CDK protein complexes that are required for the progression from the G1 phase to the S phase of the cell cycle (Box 3.19).


Irradiated cells display a complex set of responses that can include either progression or arrest of the cell cycle.

Every phase of the cell cycle has a very specific set of cyclins and cyclin-dependent kinases to perform functions associated with that particular cell cycle phase.


Irradiated cells display a complex set of responses that can include either progression or arrest of the cell cycle.


Every phase of the cell cycle has a very specific set of cyclins and cyclin-dependent kinases to perform functions associated with that particular cell cycle phase.


To study the variation of radiosensitivity with position in the cell cycle, it is necessary to synchronize the cells to get a population of cells that are all in the same cell cycle phase.


For cells in culture, there are three main techniques.1.
In fluorescence-activated cell sorting (FACS), a flow cytometer is used to sort cells based on fluorescence from a DNA-binding dye, such as Hoechst 33342, which can be used for live cells.
 2.
Chemically induced cell cycle arrest collects over time all the cells at a cell cycle checkpoint. When the drug is removed, the cells will go through the cell cycle synchronously for some time before they become more and more asynchronous. The most used drug is hydroxyl urea, which arrests cells at the border between G1 and S. The advantage of this method is that it can also be used in vivo.
 3.
Mitotic selection was introduced by Terasima and Tolmach [115] and is the most used synchronization method in cell culture in vitro. As cells enter mitosis, they round up and become less attached to the flask bottom. By then shaking the flask, the mitotic cells will detach and can be collected with the medium. The cells can then be irradiated at different time points as they go through cell cycle.


In fluorescence-activated cell sorting (FACS), a flow cytometer is used to sort cells based on fluorescence from a DNA-binding dye, such as Hoechst 33342, which can be used for live cells.


Chemically induced cell cycle arrest collects over time all the cells at a cell cycle checkpoint. When the drug is removed, the cells will go through the cell cycle synchronously for some time before they become more and more asynchronous. The most used drug is hydroxyl urea, which arrests cells at the border between G1 and S. The advantage of this method is that it can also be used in vivo.


Mitotic selection was introduced by Terasima and Tolmach [115] and is the most used synchronization method in cell culture in vitro. As cells enter mitosis, they round up and become less attached to the flask bottom. By then shaking the flask, the mitotic cells will detach and can be collected with the medium. The cells can then be irradiated at different time points as they go through cell cycle.


The first age-response curve by Terasima and Tolmach [115] using 3 Gy irradiation of HeLa cells is shown in Fig. 3.32, left panel, together with a curve showing the fraction of labeled cells after pulsed incorporation of [3H]-thymidine during S phase. Cell survival was measured as the ability to form a macroscopic colony. The data indicate four times higher survival if the dose is delivered during early G1 compared to at the start of S phase. Furthermore, there is an increase in cell survival with age during S phase; that is, the radioresistance increases as more and more of the DNA is synthesized. HeLa cells are HPV infected and do not have functional p53, which normally would give the cells time for repair before entering S phase. The cells irradiated early in G1 will have time for repair, which is reflected in a high survival, while the cells in late G1 are more sensitive, because they may enter S phase with unrepaired DNA damages. Cell lines with short G1 are sensitive throughout G1. Terasima and Tolmach also irradiated the synchronized cells with various radiation doses and thereby recorded complete dose-response curves for HeLa cells irradiated in different phases of the cell cycle (see Fig. 3.32, right panel). These curves confirmed the variation in radiosensitivity through the cell cycle, as was demonstrated by the age-response curves. In addition, they also showed that cells irradiated while in mitosis are far more radiosensitive than cells irradiated in any part of interphase.

2 line graphs. Graph 1 plots the relative number of cells and percentage of mitoses versus the hour after collection. Graph 2 plots the fraction surviving versus dose for random population, 0, 5, 14, and 19 hours.

Fig. 3.32
Age-response of cells after radiation. Left: Age-response curves for HeLa-S3 cells (open circles: synchronized cells, triangles: asynchronous cells) irradiated with 3 Gy X-rays (= 300 rad) at different time points after selection in mitosis and the fraction of cells with incorporated [3H]-thymidine in DNA after a 20-min pulse (black circles, right y-axis). Right: Dose-response curves for HeLa-S3 cells synchronized by mitotic selection and X-irradiated at different times after selection. 0 h: mitosis, 5 h: early G1 phase, 14 h: S phase, 19 h: late S/G2 phase. [Reproduced with permission from Terasima and Tolmach [115]]


2 line graphs. Graph 1 plots the relative number of cells and percentage of mitoses versus the hour after collection. Graph 2 plots the fraction surviving versus dose for random population, 0, 5, 14, and 19 hours.


Measurements of the radiosensitivity of cells in G2 are technically difficult, and it has become customary to suppose that cells are radiosensitive if irradiated in G2. However, the radiosensitivity of cells in G2 has been shown to be dose dependent to a quite different degree than in any other phase of the cell cycle. Cells are hyper-radiosensitive for small radiation doses because the mechanism for early radiation-induced G2 arrest by ATM is not activated by radiation doses in the range below about 0.3–0.5 Gy (Box 3.20).


Cells increase radioresistance throughout S phase.

Cell radiosensitivity is highest during mitosis.


Telomeres are nucleoprotein structures located at the end of each linear chromosome in the cell nucleus. They are composed by tandem repeats of the G-rich hexanucleotide TTAGGG and are typically 10–15 kb long [116]. These structures are organized into heterochromatin domains, and they play a significant role in maintaining genome stability. There are at least two very important functions of telomeres in eukaryotes. The first one is the protection of the linear DNA molecules from the DNA repair mechanisms, which may recognize these sites as double-strand breaks. Secondly, they define the maximum number of cell cycles that a cell may undergo [116]. At each cell cycle division, telomeres shorten by 50–200 bp due to the DNA end-replication problem [117]. This problem results from the inefficient copying of the last base pairs of the linear DNA molecule by DNA polymerase. After several cell divisions, the length of telomeres reaches a critical threshold, which means that the cell can no longer divide. The cell has then reached its Hayflick limit, and it proceeds to senescence. Telomere shortening is thus a very-well-known hallmark of cellular senescence and aging. A good example of the telomere shortening is the deficiency of the adaptive immune system in older individuals caused by T cells reaching their Hayflick limit [118].


The telomere attrition can be opposed by an RNA-dependent DNA polymerase known as telomerase. This enzyme can elongate the telomeres by adding 5′-TTAGGG-3′ repeats to the chromosomes 3′ terminal ends. Telomerase is connected with cells’ immortality; thus, it is present in germline and malignant cells. There is only little or no telomerase in most somatic cells [118]. This information is summarized in Fig. 3.33. An inverse correlation between the telomere length and the radiation-induced cytogenetic damage was found for lymphocytes, fibroblasts, epithelial cells, and many cancer cell lines. It was shown that telomere shortening leads to chromosome fusion, chromosome bridges, or higher frequencies of micronuclei. Thus, telomere shortening is closely linked to the cell radiosensitivity. Therefore, targeting the telomeres could be a very good radiosensitizing method in our fight with cancer during radiotherapy [116].

An illustration depicts the varying length of telomeres in the chromosome of an adult cell with a short telomere after multiple replications and of an immortal cell with telomerase and a telomere at senescence.

Fig. 3.33
Telomeres, their shortening, the senescence state, and immortal cells. An adult cell chromosome with telomeres and the enzyme telomerase, which plays a crucial role in telomere end lengthening (left). Telomere characteristics in an adult cell’s chromosome, after multiple replications, at cell senescence, and when the cell is immortal (left to right, blue box). (Adapted with permission from Aunan et al. [119])


An illustration depicts the varying length of telomeres in the chromosome of an adult cell with a short telomere after multiple replications and of an immortal cell with telomerase and a telomere at senescence.


As described in Sect. 3.7, cellular senescence is a cell state triggered by extrinsic (cellular stressors) and intrinsic (physiological processes) factors. It is characterized by a prolonged and generally irreversible cell cycle arrest, associated with secretory features, macromolecular damage, and altered metabolism, with its function to remove potentially harmful cells from the proliferative pool [120]. Senescent cells are detected at any life stage from embryogenesis (contributes to tissue development) to adulthood (to prevent the proliferation of damaged cells). Yet, senescent cells can also potentiate various aspects of tumorigenesis, including proliferation, metastasis, and immunosuppression by secreting a collection of pro-inflammatory factors collectively termed as senescence-associated secretory phenotype (SASP) [121]. It is important to clarify that senescence is a distinct form of cell cycle arrest and distinct from quiescence, where cells can reenter the cell cycle when favorable growth conditions are restored; terminal differentiation, where cells exhibit functional and morphological changes resulting in loss of original cellular identity; and cell death, where cells are being eliminated and are thus nonfunctional. The existence of multiple senescence programs and the nonspecificity of current senescence markers make it difficult to fully unveil the complex mechanism behind senescence (current understanding presented in Fig. 3.34). It is therefore recommended to apply a multi-marker approach when investigating cellular senescence [120]. Yet, it is currently accepted that two main signaling pathways initiate and maintain the cell cycle arrest: p53–p21–retinoblastoma protein (RB) and p16INK4A–RB. As a consequence, depending on the senescence program, senescent cells express a multitude of hallmarks such as morphological alterations, senescence-associated beta-galactosidase (SA-β-gal), and SASP among others [122].

A flow diagram depicts the conversion of a healthy cell into a senescent cell by senescence inducers and senescence signals. Senescent cells cause S A S P which leads to senescence programs.

Fig. 3.34
Overview of cellular senescence processes. ROS reactive oxygen species, ATM ataxia-telangiectasia mutated, ATR ATM and Rad3-related protein, Cdk2/4/6 cyclin-dependent kinase 2/4/6, RB retinoblastoma tumor suppressor gene, SASP senescence-associated secretory phenotype, SA-β-gal senescence-associated beta-galactosidase


A flow diagram depicts the conversion of a healthy cell into a senescent cell by senescence inducers and senescence signals. Senescent cells cause S A S P which leads to senescence programs.


Senescence in developmental processes, i.e., in embryogenesis and organogenesis, is induced by paracrine signaling and is mediated by the expression of the cell cycle inhibitor p21. Although SA-β-gal is highly expressed, developmental senescence is not associated with DNA damage, does not secrete the typical range of SASP cytokines, and is independent of p53 and p16INK4a. Senescence in wound healing prevents excessive fibrosis by secreting PDGFA-enriched SASP to stimulate appropriate skin repair. Senescence causes, or at least contributes to, organismal aging through the shortening of telomeres followed by the induction of p16INK4a and resulting in an accumulation of senescent cells over time. Studies by Baker et al. [123], first in BubR1-mutant mice (Cdkn2ap16 knockout mice) and then later in naturally aged mice, demonstrated that in the absence of p16INK4a, it is possible to inhibit the production of senescent cells and improve health span [123]. Also, SASP triggers multiple intercellular communication paths that also promote aging. Finally, the elimination of senescent cells improved several age-associated conditions. Senescence in cancer has shown a dual role as tumor suppressor and tumor promoter. Senescence is a key mechanism of tumor suppression via the inhibition of proliferation of cancer cells or by stimulating immune surveillance. Yet, cells induced to senescence by oncogenes or chemotherapy exhibit stemlike properties that promote cancer. Several stressors can induce cellular senescence and radiation in one of them. Thus, IR may cause cell cycle arrest resulting in a prematurely induced senescence phenotype (including SA-β-gal, p16INK4a, p21, and SASP), which is p53 dependent [121]. Unfortunately, the accumulation of these senescent cells can have a negative impact by promoting tumorigenesis. Thus, eliminating senescent cells from tumors and surrounding healthy tissues may be a successful and beneficial adjuvant strategy (Box 3.21).


Telomeres are part the ending parts of chromosomes, which protect the genome integrity

Telomeres shorten in each cell division by 50–200 bp due to the DNA end-replication problem.

Telomere shortening is closely linked to cellular radiosensitivity.

After several cell divisions, the length of telomeres reaches a critical threshold, the Hayflick limit, and the cell proceeds to senescence.

Senescence is sometimes addressed as a type of cell death. A cell in senescence cannot proliferate anymore, it lives only metabolically.

Cellular senescence is characterized by a prolonged and generally irreversible cell cycle arrest, and it functions as a process to remove potentially harmful cells from the proliferative cell pool.

Senescence is a key mechanism of tumor suppression via the inhibition of proliferation of cancer cells or by stimulating immune surveillance in cancers treated with radiotherapy.


Telomeres are part the ending parts of chromosomes, which protect the genome integrity


Telomeres shorten in each cell division by 50–200 bp due to the DNA end-replication problem.


After several cell divisions, the length of telomeres reaches a critical threshold, the Hayflick limit, and the cell proceeds to senescence.


Senescence is sometimes addressed as a type of cell death. A cell in senescence cannot proliferate anymore, it lives only metabolically.


Cellular senescence is characterized by a prolonged and generally irreversible cell cycle arrest, and it functions as a process to remove potentially harmful cells from the proliferative cell pool.


Senescence is a key mechanism of tumor suppression via the inhibition of proliferation of cancer cells or by stimulating immune surveillance in cancers treated with radiotherapy.


In response to IR, multiple, molecularly distinct forms of cell death may be initiated. Although the decision points of their initiation are not completely clear, it is known that the level of the DNA damage but also the individual signaling status of different cell death pathways in different cell types, e.g., hematological vs. epithelial cells, influence the decision regarding the cell death route.


The cellular factors that influence include cell type, position in cell cycle when irradiated, DNA repair capacity, as well as functionality of TP53 and similar DNA-damaging sensors [124]. The dose and radiation quality also contribute to the cellular IR response to cell death, and in the tissue, the oxygen levels may impact the cell death route taken [124]. In this section, an overview of four cell death mechanisms are given: (I) mitotic cell death/mitotic catastrophe, (II) apoptosis, (III) necrosis, and (IV) autophagy (Fig. 3.35), some of which are also interconnected in the cell. Furthermore, the underlying molecular mechanisms and importance of these forms of cell death following IR are also described alongside methods of assessment.

A flow diagram depicts the factors influencing the cell death response. Cell fate is divided into damage repair and cell survival and cell death. The principle cell death pathways are mitotic cell death, apoptosis, necrosis, and autophagy.

Fig. 3.35
Overview of cell death and cell death-protective mechanisms in response to radiation. Radiation-induced cell death is influenced by different factors, such as radiation factors, cell intrinsic factors, and cellular microenvironment factors (left). Cell death pathways are listed to the right. The mechanisms and importance of these principal cell death forms are described in detail in the text


A flow diagram depicts the factors influencing the cell death response. Cell fate is divided into damage repair and cell survival and cell death. The principle cell death pathways are mitotic cell death, apoptosis, necrosis, and autophagy.


Mitotic catastrophe (MC) is an important type of IR-induced cell death mechanism, which is triggered when cells enter into the mitotic phase without appropriately completing the S and G2 cell cycle phases [125]. Hence, MC controls cells that are often incapable of successfully completing mitosis. MC works by activating mitotic arrest, and later it may lead to a controlled or a regulated cell death mechanism or senescence. Therefore, MC is a controled cell death that usually follows the intrinsic apoptotic pathway route [124] (Fig. 3.37). MC is also promoted when the proteins that regulate the G2 phase like the p21CDKN1A, checkpoint kinases 1 and 2 (CHK1/2), ataxia-telangiectasia mutated (ATM) and ataxia-telangiectasia, and Rad3-related protein (ATR) are inhibited. MC basically commences with the irregular condensation of the chromatin around the nucleoli, which looks similar to early chromosome condensation. Cells may die in the same cell cycle or in the successive cell cycle progression or division after IR. The anomalous mitosis in such cases leads to unusual segregation of the chromosomes and cell division. As a consequence, this causes formation of giant cells which exhibit the uncharacteristic nuclear morphology and numerous micronuclei and nuclei. Also, it is noteworthy that MC induced by IR is accompanied with excess duplication of chromosomes and hyper-amplification, which results in a mitosis that is multipolar and later development of micronuclei. DNA damage and flaws in the DNA repair processes lead to centrosome hyper-amplification. Cyclin-dependent kinase 2 (CDK2) and cyclin A or E initiate the amplification of the centrosomes at the boundary of G1/S phase. This is often observed in cells that lack a functional TP53; however, in cells with a functional TP53 and p21CDKN1A, which is known as an inhibitor of CDK2, cellular senescence is promoted.


The outcome of MC in the form of cell death can be elicited in the mitotic phase or in the successive interphase. Some cells activate apoptotic pathways in the metaphase that results in delayed apoptosis, i.e., it can take up to 6 days after IR. Cells that get away with the mitotic arrest of the mitotic cell death are frequently observed to have an unsuccessful cytokinesis consequentially exhibiting tetraploid anomalous nuclei developing into giant cells. Giant cells that possess a functional TP53 will eventually undergo apoptosis following the mitochondrial pathway of apoptosis in the subsequent G1 phase. However, cells with mutant TP53 or deficient TP53 function go on with a few number of cell cycles and attain a growing amount of chromosomal anomalies before they finally succumb to either delayed apoptosis or necrotic form of cell death [125]. As the cells that undergo MC are usually the ones who have lost the potential to carry out any further replication, MC is frequently referred to as a genuine type of cell death. One of the most common properties exhibited by cancer cells is that of defects in cell cycle checkpoints. This lets the cells enduring IR-induced damage to hastily inscribe in the mitotic process even with the misrepaired DNA that eventually leads to MC. More than a few cell division attempts can take place before adequate genetic injuries mount up to activate mitotic death, emphasizing why solid tumors frequently display deferred reactions to IR [124]. MC is triggered after IR exhibits diverse mechanisms of action (Table 3.10) [126].Table 3.10
Examples of IR-induced MC in different tumor cell lines

Inducer of MC

Cell line

Features/signaling components of MC

Ionizing radiation

HeLa (cervical adenocarcinoma)

Increased levels of cyclin B

U2OS (osteosarcoma)

Checkpoint adaptation

HT0180 (fibrosarcoma)

Micronucleation

MOLT4 (leukemia)

Checkpoint adaptation


During MC, the mitotic damage is recognized and guides the cell into one of the three potential antiproliferative fates (Fig. 3.36). In one of them, when cyclin B levels are elevated, the malfunctioning mitotic cells recruit the cell death machinery and die during mitosis. Another cell death pathway that cells can take is by mitotic slippage. Here, cells go out from mitosis and cell death is triggered in the next G1 cell cycle transition. Lastly, cells with a MC character can also undergo senescence after exiting mitosis.

A flow diagram depicts that the D N A damage response leads to interphase cell death that leads to apoptosis and necrosis and mitotic cell death that is further divided into caspase-independent and caspase-dependent, leading to mitotic catastrophe.

Fig. 3.36
Cell death pathways operative in mitotic catastrophe. Different signaling events triggered in response to a nonfunctional mitosis are shown. Upon DNA damage, cells which lack functional p53 can go out from mitosis without commencing cytokines or initiate cell death even in mitosis. Apoptosis and necrosis signaling in the context of mitotic catastrophe are depicted


A flow diagram depicts that the D N A damage response leads to interphase cell death that leads to apoptosis and necrosis and mitotic cell death that is further divided into caspase-independent and caspase-dependent, leading to mitotic catastrophe.


MC may not at all time be accompanied by mitotic arrest. Nevertheless, the mechanism of action that dictates cell fate of subsequent MC continues to remain unclear [127]. When mitotic arrest is extended, the amount of cyclin B is decreased albeit the spindle assembly checkpoint (SAC) is functional. As a result, if cyclin B levels drop below the verge that determines mitotic exit, slippage occurs (Box 3.22).


IR-induced cell death depends on radiation quality, dose as well as cell type, cell cycle position, and functionality in DNA damage signaling.

Mitotic catastrophe is one of the principal forms of IR-induced cell death that results from early/untimely entry into mitosis, even before the fulfillment of S and G2 phases of the cell cycle.

The characteristic features of IR-induced mitotic catastrophe are altered nuclear morphology, micronucleation, and formation of multinucleated cells.


IR-induced cell death depends on radiation quality, dose as well as cell type, cell cycle position, and functionality in DNA damage signaling.


Mitotic catastrophe is one of the principal forms of IR-induced cell death that results from early/untimely entry into mitosis, even before the fulfillment of S and G2 phases of the cell cycle.


The characteristic features of IR-induced mitotic catastrophe are altered nuclear morphology, micronucleation, and formation of multinucleated cells.


Apoptosis (originally from Greek language translated “falling off”) is also known as “cellular suicide.” It is a cell death process which may be executed under normal physiology, e.g., organism development, but also in the context of disease. Apoptosis is a highly controlled pathway with distinct molecular features. Thus, some of the rapidly proliferating cells undergo apoptosis, which is an essential part of neurogenesis and tissue development in humans as well as in other mammalians. During apoptosis, cells are disposed in a complex but well-ordered fashion which involves energy-requiring molecularly defined effector mechanisms [128]. To simplify, apoptosis allows the cells to self-destruct with limited tissue damage when they are exposed to different triggers/signals which can be endogenous, e.g., formed DNA damages, telomere shortening, or encountered from the outside of the cell, e.g., cytotoxic or DNA-damaging agents, IR exposure, loss of growth factors, cytokine or glucocorticoid hormone level alterations, or hypoxia [128].


Apoptosis results in the production of apoptotic bodies, which are cell fragments, e.g., collapsed cytoskeleton, disassembled nuclear envelope, and fragments of nuclear DNA. An apoptotic cell is also marked by certain “find-me” and “eat-me” signals at the cell surface, which allow the dying cell to be recognized and rapidly engulfed by different macrophage subtypes in the near or distant tissue, thereby avoiding inflammation. A well-recognized potential “eat-me” signal is the expression of phosphatidylserine (PS) on the outer side of plasma membrane, which in turn is being used for assessing early apoptotic cells [129].


In the 1990s, studies which resulted in authors being awarded a Nobel Prize revealed that core machinery components of some apoptotic pathways are highly conserved from nematodes to humans [130]. Subsequently, research on the molecular mechanisms regulating apoptosis has established two major routes of this cell death type, namely intrinsic and extrinsic apoptosis, respectively (Fig. 3.37).

A flow diagram depicts the intrinsic and 2 extrinsic pathways leading to apoptotic membrane blebbing, the formation of apoptotic membrane protrusion, and cell fragmentation via different steps.

Fig. 3.37
The intrinsic and extrinsic route to apoptosis. Intrinsic stress signals (e.g., DNA damage, hypoxia, metabolic stress) or lethal stimuli (e.g., IR exposure) can induce intrinsic mitochondrial apoptosis (middle). Cleaved or truncated Bid (tBid) can also connect the extrinsic pathway to the intrinsic route. In the extrinsic pathway, ligands for death receptors (left) can trigger caspase activation, but the pathway can also be activated when some dependence receptors are inactivated (right). Abbreviations: FasL Fas ligand, TRAIL TNF-related apoptosis-inducing ligand, TNF tumor necrosis factor, Fas Fas cell surface death receptor, TRAILR TNF-related apoptosis-inducing ligand receptor, TNFR tumor necrosis factor receptor, TRADD TNFR1-associated death domain protein, FADD Fas-associated protein with death domain, caspase cysteine-aspartic proteases, BID BH3-interacting domain death agonist, tBID truncated BID, Bcl-2 B-cell lymphoma 2 (an apoptotic inhibitor), BCL2L1 Bcl-2-like 1, MOMP mitochondrial outer membrane permeabilization, BH3 Bcl-2 homology 3, DIABLO direct inhibitor of apoptosis-binding protein with low pI, APAF-1 apoptotic peptidase-activating factor 1, Bax Bcl2-associated X (an apoptotic regulator), Bak Bcl-2 homologous antagonist/killer, XIAP X-linked inhibitor of apoptosis protein, SMAC second mitochondria-derived activator of caspase, UNC5B Unc-5 netrin receptor B


A flow diagram depicts the intrinsic and 2 extrinsic pathways leading to apoptotic membrane blebbing, the formation of apoptotic membrane protrusion, and cell fragmentation via different steps.


Multiple perturbations may trigger intrinsic apoptotic cell death, e.g., growth factor withdrawal, cytokine alterations, endoplasmic reticulum stress, replication stress, formation of reactive oxygen species (ROS), microtubular alterations or mitotic defects, and IR-induced DNA damage. In the context of DNA damage, mitochondrial release of apoptogenic proteins is central. This commences in part via mitochondrial outer membrane permeabilization (MOMP) that allows cytochrome C and other proteins to be released to cytosol. Once there, cytochrome forms the apoptosome complex together with apoptotic peptidase-activating factor 1 (APAF-1), where pro-caspase-9 is cleaved to active caspase-9 (CASP9). Subsequently, CASP9 cleaves the effector caspases (e.g., caspase-3, caspase-6, and caspase-7), which then causes degradation of cell signaling and structural proteins resulting in an apoptotic morphology. The BCL-2 proteins are regulators of MOMP. These can either promote, e.g., BCL-2-associated X apoptosis regulator (BAX) or BCL-2 antagonist/killer (BAK) or block MOMP, e.g., BCL-2 or BCL-XL members [131]. Another set of BCL-2 members, which only have a BH3 domain, can also promote MOMP, but they act via alleviation of BCL-2 or BCL-XL function or via promotion of BAX/BAK activity. Examples thereof are BCL2-associated agonist of cell death (BAD), BH3-interacting domain death agonist (BID), BCL2-interacting mediator of cell death (BIM), NOXA, and TP53-upregulated modulator of apoptosis (PUMA).


In DNA damage-induced apoptosis, TP53 and BAX/BAK proteins are important. The BCL-2 family members also sense other cellular clues to elicit intrinsic apoptosis including alterations in growth factor receptor/PI3K signaling or microtubule disruption, both of which may have impact in the context of IR-induced cell death. In addition, the mitogen-activated protein kinases 8 and 9 (MAPK8 and MAPK9), more commonly referred to as c-jun N-terminal kinase 1/2 (JNK1/JNK2), are known to regulate the BCL-2 rheostat by phosphorylation of BCL-2 and BAD, via induction of NOXA and PUMA by TP53 transcriptional regulation as well as by association of BIM to microtubuli [132].


The extrinsic pathway starts by the activation of membrane receptors, so-called death receptors (DRs), e.g., FAS/CD95 cell surface death receptor and TNF receptor superfamily member 1A (TNFRSF1A)/TNFR1, and is driven by initiator caspases, e.g., caspase-8 (CASP8) and caspase-10 (CASP10). The extrinsic pathway is also used by various immune cells to trigger apoptotic cell death in tumor cells including TRAIL [133]. In addition, the inflammatory cytokine TNF-α produced by activated macrophages, which binds to the TNFR1 and TNFR2 receptors in most human cells, can elicit apoptotic response. Moreover, cytotoxic lymphocytes carry the FasL, which binds and activates the FAS receptor on the surface of the target cell that is followed by death-inducing signaling complex (DISC) formation. Subsequently, adapter proteins bind to the intracellular region of aggregated DISC complex, causing the accumulation of procaspase-8 molecules, which via proteolytic cleavage initiate a proteolytic cascade leading to effector caspase activation. There is also an amplification step where further release of mitochondria-localized pro-apoptotic factors takes place to amplify the initial CASP-3 activation (Box 3.23).


Apoptosis is a distinctive and highly controlled form of programmed cell death, which requires energy to hit the self-destruct button of an affected cell.

Apoptosis which can be triggered in response to endogenous or exogenous signals is a chain of sequential morphological events during which the early apoptotic cell shrinks and chromatin is irreversibly condensed and cleaved culminating into formation of apoptotic bodies.

In the mitochondria-mediated or intrinsic route to caspase activation, induction of mitochondrial outer membrane permeabilization (MOMP) is a central event that sets free pro-apoptotic factors such as cytochrome c.

The BCL-2 proteins can positively and negatively control MOMP.

The extrinsic pathway is mediated by a death ligand/signal binding to a membrane death receptor and downstream activation of CASP8.


Apoptosis is a distinctive and highly controlled form of programmed cell death, which requires energy to hit the self-destruct button of an affected cell.


Apoptosis which can be triggered in response to endogenous or exogenous signals is a chain of sequential morphological events during which the early apoptotic cell shrinks and chromatin is irreversibly condensed and cleaved culminating into formation of apoptotic bodies.


In the mitochondria-mediated or intrinsic route to caspase activation, induction of mitochondrial outer membrane permeabilization (MOMP) is a central event that sets free pro-apoptotic factors such as cytochrome c.


The extrinsic pathway is mediated by a death ligand/signal binding to a membrane death receptor and downstream activation of CASP8.


IR-induced DNA damages, e.g., unrepaired DNA SSBs or DSBs, primarily trigger apoptosis via the intrinsic pathway [134]; however, at certain IR doses and in certain cell types, the extrinsic apoptotic pathway may also be executed. IR can also initiate mitochondria-mediated signaling in response to ceramide production/formation at the plasma membrane. Moreover, IR can trigger the production of O2− and ROS (like H2O2 or OH− radicals), which via release of Ca2+ and cytochrome c from mitochondria can cause apoptosis [135].


One important signaling regulator of apoptosis in response to IR is TP53 [136] (Fig. 3.38). Thus, TP53 is phosphorylated in response to DDR signaling, accumulates in the nucleus, and binds to promoters of target genes, e.g., BAX, PUMA, NOXA, p53AIP1, and APAF-1. This results in an alteration in their transcription and hence expression levels, which is followed by mitochondria-mediated apoptosis.

A flow diagram depicts that stress or stimuli activate p 53 which binds to pro-apoptotic genes, resulting in cytochrome c release, apoptosome formation, and effector caspases with apoptosis.

Fig. 3.38
TP53-mediated intrinsic route to apoptosis. The mechanisms of TP53-induced apoptosis through the Bcl-2-regulated pathways in cells undergoing stress are shown. DNA damage triggers stress signaling, which in turn causes stabilization of the TP53 protein in the nucleus. Subsequently, TP53 as a nuclear transcription factor increases the expression of BH3-only proteins such as PUMA and NOXA and downregulation of BCL-2 or BCL-XL expression. The BH3-only proteins bind and inhibit the anti-apoptotic or pro-survival BCL-2 family proteins, so as to unleash the cell death effectors (BAX/BAK) which are often held as hallmarks of apoptosis in affected cells. Oligomerization of BAX/BAK causes MOMP, with subsequent release of cytochrome c, formation of the apoptosome complex, and activation of CASP9 and subsequently effector caspases, which causes apoptotic features of the dying cells. Abbreviations: ROS reactive oxygen species, MOMP mitochondrial outer membrane permeabilization, BH3 Bcl-2 homology 3, PUMA p53 upregulated modulator of apoptosis, BAD Bcl-2-associated agonist of cell death, CHOP CCAAT/enhancer-binding protein homologous protein, Bcl-2 B-cell lymphoma 2 (an apoptotic inhibitor), Bcl-xL B-cell lymphoma-extra-large, Bax Bcl2-associated X (an apoptotic regulator), Bak Bcl2 antagonist killer 1, APAF-1 apoptotic peptidase-activating factor 1, caspase cascade of aspartate-specific cysteine proteases


A flow diagram depicts that stress or stimuli activate p 53 which binds to pro-apoptotic genes, resulting in cytochrome c release, apoptosome formation, and effector caspases with apoptosis.


The extrinsic pathway may also play a role in IR-induced apoptosis in which TP53 may upregulate the expression of the FAS receptor and its ligands, which subsequently causes downstream transactivation of initiator CASP8 and apoptosis.


IR may moreover activate the ceramide pathway at the plasma membrane, wherein formation of ROS inflicts lipid oxidative damage in the membrane (Fig. 3.39). Subsequently, acid sphingomyelinase is activated, and second messenger ceramide is released as a result of sphingomyelin hydrolysis. IR-induced DNA damage may also trigger mitochondrial ceramide synthase resulting in the accumulation of ceramide which subsequently can induce apoptosis [137].

 A flow diagram depicts many stress factors that cause E R, Golgi, and mitochondrial stress. They cause the accumulation of misfolded proteins, altered B c l 2, and A T P depletion that leads to apoptosis.

Fig. 3.39
Overview of ceramide signaling and connection to the apoptotic machinery. IR-induced lipid oxidative damage causes sphingomyelinase activation at the plasma membrane, followed by hydrolysis of sphingomyelin and release of ceramide. High dose of IR-induced DNA DSBs can also trigger the mitochondrial ceramide synthase for de novo synthesis of ceramide. Inhibition of SERCA and calcium depletion in ER promote ER stress. Expression of downstream pro-apoptotic factor, e.g., CHOP, increases. The UPR activator proteins, ATF6, IRE1, and PERK, alter ER stress. The PERK pathway via ATF4-dependent NRF2 expression triggers the CHOP-mediated apoptotic pathway. CHOP can also be induced by spliced ATF-6 (in Golgi), which regulates the Bcl-2 protein family. CAPPs can alter the BCL-2 protein family, which determines the commitment of cells to apoptosis. Abbreviations: Cer ceramide, CerS1–6 a family of six ceramide synthases, SMase sphingomyelinase, SERCA sarco-endoplasmic reticulum calcium transport ATPase, ER endoplasmic reticulum, ATF6 activating transcription factor 6, IRE1 inositol-requiring enzyme 1, PERK protein kinase R-like ER kinase, NRF2 nuclear factor erythroid 2-related factor-2, ATF4 activating transcription factor 4, CHOP CCAAT/enhancer-binding protein homologous protein, Mt mitochondria, CAPPs ceramide-activated protein phosphatase, Bcl-2 B-cell lymphoma 2 (an apoptotic inhibitor), Bcl-xL B-cell lymphoma-extra-large, Bax Bcl-2-associated X (an apoptotic regulator), RNS reactive nitrogen species, ATP adenosine triphosphate


A flow diagram depicts many stress factors that cause E R, Golgi, and mitochondrial stress. They cause the accumulation of misfolded proteins, altered B c l 2, and A T P depletion that leads to apoptosis.


Ceramide may also activate the RAC1/mitogen-activated protein kinase kinase kinase-1 (MAP3K1) pathway by which MAPK8 and the effector CASP-1, -3, and -6 are induced and which also stimulate the DR pathway. MAPK8/JNK1 is known to be triggered in response to IR as well as other apoptotic stimuli, and depending on the duration of activity, it may induce apoptotic signaling. In summary, the rate of apoptotic events after IR may be executed via different routes and is influenced by cell type, cell cycle phase, dosage number, as well as radiation quality (Box 3.24).


IR-induced apoptosis can be executed through intrinsic, extrinsic, or membrane stress (ceramide) pathways.

IR may trigger apoptosis via mitochondria where TP53 regulation of the BCL-2 family proteins is of major importance.


IR-induced apoptosis can be executed through intrinsic, extrinsic, or membrane stress (ceramide) pathways.


IR may trigger apoptosis via mitochondria where TP53 regulation of the BCL-2 family proteins is of major importance.


The apoptotic cell features, i.e., cell morphology, and the activation of different apoptotic signaling routes giving rise to distinguishable phenotypes have been extensively studied with multiple methods at hand. The detection of apoptosis includes methods (Fig. 3.40) related to membrane alterations, e.g., PS exposure monitored by annexin V association [129]; DNA fragmentation assessment; cytotoxicity and cell proliferation assays; analyses of mitochondrial effects, i.e., cell permeabilization; loss of mitochondrial potential; BCL-2 family protein complex formation; association of the apoptosome or DISC complex in cytosol; and pro-caspase cleavage later via different antibody-based, enzymatic assays or by flow cytometry [138]. Moreover, less frequently used technologies such as light-scattering flow cytometry and time-lapse microscopy perfusion platform can be performed to avoid underestimating the extent and timing of apoptosis, temporal aspects of death, cell surface area assessment, cellular adhesion analysis, and genotoxicity-specific chromatin changes.

An illustration describes different assays to evaluate cell death based on membrane integrity assay, functional assay, D N A labeling assay, morphological mechanism-based assay, reproductive assay, and colony forming assay.

Fig. 3.40
Methods to detect cell death, in particular apoptotic cell death. The schematic diagram outlines various biological assays used to determine apoptotic cell death. Some of these assays can also be used to assess other types of cell death. These assays are based on the morphological criteria and distinguishing features of apoptotic pathways, e.g., staining for PS exposure on the outer plasma membrane (by annexin V assay) and caspase-3 activation or PARP cleavage (by, e.g., western blotting). Cell viability assays such as membrane integrity assays and reproductive assays are performed to monitor live cells in culture and measure an enzymatic activity as a marker of viable cells by using different classes of colorimetric reagents and substrates generating a fluorescent signal. Results from these assays do not always indicate apoptosis, but more about cell death in general. DNA labeling assay, functional assays, and morphological mechanism-based assays detect and quantify the cellular events, some of which are specifically associated with apoptotic cell death, such as formation of apoptotic antibodies, expression of apoptotic inhibitors, caspase activation in either intrinsic or extrinsic pathways, and DNA fragmentation. The principles for each assay are given in the respective yellow boxes. Abbreviations: MTT (3-(4, 5-dimethylthiazolyl-2)-2, 5-diphenyltetrazolium bromide), LDH lactate dehydrogenase, BrdU bromodeoxyuridine, PARP polyadenosine diphosphate-ribose polymerase, PS phosphatidylserine


An illustration describes different assays to evaluate cell death based on membrane integrity assay, functional assay, D N A labeling assay, morphological mechanism-based assay, reproductive assay, and colony forming assay.


Necrosis (from the Greek “nekros” designating “to kill”) has for long been seen upon as an uncontrolled, irreversible mode of cell death, while recent work suggests that necrosis is a tightly genetically regulated pathway yet triggering inflammatory and/or reparative reactions in the tissue [139]. Necrotic cell death can be classified into accidental cell death (ACD) and regulated necrotic cell death (RNCD). RNCD can be further classified into necroptosis, pyroptosis, ferroptosis, NETosis, and methuosis given their molecular routes [139] (Table 3.11).Table 3.11
Accidental and regulated necrosis, key features, and methods of detection

Type of cell death

Morphology

Detection methods

Accidental necrosis

Membrane disruption, mitochondria swelling (loss of organelle), cell swelling

LDH quantification, cell-impermeable DNA-binding dye, membrane integrity loss

Necroptosis

Membrane disruption, moderate chromatin condensation, cell swelling

Flow cytometry, western blot, immunohistochemistry—levels of biomarker proteins, mitochondrial depolarization detection, fluorescence microscopy for membrane loss, electron microscopy for morphology

Pyroptosis

Membrane disruption, bubbling, moderate chromatin condensation

LDH quantification, fluorescence microscopy for membrane integrity loss, western blot for GSDM D, IL-1β

Ferroptosis

Membrane disruption, iron accumulation, lipid peroxidation, diminutive mitochondria

Lipid peroxide quantification—flow cytometry and BODIPY-C11 probe

Methuosis

Membrane disruption, accumulation of large fluid-filled vacuoles, cell swelling

Electron microscopy, time-lapse fluorescence microscopy for morphology, metabolic flux analysis

NETosis

Membrane disruption, chromatin condensation

Fluorescence microscopy for morphology, flow cytometry, ELISA, western blot


Necroptosis, pyroptosis, methuosis, and ferroptosis are all triggered in response to IR [124, 140]. In the context of RT of cancer, necrosis can be induced either directly following DNA damage or indirectly by ROS formation that reacts with lipids generating lipid peroxides. IR has also been linked to lipid peroxidation and ferroptosis, and necroptosis together with ferroptosis was postulated to occur via ATM signaling.


Both ACD and RNCD trigger immunogenic cell death (ICD). In turn, ICD can stimulate an adaptive immune response after antigen is exposed by cells after RT or chemotherapeutics [141]. In case of immunogenic cell death, damage-associated molecular patterns (DAMPs) are delivered and identified by pathogen recognition receptors (PRRs) exhibited by intrinsic components of the immune system, conducting to the stimulation of an immune response [141]. ACD is an uncontrolled type of cell death which is activated by, e.g., physical damage, hypoxia, inflammatory toxins, and high doses of IR. The cells respond by morphological alterations, such as cytoplasmic swelling of the cell organelles, i.e., oncosis [142], which is a result of disturbance of ionic pumps causing Ca+ influx, plasma membrane disruption followed by the leakage of intracellular organelles with accidental deteriorated DNA, and absence of clear chromatin condensation [142]. RNCD comprises upregulation of diverse pro-inflammatory proteins and molecules such as nuclear factor-κB, leading to the rupture of the cell membrane causing leakage of the cellular debris, e.g., ATP, DNA, nuclear proteins, heat-shock proteins, and uric acid, into surrounding zones, provoking a cascade of inflammation and tissue injury. Thus, the release of proteins/molecules promotes inflammasome activation and production of pro-inflammatory cytokine interleukin-1 beta (IL1). The methods used to detect necrosis are lactate dehydrogenase (LDH) activity measurement and cell-impermeable DNA-binding dye. These techniques are based on the morphological characteristics proving the cellular release and membrane porosity (Table 3.12).Table 3.12
Examples of some oncogenes in cancer from Weinberg [143] and Gillies et al. [144]

Oncogene

General function

Major tumor type with deregulation

K-ras

Guanine nucleotide-binding protein

Lung, ovarian, colorectal, bladder carcinomas

N-ras

Guanine nucleotide-binding protein

Head and neck cancers

H-ras

Guanine nucleotide-binding protein

Colorectal carcinomas

c-myc

Transcription factor

Various leukemias, carcinomas

L-myc

Transcription factor

Lung carcinomas

EGFR/HER2

Receptor tyrosine kinase

Glioblastomas, lung cancer, breast cancer

Src

Cytoplasmic tyrosine kinase

Colon cancer, head and neck cancers, chronic myelogenous leukemia

Sis/PDGF

Growth factor

Simian sarcoma


Necroptosis, also known as a regulated necrosis, which works in a caspase-independent fashion, exhibits a necrotic morphology with membrane disruption and leakage of organelles (reviewed by Weinlich et al. (2017)). Different stimuli can elicit necroptosis: DRs, e.g., members of the TNFR superfamily, pattern recognition receptors (PRRs), Toll-like receptors (TLRs), T-cell receptors (TCRs), multiple chemotherapeutic drugs, and hypoxia. The process of necroptosis commences by the stimulation of receptor-interacting protein kinases (RIPKs) (Fig. 3.41).

Five flow diagrams depict the cellular pathways of necroptosis, pyroptosis, ferroptosis, netotic cell death, and methuosis. Cell death occurs for many reasons, such as membrane rupture and lipid peroxidation.

Fig. 3.41
Summary of regulated necrotic cell death. (a) Necroptosis elicited by DR, TLR, and viruses stimulates RIPK3 and then MLKL, which is required for membrane disruption. (b) Pyroptosis induced by GSDMD following its cleavage by CASP1 and CASP11. The main elicitors: PAMPs and DAMPs, or cytosolic LPS. (c) Ferroptosis is dependent on the balance between ROS production due to iron accumulation and antioxidant defense mechanisms that inhibit lipid peroxidation. The ACSL4–LPCAT3–ALOX15 pathway mediates lipid peroxidation, while system xc- (comprising SLC7A11, GPX4, and NFE2L2) impeded this process. (d) NETosis is triggered by NET leakage, which is mediated by ROS generation and histone citrullination. (e) Methuosis is associated with macropinocytosis. Nascent micropinosomes fused forming large vacuoles that contain late endosomal markers (LAMP1 and Rab7). These do not recycle or unify with lysosomes causing cell death. Reproduced with permission (CCBY) from Tang et al. [145]. DR death receptor, TLR Toll-like receptor, RIPK3 receptor-interacting protein kinases 3, MLKL mixed-lineage kinase domain-like protein, GSDMD gasdermin D, CASP1 caspase 1, CASP11 caspase 11, PAMPs pathogen-associated molecular patterns, DAMPs damage-associated molecular patterns, or cytosolic, LPS lipopolysaccharide, ACSL4 acyl-CoA synthetase long-chain family member 4, LPCAT3 lysophosphatidylcholine acyltransferase 3, ALOX15 arachidonate lipoxygenases (ALOXs, specifically ALOX15), SLC7A11 the catalytic subunit solute carrier family 7 member 11, GPX4 glutathione peroxidase 4, NFE2L2 nuclear factor erythroid 2-like 2, NET NETosis extracellular trap, ROS reactive oxygen species, LAMP1 lysosomal associated membrane protein 1, Rab7 lysosomal Rab protein 7. (Adapted from Tang et al. [145])


Five flow diagrams depict the cellular pathways of necroptosis, pyroptosis, ferroptosis, netotic cell death, and methuosis. Cell death occurs for many reasons, such as membrane rupture and lipid peroxidation.


RIPKs are stimulated to go into macromolecular complexes from the membrane receptors with the necrosome with RIPK1 and RIPK3 being the main components. RIPK3 subsequently stimulates mixed-lineage kinase domain-like protein (MLKL) through phosphorylation causing its oligomerization and relocalization, resulting in cell membrane permeabilization and subsequent cell death.


Different techniques can be used to identify necroptosis, e.g., flow cytometry, western blotting, and immunohistochemistry. Through these techniques, the expression levels of MLKL, RIPK3, and RIPK1 are evaluated as well as cell by electron microscopy (Table 3.11).


Pyroptosis, which is stimulated by IR as well as intracellular pathogenic factors in immune cells, follows a series of caspase-dependent events and is pro-inflammatory (reviewed by Yu et al. [146]). Thus, the NOD-like receptors (NLRs) of irradiated/infected macrophages/monocytes recognize cytoplasmic pathogen-associated molecular patterns (PAMPs) as well as DAMPs and trigger inflammasome complex production, which activates CASP1. CASP1 in turn activates gasdermin D, which mediates the plasma membrane rupture (Fig. 3.42) as well as the inflammatory cytokines interleukin 1β (IL-1β) and IL-18, which further regulate inflammation. Pyroptosis also involves cell swelling followed by disintegration of the plasma membrane and leakage of the pro-inflammatory contents, e.g., DAMPs, IL-1β, and IL-18, contributing to elimination of the immunologic challenges locally or systemically. Pyroptosis can be detected by LDH assay, fluorescence microscopy, western blot analysis (for identification of gasdermin D, IL-1β), and measurement of the cell intake of propidium iodide (Table 3.11).

A flow diagram depicts those monolayer cells in trypsin form cell suspension, which is 2 D and 3 D. Seeding, incubation, and staining give colony count, plating efficiency, and survival fraction.

Fig. 3.42
Methodology for 2D (Puck) and 3D clonogenic curves. The clonogenic assay measures the ability of single cells to form colonies. A cancer cell that is not able to form a colony can be regarded as inactivated. Cellular monolayers are dissociated into single cells and counted and diluted to the required concentration, depending on the dose. The cells are then seeded in cell flasks/dishes for colony formation or in a 3D matrix for spheroid formation. After irradiation, the cells are incubated for 1–3 weeks depending on the cell doubling time of that particular cell line, before they are fixed, stained, and counted. The surviving fraction is calculated as the number of colonies in irradiated samples relative to the plating efficiency of unirradiated control dishes


A flow diagram depicts those monolayer cells in trypsin form cell suspension, which is 2 D and 3 D. Seeding, incubation, and staining give colony count, plating efficiency, and survival fraction.


Ferroptosis is a form of caspase-independent regulated necrosis and is distinguished by excessive iron-dependent lipid peroxidation. It presents a necrotic morphology with altered mitochondria, i.e., small mitochondria, fewer cristae, rupture of outer membrane, and an electron-dense ultrastructure. Execution of ferroptosis is decided by the equilibrium between ROS production due to iron increase and antioxidant protection mechanisms that impede lipid peroxidation. Thus, ferroptosis is activated after lipid peroxidation in a process catalyzed by iron, either in a Fenton-like manner or through lipoxygenases (Fig. 3.41). Accordingly, the oxidation of polyunsaturated fatty acids (PUFAs), like arachidonic acid (AA), is necessary for lipotoxicity in ferroptosis, which takes place via a catalytic pathway comprising acyl-CoA synthetase long-chain family member 4 (ACSL4), lysophosphatidylcholine acyltransferase 3 (LPCAT3), and arachidonate lipoxygenases (ALOXs, specifically ALOX15) [147]. In addition, lipid peroxidation can be hindered by the various antioxidant systems such as the cystine/glutamate antiporter system, which consists of the catalytic subunit solute carrier family 7 member 11 (SLC7A11), glutathione peroxidase 4 (GPX4), and pro-survival proteins, like nuclear factor erythroid 2-like 2 (NFE2L2). System xc- facilitates the exchange of cystine and glutamate in and out of the cell. The cystine which is taken up is reduced to cysteine in cells, which is needed for the synthesis of glutathione GSH. GSH is used by GPX4 to stop the generation of phospholipid hydroperoxides (PLOOH), the key mediator of chain reactions in lipoxygenases. The induction of ferroptosis can be determined by measuring lipid peroxides coupled with flow cytometry (Table 3.11).


NETosis is stimulated by various pathogens or other stimuli, which release neutrophil extracellular traps of mainly DNA-protein structures [148] in a process dependent on NADPH oxidase 4 (NOX4), the principal source of ROS (Fig. 3.41). NETosis also comes along with important increase of ROS conducting to the stimulation of protein-arginine deiminase 4 (PAD4). Then, PAD4 citrullinates (converts arginine to citrulline via deamination) the histones, promoting the nuclear chromatin decondensation. Further, the NET is released into the cytosol leading to the disruption of the neutrophil membrane. Then, neutrophil breaks up and the NETs are released into the environment. NETs can be generated by other forms of immune cells, e.g., eosinophils, mast cells, basophils, macrophages, and also epithelial cells and cancer cells as a response to various injuries [145]. NETosis can be studied using various techniques: immunofluorescence, transmission electron microscopy, scanning electron microscopy, ELISA tests, flow cytometry, as well as western blot analyses of NETosis markers (Table 3.11).


Methuosis (from Greek methuo—“drink to intoxification”) is another type of caspase-independent regulated necrotic cell death that is induced by exposure to heat, trauma, and infection and which lead to cell swelling, lysis of plasma membrane, as well as inflammation. Methuosis is correlated to macropinocytosis (referred to as “cell drinking”) and is associated with the extensive accumulation of fluid-filled cytoplasmic vacuoles stemmed from macropinosomes, which for example is observed in cancer cells driven by the oncoprotein Ras [149] (Fig. 3.41). Methuosis can be detected by electron microscopy, time-lapse fluorescence microscopy, visualization of vacuoles using fluorescent dyes, and metabolic flux analyses (Table 3.11) (Box 3.25).


Necrotic cell death is classified into accidental cell death and regulated necrotic cell death with different subtypes: necroptosis, pyroptosis, ferroptosis, NETosis, methuosis, etc.

IR may stimulate necrosis via direct DNA damage response and via radical oxygen species.

All types of necrosis are immunogenic cell death types.


Necrotic cell death is classified into accidental cell death and regulated necrotic cell death with different subtypes: necroptosis, pyroptosis, ferroptosis, NETosis, methuosis, etc.


IR may stimulate necrosis via direct DNA damage response and via radical oxygen species.


Autophagy is an adaptive and catabolic process induced by various forms of cellular stress, intended to mitigate the impact of cell damage to avoid cell death, by recycling biomolecules and damaged organelles. This mechanism occurs via a self-digestion process involving the formation of double-membrane vesicles, called autophagosomes, that merge with lysosomes. Autophagy can be induced by nutrient deprivation (amino acids, in particular leucine and glutamine, and glucose) and cytotoxic insults such as IR or chemotherapy. The main function of autophagy is to provide nutrients and building blocks for vital cellular functions during different forms of stress. Therefore, this pathway is generally considered as a cytoprotective mechanism [150]. Autophagy is a complex mechanism involving several steps. First, the recruitment of autophagy-related proteins (ATG) to a specific subcellular location called the phagophore assembly site (PAS) allows phagophore nucleation (initiation and phagophore nucleation). During phagophore elongation, a portion of the cytoplasm is engulfed (cargo sequestration) and the autophagosome, a double-membrane vesicle, is being formed (autophagosome maturation). Fusion of the autophagosome with lysosome allows the degradation of the autophagic cargo.


A key regulator of autophagy is the mammalian target of rapamycin (mTOR) that exists in two distinct protein complexes, mTORC1 and mTORC2. In its active conformation, mTORC1 prevents autophagy by inhibiting the UNC51-like kinase 1 (ULK1) complex, composed of ULK1, the autophagy-related gene 13 (ATG13), ATG101, and the FAK family-interacting protein of 200 kDa (FIP200). Upon autophagic stimuli, mTORC1 is inhibited, leading to the activation of ULK1. Active ULK1 phosphorylates ATG13 and FIP200, which leads to the activation of the class III phosphoinositide 3-kinase (PI3K) complex, allowing phagophore nucleation. This triggers the production of phosphatidylinositol-3-phosphate (PIP3) at a characteristic ER structure called omegasome. PIP3 recruits WD repeat domain phosphoinositide-interacting proteins (WIPI2) and zinc finger FYVE domain-containing protein 1 (DFCP1) to the omegasome. By binding ATG16L1, WIPI2 recruits the ATG12-ATG5-ATG16L1 complex that allows the conjugation of ATG8 family proteins (including microtubule-associated protein light-chain 3 (LC3) and γ-aminobutyric acid receptor-associated proteins (GABARAPs)) to membrane-resident phosphatidylethanolamine (PE). By this process, LC3-I (diffuse form) is converted into LC3-II (membrane-anchored, lipidated form), a marker of autophagic membranes. The recruitment of ATG9-containing vesicles (coming from the plasma membrane, mitochondria, recycling endosomes, and Golgi complex), delivering additional lipids and proteins, further contributes to autophagosomal membrane expansion. Once the membrane is sealed, the autophagosome is formed and undergoes maturation. Then it can merge with the lysosome, where the autophagic cargo will be degraded by acidic hydrolases. For the molecular details, see Dikic et al. [151].


Beyond apoptosis, the commonly studied IR-induced cell death mechanism, autophagy was shown to be frequently induced in response to IR. For example, autophagy can be triggered following DNA damage inflicted by IR or other agents. Indeed, DNA damage repair (DDR) is an energy-demanding process that consumes ATP but also NAD+ via the action of polyADP-ribose polymerase 1 (PARP1). Autophagy induction allows the recycling of metabolic precursors for ATP and provides energy for the DDR. ROS was also shown to trigger and regulate autophagy [152]. The function of IR-induced autophagy is still being debated. Results of in vitro and in vivo studies provided conflicting notions whether autophagy acts as a cytoprotective mechanism, promoting cell survival responsible for radioresistance. In that respect, radiosensitization strategies based on genetic or pharmacological autophagy inhibition led to different outcomes. Several studies also pointed out the non-cytoprotective function of IR-induced autophagy where autophagy inhibition failed to alter radiosensitivity. Although autophagic functions may vary depending on both cell type and treatment regimen applied, specific characteristics able to distinguish cytotoxic, cytoprotective, or non-cytoprotective forms of IR-induced autophagy have not yet been identified. There are assumptions that autophagy duration may play a role in radiosensitivity, with radioresistance occurring in case of prolonged autophagy, while a transient form of autophagy will ultimately lead to apoptosis [150].


Outcomes of clinical trials conducted with approved autophagy inhibitors (e.g., chloroquine and hydroxychloroquine) were mitigated due to toxicity issues and unsatisfactory autophagy inhibition. Concerns were raised regarding the most probable non-tumor selectivity of autophagy inhibitors, off-target effects, effects on immune response, and difficulty to monitor autophagy inhibition in patients’ tumors [153]. Further studies on the molecular mechanisms governing IR-induced autophagy may bring additional evidence on how to optimally modulate autophagy to produce favorable outcomes (Box 3.26).


Autophagy is triggered by IR and often considered as a cytoprotective mechanism.

Autophagy inhibition as a radiosensitization strategy led to inconsistent results, suggesting an intricate role of autophagy, being regulated by many factors.


Autophagy is triggered by IR and often considered as a cytoprotective mechanism.


Autophagy inhibition as a radiosensitization strategy led to inconsistent results, suggesting an intricate role of autophagy, being regulated by many factors.


As described in the sections before, cells damaged by radiation might suffer from genetic instability and/or die through, e.g., apoptosis or other types of cell death. These consequences of radiation exposure can be used to qualify and quantify the damage and draw conclusions on its severity. In this context, it is possible to look at not only the fatal outcome of radiation damage but also the capability of cells to survive IR. It is important to distinguish between cell survival and cell viability. In radiobiology, the term cell death is used also for cells that are inactivated, i.e., have lost their proliferation ability. Cancer cells and stem cells are characterized by their capacity for sustained proliferation. A cancer cell that has lost the ability to divide is by definition dead as a cancer cell even though it may still have an intact cell membrane and retained metabolic function. While non-proliferating cells retain their function even after radiation doses as high as 50–100 Gy, cancer cells may lose the capacity for uncontrolled cell division after doses in the order of 2 Gy.


There are several assays available to measure cell viability. Some use dye exclusion, such as trypan blue, to measure the proportion of cells with intact cell membrane. Others measure metabolic function through the activity of mitochondrial enzymes, such as the MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium) assay, cellular reducing conditions such as the Alamar Blue assay, or ATP production. Even though viability measurements over time can give an indication of cell proliferation, the only direct measurement of clonogenic function is the clonogenic assay, the gold standard for cell survival measurements. These assays can be performed in vitro with cultured cells or in vivo from biopsies.


The first survival curve, i.e., the relation between survival and delivered doses, was established with HeLa cells cultivated in vitro and irradiated with X-rays by Puck and Marcus in 1956 [154]. A surviving cell is defined as a cell able to divide and form a colony composed of at least 50 cells. To find the surviving fraction, the capacity of nonirradiated and irradiated cells to form colonies is compared. Typically, in vitro cell survival is measured in adherent cells in monolayer culture. The day before the experiment, cells are trypsinized. Viable cells are counted with a hematocytometer or a cell counter. A determined number of cells in suspension is seeded in Petri dishes (or flasks) destined to be a control or irradiated before their first doubling time. Depending on the design of the experiments, the medium can be changed after irradiation. Then cells are incubated at 37 °C for 1–3 weeks according to the cell types (≥8 divisions). When the colonies grow to exceed 50 cells, observable by microscopy or visually detectable, they are fixated with methanol or ethanol and then stained with Giemsa, methylene blue, or crystal violet before several washes with water and drying [155]. After that, the clones formed are counted manually or with an automatic counter (Fig. 3.42).


All cells comprising each colony are the progeny of a single initial cell seeded, which survived irradiation. If we consider 100 untreated cells, the ideal number of colonies formed should be 100. However, this is never the case, depending on diverse factors (medium change, errors and uncertainties in counting the cell suspension, trauma of the detachment …), and in fact 50–90 colonies might be expected. Considering the outcome of the control conditions (nonirradiated), the term plating efficiency (PE) can be defined. This corresponds to the percentage of cells seeded, which grew into colonies. If 75 colonies are counted after seeding 100 cells, we talk about a PE of 75%. It must be noted that the PE may differ according to the number of cells seeded: this is the “feeder effect.” This effect is attributed to the need of some cell types to be able to cooperate with neighboring cells [156]. If this communication is missing, the cells are not able to start proliferation. Therefore, the cell density seeded might play a role in the fraction of cells able to form colonies. This might limit the robustness of the classical analysis of the colony-forming assay. In future, a different way of performing and analyzing this assay might be necessary [156].


In classical colony-forming assay parallel to the control samples, cells are irradiated, then incubated, fixed, and stained at the same time point as control cells. Different cases can therefore be observed: (1) some of the seeded cells being still single and not divided; (2) cells that managed one or two divisions to form a tiny abortive clone; and (3) cells able to form large colonies of at least 50 cells, corresponding to 5–6 cell divisions, but which can look like a little bit different from the untreated cells in terms of aspect and size. These latter cells, able to form colonies, are qualified of “survivors” and counted since they have retained their reproductive integrity. For example, if we seed 3000 cells followed by irradiation of 5 Gy, and if the PE previously determined is 0.75, then we can expect the attachment of 2250 cells (0.75 × 3000). If at 5 Gy 42 colonies grew up after incubation, the surviving fraction can be calculated at 1.9%: 42/(3000 × 0.75) = 0.019. In general, the plating efficiency (PE) and the surviving fraction (SF) are given by

 (3.1)


Survival curves for mammalian cells are usually presented in a form with dose plotted on a linear scale and surviving fraction on a logarithmic scale and can be fitted by several models, as for example the linear-quadratic model (see Chap. 1). The form of the curves, as seen in Chap. 1, depends on the linear energy transfer and allows determining important biological parameters such as the surviving fraction, the ratio α/β, or the relative biological efficiency (RBE) for example (see Chap. 1 for details). The surviving fraction at 2 Gy (SF2) is often used to approximate cell radiosensitivity.


To obtain a survival curve, several doses of irradiation have to be applied. The number of cells seeded per dish needs to be accurate and often adjusted after preliminary experiments to count a significant number of colonies since these parameters are dependent on doses, cell lines, and type of radiation. At least a triplicate of different dilutions is realized for each condition tested (here each dose delivered). If colonies are few, the statistical significance is reduced. On the opposite, if the colonies are too many, some colonies can be merged with another one, and the counting is inaccurate. In some cases, cells could be irradiated first (one flask for one dose) and then detached to be seeded at different dilutions [157]. However, precautions need to be considered since some cells are sensitive to detachment after irradiation, which affects cell survival. In addition, colony-forming assays require very accurate cell counting, since the controls come from a separate trypsinization. Clonogenic curves cannot discriminate the type of cell death, but they give information about the radiosensitivity of the cells.


More recently, the literature showed that survival curves obtained with three-dimensional (3D) cell models more reliably reflect the cell response in vivo than the results obtained with 2D cell monolayer culture [158]. 3D cell models for cell survival can be obtained by embedding single cells in an extracellular matrix, put in 96-well plates pre-coated with agarose, covered with medium, and then exposed to radiations. Cells are grown for a few days until cell clusters reach 50 cells, and the number of colonies is microscopically counted (Fig. 3.42).


An in vivo clonogenic assay allows measuring cell survival in an animal model, allowing the study of radiosensitivity of normal or tumor cells treated in vivo. These systems depend on the reproductive integrity of individual cells and allow the observation of a clone of cells regenerated in the irradiated tissue. There are assays developed for early-responding tissues, which divide rapidly and respond early to the effects of radiation, like bone marrow cells, skin, and intestinal epithelium, and assays for late-responding tissues, like lung, kidney, and spinal cord (Fig. 3.43).

An illustration depicts the approximate number of days taken for the jejunum crypt assay, skin clone assay, spleen colony assay, bone marrow assay, and kidney assay carried out in an irradiated mouse.

Fig. 3.43
In vivo assays. Four in vivo animal assays to assess clonogenic capacity after irradiation have been important for radiobiology. (1) The jejunum crypt assay measures the regenerative ability of jejunal crypts after high doses of irradiation. The animals are sacrificed 3.5 days after irradiation, and the numbers of regenerating crypts per circumference are measured. One regenerating crypt corresponds to one surviving clonogenic cell. (2) The skin clone assay used pre-irradiation with a high dose in a ring (moat) around the test skin area to avoid migration of neighboring cells into the test area. The test area is then irradiated, and the number of regrowing skin nodules per cm2 is counted. (3) The spleen colony assay uses transplants of bone marrow cells from an irradiated donor animal. These cells are transferred to recipient animals who have previously been irradiated with a high dose to kill all their own bone marrow cells. After 10–11 days, the recipient animals are sacrificed and their spleens are analyzed for colony-forming units arising from the implanted single cells. (4) The kidney assay uses the same animal for irradiation and control. One kidney of each animal is irradiated, and 60 weeks later, the animals are sacrificed. The number of intact kidney tubules is then counted in both kidneys, and the irradiated kidney can be compared to the unirradiated one


An illustration depicts the approximate number of days taken for the jejunum crypt assay, skin clone assay, spleen colony assay, bone marrow assay, and kidney assay carried out in an irradiated mouse.


The spleen colony assay, also called bone marrow stem cell assay, was first described by Till and McCulloch [159]. The basis of this assay relies on the use of one donor mouse and a group of recipient mice. Recipient mice are previously exposed to whole-body irradiation (9 Gy) to sterilize the spleen and suppress endogenous hematopoiesis. Then, from a donor mouse irradiated with a test dose, a cell suspension of bone marrow cells is taken and injected intravenously into the recipient donors. Some of these cells will lodge in the spleen, and after 10–11 days, single cell-derived clones will appear in the surface of the spleen. These colonies are usually called colony-forming units (CFUs). At this point of the experiment, the spleen of the recipient mouse is removed and the CFUs are counted. The surviving fraction is given by Eq. (3.2), similar to the one used for the in vitro assay. The experiment is then repeated for different radiation doses, enabling to trace a survival curve:

 (3.2)


The skin clone assay is based on the formation of nodules of mouse skin regrowing from a single surviving cell. In a practical way, after shaving a small area on the back of one mouse, a ring of skin is irradiated with a massive dose of 30 Gy to create a “moat” of dead cells. A small metal sphere is put in the central area to protect it from the radiation and create an isolated island of intact skin. This skin island is then irradiated with a test dose. Some days later, nodules of regrowing skin will be observed. The survival curve is obtained after repeating the experiment in different skin areas and by plotting the number of surviving cells per cm2 of skin as a function of the radiation dose (Gy).


The jejunal crypt stem cell assay is based on the self-renewal system of the jejunum. Within this system, the stem cells in the crypts divide rapidly and move up to the villi where they undergo differentiation in functioning cells. For the assay, groups of animals are subjected to increasing doses of whole-body irradiation. The jejunal crypts will begin to regenerate after 3.5 days, time at each animal is sacrificed, and sections of the jejunum are imaged. One regenerating crypt corresponds to one surviving clonogenic cell. The survival curve is obtained by plotting the number of regenerating crypts per circumference of the sectioned jejunum as a function of the radiation dose (Gy).


The kidney tubule assay includes the irradiation of one kidney per mouse with a small field. As the kidney is a late-responding tissue, the assay is finished 60 weeks later, when unirradiated and irradiated kidneys are removed, and histologic sections are imaged. The number of intact kidney tubules is compared between the unirradiated and irradiated sides. The survival curve is obtained by plotting the number of tubule-regenerating cells in a defined number of tubule cross sections counted as a function of the radiation dose (Gy).


In addition, the tumor control dose assays (TCD50) relate with tumor survival. During these assays, small parts of tumors (xenografts), which can be derived from tumor cell lines or from patient tumors, are implanted to nude mice. After they reach a desirable size, the tumors are irradiated by several doses and then the local control or recurrence is observed. A plot between the percentage of the controlled tumors versus the dose is made. TCD50 is then the dose to control 50% of the tumors [160] (Box 3.27).


A surviving cell corresponds to a cell able to divide and form a colony.

Clonogenic assay is based on the ability of a single cell to grow into a colony.

The only direct measurement of clonogenic function is the clonogenic assay, the gold standard for cell survival measurements.

Cell survival measurements allow to trace a cell dose-response curve, usually presented with dose plotted on a linear scale and surviving fraction on a logarithmic scale, and can be fitted by several models.

An in vivo clonogenic assay allows measuring cell survival in an animal model, allowing the study of radiosensitivity of normal or tumor cells treated in vivo.


A surviving cell corresponds to a cell able to divide and form a colony.


Clonogenic assay is based on the ability of a single cell to grow into a colony.


The only direct measurement of clonogenic function is the clonogenic assay, the gold standard for cell survival measurements.


Cell survival measurements allow to trace a cell dose-response curve, usually presented with dose plotted on a linear scale and surviving fraction on a logarithmic scale, and can be fitted by several models.


An in vivo clonogenic assay allows measuring cell survival in an animal model, allowing the study of radiosensitivity of normal or tumor cells treated in vivo.


Transformation of a normal cell into a cancer cell is a multistep process where mutations or other genomic alterations, e.g., copy number alterations, deletions, and gene fusions, alter the normal gene coding sequence. These alterations can occur due to mis- or unrepaired IR damage. Not all alterations lead to the transformation of a normal cell to a cancer cell, called oncogenesis, as it is associated with alterations of specific places on DNA [143]. Cell transformation is mostly related to the activation of proto-oncogenes, which are then named oncogenes and the deactivation of tumor suppressor genes [143]. Proto-oncogenes are genes associated with the activation of cell proliferation and differentiation. When they mutate or are somehow pressed to overexpression, cells proliferate out of control [143]. On the other hand, tumor suppressor genes are genes that control cell proliferation, play significant roles during DNA repair, or activate cell death pathways, when it is needed. Mutations of tumor suppressor genes cause loss of control upon important pathways, which may again lead to unregulated cell proliferation [143]. Oncogenes and tumor suppressor genes can be affected genetically by mutations on the DNA or also switched on or off epigenetically. An overview is given in Fig. 3.44.

 A flowchart depicts that proto-oncogenes and tumor suppressor genes cause cancer due to their activation and control of cell proliferation, followed by oncogene upregulation and tumor suppressor gene downregulation, respectively.

Fig. 3.44
Overview of oncogenes and tumor suppressor genes’ function and regulation


A flowchart depicts that proto-oncogenes and tumor suppressor genes cause cancer due to their activation and control of cell proliferation, followed by oncogene upregulation and tumor suppressor gene downregulation, respectively.


The discovery of proto-oncogenes came with investigation of the Rous sarcoma virus (RSV). This virus is able to transform normal chicken cells to cancer cells, and in its structure, the src gene was found, which as it was shown later was responsible for this transformation. The src gene was later also found in the normal chicken genome, but it was inactivated. These findings meant that the genomes of normal cells carry genes (proto-oncogenes) that have, under certain circumstances, the potential to induce cell transformation when activated [143]. For some time, biologists were convinced that cancer is caused by viruses which present into cells’ genes (oncogenes) that activate uncontrolled cell proliferation. It was thus strange that people around these “infected” people do not suffer from the same cancer type as well, due to the fact that viruses are infectious. Indeed, viruses can include oncogenes into a cell’s DNA, but viruses are not the main cancer cause. Viruses are responsible only for a minority of all cancers [143]. All this information led to new questions about proto-oncogenes and oncogenes. To find out if oncogenes exist in chemically or physically transformed cells, DNA from cancer cells was introduced to normal cells to see if they will be transformed. This gene transfer procedure is named transfection. Indeed, many other oncogenes were revealed using this method [143]. Another very important issue is that it is sufficient to activate only one of the alleles of a proto-oncogene to get oncogene upregulation [161]. Some of the most common oncogenes in human cancer are given in Table 3.12.


In general, when a system has an activation “button,” there has to be somewhere a deactivation “button” as well. Oncogenes are the genes activating uncontrolled cell proliferation, and on the other hand the deactivation/control of cell proliferation is associated with tumor suppressor genes. Tumor suppressor genes were discovered much later than proto-oncogenes and oncogenes. Some of the tumor suppressor genes are listed in Table 3.13. One of the most important and known tumor suppressor genes is the TP53. A mutation of TP53 is associated with various tumor types. This gene codes the p53 protein, which is also sometimes called the “Master Guardian.” p53 is responsible for activation of DNA repair as well as activation of cell cycle arrest, to enable DNA repair.Table 3.13
Examples of some tumor suppressor genes and familial cancer syndromes from Macleod [162] and Weinberg [143]

Tumor suppressor gene

General function

Types of cancer

Familial syndrome

TP53

Chromosome stability, transcriptional regulator, growth arrest, apoptosis

Many

Li–Fraumeni syndrome

p16

Cyclin-dependent kinase inhibitor

Many

Familial melanoma

BRCA1

Transcriptional regulator, DNA repair

Many, mostly breast and ovarian cancer

Familial breast cancer

BRCA2

Transcriptional regulator, DNA repair

Many, mostly breast and ovarian cancer

Familial breast cancer

RB1

Transcriptional regulator of cell cycle

Retinoblastoma, osteosarcoma

Familial retinoblastoma

E-cadherin

Cell adhesion regulator

Breast, colon, lung, skin carcinoma

Familial gastric cancer

APC

β-Catenin degradation

Colorectal, pancreatic, stomach, prostate cancer

Familial adenomatous polyposis coli

NF2

Cytoskeleton-membrane linkage

Schwannoma, meningioma, ependymoma

Neurofibroma-predisposition syndrome


To deactivate a tumor suppressor gene, both alleles have to be damaged or switched off, because only one allele is enough for the production of a specific protein. Anyhow, if one allele of a tumor suppressor gene of a germ line cell is defective, then there is much higher probability of the born individual to suffer from cancer. This is because for this person, it becomes much more probable that the second allele will be damaged during life as well [143, 161, 162]. Since the defective allele in this case is genetically transferred to offspring, many familial syndromes were identified (Box 3.28).


DNA alterations in genomic or epigenetic level may cause proto-oncogenes to become oncogenes, disrupting normal cell division and causing cancers to form.

Cell transformation is mostly related to the activation of proto-oncogenes, which are then named oncogenes, and deactivation of tumor suppressor genes.

Proto-oncogenes are genes associated with the activation of cell proliferation and differentiation.

Tumor suppressor genes are genes that control cell proliferation, play significant roles during DNA repair, or activate cell death pathways.

Mutations of tumor suppressor genes cause loss of control upon important pathways, which may again lead to unregulated cell proliferation.

Oncogenes and tumor suppressor genes can be affected genetically by mutations on the DNA or also switched on or off epigenetically.

TP53 is one of the most important tumor suppressor genes.


DNA alterations in genomic or epigenetic level may cause proto-oncogenes to become oncogenes, disrupting normal cell division and causing cancers to form.


Cell transformation is mostly related to the activation of proto-oncogenes, which are then named oncogenes, and deactivation of tumor suppressor genes.


Proto-oncogenes are genes associated with the activation of cell proliferation and differentiation.


Tumor suppressor genes are genes that control cell proliferation, play significant roles during DNA repair, or activate cell death pathways.


Mutations of tumor suppressor genes cause loss of control upon important pathways, which may again lead to unregulated cell proliferation.


Oncogenes and tumor suppressor genes can be affected genetically by mutations on the DNA or also switched on or off epigenetically.


Cells are organized in complex cellular systems such as tissues or organs; therefore, it is crucial that they are able to communicate with each other. The most rapid way of communication is directly through cell-to-cell contact. There are various ways of direct interconnectivity of cells as shown in Table 3.14.Table 3.14
Summary of the size properties of the three main direct cell connections

Type of connection

Diameter

Length

Gap junctions
Tunneling nanotubes
Epithelial bridges

2–3 nm
50–1500 nm
1–20 μm

2–4 nm
Few to >100 μm
25–1000 μm


The most famous type of cell-to-cell connection is gap junction, which is the most direct manner of cell interconnectivity and forms the fastest communication channel. Gap junctions have a pore diameter of 2–3 nm and a length of 2–4 nm and are involved in the exchange of nutrients, ions, second messengers, and small metabolites up to ~1 kDa, allowing ionic and biochemical coupling between neighboring cells. These specialized structure membranes have a short half-life of a few hours (~1–4 h), and their biosynthesis and assembly are firmly regulated [163]. These transmembrane structures are composed of connexons (Fig. 3.45) constituted of six connexin (Cx) subunits around a central pore, which allow communication between adjacent cells. These connexons could be made up of six similar Cx isoforms (homomeric) or a combination of six different Cx isoforms (heteromeric). To date, 21 Cx isoforms have been identified in human proteosome, each named according to its approximate molecular weight (in kDa), with Cx43 being the most studied till now [163]. According to electron microscopy analyses, all Cx share a common topology composed of four transmembrane proteins, with a cytoplasmatic C- and N-terminal domains, two extracellular loops, and an intracellular loop. In contrast to the transmembrane proteins and the extracellular loop which are highly conserved among the Cx family members, the intracellular loop and the C- and N-terminal showed high variability in terms of the length and amino acid sequence of each Cx. Thus, these regions play an important role in the modulation of the gap junction channel gating and in the intracellular trafficking of connexins, and consequently a variety in their biological roles and interactions [163].

A schematic representation depicts the structural organization of connexin, homomeric connexon, heteromeric connexon, homotypic, and heterotypic gap junction channels in cells.

Fig. 3.45
Connexins and gap junctions. Each connexin (a) consists of four transmembrane domains. Six connexins form a hexameric torus called connexon (b). Depending on the composition, connexons are called homomeric (six equal connexins) or heteromeric (up to six different connexins). (c) When the cells form direct contact, the connexons stick together forming gap junctions. Here, the differentiation is made between homotypic channels (both connexons are the same) and heterotypic channels (different connexons). (Reproduced with permission (CCBY) from Totland et al. [163])


A schematic representation depicts the structural organization of connexin, homomeric connexon, heteromeric connexon, homotypic, and heterotypic gap junction channels in cells.


The spatial arrangements of Cx43 in breast cancer cells, fibroblasts, and internal mammary artery endothelial cells were studied by CLSM and super-resolution localization microscopy [164]. After radiation treatment (50 min postirradiation with a dose of 4 Gy), these cells behaved differently concerning the trafficking and response of Cx43. In breast cancer cells, high accumulations of Cx43 were found in the cytosol and along the membrane. The results did not significantly differ between non-treated and irradiated cells. In contrast to that, normal fibroblasts and endothelial cells revealed differences at the membrane and in the perinuclear cytosol after radiation exposure. In endothelial cells, a significant Cx43 accumulation and condensation were observed in the perinuclear region, whereas at the membrane, a signal reduction was found. In fibroblasts, Cx43 accumulations were found in the perinuclear region but also at the membrane.


Furthermore, as the Cx are phosphoproteins, they also play an important role in modulating the physiological properties and regulation responses of the channels, such as differentiation process, neuronal activity, development, cell synchronization, and immune response. Therefore, the presence of mutation in these structures is associated with several human diseases, such as neurodegenerative and skin diseases, deafness, and developmental abnormalities [165]. Also, gap junctions have been described as having a selective permeability, dependent on the combination of Cx isoforms that are made, conferring a single gating, conductance, and permeability to specific molecules, which could allow the association of each channel to a specific disease.


Another type of intercellular communication is via membrane connections such as tunneling nanotubes (TNTs) and epithelial (EP) bridges, which can be distinguished through their structural composition. These connections serve as direct signaling path when cells are separated by greater distances, than necessary for gap junctions. A microscopic image of both connection types can be found in Fig. 3.47.


TNTs are thin cytoplasmic membrane bridges, which appear in straight lines in vitro but also with a curved shape in tissue or in vitro cultures in a three-dimensional extracellular matrix found in various mammalian cells [166]. Their diameter ranges from 50 nm up to 1.5 μm, and they can contact cells over long distances up to several cell diameter length. Even if an obstacle blocks the direct distance between two cells, TNTs, due to their flexible structure, can form a connection. The length of the TNTs dynamically varies when cells migrate up to a certain distance of several 100 μm, which is too large to keep the structure, and the tube disappears. The detailed structure of TNTs is very complex and not yet known in detail. Most TNTs consist of F-actin, and the thicker ones additionally contain microtubules and cytokeratin filaments. Further compounds are sequentially identified as more and more information about the responsibility of TNTs is gathered. TNTs are proven to serve as a highway for exchange of cellular compounds such as mitochondria, vesicles, and many more. Larger compounds are mainly transported along TNTs in so-called gondolas (see Fig. 3.46). Furthermore, TNTs play a key role in direct and active signal transduction including calcium and electric signals, which are known to occur in cells due to radiation stress. Overall, it can be said that the frequency of occurrence and also the complexity of TNT networks within a cell composite are connected to the stress this composite is exposed to. Under stress conditions, the networks are intensified, so that signal and compound exchange is enhanced and fastened. Furthermore, the TNT networks were identified to play a role in the bystander and also the rescue effect and other effects related to radiotherapy [166].

A micrograph highlights a thin intercellular membrane connected by a T N T with a gondola at its end and a thick epithelial bridge.

Fig. 3.46
Membrane connections. Microscopic image of membrane label of cells connected by a tunneling nanotube transporting a gondola and an epithelial bridge containing vesicles and cytoplasmic material. Scale bar: 10 μm. EP epithelial, TNT tunneling nanotube


A micrograph highlights a thin intercellular membrane connected by a T N T with a gondola at its end and a thick epithelial bridge.


In contrast to TNTs, EP bridges could, as also the name suggests, only be found in normal as well as cancerous human epithelial cells. They also differ from TNTs structurally, as they show a larger diameter of 1–20 μm and also a larger range from 25 μm to over a millimeter [166]. EP bridges consist of F-actin as well as microtubules, which promotes the structural stability allowing these connections to bridge such large distances. As TNTs, the EP bridges play a major role in cellular compound and signal transduction (Box 3.29).


Cells communicate through direct cell-to-cell contact and for interconnectivity networks.

Gap junctions, constituted by connexins, allow short-range ionic and biochemical coupling.

TNTs and EP bridges are responsible for long-range signal and molecule transduction.

Direct cellular communication plays a role in various diseases, spreading of pathogen and health signals, as well as stress and radiation response of cell composites.


Gap junctions, constituted by connexins, allow short-range ionic and biochemical coupling.


TNTs and EP bridges are responsible for long-range signal and molecule transduction.


Direct cellular communication plays a role in various diseases, spreading of pathogen and health signals, as well as stress and radiation response of cell composites.


Since inflammation that can be induced by microbial infections and tissue damage is an essential mechanism of innate immune response, the terms “inflammation and immunity” are intrinsically linked [167]. The process of inflammation includes several biochemical events and multi-level cellular interrelationships. In a concerted action, inflammation is initiated, propagated, matured (effector phase), and finally resolved. This implies that radiation exposure under inflamed conditions affects several cell types including many immune cell (sub)types. Macroscopically, vasodilatation and extravasation of immune cells into the inflamed tissue occur that in sum results in the key characteristics of inflammation, namely swelling, redness, pain, loss of function, and increased temperature. The major immune cells involved in the inflammatory process are polymorphonuclear neutrophils (PMNs), which are the most abundant leukocytes in peripheral blood and are very quickly recruited to sites of inflammation, mononuclear monocytes that can differentiate into dendritic cells (DCs) and macrophages, and different subtypes of B and T lymphocytes mediating an antigen-specific adaptive immune response.