Telomeres, interstitial telomeric repeat sequences, and chromosomal aberrations

Mutation Research 612 (2006) 189–214 www.elsevier.com/locate/reviewsmr Community address: www.elsevier.com/locate/mutres

Review

Telomeres, interstitial telomeric repeat sequences, and chromosomal aberrations Alejandro D. Bolza´n *, Martha S. Bianchi Laboratorio de Citogene´tica y Mutage´nesis, Instituto Multidisciplinario de Biologı´a Celular (IMBICE), C.C. 403, 1900 La Plata, Argentina Received 17 October 2005; received in revised form 29 December 2005; accepted 30 December 2005 Available online 21 February 2006

Abstract Telomeres are specialized nucleoproteic complexes localized at the physical ends of linear eukaryotic chromosomes that maintain their stability and integrity. The DNA component of telomeres is characterized by being a G-rich double stranded DNA composed by short fragments tandemly repeated with different sequences depending on the species considered. At the chromosome level, telomeres or, more properly, telomeric repeats – the DNA component of telomeres – can be detected either by using the fluorescence in situ hybridization (FISH) technique with a DNA or a peptide nucleic acid (PNA) (pan)telomeric probe, i.e., which identifies simultaneously all of the telomeres in a metaphase cell, or by the primed in situ labeling (PRINS) reaction using an oligonucleotide primer complementary to the telomeric DNA repeated sequence. Using these techniques, incomplete chromosome elements, acentric fragments, amplification and translocation of telomeric repeat sequences, telomeric associations and telomeric fusions can be identified. In addition, chromosome orientation (CO)-FISH allows to discriminate between the different types of telomeric fusions, namely telomere–telomere and telomere–DNA double strand break fusions and to detect recombination events at the telomere, i.e., telomeric sister-chromatid exchanges (T-SCE). In this review, we summarize our current knowledge of chromosomal aberrations involving telomeres and interstitial telomeric repeat sequences and their induction by physical and chemical mutagens. Since all of the studies on the induction of these types of aberrations were conducted in mammalian cells, the review will be focused on the chromosomal aberrations involving the TTAGGG sequence, i.e., the telomeric repeat sequence that ‘‘caps’’ the chromosomes of all vertebrate species. # 2006 Elsevier B.V. All rights reserved. Keywords: Telomere; Interstitial telomeric repeat sequences; Telomere dysfunction; Chromosomal aberrations; Fluorescence in situ hybridization (FISH); Chromosome orientation-FISH (CO-FISH)

Contents 1.

What are telomeres?. . . . . . . . . . . . . . . . . 1.1. Definition . . . . . . . . . . . . . . . . . . . 1.2. Structure . . . . . . . . . . . . . . . . . . . . 1.2.1. Basic structure of telomeres. 1.2.2. Telomere chromatin . . . . . .

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* Corresponding author. Tel.: +54 221 4210112; fax: +54 221 4253320. E-mail address: [email protected] (A.D. Bolza´n). 1383-5742/$ – see front matter # 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.mrrev.2005.12.003

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1.2.3. Telomere replication . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.4. Telomere length . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.5. Telomere loss. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.6. Telomere loss is usually prevented by a specialized enzyme . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.7. Chromosome healing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.8. ‘‘Telomere capture’’ and beyond . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.9. Alternatives to telomerase: the ‘‘ALT’’ mechanisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.10. The ‘‘other’’ telomeres . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3. Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3.1. Main role of telomeres . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3.2. Additional functions of telomeres . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3.3. Telomere function and DNA repair factors are associated . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3.4. Sometimes telomeric repeats are located at interstitial sites of the chromosomes . . . . . . . . . . . . Telomeres and FISH technologies: detection of telomeric sequences . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Techniques for telomere detection and telomere length assessment. . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Conventional FISH to detect telomeric repeats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. The ‘‘PRINS’’ alternative . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. PNA-FISH. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5. PRINS versus PNA-FISH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6. The ‘‘CO-FISH’’ technique . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7. Some cells show no telomeric signals at terminal regions of chromosomes . . . . . . . . . . . . . . . . . . . . . . Telomeres, ITRs, and chromosomal aberrations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Different types of chromosomal aberrations involving telomeres and ITRs . . . . . . . . . . . . . . . . . . . . . . 3.1.1. Types of chromosomal aberrations involving telomeres and ITRs. . . . . . . . . . . . . . . . . . . . . . . 3.1.2. Aberrations involving telomeres . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.3. Aberrations involving ITRs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Induction of chromosomal aberrations involving telomeres and ITRs by physical and chemical mutagens. 3.2.1. Induction of chromosomal aberrations involving telomeres . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.2. Induction of chromosomal aberrations involving ITRs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Future prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. What are telomeres? 1.1. Definition Telomeres or the ends of linear eukaryotic chromosomes, were first described almost 70 years ago since the pioneering studies of the geneticists Hermann Joseph Muller and Barbara McClintock in the fruit fly Drosophilia melanogaster and maize, respectively [1,2]. Muller observed that the ends of chromosomes rarely interacted with breaks that resulted from ionizing radiation, i.e., X-ray-induced chromosomal aberrations never included deletions or inversions involving the terminal regions of the chromosomes. Thus, he proposed that chromosome ends are specialized structures that he coined ‘‘telomeres’’, from the Greek, telo = end, and mere = part [1]. The concept of ‘‘telomere’’ meant not only the physical ends of the chromosome itself but also in Muller’s words ‘‘a terminal gene with a special function, that of sealing the

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end of the chromosome’’ [1]. Summarizing all of his work in a classic lecture in 1938 at the Woods Hole Marine Biological Laboratory, he also concluded that ‘‘. . . for some reason a chromosome cannot persist indefinitely without having its ends thus sealed’’ [1]. Also by the end of 1930s, Barbara McClintock, studying chromosomal aberrations induced by X-rays in maize, found that broken chromosomes frequently fused to their sister chromatids, creating breakagefusion-bridge (BFB) cycles, which were always accompanied by the loss of the terminal regions at the fusion site, demonstrating that broken chromosomes (i.e., without ‘‘end caps’’) were subject to fusion events ([2–4], see also Refs. [5,6] for a more detailed description of Muller and McClintock’s experiments). In BFB cycles, a chromatid break occurs, exposing an unprotected chromosomal end that, after replication, is thought to fuse with either another broken chromatid or its sister chromatid to produce a dicentric chromosome. The fused chromatids then form a bridge that breaks

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during the anaphase stage of mitosis, when the two centromeres are pulled to opposite poles. Following DNA replication in the next cell cycle, the sister chromatids fuse once again, and thus the cycle continues [2–4,7]. Barbara McClintock distinguished normal chromosome ends from abnormal ends by their absence in BFB cycles. In conclusion, Muller and McClintock’s studies demonstrated that chromosomes with broken ends were unstable, leading to BFB cycles, and revealed resistance of intact chromosomes to fusion with either chromosomal fragments or other chromosome ends. These observations lead to the idea that the ends of chromosomes or telomeres were ‘‘capped’’ and therefore protected from fusion reactions characteristic of ends created by chromosome breakage events. In this way, telomeres were defined as the terminal regions or physical ends of eukaryotic chromosomes, which protected them from fusion with either broken chromosomal fragments or other telomeres. Nowadays, in the light of molecular biology studies, telomeres are defined as specialized nucleoproteic complexes localized at the physical ends of linear eukaryotic chromosomes that maintain their stability and integrity [8]. Recently, the term ‘telomere capping’ emerged to describe the protective role of telomeres [9–12], i.e., telomeres provide a protective ‘‘cap’’ for chromosomal DNA against illegitimate recombination, exonucleolytic attack and degradation, and oxidative damage. Thus, ‘‘capping’’ refers to the ability of telomeres to protect chromosome ends from DNA damage responses. It is now well established that telomere capping function is facilitated by several telomere-repeat binding factor proteins [13–15] characterized by their affinity for telomeric DNA, and DNA damage response proteins which do not directly interact with telomeric DNA but rather with telomere binding proteins [14–18]. We will further analyze telomere functions later in this section. 1.2. Structure 1.2.1. Basic structure of telomeres As mentioned before, telomeres are nucleoproteic complexes [10,15,19,20]. Molecular dissection of telomeres started with the discovery of the telomeric DNA sequence of the ciliated protozoan Tetrahymena thermophila, (TTGGGG)n, by Blackburn and Gall in 1978 [21]. Telomeric DNA is characterized by being a G-rich double stranded DNA composed by short fragments tandemly repeated with different sequences depending on the species considered [22,23]. In all vertebrates, telomeres consist of tandem repeats of the hexanucleotide sequence (TTAGGG/

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CCCTAA)n and associated proteins [15–17,24–28]. The telomere-specific proteins complex associated to this telomeric sequence has been termed ‘‘shelterin’’ by de Lange [15] and includes six subunits, three of them that directly recognize TTAGGG repeats, i.e., TRF1, TRF2 and POT1, interconnected by three additional proteins named TIN2, TPP1 and Rap1. The arrays of TTAGGG repeats, first identified at human chromosome ends by Moyzis et al. in 1988 [28], are oriented 50 ! 30 towards the end of chromosomes [25,29] and form a 30 single-stranded G-rich overhang found at both chromosomal ends [30,31]. The C-rich telomere strand is at the 50 end and the G-rich telomere is at the 30 end of each chromosomal DNA strand. These single-stranded, G-rich 30 overhangs result from both the ‘‘end replication problem’’, i.e., the inability of DNA polymerase to replicate the very end of the telomeres, and postreplication processing [32,33]. Thus, removal of the most distal RNA that primes lagging-strand synthesis leaves an 8- to 12base gap at the 50 end that, if not filled in, leads to a small loss of DNA in each round of DNA replication (see Ref. [34] for review). Following replication, telomeres created by leadingstrand synthesis are either blunt-ended, or posses a small 50 overhang [35], whereas those created by the lagging-strand synthesis have a 30 overhang with a length determined by the position of the most terminal RNA primer [36]. The physical structure of the telomere was revealed by electron microscopy to be a large duplex loop [37– 39], which is created when a telomere’s end loops back on itself and the single-stranded overhang invade an interior segment of the duplex telomeric DNA. The t-loop structure of mammalian telomeres is thought to repress the nonhomologous end-joining (NHEJ) DNA double-strand breaks (DSBs) repair process at chromosomal ends, thus rendering telomeres nonrecombinogenic, since upon loss of theTRF2 protein NHEJ occurs [40]. The TRF2 protein binds the duplex telomere tract as a homodimer [16,17,41], protecting chromosome ends from fusion [13] and is required to remodel linear DNA into t-loops in vitro, perhaps facilitating invasion of the overhang [37]. Moreover, it has been recently demonstrated that the single-stranded telomeric DNA-binding protein POT1 determines the structure of the 30 and 50 ends of human chromosomes and its inhibition generates a novel combination of telomere dysfunction phenotypes in which chromosome ends behave transiently as sites of DNA damage, yet remain protected from NHEJ [42].

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1.2.2. Telomere chromatin Telomere chromatin contains nucleosomes but differs from bulk nuclear chromatin in having a nucleosome repeat length that is about 40 bp shorter [30,43,44]. It has been shown that, in nuclei, telomere DNA occurs in distinct circles or rings [45,46]. In yeast, telomeres are localized on the nuclear periphery [47], whereas in human cells, telomeres seem to be randomly positioned throughout the cell nucleus [45] or, as observed in T-lymphocytes, positioned within the interior 50% of the nuclear volume [48]. Recently, Amrichova´ et al. [49] analyzed the nuclear and territorial topography of chromosome telomeres in human lymphocytes and found that a majority of the polar chromosome territories are partially oriented with the centromere localized near the nuclear periphery and both telomeres placed in the interior of the cell nucleus. Little is known about the chromatin context of telomeres, because, in most cells, telomeres are strongly anchored within the nucleus, and remain with the insoluble ‘‘matrix’’ fraction after nuclease digestion [45,50]. However, Nikitina and Woodcock [51] reported very recently the successful isolation of telomere chromatin from chicken erythrocytes and quiescent mouse lymphocytes, and showed by electron microscopy that telomere chromatin fibers are organized in the form of closed terminal loops, which correspond to the ‘‘tloop’’ structure adopted by telomere DNA. Furthermore, Molenaar et al. [52] analyzed the telomere dynamics in living mammalian cells using a peptide nucleic acid (PNA) probe and showed that telomeres have the ability to associate with each other in a dynamic manner. This tendency to physically associate may account for the high incidence of chromosomal rearrangements in (sub)-telomeric regions found in somatic cells [53,54] and for the recombination-based interchromosomal telomeric DNA exchanges observed in cells which maintain telomeres by mechanisms not involving the enzyme telomerase [55] (see Section 1.2.9). 1.2.3. Telomere replication In mammalian cells, telomeres are early replicating and replicate throughout the S phase of the cell cycle [56–59]. Moreover, Zou et al. [60] using Indian muntjac fibroblast cell lines demonstrated that each telomere of a mammalian cell has a characteristic timing for replication during S phase, i.e., some specific telomeres replicate early in S phase, and p- and q-arm telomeres of the same chromosome replicate asynchronously. Thus, the timing and regulation of replication of mammalian

telomeres seems to be very different from that found in yeast, in which all telomeres replicate late in S phase, and the two telomeres at each end of the same chromosome replicate synchronously [61–63]. 1.2.4. Telomere length The length of the double stranded telomeric repeat varies greatly among species [8]. For example, in the ciliate Oxytricha, it is only 20 base pairs (bp) long [64] in S. cerevisiae, it is a few hundred bp long [65] while, in vertebrates, individual telomeres may extend to more than 100 kb, such as in some mouse cells. [66]. In normal human cells, the DNA at each chromosome terminus spans 5–20 kb in length [28], terminating in a 30 single-stranded overhang 100–400 nt in length [36]. In human tumor cells which use telomerase for telomere maintenance (see below), telomere length varies from 1 to 20 kb [67–69]. It has been shown that in humans and mice, the length of telomere repeats at individual chromosome ends in individual cells is highly variable [70–74] and that mouse and human cell lines exhibit subpopulations of cells with different telomere lengths [75]. The longest human telomere is found at the long arm of chromosome 4, whereas the shortest one is at the short arm of chromosome 17 [76]. It was shown that a stable hierarchy exists in the form of a telomere length profile of the human karyotype [77]. This rank order is conserved between different human cell types and individuals, maintained throughout a lifetime, and seems to be genetically determined [77,78]. The telomere length profile seems to be stable, even under the influence of mutagens like bleomycin (BLM) or mitomycin C [77]. 1.2.5. Telomere loss The telomeres lose approximately 50–200 bp of repeat sequences every cell division mainly due to the so-called ‘‘end replication problem’’ [32,33] in which the DNA replication machinery is not able to completely replicate the ends of linear chromosomes (telomere erosion) because the overhang has no template strand to guide its synthesis. Oxidative damage within telomeric DNA [79,80], chromosome endspecific exonuclease activity [30] and the lack of telomerase activity [81] may also contribute to telomere loss. Accordingly, it has been observed that radiosensitive cells exhibit accelerated telomere shortening and telomere abnormalities [82]. Telomere loss can generate chromosome instability through the abovementioned BFB cycles [2] producing several types of chromosome rearrangements, including terminal deletions, inverted duplications, DNA amplification, dupli-

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cative and non-reciprocal translocations, and dicentric chromosomes, all of which have been associated with human cancer (see Refs. [7,83] for review). Recently, Lo et al. [84] demonstrated that even DSBs occurring near telomeres can result in complex chromosome aberrations and chromosome instability, which can be prevented or terminated by the addition of telomeric repeats to the end of the broken chromosome. 1.2.6. Telomere loss is usually prevented by a specialized enzyme The loss of telomeric repeats is usually prevented by telomerase, a specialized reverse transcriptase-like enzyme, containing a RNA subunit and a catalytic protein subunit called telomerase reverse transcriptase (TERT) which is the rate-limiting factor for the enzyme activity. This enzyme, first discovered by Greider and Blackburn in 1985 in Tetrahymena [85], works via an RNA template – using exclusively single-strand 30 telomeric overhangs as primers [10,19,24,30,86,87] – by adding TTAGGG repeats to the telomere. Although repressed in the majority of normal somatic cells (with the exception of a transient S phase activity thought to maintain the single-stranded overhang [88]), telomerase is present in immortal cell lines, germline cells, stem cells, activated lymphocytes, and most of the tumor cells analyzed so far [67–69,89–91]. Telomerase activity favors 30 overhangs over blunt DNA ends for addition of telomere sequence, at least in vitro [92,93]. Telomere length is controlled by a mechanism involving telomerase and the telomere-binding proteins [24,94]. Loss of telomerase enzymatic function leads to progressive telomere shortening over time, eventually resulting in the disappearance of detectable telomeric DNA and the formation of end-to-end chromosome fusions, followed by growth arrest or cell death [19,95]. It has been shown that overexpression of the TRF2 protein favors telomere shortening in cultured human and murine cells [96,97], and that overexpression of telomerase activity does not prevent proliferation-associated telomere shortening in human hematopoietic cells [98]. This latter finding points to the existence of cell-specific differences in telomere maintenance. 1.2.7. Chromosome healing Besides maintaining pre-existing telomeres, telomerase can catalyze the addition of telomeric sequences directly on to non-telomeric DNA [99]. This process of direct addition of telomeric repeats to the ends of broken chromosomes by telomerase is called ‘‘chromosome healing’’ (a term coined by Barbara McClintock

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in 1941 to describe the phenomenon that halted the BFB cycles in the embryo of plants [2]), and has been observed in protozoans, yeast, plants, insects, and mammals [100–107]. Repair function of telomerase would lead to uncontrolled chromosomal fragmentation and karyotypic instability, because chromosome healing prevents repair of broken ends. Therefore, telomerase must be prevented from accessing internal DSBs. Slijepcevic and Al-Wahiby [108] proposed that Ku, a DSB protein which has a high affinity for DNA ends, acts to prevent telomerase from accessing internal DSBs. This model is supported by the fact that the efficiency of chromosome healing is extremely low, about 1% [109]. In fact, no evidence of ‘‘chromosome healing’’ was found in normal human lymphocytes [110] or in ataxia–telangiectasia cells [111], which have difficulties in rejoining the ends of broken chromosomes, exposed to ionizing radiation. The failure to recruit and/or activate telomerase at sites of DSB may contribute to the paucity of chromosome healing events. 1.2.8. ‘‘Telomere capture’’ and beyond Lost telomeres in broken chromosomes can also be acquired by ‘‘telomere capture’’ and break-induced replication. Telomere capture is a process which involves the addition of telomeres at the site of DSB by subtelomeric cryptic translocations, undetectable by classical cytogenetic techniques [84,106–108,112]. In telomere capture, broken chromosomes are stabilized by the transfer of telomeres from normal chromosomes. This phenomenon was first reported in human malignant melanoma cells [112], and has been recently observed in the leukocytes of chronic lymphocytic leukemia and chronic myeloid leukemia patients [113]. Telomere capture is essentially a non-reciprocal process, producing a chromosome with only one telomere (donor chromosome) and another one with a new telomere (recipient chromosome) plus an acentric fragment (a terminal fragment or terminal deletion, as seen by telomeric FISH – see Section 3.1 – if the chromosome break occurs at G0/ G1/early S phase, or a chromatid-type fragment, if the break takes place in late S/G2). However, the donor chromosome may be involved in secondary recombination events [107,112,114] by the initiation of BFB cycles in this chromosome. Slijepcevic et al. [114] showed that only a small percentage of radiation-induced chromosome/chromatid breaks may be modified by ‘‘telomere capture’’. In break-induced replication [115,116], the broken end of a chromosome invades a region of homology and initiates replication, thereby duplicating the end of that chromosome. Although not involving telomere restora-

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tion, the formation of ring or dicentric chromosomes can also compensate for telomere loss [117–119].

are composed of short tandem repeats have been found in different species [23].

1.2.9. Alternatives to telomerase: the ‘‘ALT’’ mechanisms An alternative method of telomere elongation in the absence of telomerase has been described in several tumor cells and immortalized cell lines and named ‘‘ALT’’ (for ‘‘alternative lengthening of telomeres’’) [120–133], although some tumors were found to posses neither telomerase activation nor ALT mechanisms for telomere length maintenance [121,134]. ALT cells are characterized by absence of telomerase, extreme heterogeneity of telomere length [125], high rates of telomeric exchange [135], and the presence of ALTassociated promyelocytic leukemia bodies [52,122] that contain extrachromosomal telomeric circles (t-circles) [131], telomere-specific binding proteins, and proteins involved in DNA recombination and replication [136– 138]. It has been recently shown that ALT-associated promyelocytic leukemia bodies are not always essential for ALT-mediated telomere maintenance [132,133]. Moreover, lack of expression of the telomerase gene in ALT cells is associated with histone H3 and H4 hypoacetylation and methylation of Lys9 histone H3 [139]. ALT is thought to occur as a recombinationmediated lengthening of telomeres [55,122,123,135,140, 141]. Very recently, Nosek et al. [142] presented evidence that t-circles in a natural telomerase-deficient system of mitochondria of the yeast Candida parapsilosis replicate independently of the linear chromosome via a rollingcircle mechanism and suggested that extrachromosomal t-circles observed in a wide variety of organisms, including yeasts, plants, Xenopus laevis, and certain human cell lines, may represent independent replicons generating telomeric sequences and, thus, actively participating in telomere dynamics.

1.3. Function

1.2.10. The ‘‘other’’ telomeres The findings that Drosophila melanogaster telomeres comprise head-to-tail arrays of non-LTR retrotransposons (see Ref. [143] for a review) instead of simple DNA repetitive sequences, and that in the silkworm Bombyx mori – which expresses very low levels of telomerase activity in all stages of various tissues – the characteristic TTAGG motifs of insect telomere are associated to non-LTR retrotransposons [144], showed that in some organisms there are alternatives to telomerase for telomere elongation and that end protection is not always dependent on the presence of short repeats. Since this discovery many exceptions to the general rule that the chromosome ends

1.3.1. Main role of telomeres The main role of telomeres is to preserve the integrity of the chromosomes, protecting them from degradation, recombination or fusion [145] by preventing the ends of linear chromosomes from being recognized as DSB by the DNA repair machinery, i.e., they distinguish natural DNA ends from DNA ends resulting from breakage events [1,2]. Thus, telomeres prevent inappropriate repair and recombination between internal DNA breaks and native chromosomal or the ligation of chromosomal ends. Accordingly, when telomeres become dysfunctional, fusions between two telomeres and a telomere and a DSB occur, as shown by recent studies using the chromosome orientation (CO)FISH technique [146,147]. In mice, one of the consequences of impaired telomere function is the formation of Robertsonian-like chromosome fusions [148–150] (see below). Therefore, the maintenance of telomere function is crucial for genomic stability and cell viability. Cells respond to dysfunctional telomeres by undergoing senescence, cell death, or genomic instability [150–163]. Cellular response to dysfunctional telomeres is governed by proteins that also control the DNA damage response [15,17,23,164,165]. The domain of telomere-associated DNA damage factors has been termed telomere dysfunction-induced focus or ‘‘TIF’’ by Takai et al. [164] who showed that DNA damage foci form at telomeres uncapped by TRF2 inhibition, and that uncapping of telomeres occurs in late S/G2. Several studies point to the possibility that telomere maintenance may constitute a potential genetic marker of radiosensitivity, i.e., it seems that telomere maintenance and radiosensitivity are linked phenomena (see Ref. [166] for a review). Thus, telomere integrity depends on the ability to maintain telomere length and/ or the ability to mask telomeres from being recognized as damaged DNA. 1.3.2. Additional functions of telomeres Besides the above-mentioned functions, telomeres regulate the replicative life span of somatic cells, contribute to maintenance of chromosome topology in the cell nucleus and play a fundamental role in the proper alignment of chromosomes for recombination during the first meiotic prophase [8,30,45,49,50,151, 167].

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An outstanding feature of telomeres is that they silence genes flanking the telomere repeat sequence [168–170]. This phenomenon, called the ‘‘telomere position effect’’, is thought to be modulated by telomere length and local heterochromatin structure [171–173]. It has been shown that mutations in the encoding genes for Ku proteins lead to disruption of nuclear organization of telomeres and loss of telomeric silencing [174,175]. Moreover, in mammalian cells, loss of Ku leads to aberrant telomere–telomere fusions [176,177]. On the other hand, a new mechanism by which short telomeres inhibit tumorigenesis through interference with oncogenic translocations has been recently described in mice which are null for both ataxia– telangiectasia-mutated and telomerase RNA [178]. 1.3.3. Telomere function and DNA repair factors are associated Several studies have shown that DNA repair factors and telomere function are associated in a way that loss of telomere function – either by loss of function of telomere-binding proteins or by loss of telomeric repeats – causes activation of DNA damage response [14,108,164,179–182]. In effect, to maintain their integrity, telomeres bind many proteins involved in DSB repair (the DNA-PK NHEJ complex including DNA-PKcs, Ku70-86 and Artemis; ATM, the MRN complex, and RAD51D) and mutations in genes involved in signaling DNA damage affect telomere stability [10,12,16–18,108,149,183–185]. Thus, the presence of uncapped telomeres induces a response similar to that observed with DNA breaks [96,108,156,164,177,186–194] and growing evidence suggests a dual role of DNA damage response proteins in the protection of chromosome ends and the ability to promote cell-cycle arrest in response to dysfunctional telomeres [24,108,176,195,196]. 1.3.4. Sometimes telomeric repeats are located at interstitial sites of the chromosomes By definition, telomeric repeats are located at the very ends or terminal regions of chromosomes. However, some vertebrate species show large blocks of (TTAGGG)n repeats present in non-terminal regions of most chromosomes, the so-called interstitial telomeric repeat sequences or ITRs, which includes those repeats located close to the centromeres and those found at interstitial sites, i.e., between the centromere and the telomere [27,197–201]. In Chinese hamster, ITRs comprises 250–500 kb of DNA on each chromosome [202]. Several studies revealed that even human chromosomes contain different kinds of ITRs [203–

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205], most of them probably originated through the repair of interstitial DSBs that occurred in the germ line during evolution, as suggested by Azzalin et al. [205]. Therefore, the location of telomeric sequences does not always correspond to that of the very ends of chromosomes, i.e., the authentic telomeres. The presence of (TTAGGG)n repeats on non-telomeric regions of the chromosomes has been assumed to be the result of tandem chromosome fusion (telomere– telomere fusions) during evolution [27,206,207] or the insertion of telomeric DNA within unstable sites during the repair of DSB [205]. This later possibility is supported by the finding that some relatively small ITRs appear to be flanked by AT-rich DNA sequences, which could be unstable [208]. It has been shown that ITRs do not represent a functional telomere [27]. An exception is represented by an Indian Muntjac cell line, where in a small percentage of cells, ITRs get amplified and chromosomes fall apart into many small fragments with functional telomeres on most chromosome ends, as demonstrated by Zou et al. [209]. Moreover, unlike terminal telomeric sequences, ITRs seem not to be associated with the nuclear matrix [210]. 2. Telomeres and FISH technologies: detection of telomeric sequences 2.1. Techniques for telomere detection and telomere length assessment At the chromosome level, telomeres or, more properly, telomeric repeats – the DNA component of telomeres – can be detected either by using the Fluorescence in situ hybridization (FISH) technique with a DNA or a peptide nucleic acid (PNA) (pan)telomeric probe, i.e., which identifies simultaneously all of the telomeres in a metaphase cell, or by the primed in situ labeling (PRINS) reaction using an oligonucleotide primer complementary to the telomeric DNA repeated sequence. 2.2. Conventional FISH to detect telomeric repeats The conventional FISH technique has been successfully employed for in situ detection of telomeric repeat sequences in chromosomes of various vertebrate species, using synthetic oligonucleotide probes (TTAGGG or CCCTAA), but the efficiency of these probes has not been sufficient to extend this procedure beyond qualitative analysis of telomeric repeat sequences or even to detect all human telomeres. In

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effect, the use of conventional FISH with a telomeric DNA probe alone or in combination with whole chromosome-specific DNA probes allowed for the detection of 75–82% of telomeres in human lymphocytes [211,212]. 2.3. The ‘‘PRINS’’ alternative The PRINS technique [213–215] has offered a fast alternative method than conventional FISH for the in situ labeling of nucleic acids, including telomeric sequences [216–223]. This technique is based on the in situ annealing of synthetic oligonucleotides (CCCTAA)7 for telomere detection [216,217] to complementary nucleic acid sequences followed by primer extension in the presence of a hapten- or fluorochrome-labeled nucleotide [213]. The telomeric fluorescent signal is produced in situ following the incorporation of a labeled nucleotide into the newly synthesized DNA strand. It has been demonstrated that the omission of dGTP in the PRINS reaction solution prevents non-specific elongation of the primer, since the elongated telomeric DNA sequence lacks this base, and thus only the telomeric sequences are detected [216–221]. PRINS provided superior efficiency to conventional FISH for the detection of telomeric sequences [222,223] since, unlike PRINS reaction, the conventional FISH approach uses a fluorescently labeled probe to hybridize to the TTAGGG telomeric sequence, generating a high level of non-specific hybridization, sometimes making the results difficult to analyze. 2.4. PNA-FISH PNAs are synthetic DNA mimics in which the sugar phosphate backbone is replaced by an uncharged and flexible polyamide backbone, and are particularly resistant to protease and nuclease degradation [224– 226]. Because of its neutral backbone, which allows the probe to penetrate into the chromosome rather than only bind to the surface of it, a telomeric PNA probe provides a higher and much better efficiency in the detection of (TTAGGG)n repeats than conventional FISH with a DNA probe [70,224–226]. In addition, PNAs are highly resistant to degradation by DNases, RNases, proteinases and peptidases [224–226]. The PNA-FISH technique has been extensively used to detect telomeric repeats in human and other vertebrate cells, yielding detection efficiencies of 100% of human telomeres [70,71,110,201,227–229]. In addition, PNA-FISH can be used as an alternative to Southern Blot (terminal restriction fragment or TRF analysis) (see [230,231] for review) to assess the length of telomeric repeats at

individual chromosomes, i.e., quantitative telomere analysis using the quantitative (Q)-FISH technique [70,71,73,76,77,80,231–235]. The Q-FISH technique provides estimate of telomere length in each individual chromosome with a resolution of 200 bp and may be used to estimate telomere length in species containing ITRs, like Chinese hamster [200,233]. Other recently developed techniques to analyze telomere length include flow-FISH (flow cytometry + FISH) and single telomere length analysis or ‘‘STELA’’ [235–240]. It has been suggested by Lavoie et al. [220] that PRINS alone or in combination with FISH may also be used for quantitative analysis of telomeres [238,241,242]. 2.5. PRINS versus PNA-FISH Since both, PRINS and PNA-FISH techniques were shown to be much more specific and sensitive than conventional FISH for in situ detection and sizing of telomeric repeated sequences, they were compared each other in their efficiency for telomere detection on mouse, hamster and human cell lines. Both staining efficiency and sensitivity appeared to be identical [243], although for the detection of human telomeres PNAFISH is more appropriate [223], whereas for the detection of mouse telomeres the PRINS technique seems to be more suitable than PNA-FISH [220]. As pointed out by Lavoie et al. [220], the key advantages of PRINS for telomere detection are the intensity and clarity of the signals and the speed of the protocol. Also, the PRINS technique is more cost effective than PNAFISH, and when using a fluorochrome-labeled nucleotide, PRINS takes less than 30 min to be completed, and the conditions used for the PRINS reaction prevent the telomeric DNA from renaturing. Nevertheless, the availability of commercial kits to perform a fast (about 2 h) and very sensitive PNA-FISH procedure with a directly labeled TTAGGG probe, where the fluorescence intensity of the spots is directly correlated to the length of the telomeres [70,71,76,77,244,245] makes this technique the preferred method to currently analyze the chromosome ends of vertebrate cells, specially human cells, which possess short telomeres compared with other vertebrate species [70,76]. 2.6. The ‘‘CO-FISH’’ technique To discriminate between the different types of telomeric fusions and to detect telomeric sisterchromatid exchanges (T-SCE) (see Section 3.1) the CO-FISH technique must be used [141,146,246]. This technique, first described by Goodwin and Meyne in

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1993, is a strand-specific hybridization technique commonly used to deduce the orientation of sequences along the chromosomes [247]. In the CO-FISH procedure, cells are grown in the presence of 5bromodeoxyuridine (BrdUrd) and/or bromodeoxycytidine (BrdC) for a single round of replication (a single S phase), so that sister chromatids are unifilarly substituted. After culture, routine chromosome preparations made by standard cytogenetic techniques are exposed to UV light and exonuclease III treatment. Exposure of cells to UV light in the presence of the photosensitizing DNA dye Hoechst, results in numerous strand breaks that occur preferentially at the sites of BrdU/BrdC incorporation. Nicks produced in the chromosomal DNA by this treatment then serve as selective substrates for enzymatic digestion by Exo III. This results in the specific removal of the newly replicated strands while leaving the original (parental) strands largely intact. Thus, at the end of the procedure, chromatids are single-stranded and, when using a single-stranded DNA or PNA probe, only one chromatid will show a hybridization signal if the tandem repeats are oriented head-to-tail along the DNA strand [246,247]. Accordingly, while standard FISH or PRINS with a telomeric probe produces four signals, one on each end of the two chromatids of a mitotic chromosome, the strand-specific nature of CO-FISH typically yields just two signals, one at each end of the chromosome (Fig. 1I). Because replication of telomeric DNA begins at internal origins and proceeds towards the ends of the chromosomes, CO-FISH hybridization patterns produced with G-rich telomere probes [i.e., labeled (TTAGGG)n oligomeres] specifically identify telomeres produced by leading-strand synthesis, whereas hybridization with the complementary C-rich probes [i.e., labeled (CCCTAA)n oligomeres] identify ‘‘lagging-strand telomeres’’ [246]. 2.7. Some cells show no telomeric signals at terminal regions of chromosomes Some immortalized cells lines like CHO, show no telomeric signals at terminal regions of chromosomes [199,248], even when a technique as sensitive as PNAFISH is applied to detect the telomeric repeats in these cells [201] The absence of signal at the terminal regions of CHO chromosomes has been ascribed to the fact that the telomeric sequences are either completely lost from the chromosomal ends or, more likely, that these sequences are present in a copy number that is too low to be effectively detected by FISH [197–199], due to extensive loss of telomeric sequences occurring during proliferation

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and immortalization of these cells in vitro. In fact, all immortalized Chinese hamster cell lines that have been analyzed to date have extremely short telomeres, estimated to be about 1 kb long [199,200,248], whereas the average telomere length in vivo in the Chinese hamster is about 38 kb [200]. It is noteworthy to mention that, despite their very short telomeres, chromosomes from immortalized Chinese hamster cell lines do not show a high frequency of end-to-end chromosome fusions [248]. This finding suggests that a few telomeric repeats present at the chromosomes from immortalized Chinese hamster cell lines are sufficient for telomere function. 3. Telomeres, ITRs, and chromosomal aberrations 3.1. Different types of chromosomal aberrations involving telomeres and ITRs 3.1.1. Types of chromosomal aberrations involving telomeres and ITRs There are several types of chromosomal aberrations involving telomeres and ITRs that can be identified using molecular cytogenetics techniques, as listed in Table 1. In order to identify each of these aberrations, conventional FISH or PRINS with a telomeric DNA or PNA probe alone or in combination with a pancentromeric and/or a painting probe can be used, depending on the type of aberration to be scored and the cell type to be analyzed (Table 1). For instance, to accurately identify incomplete chromosomes and acentric fragments in human cells – whose centromeres are sometimes difficult to visualize after a FISH procedure without the use of a pancentromeric probe – a combination of telomere and centromere probes must be used [228,249]. In addition, to further discriminate between terminal deletions and incomplete exchanges (see Section 3.1.2.1) a combination of telomeric, centromeric, and painting probes (for a few chromosomes or the entire set of human chromosomes in a multicolor FISH procedure) is needed [110,111,229,250,251]. Furthermore, to discriminate between the different types of telomeric fusions and to detect T-SCE, the above-mentioned CO-FISH technique with a pantelomeric DNA or PNA probe must be used (Table 1) [141,146,246]. A briefly description of each type of chromosomal aberrations involving telomeres that can be recognized using current molecular cytogenetics techniques follows. 3.1.2. Aberrations involving telomeres 3.1.2.1. Incomplete chromosome elements and acentric fragments. Lack of repair or incomplete repair of

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Fig. 1. Schematic representation of chromosomal aberrations involving telomeres (A–G) and interstitial telomeric repeat sequences (H and I) as seen in metaphase cells using FISH techniques (see text for explanation). Black circles represent telomeric repeat sequences; grey circles represents the centromere of each chromosome, and empty circles indicate the region of the chromosomes where the telomeric association or fusion has taken place. (A) Incomplete chromosome elements (ICE). (1) An incomplete chromosome (IC) accompanied by a terminal fragment (TF). (2) A pair of incomplete chromosomes (IC) accompanied by a compound fragment (CF), also known as a ‘‘proximal incomplete dicentric’’. (3) A pair of terminal fragments accompanied by a dicentric (DIC), also known as a ‘‘distal incomplete dicentric’’, or a ring chromosome (incomplete ring). (B) Acentric fragments. Terminal (TF), interstitial (IF) and compound (CF) fragments. (C) Telomeric association. (D) Telomeric fusion (as seen by conventional FISH, i.e., without discrimination of the type of fusion), giving rise to dicentric chromosomes. (1) Chromosome-type fusion (end-to-end fusion of two metacentric chromosomes). (2) Chromosome-type fusion (metacentric chromosome resulting from two acrocentric or telocentric chromosomes fused by their p-arms) (see text for explanation). (3) Chromatid-type fusion (chromatid dicentric). (E) Telomere–telomere fusion (as seen by COFISH), giving rise to dicentric chromosomes. (1) Chromosome-type; (2) chromatid-type (chromatid dicentric). (F) Telomere–DSB fusion. Two examples of chromosome-type (1 and 2) and one example of chromatid-type (3) telomere–DSB fusions are shown: (1) telomere–DSB fusion

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Table 1 Chromosomal aberrations involving telomeres (types 1–6) and interstitial telomeric repeat sequences (types 7 and 8) Type of aberration

Technique employed to detect the aberrationa

1. Incomplete chromosome elements (ICE)

Conventional FISH with a pantelomeric probe, which, if necessary, can be combined with a pancentromeric and/or a painting probe (see text) Conventional FISH with a pantelomeric probe

2. Acentric fragments: terminal (TF), interstitial (IF) and compound (CF) fragments 3. Telomeric association 4. Telomeric fusion (end-to-end chromosome fusion) 5. Telomere–telomere and telomere–DSB fusions 6. Telomeric sister-chromatid exchange (T-SCE) 7. Amplification of telomeric sequences 8. Translocation of telomeric sequences a

Conventional FISH Conventional FISH CO-FISH [146] CO-FISH [141] Conventional FISH Conventional FISH

with a pantelomeric probe with a pantelomeric probe

with a pantelomeric probe with a pantelomeric probe

The primed in situ labeling or PRINS technique [216–223] can be used as an alternative to conventional FISH.

chromosome breaks produced by chromosome breaking agents (clastogens) leads to terminal deletions (i.e., a truncated chromosome together with a terminal fragment derived from the same chromosome) or incomplete exchanges (i.e., incomplete dicentrics, rings or translocations) [227,228]. Hence, unrepaired chromosome breaks lead to unjoined or ‘‘open’’ chromosome elements, which represent the fraction of unrepaired chromosome damage in a given cell population exposed to a clastogenic agent. Unjoined chromosome elements are characterized by the presence of telomeric signals at only one end and are thus termed incomplete chromosome elements (ICE) [227,228,252–255]. Two types of ICE can be distinguished, namely, centric (incomplete chromosomes) or acentric (terminal fragments). In complete cells (i.e., containing all telomeres and centromeres) ICE are always observed in pairs, which can be constituted by an incomplete chromosome plus a terminal fragment, two incomplete chromosomes (accompanied by a compound fragment) or two terminal fragments (accompanied by an ‘‘incomplete’’ dicentric or ring chromosome) (Fig. 1A). In some cases, a ring-like structure exhibiting two telomeric signals can be formed, possibly due to the telomere loss in only one chromosome arm and fusion between the sister chromatids (see Fig. 3 in Ref. [179] and Fig. 1 in Ref. [256]). Besides terminal fragments, two other types of acentric fragments can be distinguished by FISH with a telomeric probe [257]: compound fragments, which results from the fusion of two terminal fragments and are characterized by the presence of telomeric signals at

both ends, and interstitial fragments (also termed acentric rings or double minutes, depending on their size) which can be easily identified because they lack telomeric signals [201,228,253–255,257] (Fig. 1B). All acentric fragments minus those accompanying a dicentric or ring chromosome are usually considered as ‘‘excess’’ acentric fragments. Although this class of aberrations includes terminal and interstitial fragments, compound fragments accompanying pairs of incomplete chromosomes should also be considered as excess acentric fragments. Excess acentric fragments might arise either from a complete exchange (an intra-arm intrachange, leading to an interstitial fragment or deletion) or from an incomplete exchange or breakage (terminal deletions, leading to terminal or compound fragments) and their scoring provides additional information regarding chromosomal incompleteness. 3.1.2.2. Telomeric associations and fusions. Telomeric associations and fusions are common cytogenetic findings that have been implicated in the initiation of chromosome instability and tumorigenesis [7,258– 261]. Also, the occurrence of telomere–telomere associations has been suggested to play a role in nuclear organization [262]. Telomere shortening in human chromosomes usually leads to increased frequencies of telomeric associations in metaphase [263], which implies that a minimum telomere length is required for proper telomere function, i.e., when telomeres become critically short, telomere separation at mitosis cannot be performed

resulting in the formation of a dicentric chromosome (DIC); (2) chromosome-type and (3) chromatid-type telomere–DSB fusions resulting from an acentric chromosomal fragment (terminal type) fused to an intact chromosome. The reactive or ‘‘open’’ end resulting from the telomere–DSB chromosome-type fusion comes from the terminal fragment (TF) (case 1) or the incomplete chromosome (IC) (case 2) formed. (G) Telomeric sisterchromatid exchange (T-SCE). (H) Amplification of telomeric sequences. Increase in the number (A) or in the size (B) of the telomeric signals. (I) Translocation of telomeric sequences. Two examples are shown: a translocation of telomeric sequences located at the centromeric region of a chromosome, which results in a chromosome with interstitial telomeric repeats (ITR) (case A) and a translocation of telomeric sequences located at the terminal region of a chromosome, which results also in a chromosome with interstitial telomeric repeats (ITR) (case B).

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properly, leading to telomeric association at metaphase and chromosome instability, as observed in several human pathologies [264–266]. However, the presence of telomeric association at metaphase in mammalian cells does not always correlate with short telomeres [248,267,268]. Although telomeric associations and fusions cannot be clearly distinguished using conventional staining techniques – and thus for scoring purposes they are considered as the same phenomenon, i.e., endchromosome fusion [269,270] – thanks to the recent developments in the field of molecular cytogenetics each of these events can be properly identified. Telomeric associations are recognized by the presence of two pairs of very close telomeric signals after hybridization with a telomeric probe, each pair corresponding to a different chromosome [147] (Fig. 1C). Therefore, to be scored as a telomeric association, the telomeres of adjoining chromosomes had to be clearly distinguished from one another, i.e., four telomeric signals must be present, and the DAPI signal had to be discontinuous through the point of fusion [147]. 3.1.2.3. Telomeric fusions: interactions between two telomeres and between telomeres and DSBs. Telomeric fusions are the result of telomere dysfunction due to unprotected or eroded chromosome ends [16], and they are usually found in repair- and/or telomerase-deficient cells [81,84,149,179,195,256]. Mammalian cells with a variety of mutations that affect telomere function, including those ones occurring in DNA-PKcs [195,271], Ku [195,176,177] and TRF2 [13,146,164,272] proteins, show a high frequency of telomeric fusions (end-to-end) and chromosome instability. In particular, Smogorzewska et al. [272] showed that telomere fusions resulting from TRF2 inhibition are generated by DNA ligase IV-dependent NHEJ and that these aberrations are accompanied by active degradation of the 30 overhangs. Moreover, chromosome fusion resulting from telomere loss appears to play an important role in chromosomal rearrangements associated with cancer [7]. True chromosome-type telomere–telomere or endto-end chromosome fusions can be visualized using conventional FISH with a telomeric probe [147,179, 195,256,257] as events where the telomeres of adjoining chromosomes had fused into a single signal (Fig. 1D, cases 1 and 2) and the DAPI signal had to be continuous through the point of fusion [146,147]. There are two types of chromosome-type telomeric fusions which can be identified by conventional FISH. One type involves the fusion of two metacentric or submetacentric chromosmes (see for example Fig. 1D, case 1), whereas the other involves the fusion of two acrocentric/

telocentric chromosomes, and has been found in murine cells (Fig. 1D, case 2). In the latter case, the acrocentric/telocentric chromosomes fuse to each other to generate a metacentric Robertsonian-like configuration with telomeres at the fusion point (Fig. 1D, case 2). This could be generated by the loss of telomere equilibrium and the subsequent loss of capping function – telomere inactivation – in the cells possessing these aberrations [179,273]. Alternatively, the two chromosomes can be fused by the q-arm, and thus a long dicentric chromosome results [179]. It is important to note that not all chromosome fusions are telomeric fusions, since in most cases, the formation of a metacentric chromosome through a Robertsonian translocation involves the fusion of chromosomes that had lost their telomeres, i.e., there is a complete loss of telomeres or telomere erosion at the chromosome arm involved in the fusion (the p-arm), and thus after telomere FISH the metacentric chromosome shows no telomeric signal at the point of fusion [179,273,274]. The Robertsonian translocation involving telomere loss at the point of fusion is one of the most frequent chromosomal rearrangements in mammalian karyotype evolution, especially rodents [274–277]. It has been shown that after a Robertsonian translocation, all telomeric and many minor satellite DNA sequences are lost, so that in the newly formed centromeric region of every Robertsonian chromosome about 20–60 kb of minor satellite DNA are retained, flanked by two blocks of about 6 megabases each of major satellite DNA [278,279]. In conclusion, there are two types of chromosome fusions: classical Robertsonian translocations, involving the fusion of two acrocentric/telocentric chromosomes by their centromeres [274] and end-to-end chromosome fusions (true telomeric fusions) involving the telomeres of the p- or the q-arms of metacentric, submetacentric, or acrocentric/telocentric chromosomes (this latter termed Robertsonian-like fusions), giving rise to dicentric chromosomes [179,195] (Fig. 1D). Although end-to-end chromosome fusions can be detected using conventional FISH with a telomere probe, the CO-FISH technique allows discriminate two different types of telomere fusions [146,147]. One type comprises the interaction of two telomeres, which join to each other (Fig. 1E), whereas the other one consists of a fusion of a telomere with a DSB (Fig. 1F). The two types of chromosome-type telomeric fusions can be clearly distinguished by simply looking at the CO-FISH hybridization pattern [146] (Fig. 1E1 and F1 and 2). In the case of a telomere– telomere fusion, two blocks of telomeric DNA join in opposite orientations as a result of maintaining polarity

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(Fig. 1E, empty circle). Therefore, telomeric signals are detected on both chromatids at the point of fusion. In contrast, a telomere–DSB fusion produces a signal on just one chromatid (Fig. 1F). In conclusion, a chromosome-type telomere–telomere fusion produces two interstitial signals, whereas a telomere–DSB fusion gives only one interstitial signal at the fusion point. Telomeric fusions can also be of chromatid-type (see Fig. 1D3, E2 and F3), if just one chromatid per chromosome is involved in the fusion [146]. Some chromatid-type (like chromatid dicentrics, see Fig. 1D, case 3) and chromosome-type (see Fig. 1D, cases 1 and 2) telomeric fusions can be distinguished by conventional FISH [195], whereas, as stated above, only COFISH allows discriminate between the different types of chromosome-type telomeric fusions [146]. The presence of one or both types of telomeric fusions in a cell is a clear indication of impaired end-capping, i.e., dysfunctional telomeres [108,146,147]. When cells suffering telomeric end-capping dysfunction are exposed to a clastogenic agent, uncapped telomeres and DSBs interact in different ways, allowing for the formation of three types of fusions, DSB–DSB – leading to ordinary chromosome-type aberrations – telomere–telomere and telomere–DSB. True telomere–telomere fusions results in the formation of a dicentric chromosome without an accompanying acentric fragment (Fig. 1D and E), in contrast to classical dicentric chromosomes derived from the interaction of DSB from two chromosomes and which involves the formation of an additional acentric fragment (a compound fragment or two terminal fragments, see Fig. 1A). As a result, a cell containing a dicentric chromosome derived from a DSB–DSB interaction may die because of the loss of the genetic information contained in the acentric fragment. By contrast, cells with telomere–telomere fusions are more likely to survive and undergo further chromosomal instability through the formation of BFB cycles. On the other hand, telomere–DSB fusions allow the formation of several types of chromatid- and chromosome-type aberrations, including dicentrics (Fig. 1F, case 1) [146], but each has one key feature in common—the fusion heals just one end of the DSB; the other end (corresponding to a terminal fragment or an incomplete chromosome) remains reactive (open) and thus capable of promoting further chromosomal instability (Fig. 1F). 3.1.2.4. Telomeric sister-chromatid exchanges (TSCE). Recombination events at telomeres can be detected cytogenetically in the form of sister-chromatid

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exchanges (SCEs) that involve telomeric sequences – telomeric sister-chromatid exchanges or T-SCE – using the CO-FISH technique [123,135,146,246,273] (Fig. 1G). As previously described, with telomeric CO-FISH a single stranded telomeric probe hybridizes to complementary telomeric DNA on one chromatid of each chromosome arm producing a two-signal pattern instead of the four signals seen with conventional FISH (Fig. 1G, case 1). However, if a T-SCE has taken place, a three-signal CO-FISH hybridization pattern is expected, since the effect of an SCE within telomeric DNA is to split the hybridization signal (Fig. 1G, case 2). The sequential use of a C-rich telomere probe and a G-rich telomere probe produces a reciprocal pattern of hybridization when a true T-SCE has taken place [246]. As noted by Bailey et al. [146], other types of recombination could conceivably produce also a threetelomere hybridization pattern like that of a T-SCE – for example, a telomeric fusion followed by breakage within telomeric DNA on one side of the fusion point, a large extrachromosomal telomeric DNA fragment joined to one telomere, or a break in one sister telomere producing a fragment that then joins to the other sister telomere – but in this case, the three-signals pattern is produced with only one of the telomeric probes (i.e., is not reciprocal) and the mechanisms involved require ligation to a chromosome end, an event that can be prevented by functional telomeres. Therefore, in cells with functional telomeres, if a threetelomere hybridization pattern occurs, this can be ascribed to a T-SCE. CO-FISH revealed high rates of mitotic recombination within sub-telomeric regions in human cells [280]. Moreover, the ‘‘spontaneous’’ occurrence of T-SCE has been documented in different human and murine repairor telomerase-deficient cell lines [135,141,273,281,282]. T-SCEs have been implicated as a marker of certain human cancers whose cells are sometimes forced to utilize telomerase-independent pathways to maintain telomere length, the above-mentioned ALT pathway [123,135] (see Section 1.2.9). Moreover, it has been demonstrated that in murine telomerase-deficient cells possessing telomere signals-free ends, telomere can be maintained through the occurrence of T-SCE [282]. It should be pointed out that high rates of T-SCE are characteristic of all established ALT cells analyzed so far [123]. Reactivation of telomerase completely abolished the T-SCE pathway in these cells, suggesting that T-SCE is a special type of ALT that does not coexist with telomerase [123]. It has been suggested that unequal TSCE delays clonal senescence and may extend the proliferative life of telomerase-negative cells [141]. In

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some cells, telomere shortening coincides with reduction in recombination at telomeres [273]. 3.1.3. Aberrations involving ITRs 3.1.3.1. Amplification and translocation of telomeric sequences. Telomeric repeats amplification is visualized as an increase in the number and/or the size and intensity of the hybridization signals after FISH with a telomeric probe compared with the normal telomeric hybridization pattern of the cell type or species being studied [201] (Fig. 1H), whereas a translocation of telomeric sequences means that a relocation of one or more pairs of telomeric repeats compared to the normal telomeric hybridization pattern of the species or cell type being studied has taken place (Fig. 1I). Alternatively, in cells which ordinarily do not contain cytogenetically visible blocks of telomeric repeats at internal sites, like mouse or human cells, the presence of ITRs would indicate that a telomeric (end-to-end) fusion has taken place [195]. A special case of telomeric repeats translocation is the above-mentioned process termed ‘‘telomere capture’’ (see Section 1.2.8), in which a break in one chromosome is ‘‘healed’’ by the addition of the telomere of another chromosome [107,112]. The coexistence of chromosomes with duplicated telomere signals at one end (‘‘telomere duplication’’) and chromosomes lacking telomeric signals at one of the chromatids (‘‘chromatid telomere loss’’) in the same metaphase – as observed in normal human fibroblasts exposed to ionizing radiation [283] – could be also explained by telomeric repeats translocation. Amplification of telomeric repeats may arise due to unequal SCEs [141,284], BFB cycles [7,197,285] or excision and reintegration events, i.e., the ‘‘rolling circle’’ mechanism [142,286]. In cells exhibiting telomeric signals exclusively or predominantly at the terminal regions of chromosomes, both events, i.e., amplification and translocation of telomeric repeats are visualized most of the time as ITRs [201]. 3.2. Induction of chromosomal aberrations involving telomeres and ITRs by physical and chemical mutagens In this section, we will summarize our current knowledge of the induction of chromosomal aberrations involving telomeres and ITRs by physical and chemical mutagens. The study of these aberrations has given new insights into the mechanisms underlying the formation of chemically and radiation-induced chromosomal aberrations.

3.2.1. Induction of chromosomal aberrations involving telomeres 3.2.1.1. Incomplete chromosome elements (ICE) and acentric fragments (AF). Several studies were conducted in the last few years using FISH with a telomeric probe in order to obtain information concerning the induction of ICE and interstitial fragments by ionizing radiation and chemical mutagens [212,227–229,250, 252–255,287]. These studies included the analysis of all types of asymmetrical chromosome aberrations induced by ionizing radiation in human lymphocytes and those ones induced by the radiomimetic agents BLM and streptonigrin (SN) and the methylating compound streptozotocin (STZ) in Chinese hamster cells. It was found that about 70% of the ICE induced by X-rays and 99–100% of those induced by the chemical mutagens BLM, SN, and STZ, consisted of pairs of an incomplete chromosome and a terminal fragment [228,253–255]. This means that it is much more likely that a terminal fragment is accompanied by an incomplete chromosome than a compound fragment by two incomplete chromosomes or a dicentric or ring chromosome by two terminal fragments. Thus, the main form of ICE induced by clastogenic agents – irrespective of their mode of action – consists of pairs of an incomplete chromosome and a terminal fragment (see Fig. 1A, case 1). The predominance of some form of ICE over the other ones might be explained by the attachment of the telomeres to the nuclear matrix [50,228,254,255]. In effect, movement of interphase chromatin after the induction of chromosome breaks might result in dislocation of break-ends that were in close proximity, giving rise to ICE. Predominance of pairs of incomplete chromosomes over pairs of terminal fragments might be explained by the fact that dislocation of the centric chromosome element (i.e., a dicentric chromosome) might be more likely and therefore more prone to remain unjoined. Likewise, predominance of pairs consisting of an incomplete chromosome and a terminal fragment over pairs of incomplete chromosomes is probably due to the fact that this latter type of ICE implies the formation of a dicentric chromosome, whereas only a single isochromatid-type or chromosome-type break is needed to originate a pair of incomplete elements consisting of an incomplete chromosome and a terminal fragment. In the above-mentioned studies no discrimination could be made between incompleteness derived from incomplete exchanges or terminal deletions because to correctly discriminate between these types of aberrations a combination of telomere FISH or PRINS with chromosome painting is needed [110,229,250,287].

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However, following Boei et al. [228], a rough estimate of the frequency of terminal deletions can be made assuming that incompleteness is equally likely to occur for dicentrics and translocations. Thus, subtracting the frequency IC + TF from IC + IC + TF + TF estimates the number of terminal deletions [228]. Therefore, the estimated frequency of terminal deletions in CHE cells induced by the highest concentration of BLM (7.5 mg/ ml), SN (250 ng/ml) and STZ (4 mM) employed is three to eight times higher (0.61 for BLM, 1.70 for SN, and 0.70 for STZ) [253–255] than that observed in human lymphocytes exposed to 6 Gy of X-rays (0.22) [228]. Furthermore, ICE were found to be the most frequent type of asymmetrical chromosome aberrations induced by chemical mutagens [253–255]. By contrast, in low and high LET ionizing radiation-exposed cells, dicentric/multicentric chromosomes were observed with a higher frequency than ICE [228,249]. The above findings allow one to conclude that chromosomal incompleteness or the formation of ‘‘true’’ incomplete exchanges is a rare event following exposure to low and high LET radiation [212,227–229,249,250,252,287– 289], but a common one after exposure to chemical mutagens [253–255]. It must be noted that, in some studies, the percentage of incomplete rejoinings was found to be higher after high-LET than after low-LET radiation exposures [249,251,288], while in others, a significant difference between these two types of radiation regarding the induction of incomplete rejoinings could not be found [250,252,289]. As suggested by Mestres et al. [249], an increase in the incompleteness for high-LET radiation could be due to the complexity of DNA damage after a high-LET exposure that can include one or more DSBs as well as associated SSBs, damaged sites, and DNA–DNA and DNA–protein cross-links [290–292]. The studies about ICE and interstitial fragments also demonstrated that interstitial fragments form a major class of low-LET radiation-induced chromosomal aberrations, corresponding to about 80% of the excess acentric fragments induced by X-rays in human lymphocytes [227,228]. Therefore, most of the excess acentric fragments induced by low LET ionizing radiation originate from complete exchanges (intraarm intrachanges leading to an interstitial fragment). By contrast, Mestres et al. [249] analyzed the induction of chromosome aberrations by a-particle (high LET radiation) in human lymphocytes using a combination of pancentromeric and pantelomeric probes and found that most of the excess acentric fragments are of terminal type. Moreover, about 50, 80, and 100% of excess acentric fragments induced by BLM, SN, and

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STZ, respectively, originate from incomplete exchanges or terminal deletions [253–255]. Hence, ionizing radiation and chemical mutagens exhibit a different pattern of induction of ICE and excess acentric fragments. These findings suggest that chromosome breaks induced by S-independent agents (i.e., ionizing radiation and BLM) are better repaired than those induced by clastogens acting partially (i.e., SN) or mainly (i.e., STZ) in an S-dependent manner. However, further studies are needed to validate this hypothesis. On the other hand, we found that some of the acentric fragments induced by BLM and SN in Chinese hamster cells were labeled along their entire length [201]. This finding seems to indicate that these fragments were very likely derived from breaks occurring at the centromeric regions of the chromosomes, suggesting that centromeric regions containing ITRs are prone to breakage and recombination by radiomimetic compounds (see Section 3.2.2.1). The dose–response relationship regarding the induction of ICE and other types of asymmetrical or unstable chromosome aberrations was found to be of linearquadratic type for low LET radiation [228], of linear type for high LET radiation [250,252,293], BLM [253] and SN [254] and of curvilinear type for STZ [255]. Therefore, the clastogenic effects of BLM and SN in terms of ICE are not comparable to those obtained with low LET (X-rays) but to those of high LET (neutrons) radiations instead. Moreover, the comparable trend of the dose/concentration–response relationship for the different aberrations strongly suggests that all X-rays, BLM- and SN-induced asymmetrical aberrations are formed by a similar underlying mechanism. The linear relationship between the yield of chromosomal aberrations and ionizing radiation dose or BLM/SN concentration suggests that these aberrations are formed by a single track mechanism, i.e., only one ionizing radiation-, BLM- or SN-induced DNA lesion (presumably a DSB) initiates the formation of exchanges. Conversely, the fact that STZ gave curvilinear concentration–response curves for induction of ICE and terminal fragments, suggests that ICE are formed by a mechanism involving two-hit events, i.e., these aberrations are caused by the interaction of two STZinduced DNA lesions (probably DSBs). 3.2.1.2. Telomeric associations. Most of the studies performed so far regarding induced telomeric associations have been made using conventional staining techniques [269,270,294]. FISH studies reporting the induction of ‘‘true’’ telomeric associations by ionizing radiation are scarce [231,295] and there is no FISH data

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available regarding the induction of these events by chemical mutagens. This is very likely due to the fact that, as previously stated, telomeric associations and fusions have been usually considered as the same phenomenon and, therefore, these two distinct events were scored as a single one. However, the most recent studies regarding the scoring of chromosomal aberrations involving telomeres make a clear and proper distinction between telomeric associations and telomeric fusions, although the former are not always scored as chromosomal aberrations [147,195].

lines having normal telomere function [211,212,299] and that dysfunctional telomeres join to the ends of DSBs induced by gamma irradiation providing the first conclusive evidence of telomere–DSB fusions in mammalian cells [146]. In the other study, Al-Wahiby and Slijepcevic [147] analyzed with conventional FISH and CO-FISH the chromosomal aberrations involving telomeres in BRCA1 deficient human and mouse cell lines. They found that bleomycin induces chromosometype telomere–DSB but not telomere–telomere (end-toend) fusions.

3.2.1.3. Telomeric fusions. The induction of end-toend chromosome fusions has been demonstrated in different cell types by using conventional FISH with a telomeric probe. For instance, Fimon et al. [296] showed that defective telomere function – evaluated by the presence of Robertsoninan-like telomeric fusions – correlates with chromosomal radiosensitivity in a mouse tumor cell line and Undarmaa et al. [297] showed that ionizing radiation induces end-to-end chromosome fusions in ATM-deficient mouse cells (ATM is a repair protein encoded by the gene responsible for ataxia telangiectasia). Moreover, McHugh et al. [298] recently showed that the antitumor radiomimetic enediyne antibiotic C-1027 induces telomere dysfunction in the form of chromosome fusions (telomere–telomere fusions following Bailey et al. [146]) in cultured human colon carcinoma HCT116 cells. However, no telomeric fusions were observed in normal human fibroblasts exposed to gamma radiation [211]. Since only after the very recent development of telomeric CO-FISH [146,246] it was possible to distinguish between the different types of telomeric fusions, data about the induction of telomere–telomere or telomere–DSB fusions by chemical or physical mutagens are very scarce [146,147,273]. The only data currently available were provided by two studies with human and mouse cells exposed to ionizing radiation [146] and BLM [147]. In one of these studies, Bailey et al. [146] analyzed the induction of chromatid- and chromosome-type telomeric fusions by gamma irradiation in cells derived of a wild-type mouse, a mutant scid mouse and a HTC75 human fibrosarcoma cell line expressing a TRF2 dominant-negative allele (i.e., with uncapped telomeres). The only chromosome-type telomeric fusion present in these cell types was telomere–DSB fusions. This study demonstrated that functional telomeres protect chromosome ends from joining to radiation-induced DSB ends, a finding consistent with previously published work in other cell

3.2.1.4. Telomeric sister-chromatid exchanges (TSCE). At present, there is no published data regarding the induction of T-SCE by chemical or physical mutagens. Undoubtedly, this is due to the very recent development of the CO-FISH technique to detect TSCE [141]. Although there is a recent study [300] reporting the frequent occurrence of ultraviolet light B and cisplatin-induced sister-chromatid exchanges in telomere regions of Chinese hamster chromosomes, these events were scored using conventional staining techniques and thus no specific data on true T-SCE are available. Nevertheless, results presented at the ‘‘Telomere sessions’’ of the XV International Chromosome Conference (London, September 2004) by W. Wright (University of Texas) indicate that DNA damaging agents, such as ionizing radiation, can induce SCEs at telomeres (cited in Ref. [108]). Since ionizing radiation normally does not induce SCEs, this finding suggests that, in telomeres, homologous recombination mechanisms behave differently in comparison with the rest of the genome. 3.2.2. Induction of chromosomal aberrations involving ITRs 3.2.2.1. Amplification and translocation of telomeric sequences. Studies with Chinese hamster cell lines exposed to ionizing radiation or chemical mutagens have demonstrated that telomeric sequences can undergo both, amplification and translocation events, and that chromosomal regions containing ITRs are prone to chromosome breakage, fragility and recombination [107,197,198,201,207,301–306]. It has been shown that following treatment with ionizing radiation [107,197,210,303–309], restriction endonucleases [198], mitomycin C and teniposide (VM-26) [304] or the radiomimetic antibiotics BLM and SN [201] there is an increased frequency of aberrations involving ITRs in Chinese hamster cells, including chromatid-type aberrations like trirradials and quadrirradials [201,304]. In some cases, it could be demonstrated that the

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percentage of chromosomal aberrations involving ITRs was higher than expected based on the percentage of the genome composed by telomeric sequences [201,303, 304]. Nevertheless, it has been demonstrated that not all ITRs are hot spots for rearrangement or recombination [301], and the sensitivity to breakage of short ITRs like those present in Chinese hamster chromosomes at noncentromeric sites [310] is not clear [311,312]. In fact, studies by Desmaze et al. showed that ITRs are not responsible for the chromosomal instability observed in human cells [311,313]. Azzalin et al. [205] proposed that short telomeric arrays – like those found in human cells – may not be fragile sites but simply mark sites of double-strand breaks that occurred within unstable regions. In this way, short ITRs could be envisaged as relics of ancient breakage within fragile sites, rather than fragile sites themselves [205]. This hypothesis does not rule out the possibility that very extended blocks of ITRs, such as those present in Chinese hamster cells could be prone to breakage and recombination. Some of the above-mentioned studies showed amplification of telomeric sequences at breakpoints and fragile sites [197,198,201]. Although no data were available on the activity of telomerase in the Chinese hamster cells exposed to ionizing radiation [197] or restriction enzymes [198], we found that the amplification of telomeric repeats induced by BLM and SN in Chinese hamster embryo cells (CHE cell line) is not accompanied by an increase in the activity of telomerase [201]. Spontaneous amplification of interstitial telomeric bands on specific marker chromosomes has been observed in different sub-clones of the CHO cell line by several authors [197,302,303,314]. However, Pandita and DeRubeis [314] exposed CHO cells to different DNA-damaging agents (including BLM) and DNA synthesis inhibitors and found that none of these treatments increased the acquisition of interstitial telomeric bands on marker chromosomes in these cells. On the other hand, telomeric sequences were found to be involved in terminal as well as interstitial translocation events induced by BLM and STN in CHE cells [201]. As suggested by Ferna´ndez et al. [304] and Balajee et al. [198], it is likely that the capacity of telomeric repeat sequences to form secondary structures within and between chromosomes [315] could account for their fragility and recombination. 4. Future prospects As reviewed here, studies performed mainly during the last decade have shown that telomeres play a

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significant role in the formation of chromosomal aberrations, and have provided some important clues regarding the origin of aberrations induced by physical and chemical mutagens. However, the scarcity of data available on the induction of some types of chromosomal aberrations involving telomeres, like telomere– telomere and telomere–DSB fusions and also T-SCEs, indicates that a more intensive work on this subject is needed. Some interesting and very important questions that emerged from the studies reviewed here are: Why chromosomal incompleteness is a common event following exposure to chemical mutagens but not ionizing radiation? Should BLM and SN be considered true radiomimetic clastogens, since the spectrum of chromosomal aberrations induced by these antibiotics and ionizing radiation is very different? Why ionizing radiation and BLM induce telomere–DSB but not telomere–telomere fusions? Why ionizing radiation induces T-SCE, since it normally does not induce SCE? Should telomeric associations be considered as true chromosomal aberrations? These represent only a small sample of the questions regarding the induction of chromosomal aberrations involving telomeres that remain to be answered. In addition, further research on the induction of amplification and translocation of telomeric repeats and the mechanisms involved in these events may provide some new insights on tumorigenesis, since both gene amplification and translocation are involved in this process. It must be noted that most of the studies regarding chromosomal aberrations involving telomeres were made on a few cell types and using ionizing radiation as the clastogenic agent. Since different cell types show different chromosomal sensitivity to physical and chemical clastogens and most chemical mutagens act in a different way than ionizing radiation, in order to establish general conclusions regarding the induction of these aberrations, additional studies with chemical mutagens and using other cell types are clearly needed. On the other hand, although current molecular cytogenetics techniques allow to identify several types of aberrations involving telomeres, improvements in the resolution of these techniques to a degree which allows to detect very short telomeres – like those present in some mammalian cell lines – will allow a more extensive and detailed analysis of these aberrations. Undoubtedly, further research on the topic outlined in the present review will provide an answer to the above questions and lead us to a better understanding of the mechanisms of induction of chromosomal aberrations by physical and chemical mutagens.

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