Telomeres and Their Neighbors: History
View Latest Version
Please note this is an old version of this entry, which may differ significantly from the current revision.
Contributor:
Leon P. Jenner
,
Vratislav Peska
,
Jana Fulnečková
,
Eva Sýkorová

Telomeres are essential structures formed from satellite DNA repeats at the ends of chromosomes in most eukaryotes. Satellite DNA repeat sequences are useful markers for karyotyping, but have a more enigmatic role in the eukaryotic cell. More research is needed until there is a complete picture of the biological function of telomere and other DNA satellite sequences, including chromatin structure, chromosome end-protection and species evolution with a particular focus on non-model organisms. The first problem to solve is the identification of telomere repeats, because telomere repeat identity is the foundation for any hypothesis about telomere maintenance and structure, or binding of specific  proteins. Celebrating Gregor Mendel’s anniversary by going to the principles behind the experiments, a selection of recent developments and underexplored areas of research from the past are illustrated in plants and insects. Indeed, much recent work has expanded beyond the human and yeast models traditional in telomere research. Classic methods from the past, and cutting-edge in silico methods are described. These do not require specialized equipment or expensive materials and can be used, often in combination, to aid research into telomeres and satellites. This can both enrich the general understanding of chromosome maintenance mechanisms and further explore the evolution of telomeres and telomerases.

  • satellite
  • telomere evolution
  • interstitial telomere sequences
  • telomerase RNA
  • NGS
  • eukaryotic tree of life

1. Introduction

The essential DNA structures that form eukaryotic chromosomes are centromeres, telomeres and origins of replication. Centromeres are vital for proper nuclear division and telomeres protect the ends of linear chromosomes from attrition during DNA replication. Each chromosome also possesses genes which populate the chromatin regions lying between centromeres and telomeres. Genes are sequence-based quanta of information, Gregor Mendel’s “elements”, that are the foundation of organism identity when realized. A simplified picture of genetics was initially recognized by Mendel without any deep knowledge of DNA, RNA and proteins; molecules that we now know realize organism function. When thinking about Mendel 200 years after his birth, it is obvious that simple methods and an open mind are the solution to many scientific questions, even those related to gene function, genome structure and evolution. Of course, there is also a little bit of luck in choosing a model. Mendel’s choice, the garden pea (Pisum sativum) has a large genome that has only recently been sequenced [1] and is not a popular organism for modern plant genetics due to the abundance of repetitive elements in its genome. Conversely, another one of Mendel’s favorite organisms, the honeybee (Apis melifera, [2]) is still intensively studied as it offers a chance to understand the phenomena of social life and cooperation in the insect world. It is an extraordinary coincidence that the dawn of telomere biology in 1938 is linked to maize and Drosophila, models which were used in the pioneering works of McClintock and Muller, who showed breakage-fusion-bridge cycles and chromosome healing after X-ray damage [3][4]. At that time, it was assumed that the entire chromosome was filled with genes, and Muller’s original definition of the telomere speaks about it as a special terminal gene. Today, the definition of a gene has changed and we know more about chromosomes and telomeres, but there is still much yet to be discovered (e.g., [5][6][7][8][9][10][11][12][13] and references herein). Telomeres and satellites are often neighbors at chromosome termini. Much telomere research derives from human, ciliate and yeast models. Knowledge of telomere evolution has increased enormously in recent years, however, thanks to discoveries in plant and insect models.

2. Telomeres as Steps in Species Evolution

To begin with, telomere DNA sequences were assigned as a trait of a large group of organisms, e.g., TTAGGG in vertebrates, TTTAGGG in plants, TTAGG in insects/arthropods [14][15][16][17][18] (see [19] for review). Thus, the majority of identified telomere repeats are of minisatellite size and maintained by a special enzyme, telomerase telomerase ([20][21][22], reviewed in, e.g., [7][11][23]), except well-known examples of non-telomerase alternatives from Diptera ([24][25][26][27][28], reviewed in [11][19][29][30]). This conservation has proven advantageous in microscopy studies and telomeric probes are second only to rDNA probes [31][32][33], e.g., to distinguish and study telocentric chromosomes, to recognize Rabl-like or bouquet organization or various chromosomal aberrations [34][35][36][37][38][39][40][41]. Numerous reports that characterized typical telomeric sequences in an increasing number of species seemed to confirm the telomere consensus TxAyGz. Telomeric sequences in yeast models, e.g., TG1–3 in budding yeast [42][43], T1–2ACA0–1C0–1G1–6 in fission yeast [44], 8–25 bp-long repeats in Kluyveromyces and Candida [45][46] were treated as an interesting variety from the general repeat unit TxAyGz and special only to yeast. Current research on Saccharomycotina [47][48] has revealed even more telomeric variants, although despite their considerable divergence, all of these telomere sequences have guanines (Gs) as one of their most conserved features [46][47][48][49].
Missing signals using telomere probes in in situ hybridization experiments were the first hints towards identifying organisms that do not possess typical telomeres formed by the expected repeat, e.g., plants Allium (Asparagales, [15]), Cestrum (Solanales, [50]), some beetles and the spider Tegenaria ferruginea [18]. In the next few years, detailed studies revealed gradually more species with unknown telomeres from plants [51][52] and insects [53]. This led to a breakthrough in the general view of telomeres. Studies that mapped telomere sequences in plants, animals and algae identified evolutionary switchpoints in which sequences typical to one group were replaced by other variants [50][54][55][56][57][58][59][60][61]. For example, a group of species from the plant order Asparagales changed their telomeric sequence from the Arabidopsis-type repeat TTTAGGG to the human-type TTAGGG. An elusive, highly divergent telomere repeat was finally identified in Allium (Amaryllidaceae, Alloidae, [62], see Section 7), one of the largest monocotyledonous genera with an estimated 800–900 species [63]. Similar step changes were found in green algae, in which the transitions from TTTAGGG to novel types TTAGGG, TTTTAGGG or TTTTAGG allowed the grouping of species with the same telomere in distinct phylogeny clades [54][58][64]. A similar switch was identified in beetles where the repeat TTAGG was replaced with the TCAGG repeat [59]. A broad experimental study of algal telomeres, accompanied with the identification of candidate telomeric sequences from genomic databases of various species across the eukaryotic tree of life, showed TTAGGG and TTTAGGG telomeres as being the predominant telomeric types [54]. Fulneckova and colleagues [54] mapped the occurrence of telomeric sequences in phylogeny revealing the TTAGGG repeat as an ancestral eukaryotic telomere and current phylogeny [65][66] still supports this hypothesis (see [7] for review). Interestingly, just as many telomere variants were experimentally verified, many more species and groups with unknown telomeres were discovered [11][54][58][64][67]. Telomere sequence variants and their evolution in plants and algae are described in detail in a review by Peska and Garcia [7]. Progress in insect telomere identification is reviewed in Mason et al. [11] and recent findings are mapped in [67][68][69][70].

3. Telomere Minisatellites Are Much like Any Other DNA Sequences

When exploring the occurrence of telomere minisatellite repeats in the genome, keep in mind that telomere-like sequences can occur in locations other than in the telomere. Such sequences are called interstitial telomeric sequences (ITSs) and can be classified as part of several groups according to their length, occurrence and structure (recently reviewed in [71], Figure 1). ITSs can have the same sequences as telomeres or they can have variant telomere-like repeats. For example, budding yeast has the telomeric sequence TG1–3 and interstitial tracts of TTAGGG repeats are present in subtelomeric and other regions [72]. ITSs can occur as a few copies across the genome, including regions that are proximal to genes, but also in clusters found frequently in pericentromeric or subtelomeric regions. The arrangement of ITS sites can also be classified in respect to the orientation and composition of telomere-like sequences as head-to-tail or head-to-head, homogeneous or degenerated tandem repeats and with or without linker sequence(s) (Figure 1b). When ITSs occur in a head-to-head orientation with a linker sequence, these can be amplified using a single-primer PCR reaction [73] (Figure 1c). ITSs can be unique or part of longer repetitive sequences and are a suitable genetic marker for mapping [73][74][75]. ITSs in clusters usually contain a large portion of degenerate telomeric motifs and could be interspersed with other repetitive sequences [74][75][76].
Genes 13 01663 g002 550
Figure 1. Experimental examination of ITS and telomeric repeats. (a) Telomere repeats are strand oriented. (b) Telomere-like repeats in telomeres or internal sites may form clusters or short stretches. Single-primed PCR distinguishes between these using an extension reaction with a single telomeric oligonucleotide primer (C-rich primer is shown, triangles). Telomeric sequences, short and clustered ITSs produce a smear of ssDNA products visible after hybridization with a radioactively (*) labeled probe (right, e.g., from Chlorela vulgaris, experiment performed as in [50]). Cloneable dsDNA products visible in an ethidium-bromide stained agarose gel (etd) are produced when ITSs occur in head-to-head orientation. When dGTP is omitted, bands are not produced by ssDNA or short ITSs, but ssDNA from a telomere is elongated until primer extension stops at the first G in the subtelomere. This reaction showed the Arabidopsis- and human-type telomere repeats are absent in Allium and Cestrum [50][56][77]. (c) Different patterns of ITSs amplified from four Cestrum species in single-primed PCR using C-rich and G-rich primers for the Arabidopsis-type telomeric repeat [50][73]. (d) The specific pattern of ITS-associated sequence BR23 (green) was visualized on Cestrum elegans chromosomes using FISH. The high-copy repeat BR23 shows dispersed and clustered signals (5S rDNA in red, counterstained with DAPI; adapted from [73]). (e) Allotetraploid Cardamine scutata, a hybrid of C. parviflora and C. amara with the parental origin of chromosomes visualized by GISH (left panel, GISH) and telomeric probe (TEL) that detects differing pericentromeric ITS clusters (adapted from [75]; modified). (f) FISH of the 180-bp centromeric satellite (CEN180), retroelement ATHILA and TEL on pachytene chromosomes of A. thaliana. Interstitial telomeric locus in the pericentromeric region of chromosome Ch1 is marked by an arrow (adapted from [76]; modified). (g) TRF (terminal restriction fragment) method visualizes telomeric and ITS fragments from A. thaliana after restriction digestion of gDNA with MseI. (h) Schema illustrating the effect of Bal31 nuclease digestion on telomeric, subtelomeric, ITS and internal genomic sequences. After DNA isolation, DNA is fragmented and Bal31 nuclease gradually shortens these fragments from the end. Bal31-digested samples can be used for specific telomere-subtelomere PCR (left, see below). Further restriction digestion (right, H) results in the visualization of TRF signal (h-tel) shortening and verification of the terminal position of a candidate sequence. (Left) PCR/qPCR investigation of genomes with short telomeres (e.g., A. thaliana, see results in (i–k) adapted from [78]) proving subtelomeric position of candidate sequences (A,B). When the telomere is completely digested, PCR with a C-rich primer cannot amplify the product (tel-a, tel-b), and further digestion results in a loss of amplification signal from subtelomere regions proximal to telomeres (A) in contrast to ITS (C, pericentromeric ITS in A. thaliana, see schemas in (j)) or control sequences (D,E). Bal31 nuclease also degrades ssDNA (F) and some dsDNA sites with altered structures (G). (i) Dynamics of Bal31 digestion monitored by qPCR. Short gDNA exposure to Bal31 results in a sudden, seemingly non-specific decrease in gDNA amount followed by a gradual decrease over a prolonged time. (j) Bal31-sensitivity of specific subtelomeric sequences from chromosome arm 2R (pat and gal2) and the resistance of the centromeric ITS region to Bal31 digestion resolved by PCR. gDNA integrity was monitored by amplification of 5 kb-long fragments of the TERT gene. (k) qPCR analysis of specific subtelomere (gal2, pat, gal5), ITS and control sequences documented a decrease of subtelomeric sequences in relation to their position in the subtelomere. Relative DNA levels were calculated by the ΔCt method (i) or ΔΔCt method [79] using ubiquitine-10 as a reference gene relative to the nontreated DNA sample (k). Color coding is the same for (h–k). Pictures were adapted by courtesy of Dr. Terezie Mandáková (e,f) and Prof. Andrew Leitch (d), scale bars are 10 µm.
When such clusters are big enough, these can be detected by FISH (Figure 1d–f) and distinguished from telomeres (e.g., [73][75][80][81][82][83][84][85][86]). If they are shorter than the detection limit of this method, they can still show a positive signal when investigated by Southern hybridization or primer extension (Figure 1g). The origin, evolution and function of ITSs are still subject to much discussion [35][71][86][87][88][89][90]. The massive areas of ITSs often found in pericentromeric regions can be explained as the result of mechanisms such as unequal gene conversion, crossing-over, DNA replication slippage and rolling circle replication of extrachromosomal circular DNA. Some ITSs co-localize with sites of chromosomal breakage and are described as remnants of ancient chromosomal rearrangements, such as during primate evolution [91]. A similar view holds for human ITSs arranged as head-to-head blocks of telomeric repeats that seem to result from the terminal fusion of ancestor chromosomes [41][92].
Researchers are still far from understanding the interplay of mechanisms that are activated during genome instability. It has long been considered that overall change in chromosome architecture can result from breakage-fusion-bridge cycles, a phenomenon first described in maize ([93], reviewed in [94]). The classic theory behind this is that a chromosome with one end broken during meiotic crossing-over can fuse with another such broken chromosome, leading to the formation of a “bridge“ conformation chromosome with two centromeres during the subsequent cycle of meiosis. This bridged chromosome is then ultimately cleaved into two daughter chromatids, but not necessarily at the site of the original breakage. This can lead to sequence deletion or replication on subsequently-healed daughter chromatids [93]. Experimental examination of this theory in Caenorhabditis elegans revealed evidence of such cycles, but also suggested more complex chromatin rearrangements can arise [95]. These more extensive rearrangements are proposed to arise from stalled replication events followed by template switching as may occur in areas with high-homology satellite sequences [95]. A simpler phenomenon is where non-reciprocal translocations can occur during break-induced DNA replication ([96], reviewed in [97]). Broken chromosomes are proposed to invade intact chromosomes with areas of homology during the G1 or G2 phase of the cell cycle, initiating DNA repair with the sequence from the other chromosome arm, possibly acquiring new genes and a telomere in the process [96]. Similar genome instability is also possible when telomeres are lost, making chromosome ends indistinguishable from double-strand breaks [98].
It is clear that telomerase and possible ITSs could have an important role in chromosome rearrangement. For example, when tobacco cells recovered to full cell viability after extensive chromatin fragmentation induced by cadmium stress, this was accompanied by a concomitant increase in telomerase activity [99]. Wheat chromosome end healing after gametocidal gene-induced breakage, efficient telomere healing by telomerase and stabilization of holocentric chromosomes in irradiated Luzula elegans plants were also previously reported [100][101]. Interestingly, when constructs containing telomeric arrays are introduced into mammalian or plant cells, the sites of integration become fragile, chromosomal breakage is induced and the new ends are stabilized [102][103][104]. Telomere-mediated chromosomal truncation has even been employed as a chromosome engineering technique [105][106][107][108]. All this supports the hypothesis that ITSs are preferred sites for breakage and that telomere-like repeats at a break site may favor chromosomal healing [87].

4. Telomere Proteins

Chromosomal DNA in cells associates with proteins that fold these long polymeric molecules into condensed, ordered forms. Most of the DNA sequence, including genes, subtelomeric satellites and the proximal sections of telomeres is folded into a series of compact but dynamic protein-DNA complexes called nucleosomes [109]. In 2001, Fajkus and Trifonov [110] proposed telomeric nucleosomes are packed in a variant, columnar chromatin structure. Recently, the formation of this structure was confirmed experimentally using cryoelectron microscopy [111]. The ends of telomeres associate with a more diverse set of proteins depending on organism that maintain a 3′ single-stranded overhang (aka G-overhang), recruiting enzymes to lengthen the 3′ strand and shorten the 5′ strand which induce and stabilize t-loop formation (reviewed in [9][112]). These mechanisms protect telomeric DNA, prevent aberrant DNA repair and mediate interactions with telomerase (see above, [112]). In Arabidopsis and Chlamydomonas some telomere ends are instead blunt, with no or little 3′ overhang, although it is unknown whether this is a special feature of these organisms or a more widespread characteristic [113][114][115].
Of principal interest to telomere researchers are the specialist proteins that interact with the distal sections of telomeres at the ends of chromosomes. Two major telomere protecting complexes have been described, CST and shelterin. These were initially thought to be alternative mutually exclusive systems, but the search for homologues revealed that many eukaryotes, including humans, had both systems able to work in parallel [116][117][118][119]. Continuing research focused on looking for homologues of human systems across all eukaryotes, however this approach has had only partial success (reviewed in [120]). The CST complex is largely conserved in eukaryotes [121] in terms of function, if not necessarily the sequence of its components [122][123][124]. CST binds ssDNA and recruits Pol1α primase for C-rich strand synthesis and also has a role in preventing stalled replication forks (for recent advancements see [119] and references herein). In comparison, shelterin (reviewed in [112]) coats telomeric DNA generally and interacts with telomerase for G-rich strand synthesis. Shelterin is not present in all eukaryotes, although most have an identifiable protein family that occupies the same role (Figure 2). In addition to these larger end-protection protein complexes, there is a highly conserved heterodimer of proteins called Ku70/Ku80 that is normally involved in non-homologous end-joining events, but which also has an enigmatic role in telomeres. This complex binds dsDNA ends non-specifically, but is known to interact with components of shelterin in mammals and telomerase RNA in yeast (reviewed in [125][126]).
Genes 13 01663 g005 550
Figure 2. Telomere protection by protein complexes. (a) The six core units of shelterin [112] form a complex coating distal telomeres, although stoichiometries of subunits may vary. TRF2 forms T-loops and binds the double-stranded vertebrate telomeric sequence. TRF1 assists this binding, TIN2 and TPP1 form the core of the complex and control other protein-protein interactions and POT1 can bind single-stranded telomeric repeats to stabilize the T-loop. (b) Fission yeast shelterin [127] is analogous to vertebrates but differs in stoichiometries of proteins. (c) Drosophila terminin [128] has a similar function to shelterin although the precise roles of components that share little homology with shelterin components are speculative. (d) Budding yeast telosomes [129]. Rap1 binds telomeric DNA and can be complexed into dimers by Rif2 or tetramers by Rif1. The entire assembly is proposed to form a velcro-like coating of telomeres although to date structural studies of this complex are on dsDNA only, so any interaction with 3′ overhangs is speculative.
Shelterin has a dynamic composition and variant complexes bind the entire length of distal telomeres, there are six core protein components in humans which are more-or-less thought to be conserved in mammals [112][130]. Telomeric repeat binding factor 2 (TRF2) binds the telomeric DNA motif with nanomolar affinity via a SANT/Myb domain sometimes termed the telobox in older literature [131], not to be confused with interstitial telomeric motifs, which are also called teloboxes [89]. TRF2 binds dsDNA and homodimerization enhances this process. It is also proposed to have helicase-like activity where it can wrap dsDNA from near to the telomere end around itself causing steric torsion in the telomere end that encourages T-loop formation. Consistent with this, TRF2 is both necessary for T-loop formation by shelterin and capable of forming T-loops in the absence of any other shelterin components [132]. TRF1 is a highly homologous protein to TRF2 which only binds telomeric repeats and lacks T-loop forming ability. Both proteins (possibly as homodimers) bind TRF1-interacting protein (TIN2) to form the core dsDNA binding subunit of shelterin [133]. TIN2 binds TINT1/PIP1/PTOP1 (TPP1) which in turn binds protection of telomeres 1 (POT1), a protein with multiple OB-fold domains that can bind ssDNA and which is thought to be the main interactor with the 3′ overhang in the complete shelterin complex. TRF2 alone can also recruit repressor/activator protein 1 (RAP1) as the sixth member of core shelterin and the interactions between shelterin subunits can generally occur across multiple protein surfaces [133]. TPP1 in complex with POT1 interacts with telomerase as part of the coordination of telomerase and shelterin protein complexes [119][134].
Unsurprisingly, Drosophila has evolved a separate group of proteins in a complex called terminin to protect the retrotransposon-derived sequences at the ends of its chromosomes. Terminin was identified from the larval brain cells of mutant flies with end-fused chromosomes and consists of a core of heterochromatin protein 1/origin recognition complex-associated protein (HOAP), Modigliani (Moi) and an OB-fold protein called Verrocchio (Ver) [135]. Whilst fission yeast has a shelterin complex made from paralogues of human proteins [136][137], budding yeast, instead has a velcro-like network of proteins called the telosome. This consists of Rap1, a general transcription factor which coats double-stranded telomeric DNA, Rif1 which binds DNA via a Myb domain and Rif2 which binds DNA via an AAA+ domain. Rif 1 and Rif2 can bind four or two molecules of Rap1 respectively through binding domains attached to long disordered chains to form a dense protein network ([138], reviewed in [139]). The system in plants is not yet clear (reviewed in [140]).
Although plant proteins that share some sequence homology to human shelterin proteins have been identified (summarized and reviewed in [141][142]), including those with C-terminal Myb domains similar to TRF1 and TRF2, these do not have any obvious end-protection role [143]. The only definitive double-stranded telomeric DNA binding proteins so far characterized in plants are the telomere repeat binding proteins (TRB1–3) [144][145][146]. These proteins bind to Arabidopsis telomeres in vivo [146][147], and TRB1 colocalizes with telomeres when introduced to Nicotiana benthamiana in live cell imaging studies, suggesting a general role for these proteins at plant telomeres [148][149]. TRBs have histone-like domains that allow multimerization and binding to telobox-related DNA motifs in a multitude of chromosome sites and N-terminal Myb domains that specifically bind double stranded telomeric DNA [144][145][146][150][151]. Similar to TPP1 in human shelterin, TRBs can interact with telomerase and so together with DNA binding and multimerization it is easy to draw parallels with other end-protecting proteins [131][141][145][146]. It can be speculated that in addition to their other regulatory DNA-binding roles [150][151][152], TRBs could form some sort of end-protection framework, similar to the telosome in yeast. Alternatively, it could simply be that any end-protection proteins in plants are sufficiently variant from other organisms to have eluded discovery so far.
One final quirk in plant telomere biology is the occurrence of blunt-ended telomeres. Some blunt DNA ends in Arabidopsis [153] are known to at least temporarily bind Ku70/80, a ubiquitous DNA end-protecting protein complex that is part of the normal double-strand break maintenance mechanism. Studies in budding yeast and human cells revealed that Ku can interact with telomeric chromatin either by directly binding to telomeric DNA or via interaction with telomere associated proteins, including the shelterin subunits such as TRF1, TRF2 and Rap1 [154][155][156]. Studies using mice revealed considerable telomere abnormalities where Ku is knocked out, but phenotypes are complex enough that a specific role is difficult to ascertain [157][158]. In yeast, Ku also binds the telomerase RNA TLC1 separately from telomere ends in a mutually exclusive fashion, and is required to maintain levels and nuclear localization of TLC1. YKu association with telomeres is independent of its association with TLC1 RNA and occurs throughout the cell cycle [159][160]. As with other eukaryotic systems, the Ku heterodimer in Arabidopsis forms a tube that slides onto and encircles the double-stranded telomere from one free end, providing simple end-protection without translocating inward [115][153]. It is so far unknown whether Ku-protected blunt ends in Arabidopsis and Chlamydomonas are unique to these organisms or whether a more widespread phenomenon is yet to be found in other eukaryotes. It is possible that these are an evolutionary step that limits the amount of work that telomerase has to conduct or provides cells without telomerase more stability during proliferation [161].

5. How to Find a Telomere Candidate

Experimental approaches which have been used successfully in the past to characterize telomeres de novo (summarized in [162]) comprise proof of the end-protection function of newly-discovered sequence in vivo [163][164], genomic DNA library screening with verification of terminal position by BAL31 digestion and Southern hybridization [16][17][62][80],[165],[166] cloning of telomerase products [54][55][58][64][167][168] and a novel combination of genomic and transcriptomic studies with classical methods [62][162][166][169]. Today raw data or assembled contigs generated by researchers or from public NGS (next generation sequencing) datasets can be mined for repetitive sequences using, e.g., Tandem Repeats Finder [170] and/or RepeatExplorer [171]. New ways of in silico analysis in combination with experimental approaches for the identification and verification of novel telomere sequences were used e.g., in yeast Lachancea sp. [47] beetle Anoplotrupes stercorosus [67], a plant with human-like telomere sequence Zostera marina (Alismatales) [172] and also a plant with unusual telomere type A. cepa [62]. Moreover, comparative transcriptome study led to identification of telomerase RNA (TR) subunits and telomeric repeats across the entire land plant phylogeny [169]. Subsequently, a new bioinformatic approach based on prediction of TR subunits in combination with results from Tandem Repeats Finder resulted in a broad identification of telomere sequences in green algae, ciliates and Stramenopiles including novel types TATAGGG, TGTTAGGG, TGTAAGGG and demonstrated the deep evolutionary TR origin in the megagroup Diaphoretickes  [173].

This entry is adapted from the peer-reviewed paper 10.3390/genes13091663

References

  1. Kreplak, J.; Madoui, M.A.; Capal, P.; Novak, P.; Labadie, K.; Aubert, G.; Bayer, P.E.; Gali, K.K.; Syme, R.A.; Main, D.; et al. A reference genome for pea provides insight into legume genome evolution. Nat. Genet. 2019, 51, 1411–1422.
  2. The Honeybee Genome Sequencing Consortium. Insights into social insects from the genome of the honeybee Apis mellifera. Nature 2006, 443, 931–949.
  3. McClintock, B. The fusion of broken chromosome ends of sister half-chromatids following chromatid breakage at meiotic anaphases. MO Agric. Exp. Stn. Res. Bull. 1938, 290, 1–48.
  4. Muller, H. The remaking of chromosomes. Collect. Net. 1938, 13, 182–198.
  5. Biscotti, M.A.; Olmo, E.; Heslop-Harrison, J.S. Repetitive DNA in eukaryotic genomes. Chromosome Res. 2015, 23, 415–420.
  6. Thakur, J.; Packiaraj, J.; Henikoff, S. Sequence, Chromatin and Evolution of Satellite DNA. Int. J. Mol. Sci. 2021, 22, 4309.
  7. Peska, V.; Garcia, S. Origin, Diversity, and Evolution of Telomere Sequences in Plants. Front. Plant Sci. 2020, 11, 117.
  8. Mikhail, F.M. Chromosomal Basis of Inheritance. In Emery and Rimoin’s Principles and Practice of Medical Genetics and Genomics, 7th ed.; Pyeritz, R.E., Korf, B.R., Grody, W.W., Eds.; Academic Press: Cambridge, MA, USA, 2019; pp. 237–265.
  9. Tomaska, L.; Cesare, A.J.; AlTurki, T.M.; Griffith, J.D. Twenty years of t-loops: A case study for the importance of collaboration in molecular biology. DNA Repair 2020, 94, 102901.
  10. Bryan, T.M. G-Quadruplexes at Telomeres: Friend or Foe? Molecules 2020, 25, 3686.
  11. Mason, J.M.; Randall, T.A.; Capkova Frydrychova, R. Telomerase lost? Chromosoma 2016, 125, 65–73.
  12. Muyle, A.; Marais, G.A.B.; Bacovsky, V.; Hobza, R.; Lenormand, T. Dosage compensation evolution in plants: Theories, controversies and mechanisms. Philos. Trans. R. Soc. Lond. B Biol. Sci. 2022, 377, 20210222.
  13. Houben, A.; Banaei-Moghaddam, A.M.; Klemme, S.; Timmis, J.N. Evolution and biology of supernumerary B chromosomes. Cell Mol. Life Sci. 2014, 71, 467–478.
  14. Meyne, J.; Ratliff, R.L.; Moyzis, R.K. Conservation of the human telomere sequence (TTAGGG)n among vertebrates. Proc. Natl. Acad. Sci. USA 1989, 86, 7049–7053.
  15. Fuchs, J.; Brandes, A.; Schubert, I. Telomere sequence localization and karyotype evolution in higher plants. Plant Syst. Evol. 1995, 196, 227–241.
  16. Okazaki, S.; Tsuchida, K.; Maekawa, H.; Ishikawa, H.; Fujiwara, H. Identification of a pentanucleotide telomeric sequence, (TTAGG)n, in the silkworm Bombyx mori and in other insects. Mol. Cell. Biol. 1993, 13, 1424–1432.
  17. Richards, E.J.; Ausubel, F.M. Isolation of a higher eukaryotic telomere from Arabidopsis thaliana. Cell 1988, 53, 127–136.
  18. Sahara, K.; Marec, F.; Traut, W. TTAGG telomeric repeats in chromosomes of some insects and other arthropods. Chromosome Res. 1999, 7, 449–460.
  19. Fajkus, J.; Sykorova, E.; Leitch, A.R. Telomeres in evolution and evolution of telomeres. Chromosome Res. 2005, 13, 469–479.
  20. Blackburn, E.H.; Gall, J.G. Tandemly Repeated Sequence at Termini of Extrachromosomal Ribosomal-Rna Genes in Tetrahymena. J. Mol. Biol. 1978, 120, 33–53.
  21. Greider, C.W.; Blackburn, E.H. Identification of a specific telomere terminal transferase activity in Tetrahymena extracts. Cell 1985, 43, 405–413.
  22. Greider, C.W.; Blackburn, E.H. The telomere terminal transferase of Tetrahymena is a ribonucleoprotein enzyme with two kinds of primer specificity. Cell 1987, 51, 887–898.
  23. Shay, J.W.; Wright, W.E. Telomeres and telomerase: Three decades of progress. Nat. Rev. Genet. 2019, 20, 299–309.
  24. Lopez, C.C.; Nielsen, L.; Edstrom, J.E. Terminal long tandem repeats in chromosomes form Chironomus pallidivittatus. Mol. Cell. Biol. 1996, 16, 3285–3290.
  25. Nielsen, L.; Edstrom, J.E. Complex telomere-associated repeat units in members of the genus Chironomus evolve from sequences similar to simple telomeric repeats. Mol. Cell. Biol. 1993, 13, 1583–1589.
  26. Roth, C.W.; Kobeski, F.; Walter, M.F.; Biessmann, H. Chromosome end elongation by recombination in the mosquito Anopheles gambiae. Mol. Cell. Biol. 1997, 17, 5176–5183.
  27. Madalena, C.R.; Fernandes, T.; Villasante, A.; Gorab, E. Curiously composite structures of a retrotransposon and a complex repeat associated with chromosome ends of Rhynchosciara americana (Diptera: Sciaridae). Chromosome Res. 2010, 18, 587–598.
  28. Rossato, R.M.; Madalena, C.R.; Gorab, E. Unusually short tandem repeats in the chromosome end structure of Rhynchosciara (Diptera: Sciaridae). Genetica 2007, 131, 109–116.
  29. Belfort, M.; Curcio, M.J.; Lue, N.F. Telomerase and retrotransposons: Reverse transcriptases that shaped genomes. Proc. Natl. Acad. Sci. USA 2011, 108, 20304–20310.
  30. Markova, D.N.; Christensen, S.M.; Betran, E. Telomere-Specialized Retroelements in Drosophila: Adaptive Symbionts of the Genome, Neutral, or in Conflict? Bioessays 2020, 42, e1900154.
  31. Garcia, S.; Garnatje, T.; Kovarik, A. Plant rDNA database: Ribosomal DNA loci information goes online. Chromosoma 2012, 121, 389–394.
  32. Sochorova, J.; Garcia, S.; Galvez, F.; Symonova, R.; Kovarik, A. Evolutionary trends in animal ribosomal DNA loci: Introduction to a new online database. Chromosoma 2018, 127, 141–150.
  33. Hemleben, V.; Grierson, D.; Borisjuk, N.; Volkov, R.A.; Kovarik, A. Personal Perspectives on Plant Ribosomal RNA Genes Research: From Precursor-rRNA to Molecular Evolution. Front. Plant Sci. 2021, 12, 797348.
  34. Aguilar, M.; Prieto, P. Telomeres and Subtelomeres Dynamics in the Context of Early Chromosome Interactions During Meiosis and Their Implications in Plant Breeding. Front. Plant Sci. 2021, 12, 672489.
  35. Bolzan, A.D. Chromosomal aberrations involving telomeres and interstitial telomeric sequences. Mutagenesis 2012, 27, 1–15.
  36. Carlton, P.M.; Cande, W.Z. Telomeres act autonomously in maize to organize the meiotic bouquet from a semipolarized chromosome orientation. J. Cell Biol. 2002, 157, 231–242.
  37. Cowan, C.R.; Carlton, P.M.; Cande, W.Z. The polar arrangement of telomeres in interphase and meiosis. Rabl organization and the bouquet. Plant Physiol. 2001, 125, 532–538.
  38. Krejci, K.; Stentoft, J.; Koch, J. Molecular cytogenetics investigation of the telomeres in a case of Philadelphia positive B-ALL with a single telomere expansion. Neoplasia 1999, 1, 492–497.
  39. Samassekou, O.; Li, H.; Hebert, J.; Ntwari, A.; Wang, H.; Cliche, C.G.; Bouchard, E.; Huang, S.; Yan, J. Chromosome arm-specific long telomeres: A new clonal event in primary chronic myelogenous leukemia cells. Neoplasia 2011, 13, 550–560.
  40. Schober, H.; Kalck, V.; Vega-Palas, M.A.; Van Houwe, G.; Sage, D.; Unser, M.; Gartenberg, M.R.; Gasser, S.M. Controlled exchange of chromosomal arms reveals principles driving telomere interactions in yeast. Genome Res. 2008, 18, 261–271.
  41. Slijepcevic, P. Telomeres and mechanisms of Robertsonian fusion. Chromosoma 1998, 107, 136–140.
  42. Shampay, J.; Szostak, J.W.; Blackburn, E.H. DNA sequences of telomeres maintained in yeast. Nature 1984, 310, 154–157.
  43. Greider, C.W. Telomere length regulation. Annu. Rev. Biochem. 1996, 65, 337–365.
  44. Matsumoto, T.; Fukui, K.; Niwa, O.; Sugawara, N.; Szostak, J.W.; Yanagida, M. Identification of healed terminal DNA fragments in linear minichromosomes of Schizosaccharomyces pombe. Mol. Cell. Biol. 1987, 7, 4424–4430.
  45. McEachern, M.J.; Hicks, J.B. Unusually large telomeric repeats in the yeast Candida albicans. Mol. Cell. Biol. 1993, 13, 551–560.
  46. McEachern, M.J.; Blackburn, E.H. A conserved sequence motif within the exceptionally diverse telomeric sequences of budding yeasts. Proc. Natl. Acad. Sci. USA 1994, 91, 3453–3457.
  47. Peska, V.; Fajkus, P.; Bubenik, M.; Brazda, V.; Bohalova, N.; Dvoracek, V.; Fajkus, J.; Garcia, S. Extraordinary diversity of telomeres, telomerase RNAs and their template regions in Saccharomycetaceae. Sci. Rep. 2021, 11, 12784.
  48. Cervenak, F.; Jurikova, K.; Devillers, H.; Kaffe, B.; Khatib, A.; Bonnell, E.; Sopkovicova, M.; Wellinger, R.J.; Nosek, J.; Tzfati, Y.; et al. Identification of telomerase RNAs in species of the Yarrowia clade provides insights into the co-evolution of telomerase, telomeric repeats and telomere-binding proteins. Sci. Rep. 2019, 9, 13365.
  49. Cervenak, F.; Sepsiova, R.; Nosek, J.; Tomaska, L. Step-by-Step Evolution of Telomeres: Lessons from Yeasts. Genome Biol. Evol. 2021, 13, evaa268.
  50. Sykorova, E.; Lim, K.Y.; Chase, M.W.; Knapp, S.; Leitch, I.J.; Leitch, A.R.; Fajkus, J. The absence of Arabidopsis-type telomeres in Cestrum and closely related genera Vestia and Sessea (Solanaceae): First evidence from eudicots. Plant J. 2003, 34, 283–291.
  51. Adams, S.P.; Hartman, T.P.; Lim, K.Y.; Chase, M.W.; Bennett, M.D.; Leitch, I.J.; Leitch, A.R. Loss and recovery of Arabidopsis-type telomere repeat sequences 5′-(TTTAGGG)(n)-3′ in the evolution of a major radiation of flowering plants. Proc. Biol. Sci. 2001, 268, 1541–1546.
  52. Adams, S.P.; Leitch, I.J.; Bennett, M.D.; Leitch, A.R. Aloe L.—A second plant family without (TTTAGGG)n telomeres. Chromosoma 2000, 109, 201–205.
  53. Frydrychova, R.; Marec, F. Repeated losses of TTAGG telomere repeats in evolution of beetles (Coleoptera). Genetica 2002, 115, 179–187.
  54. Fulneckova, J.; Sevcikova, T.; Fajkus, J.; Lukesova, A.; Lukes, M.; Vlcek, C.; Lang, B.F.; Kim, E.; Elias, M.; Sykorova, E. A broad phylogenetic survey unveils the diversity and evolution of telomeres in eukaryotes. Genome Biol. Evol. 2013, 5, 468–483.
  55. Sykorova, E.; Lim, K.Y.; Kunicka, Z.; Chase, M.W.; Bennett, M.D.; Fajkus, J.; Leitch, A.R. Telomere variability in the monocotyledonous plant order Asparagales. Proc. R. Soc. Lond. B Biol. Sci. 2003, 270, 1893–1904.
  56. Sykorova, E.; Fajkus, J.; Meznikova, M.; Lim, K.Y.; Neplechova, K.; Blattner, F.R.; Chase, M.W.; Leitch, A.R. Minisatellite telomeres occur in the family Alliaceae but are lost in Allium. Am. J. Bot. 2006, 93, 814–823.
  57. Bombarova, M.; Vitkova, M.; Spakulova, M.; Koubkova, B. Telomere analysis of platyhelminths and acanthocephalans by FISH and Southern hybridization. Genome 2009, 52, 897–903.
  58. Fulneckova, J.; Hasikova, T.; Fajkus, J.; Lukesova, A.; Elias, M.; Sykorova, E. Dynamic evolution of telomeric sequences in the green algal order Chlamydomonadales. Genome Biol. Evol. 2012, 4, 248–264.
  59. Mravinac, B.; Mestrovic, N.; Cavrak, V.V.; Plohl, M. TCAGG, an alternative telomeric sequence in insects. Chromosoma 2011, 120, 367–376.
  60. Vitkova, M.; Kral, J.; Traut, W.; Zrzavy, J.; Marec, F. The evolutionary origin of insect telomeric repeats, (TTAGG)n. Chromosome Res. 2005, 13, 145–156.
  61. Weiss, H.; Scherthan, H. Aloe spp.—Plants with vertebrate-like telomeric sequences. Chromosome Res. 2002, 10, 155–164.
  62. Fajkus, P.; Peska, V.; Sitova, Z.; Fulneckova, J.; Dvorackova, M.; Gogela, R.; Sykorova, E.; Hapala, J.; Fajkus, J. Allium telomeres unmasked: The unusual telomeric sequence (CTCGGTTATGGG)n is synthesized by telomerase. Plant J. 2016, 85, 337–347.
  63. Friesen, N.; Fritsch, R.M.; Blattner, F.R. Phylogeny and new intrageneric classification of Allium (Alliaceae) based on nuclear rDNA ITS sequences. Aliso 2006, 22, 372–395.
  64. Fulneckova, J.; Sevcikova, T.; Lukesova, A.; Sykorova, E. Transitions between the Arabidopsis-type and the human-type telomere sequence in green algae (clade Caudivolvoxa, Chlamydomonadales). Chromosoma 2016, 125, 437–451.
  65. Derelle, R.; Torruella, G.; Klimes, V.; Brinkmann, H.; Kim, E.; Vlcek, C.; Lang, B.F.; Elias, M. Bacterial proteins pinpoint a single eukaryotic root. Proc. Natl. Acad. Sci. USA 2015, 112, E693–E699.
  66. Elias, M. Protist diversity: Novel groups enrich the algal tree of life. Curr. Biol. 2021, 31, R733–R735.
  67. Prusakova, D.; Peska, V.; Pekar, S.; Bubenik, M.; Cizek, L.; Bezdek, A.; Capkova Frydrychova, R. Telomeric DNA sequences in beetle taxa vary with species richness. Sci. Rep. 2021, 11, 13319.
  68. Kuznetsova, V.; Maryańska-Nadachowska, A.; Anokhin, B.; Shapoval, N.; Shapoval, A. Chromosomal analysis of eight species of dragonflies (Anisoptera) and damselflies (Zygoptera) using conventional cytogenetics and fluorescence in situ hybridization: Insights into the karyotype evolution of the ancient insect order Odonata. J. Zool. Syst. Evol. Res. 2021, 59, 387–399.
  69. Gorab, E. Chromosome End Diversification in Sciarid Flies. Cells 2020, 9, 2425.
  70. Zhou, Y.; Wang, Y.; Xiong, X.; Appel, A.G.; Zhang, C.; Wang, X. Profiles of telomeric repeats in Insecta reveal diverse forms of telomeric motifs in Hymenopterans. Life Sci. Alliance 2022, 5.
  71. Aksenova, A.Y.; Mirkin, S.M. At the Beginning of the End and in the Middle of the Beginning: Structure and Maintenance of Telomeric DNA Repeats and Interstitial Telomeric Sequences. Genes 2019, 10, 118.
  72. Flint, J.; Bates, G.P.; Clark, K.; Dorman, A.; Willingham, D.; Roe, B.A.; Micklem, G.; Higgs, D.R.; Louis, E.J. Sequence comparison of human and yeast telomeres identifies structurally distinct subtelomeric domains. Hum. Mol. Genet. 1997, 6, 1305–1313.
  73. Sykorova, E.; Lim, K.Y.; Fajkus, J.; Leitch, A.R. The signature of the Cestrum genome suggests an evolutionary response to the loss of (TTTAGGG)n telomeres. Chromosoma 2003, 112, 164–172.
  74. Majerova, E.; Mandakova, T.; Vu, G.T.; Fajkus, J.; Lysak, M.A.; Fojtova, M. Chromatin features of plant telomeric sequences at terminal vs. internal positions. Front. Plant Sci. 2014, 5, 593.
  75. Mandakova, T.; Zozomova-Lihova, J.; Kudoh, H.; Zhao, Y.; Lysak, M.A.; Marhold, K. The story of promiscuous crucifers: Origin and genome evolution of an invasive species, Cardamine occulta (Brassicaceae), and its relatives. Ann. Bot. 2019, 124, 209–220.
  76. Naish, M.; Alonge, M.; Wlodzimierz, P.; Tock, A.J.; Abramson, B.W.; Schmucker, A.; Mandakova, T.; Jamge, B.; Lambing, C.; Kuo, P.; et al. The genetic and epigenetic landscape of the Arabidopsis centromeres. Science 2021, 374, eabi7489.
  77. Pich, U.; Schubert, I. Terminal heterochromatin and alternative telomeric sequences in Allium cepa. Chromosome Res. 1998, 6, 315–321.
  78. Sykorova, E.; Fojtova, M.; Peska, V. A polymerase chain reaction-based approach for evaluation of telomere-associated sequences and interstitial telomeric sequences. Anal. Biochem. 2013, 439, 8–10.
  79. Pfaffl, M.W. Quantification strategies in real-time PCR. In A–Z of Quantitative PCR; Bustin, S.A., Ed.; International University Line: La Jolla, CA, USA, 2004; pp. 87–112.
  80. Moyzis, R.K.; Buckingham, J.M.; Cram, L.S.; Dani, M.; Deaven, L.L.; Jones, M.D.; Meyne, J.; Ratliff, R.L.; Wu, J.R. A highly conserved repetitive DNA sequence, (TTAGGG)n, present at the telomeres of human chromosomes. Proc. Natl. Acad. Sci. USA 1988, 85, 6622–6626.
  81. Meyne, J.; Baker, R.J.; Hobart, H.H.; Hsu, T.C.; Ryder, O.A.; Ward, O.G.; Wiley, J.E.; Wurster-Hill, D.H.; Yates, T.L.; Moyzis, R.K. Distribution of non-telomeric sites of the (TTAGGG)n telomeric sequence in vertebrate chromosomes. Chromosoma 1990, 99, 3–10.
  82. Maravilla, A.J.; Rosato, M.; Alvarez, I.; Nieto Feliner, G.; Rossello, J.A. Interstitial Arabidopsis-Type Telomeric Repeats in Asteraceae. Plants 2021, 10, 2794.
  83. Richards, E.J.; Goodman, H.M.; Ausubel, F.M. The centromere region of Arabidopsis thaliana chromosome 1 contains telomere-similar sequences. Nucleic Acids Res. 1991, 19, 3351–3357.
  84. Luo, X.; He, Z.; Liu, J.; Wu, H.; Gong, X. FISH Mapping of Telomeric and Non-Telomeric (AG3T3)3 Reveal the Chromosome Numbers and Chromosome Rearrangements of 41 Woody Plants. Genes 2022, 13, 1239.
  85. Rovatsos, M.; Kratochvil, L.; Altmanova, M.; Johnson Pokorna, M. Interstitial Telomeric Motifs in Squamate Reptiles: When the Exceptions Outnumber the Rule. PLoS ONE 2015, 10, e0134985.
  86. Rego, A.; Marec, F. Telomeric and interstitial telomeric sequences in holokinetic chromosomes of Lepidoptera: Telomeric DNA mediates association between postpachytene bivalents in achiasmatic meiosis of females. Chromosome Res. 2003, 11, 681–694.
  87. Azzalin, C.M.; Nergadze, S.G.; Giulotto, E. Human intrachromosomal telomeric-like repeats: Sequence organization and mechanisms of origin. Chromosoma 2001, 110, 75–82.
  88. Chirino, M.G.; Dalikova, M.; Marec, F.R.; Bressa, M.J. Chromosomal distribution of interstitial telomeric sequences as signs of evolution through chromosome fusion in six species of the giant water bugs (Hemiptera, Belostoma). Ecol. Evol. 2017, 7, 5227–5235.
  89. Gaspin, C.; Rami, J.F.; Lescure, B. Distribution of short interstitial telomere motifs in two plant genomes: Putative origin and function. BMC Plant Biol. 2010, 10, 283.
  90. Aksenova, A.Y.; Han, G.; Shishkin, A.A.; Volkov, K.V.; Mirkin, S.M. Expansion of Interstitial Telomeric Sequences in Yeast. Cell Rep. 2015, 13, 1545–1551.
  91. Nergadze, S.G.; Rocchi, M.; Azzalin, C.M.; Mondello, C.; Giulotto, E. Insertion of telomeric repeats at intrachromosomal break sites during primate evolution. Genome Res. 2004, 14, 1704–1710.
  92. Ijdo, J.W.; Baldini, A.; Ward, D.C.; Reeders, S.T.; Wells, R.A. Origin of human chromosome 2: An ancestral telomere-telomere fusion. Proc. Natl. Acad. Sci. USA 1991, 88, 9051–9055.
  93. McClintock, B. The stability of broken ends of chromosomes in Zea mays. Genetics 1940, 26, 234–282.
  94. Bailey, S.M.; Murnane, J.P. Telomeres, chromosome instability and cancer. Nucleic Acids Res. 2006, 34, 2408–2417.
  95. Lowden, M.R.; Flibotte, S.; Moerman, D.G.; Ahmed, S. DNA synthesis generates terminal duplications that seal end-to-end chromosome fusions. Science 2011, 332, 468–471.
  96. Bosco, G.; Haber, J.E. Chromosome break-induced DNA replication leads to nonreciprocal translocations and telomere capture. Genetics 1998, 150, 1037–1047.
  97. Kramara, J.; Osia, B.; Malkova, A. Break-Induced Replication: The Where, The Why, and The How. Trends Genet. 2018, 34, 518–531.
  98. Hackett, J.A.; Feldser, D.M.; Greider, C.W. Telomere dysfunction increases mutation rate and genomic instability. Cell 2001, 106, 275–286.
  99. Fojtova, M.; Fulneckova, J.; Fajkus, J.; Kovarik, A. Recovery of tobacco cells from cadmium stress is accompanied by DNA repair and increased telomerase activity. J. Exp. Bot. 2002, 53, 2151–2158.
  100. Jankowska, M.; Fuchs, J.; Klocke, E.; Fojtova, M.; Polanska, P.; Fajkus, J.; Schubert, V.; Houben, A. Holokinetic centromeres and efficient telomere healing enable rapid karyotype evolution. Chromosoma 2015, 124, 519–528.
  101. Tsujimoto, H.; Yamada, T.; Sasakuma, T. Molecular structure of a wheat chromosome end healed after gametocidal gene-induced breakage. Proc. Natl. Acad. Sci. USA 1997, 94, 3140–3144.
  102. Farr, C.; Fantes, J.; Goodfellow, P.; Cooke, H. Functional reintroduction of human telomeres into mammalian cells. Proc. Natl. Acad. Sci. USA 1991, 88, 7006–7010.
  103. Wilkie, A.O.; Lamb, J.; Harris, P.C.; Finney, R.D.; Higgs, D.R. A truncated human chromosome 16 associated with alpha thalassaemia is stabilized by addition of telomeric repeat (TTAGGG)n. Nature 1990, 346, 868–871.
  104. Yu, W.; Lamb, J.C.; Han, F.; Birchler, J.A. Telomere-mediated chromosomal truncation in maize. Proc. Natl. Acad. Sci. USA 2006, 103, 17331–17336.
  105. Kapusi, E.; Ma, L.; Teo, C.H.; Hensel, G.; Himmelbach, A.; Schubert, I.; Mette, M.F.; Kumlehn, J.; Houben, A. Telomere-mediated truncation of barley chromosomes. Chromosoma 2012, 121, 181–190.
  106. Tamar, S.; Papadopoulou, B. A telomere-mediated chromosome fragmentation approach to assess mitotic stability and ploidy alterations of Leishmania chromosomes. J. Biol. Chem. 2001, 276, 11662–11673.
  107. Teo, C.H.; Ma, L.; Kapusi, E.; Hensel, G.; Kumlehn, J.; Schubert, I.; Houben, A.; Mette, M.F. Induction of telomere-mediated chromosomal truncation and stability of truncated chromosomes in Arabidopsis thaliana. Plant J. 2011, 68, 28–39.
  108. Tada, S.; Ohkuchi, H.; Matsushita-Morita, M.; Furukawa, I.; Hattori, R.; Suzuki, S.; Kashiwagi, Y.; Kusumoto, K. Telomere-mediated chromosomal truncation in Aspergillus oryzae. J. Biosci. Bioeng. 2015, 119, 43–46.
  109. Kornberg, R.D. Structure of chromatin. Annu. Rev. Biochem. 1977, 46, 931–954.
  110. Fajkus, J.; Trifonov, E.N. Columnar packing of telomeric nucleosomes. Biochem. Biophys. Res. Commun. 2001, 280, 961–963.
  111. Soman, A.; Wong, S.Y.; Korolev, N.; Surya, W.; Lattman, S.; Vogirala, V.K.; Chen, Q.; Berezhnoy, N.V.; van Noort, J.; Rhodes, D.; et al. The columnar structure of human telomeric chromatin suggests mechanisms for telomere maintenance. bioRxiv 2022.
  112. De Lange, T. Shelterin-Mediated Telomere Protection. Annu. Rev. Genet. 2018, 52, 223–247.
  113. Riha, K.; McKnight, T.D.; Fajkus, J.; Vyskot, B.; Shippen, D.E. Analysis of the G-overhang structures on plant telomeres: Evidence for two distinct telomere architectures. Plant J. 2000, 23, 633–641.
  114. Eberhard, S.; Valuchova, S.; Ravat, J.; Fulnecek, J.; Jolivet, P.; Bujaldon, S.; Lemaire, S.D.; Wollman, F.A.; Teixeira, M.T.; Riha, K.; et al. Molecular characterization of Chlamydomonas reinhardtii telomeres and telomerase mutants. Life Sci. Alliance 2019, 2, e201900315.
  115. Kazda, A.; Zellinger, B.; Rossler, M.; Derboven, E.; Kusenda, B.; Riha, K. Chromosome end protection by blunt-ended telomeres. Genes Dev. 2012, 26, 1703–1713.
  116. Miyake, Y.; Nakamura, M.; Nabetani, A.; Shimamura, S.; Tamura, M.; Yonehara, S.; Saito, M.; Ishikawa, F. RPA-like mammalian Ctc1-Stn1-Ten1 complex binds to single-stranded DNA and protects telomeres independently of the Pot1 pathway. Mol. Cell 2009, 36, 193–206.
  117. Wellinger, R.J. The CST complex and telomere maintenance: The exception becomes the rule. Mol. Cell 2009, 36, 168–169.
  118. He, Y.; Song, H.; Chan, H.; Liu, B.; Wang, Y.; Susac, L.; Zhou, Z.H.; Feigon, J. Structure of Tetrahymena telomerase-bound CST with polymerase alpha-primase. Nature 2022, 608, 813–818.
  119. Zaug, A.J.; Goodrich, K.J.; Song, J.J.; Sullivan, A.E.; Cech, T.R. Reconstitution of a telomeric replicon organized by CST. Nature 2022, 608, 819–825.
  120. Rice, C.; Skordalakes, E. Structure and function of the telomeric CST complex. Comput. Struct. Biotechnol. J. 2016, 14, 161–167.
  121. Linger, B.R.; Price, C.M. Conservation of telomere protein complexes: Shuffling through evolution. Crit. Rev. Biochem. Mol. Biol. 2009, 44, 434–446.
  122. Chandra, A.; Hughes, T.R.; Nugent, C.I.; Lundblad, V. Cdc13 both positively and negatively regulates telomere replication. Genes Dev. 2001, 15, 404–414.
  123. Pennock, E.; Buckley, K.; Lundblad, V. Cdc13 delivers separate complexes to the telomere for end protection and replication. Cell 2001, 104, 387–396.
  124. Wan, B.; Tang, T.; Upton, H.; Shuai, J.; Zhou, Y.; Li, S.; Chen, J.; Brunzelle, J.S.; Zeng, Z.; Collins, K.; et al. The Tetrahymena telomerase p75-p45-p19 subcomplex is a unique CST complex. Nat. Struct. Mol. Biol. 2015, 22, 1023–1026.
  125. Fell, V.L.; Schild-Poulter, C. The Ku heterodimer: Function in DNA repair and beyond. Mutat. Res. Rev. Mutat. Res. 2015, 763, 15–29.
  126. Sui, J.; Zhang, S.; Chen, B.P.C. DNA-dependent protein kinase in telomere maintenance and protection. Cell Mol. Biol. Lett. 2020, 25, 2.
  127. Chen, Y. The structural biology of the shelterin complex. Biol. Chem. 2019, 400, 457–466.
  128. Cacchione, S.; Cenci, G.; Raffa, G.D. Silence at the End: How Drosophila Regulates Expression and Transposition of Telomeric Retroelements. J. Mol. Biol. 2020, 432, 4305–4321.
  129. Shi, T.; Bunker, R.D.; Mattarocci, S.; Ribeyre, C.; Faty, M.; Gut, H.; Scrima, A.; Rass, U.; Rubin, S.M.; Shore, D.; et al. Rif1 and Rif2 shape telomere function and architecture through multivalent Rap1 interactions. Cell 2013, 153, 1340–1353.
  130. De Lange, T. Shelterin: The protein complex that shapes and safeguards human telomeres. Genes Dev. 2005, 19, 2100–2110.
  131. Bilaud, T.; Koering, C.E.; Binet-Brasselet, E.; Ancelin, K.; Pollice, A.; Gasser, S.M.; Gilson, E. The telobox, a Myb-related telomeric DNA binding motif found in proteins from yeast, plants and human. Nucleic Acids Res. 1996, 24, 1294–1303.
  132. Stansel, R.M.; de Lange, T.; Griffith, J.D. T-loop assembly in vitro involves binding of TRF2 near the 3’ telomeric overhang. EMBO J. 2001, 20, 5532–5540.
  133. Erdel, F.; Kratz, K.; Willcox, S.; Griffith, J.D.; Greene, E.C.; de Lange, T. Telomere Recognition and Assembly Mechanism of Mammalian Shelterin. Cell Rep. 2017, 18, 41–53.
  134. Zaug, A.J.; Podell, E.R.; Nandakumar, J.; Cech, T.R. Functional interaction between telomere protein TPP1 and telomerase. Genes Dev. 2010, 24, 613–622.
  135. Raffa, G.D.; Raimondo, D.; Sorino, C.; Cugusi, S.; Cenci, G.; Cacchione, S.; Gatti, M.; Ciapponi, L. Verrocchio, a Drosophila OB fold-containing protein, is a component of the terminin telomere-capping complex. Genes Dev. 2010, 24, 1596–1601.
  136. Miyoshi, T.; Kanoh, J.; Saito, M.; Ishikawa, F. Fission yeast Pot1-Tpp1 protects telomeres and regulates telomere length. Science 2008, 320, 1341–1344.
  137. Xue, J.; Chen, H.; Wu, J.; Takeuchi, M.; Inoue, H.; Liu, Y.; Sun, H.; Chen, Y.; Kanoh, J.; Lei, M. Structure of the fission yeast S. pombe telomeric Tpz1-Poz1-Rap1 complex. Cell Res. 2017, 27, 1503–1520.
  138. Shi, K.; Huang, W.M.; Aihara, H. An enzyme-catalyzed multistep DNA refolding mechanism in hairpin telomere formation. PLoS Biol. 2013, 11, e1001472.
  139. Wellinger, R.J.; Zakian, V.A. Everything you ever wanted to know about Saccharomyces cerevisiae telomeres: Beginning to end. Genetics 2012, 191, 1073–1105.
  140. Prochazkova Schrumpfova, P.; Fojtova, M.; Fajkus, J. Telomeres in Plants and Humans: Not So Different, Not So Similar. Cells 2019, 8, 58.
  141. Peska, V.; Schrumpfova, P.P.; Fajkus, J. Using the telobox to search for plant telomere binding proteins. Curr. Protein Pept. Sci. 2011, 12, 75–83.
  142. Prochazkova Schrumpfova, P.; Schorova, S.; Fajkus, J. Telomere- and Telomerase-Associated Proteins and Their Functions in the Plant Cell. Front. Plant Sci. 2016, 7, 851.
  143. Fulcher, N.; Riha, K. Using Centromere Mediated Genome Elimination to Elucidate the Functional Redundancy of Candidate Telomere Binding Proteins in Arabidopsis thaliana. Front. Genet. 2015, 6, 349.
  144. Schrumpfova, P.; Kuchar, M.; Mikova, G.; Skrisovska, L.; Kubicarova, T.; Fajkus, J. Characterization of two Arabidopsis thaliana myb-like proteins showing affinity to telomeric DNA sequence. Genome 2004, 47, 316–324.
  145. Mozgova, I.; Schrumpfova, P.P.; Hofr, C.; Fajkus, J. Functional characterization of domains in AtTRB1, a putative telomere-binding protein in Arabidopsis thaliana. Phytochemistry 2008, 69, 1814–1819.
  146. Schrumpfova, P.P.; Vychodilova, I.; Dvorackova, M.; Majerska, J.; Dokladal, L.; Schorova, S.; Fajkus, J. Telomere repeat binding proteins are functional components of Arabidopsis telomeres and interact with telomerase. Plant J. 2014, 77, 770–781.
  147. Dvorackova, M.; Rossignol, P.; Shaw, P.J.; Koroleva, O.A.; Doonan, J.H.; Fajkus, J. AtTRB1, a telomeric DNA-binding protein from Arabidopsis, is concentrated in the nucleolus and shows highly dynamic association with chromatin. Plant J. 2010, 61, 637–649.
  148. Dreissig, S.; Schiml, S.; Schindele, P.; Weiss, O.; Rutten, T.; Schubert, V.; Gladilin, E.; Mette, M.F.; Puchta, H.; Houben, A. Live-cell CRISPR imaging in plants reveals dynamic telomere movements. Plant J. 2017, 91, 565–573.
  149. Khosravi, S.; Dreissig, S.; Schindele, P.; Wolter, F.; Rutten, T.; Puchta, H.; Houben, A. Live-Cell CRISPR Imaging in Plant Cells with a Telomere-Specific Guide RNA. Methods Mol. Biol. 2020, 2166, 343–356.
  150. Schrumpfova, P.P.; Vychodilova, I.; Hapala, J.; Schorova, S.; Dvoracek, V.; Fajkus, J. Telomere binding protein TRB1 is associated with promoters of translation machinery genes in vivo. Plant Mol. Biol. 2016, 90, 189–206.
  151. Zhou, Y.; Hartwig, B.; James, G.V.; Schneeberger, K.; Turck, F. Complementary Activities of TELOMERE REPEAT BINDING Proteins and Polycomb Group Complexes in Transcriptional Regulation of Target Genes. Plant Cell 2016, 28, 87–101.
  152. Zhou, Y.; Wang, Y.; Krause, K.; Yang, T.; Dongus, J.A.; Zhang, Y.; Turck, F. Telobox motifs recruit CLF/SWN-PRC2 for H3K27me3 deposition via TRB factors in Arabidopsis. Nat. Genet. 2018, 50, 638–644.
  153. Valuchova, S.; Fulnecek, J.; Prokop, Z.; Stolt-Bergner, P.; Janouskova, E.; Hofr, C.; Riha, K. Protection of Arabidopsis Blunt-Ended Telomeres Is Mediated by a Physical Association with the Ku Heterodimer. Plant Cell 2017, 29, 1533–1545.
  154. Hass, E.P.; Zappulla, D.C. The Ku subunit of telomerase binds Sir4 to recruit telomerase to lengthen telomeres in S. cerevisiae. eLife 2015, 4, e07750.
  155. Hsu, H.L.; Gilley, D.; Blackburn, E.H.; Chen, D.J. Ku is associated with the telomere in mammals. Proc. Natl. Acad. Sci. USA 1999, 96, 12454–12458.
  156. Hsu, H.L.; Gilley, D.; Galande, S.A.; Hande, M.P.; Allen, B.; Kim, S.H.; Li, G.C.; Campisi, J.; Kohwi-Shigematsu, T.; Chen, D.J. Ku acts in a unique way at the mammalian telomere to prevent end joining. Genes Dev. 2000, 14, 2807–2812.
  157. D’Adda di Fagagna, F.; Hande, M.P.; Tong, W.M.; Roth, D.; Lansdorp, P.M.; Wang, Z.Q.; Jackson, S.P. Effects of DNA nonhomologous end-joining factors on telomere length and chromosomal stability in mammalian cells. Curr. Biol. 2001, 11, 1192–1196.
  158. Samper, E.; Goytisolo, F.A.; Slijepcevic, P.; van Buul, P.P.; Blasco, M.A. Mammalian Ku86 protein prevents telomeric fusions independently of the length of TTAGGG repeats and the G-strand overhang. EMBO Rep. 2000, 1, 244–252.
  159. Fisher, T.S.; Taggart, A.K.; Zakian, V.A. Cell cycle-dependent regulation of yeast telomerase by Ku. Nat. Struct. Mol. Biol. 2004, 11, 1198–1205.
  160. Larcher, M.V.; Pasquier, E.; MacDonald, R.S.; Wellinger, R.J. Ku Binding on Telomeres Occurs at Sites Distal from the Physical Chromosome Ends. PLoS Genet. 2016, 12, e1006479.
  161. Nelson, A.D.; Shippen, D.E. Blunt-ended telomeres: An alternative ending to the replication and end protection stories. Genes Dev. 2012, 26, 1648–1652.
  162. Peska, V.; Sitova, Z.; Fajkus, P.; Fajkus, J. BAL31-NGS approach for identification of telomeres de novo in large genomes. Methods 2017, 114, 16–27.
  163. Szostak, J.W.; Blackburn, E.H. Cloning yeast telomeres on linear plasmid vectors. Cell 1982, 29, 245–255.
  164. Perrot, M.; Barreau, C.; Begueret, J. Nonintegrative transformation in the filamentous fungus Podospora anserina: Stabilization of a linear vector by the chromosomal ends of Tetrahymena thermophila. Mol. Cell. Biol. 1987, 7, 1725–1730.
  165. Petracek, M.E.; Lefebvre, P.A.; Silflow, C.D.; Berman, J. Chlamydomonas telomere sequences are A+T-rich but contain three consecutive G-C base pairs. Proc. Natl. Acad. Sci. USA 1990, 87, 8222–8226.
  166. Peska, V.; Fajkus, P.; Fojtova, M.; Dvorackova, M.; Hapala, J.; Dvoracek, V.; Polanska, P.; Leitch, A.R.; Sykorova, E.; Fajkus, J. Characterisation of an unusual telomere motif (TTTTTTAGGG)n in the plant Cestrum elegans (Solanaceae), a species with a large genome. Plant J. 2015, 82, 644–654.
  167. Schumpert, C.; Nelson, J.; Kim, E.; Dudycha, J.L.; Patel, R.C. Telomerase activity and telomere length in Daphnia. PLoS ONE 2015, 10, e0127196.
  168. Uzlikova, M.; Fulneckova, J.; Weisz, F.; Sykorova, E.; Nohynkova, E.; Tumova, P. Characterization of telomeres and telomerase from the single-celled eukaryote Giardia intestinalis. Mol. Biochem. Parasitol. 2017, 211, 31–38.
  169. Fajkus, P.; Peska, V.; Zavodnik, M.; Fojtova, M.; Fulneckova, J.; Dobias, S.; Kilar, A.; Dvorackova, M.; Zachova, D.; Necasova, I.; et al. Telomerase RNAs in land plants. Nucleic Acids Res. 2019, 47, 9842–9856.
  170. Benson, G. Tandem repeats finder: A program to analyze DNA sequences. Nucleic Acids Res. 1999, 27, 573–580.
  171. Novak, P.; Neumann, P.; Macas, J. Global analysis of repetitive DNA from unassembled sequence reads using RepeatExplorer2. Nat. Protoc. 2020, 15, 3745–3776.
  172. Peska, V.; Matl, M.; Mandakova, T.; Vitales, D.; Fajkus, P.; Fajkus, J.; Garcia, S. Human-like telomeres in Zostera marina reveal a mode of transition from the plant to the human telomeric sequences. J. Exp. Bot. 2020, 71, 5786–5793.
  173. Fajkus, P.; Kilar, A.; Nelson, A.D.L.; Hola, M.; Peska, V.; Goffova, I.; Fojtova, M.; Zachova, D.; Fulneckova, J.; Fajkus, J. Evolution of plant telomerase RNAs: Farther to the past, deeper to the roots. Nucleic Acids Res. 2021, 49, 7680–7694.
More
© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license; additional terms may apply. By using this site, you agree to the Terms and Conditions and Privacy Policy.
This entry is offline, you can click here to edit this entry!
ScholarVision Creations
Feedback