Researchers discovery that coronaviruses rely on the enzyme Topoisomerase III-ß (TOP3B) as a host factor

0
264

In an effort to help stop the spread of COVID-19, a team of researchers is trying to block a key enzyme in the human cell that the virus needs to thrive.

FIU Biomolecular Sciences Institute Director Yuk-Ching Tse-Dinh and Associate Director Prem Chapagain have teamed up with researchers at the University of Texas Medical Branch (UTMB) at Galveston and National Cancer Institute.

The team is searching for potential treatment leads, hoping at least one of them will reduce the spread of the virus in infected individuals.

Currently there are no evidence-based treatments for COVID-19 or any other coronaviruses.

Last month, Dr. Mariano Garcia-Blanco from UTMB reported in BioRxiv his discovery that coronaviruses rely on the enzyme Topoisomerase III-ß (TOP3B) as a host factor. Normally, topoisomerases aid in DNA replication within human cells.

But when certain viruses, including Dengue, Zika and COVID-19, latch on to TOP3B, they use the enzyme to help assemble new copies of the virus, turning infected cells into virus factories.

Upon this discovery, Garcia-Blanco enlisted the help of Tse-Dinh, an internationally known expert in topoisomerases, to start screening for a drug that could block the enzyme and prevent COVID-19 from multiplying in high numbers in a person’s body.

“We approached FIU as a place where the best inhibitors can come from,” Garcia-Blanco said.

Tse-Dinh is starting with drugs already approved by the Food and Drug Administration because she is looking for a treatment that can be tested quickly on patients.

“We are targeting a host factor, something already in the human cell that the virus is hijacking,” Tse-Dinh said.

“If the cell’s TOP3B is not working, the virus would not do so well.”

The researchers believe antiviral medications could be administered in tandem with a TOP3B inhibitor to deliver a double blow to the virus and improve chances of recovery for those who are critically ill.

“Our researchers are focused on real, tangible solutions for the greatest challenges of our time,” said Mike Heithaus, dean of FIU’s College of Arts, Sciences & Education.

“This effort, if successful, has potential beyond the current pandemic. It could provide a new approach to treating other viruses and even emerging ones.”

But with thousands of FDA-approved drugs, knowing where to begin is a real challenge. That’s where Chapagain comes in.

“A lot of researchers have shifted attention from their own projects to this because COVID-19 is hitting us like a hammer all of a sudden,” Chapagain said.

Using artificial intelligence and molecular modeling, the physicist was able to quickly identify top drug candidates that could possibly serve as TOP3B inhibitors.

Tse-Dinh and Ph.D. students Ahmed Seddek and Tumpa Dasgupta have been testing those drugs against a purified version of TOP3B and related topoisomerases.

Tse-Dinh is referring drugs that show potential to Garcia-Blanco’s team who is now testing them against live cell cultures and viruses including COVID-19.

“It’s a very simple story,” Garcia-Blanco said. “We want to inhibit this enzyme. That’s it. If we can do that and combine it with other anti-viral treatment, it could enhance the chances to help people who are very ill.”

The collaborators, which also includes Dr. Yves Pommier from the National Cancer Institute, are hyper-focused, working long hours in their labs across the country to try and block TOP3B together.

Garcia-Blanco even separated his research team into two smaller groups to allow for social distancing and ensure the work will keep going even if one of those groups had to quarantine. Meanwhile Chapagain is working remotely at his home, refining his modeling and continuing the search for other possible drug candidates.

“We really feel the urgency,” Tse-Dinh said. “But we have to do things with a scientific basis.”


Cell proliferation requires that the genetic information from a starting mother cell be copied and partitioned into two daughter cells. The double-stranded nature of DNA enables semiconservative replication [1]; however, unwinding of the DNA helix produces superhelical tension that poses a topological challenge to replicative processes [2] and leads to entanglement of newly-replicated sister chromosomes [3]. In all branches of life, enzymes known as topoisomerases help cells circumvent these and other topological challenges. Topoisomerases broadly fall into two principal categories, termed type I or type II, based on whether they respectively form single- or double-strand DNA breaks during catalysis [4]. Although both type I and type II topoisomerases can alter DNA superhelicity [5,6,7], type II enzymes are uniquely able to decatenate intact sister chromosomes by an ability to pass one double-stranded segment of DNA through a transient, enzyme-mediated break in a second DNA duplex [8,9].

Most eukaryotic organisms express a single type II topoisomerase known as DNA topoisomerase II (topo II or Top2). Vertebrates carry two isoforms, topo IIα and topo IIβ [10,11]. Topo IIα is primarily active during DNA replication and mitosis [12], whereas topo IIβ has roles in transcriptional regulation of gene expression [13,14,15]. Topo II (or topo IIα) activity during mitosis is essential for cell viability [16]. Upon entry into S-phase, topo II is critical for organizing genome structure, promoting chromosome segregation, and preventing aberrant entry into anaphase with partially decatenated sister chromatids. Topo II activity is modulated by various means in response to the needs of the cell. For example, the abundance of topo IIα in vertebrates is cell cycle-regulated to accommodate the substantial changes in chromosome copy number and structure that occur during DNA replication and mitosis [17,18,19]. The affinity of topo II for DNA substrates and catalytic activity can also be directly controlled through cell cycle-dependent changes in DNA topology, protein–protein interactions, and post-translational modifications [20,21,22]. Missteps in topo II-dependent processes activate a decatenation checkpoint that stalls the cell cycle at the G2-to-M transition and is protective against genomic damage [23]; loss of this checkpoint has been linked to tumorigenesis [24]. The present review seeks to provide a brief overview of the various roles of topo II and of known cell-cycle regulatory mechanisms that help monitor and coordinate topo II activity with the dynamic genomic landscape. Due to limitations on space, recent developments concerning the action and consequences of topo IIβ in supporting gene expression will not be discussed.

Cell Cycle-Dependent Expression of Topo IIα in Vertebrates

In vertebrates, production and degradation of topo IIα is cell cycle-regulated and serves to link topo II functions with proliferation. Topo IIα levels increase in mid-S phase through mitosis and rapidly decrease upon mitotic completion (Figure 1) [18,19,25]. Evidence suggests that accumulation of topo IIα is due to upregulation of TOP2A transcription and mRNA stabilization.

For example, the promoter region of TOP2A contains multiple inverted CCAAT boxes (ICBs) that have been proposed to interact with the NF-Y, SP1, and ICBP90 transcription factors to regulate the cell cycle-dependent expression of topo IIα [26,27,28,29]. Stabilization of TOP2A mRNA by the 3′UTR from S phase through mitosis has also been observed in human cells, though not in mouse cells. Studies have suggested that the 3′UTR of human TOP2A mRNA binds to cell cycle-regulated proteins to protect it from degradation [30,31]. At the protein level, topo IIα abundance is controlled by ubiquitin-mediated proteasomal degradation [32].

Depletion of ubiquitin specific protease 15 (USP15), a deubiquitylase (DUB), has been associated with a failure to appropriately accumulate topo IIα, while USP15 expression correlates with topo IIα expression [33]. These observations suggest that USP15 stabilizes topo IIα by removing ubiquitin modifications. After mitosis, topo IIα appears to be ubiquitylated by the anaphase promoting complex (APC) and Cdh-1 in the absence of USP15, resulting in a rapid decrease in enzyme concentration [34].

Cell cycle-dependent expression of topo II has not been observed in Drosophila, which carry only one isoform [35]. In vertebrates, topo IIβ is present at uniform levels throughout the cell cycle [19]. Expression of topo II in organisms that carry a single isoform may not be proliferation-dependent because a single enzyme must assume the roles of both topo IIα and topo IIβ.

An external file that holds a picture, illustration, etc.
Object name is genes-10-00859-g001.jpg
Figure 1
Regulation of topo IIα abundance during the cell cycle. The color gradient bar indicates how levels of topo IIα vary from low (white) to high (red) in different cell cycle stages. The cartoon depicts the current understanding of how topo IIα transcription (DNA), translation (mRNA), and stability (protein) are regulated. Transcription of topo IIα occurs in S phase, through to mitosis, and is regulated by transcription factor binding to ICBs in the promoter region of the TOP2A gene. Translation is controlled by cell cycle-dependent mRNA stability, and potentially by cell cycle-dependent proteins that bind and stabilize the 3′-UTR. Topo IIα protein is protected from ubiquitin-mediated degradation during S phase through metaphase by the deubiquitylase USP15. Cell cycle-dependent phosphorylation also occurs from S phase to metaphase and SUMOylation occurs upon entry into mitosis. SUMO modifications are lost upon anaphase onset. At this stage, topo IIα is ubiquitylated by APC/Cdh1 and is subject to proteasomal degradation.

DNA Replication
During replication, origin unwinding and fork progression generate positive writhe ahead of the replisome. The accumulation of superhelical tension, if not relieved, can lead to fork stalling, the formation of precatenanes, and DNA damage [2]. In eukaryotes, supercoil relaxation by topoisomerase I (topo IA) is generally sufficient to support replication, as the loss of topo II does not appear to appreciably lower rates of DNA synthesis [36,37].

However, in S. cerevisiae at least, topo II is able to compensate for the loss of topo I, indicating that topo II is active at these early stages of DNA replication [36]. The simultaneous deletion of topo I and topo II causes genomic instability [36].

To prevent DNA damage from the accrual of superhelical tension, both topo I and topo II are rapidly recruited to sites of origin firing during replication initiation [36]. While topo II is not required for DNA replication initiation in a G1-to-S phase transition, inhibition of topo II has been observed to prevent entry into S phase from quiescence (G0) in mammalian cells and Drosophila, but not in yeast [38,39].

After initiation, topo II assists with supercoil relaxation and the disentanglement of newly replicated segments of DNA during replication elongation, as most decatenation is complete by entry into prophase (Figure 2A) [40,41].

An external file that holds a picture, illustration, etc.
Object name is genes-10-00859-g002.jpg
Figure 2
Roles of topo II throughout the cell cycle. (A) During DNA replication, topo II decatenates newly-replicated sister chromosomes and aids in relaxing positive supercoils that accumulate ahead of replication forks. Most decatenation is complete upon entry into prophase. Topo II can operate bidirectionally and is still present in high concentrations during G2; it has been suggested that the enzyme may contribute to maintaining a catenated state of sister chromosomes to assist chromosome condensation and cohesion without subsequently interfering with segregation. (B) Condensin and topo II are both required for efficient condensation of chromosomes in prophase. (C) In metaphase, topo II localizes to centromeres but does not complete decatenation of these regions. It has been suggested that cohesin protects these regions from topo II activity. (D) Upon anaphase entry, cohesin is released by separase and topo II rapidly decatenates sister chromatids to allow for chromosome segregation.

The role of topo II in replication termination is less well-understood. ChIP studies in S. cerevisiae have indicated that topo II preassociates with common replication termination sites [42]. However, unlike prokaryotic systems [43,44], topo II activity is not required for fork convergence or the dissolution of replication forks [45,46].

Topo I and topo II alone are insufficient for efficient termination; instead, recent work has suggested that replication termination may primarily rely on Pif1-family helicases [46]. Topo II activity may be more critical in telomeric regions that are prone to replication stalling due to highly repetitive sequences [47].

Stalled replication forks generate telomere-specific extrachromosomal structures that may require topo II to facilitate fork progression and replication termination [48,49]. This observation suggests that topo II may expedite termination in regions prone to replication stress, but that it is not generally essential for replication termination.

In eukaryotes, DNA replication is initiated asynchronously from multiple origins and some origins remain dormant throughout S-phase [50,51]. To prevent re-entry into S-phase, cells must dismantle replication machineries.

Interestingly, in Xenopus, depletion of topo IIα has been observed to prevent the complete degradation of origin recognition complex subunit 1 (ORC1) and to block the dissociation of ORC1/2 from replication protein A (RPA) following the completion of DNA synthesis [52]. Thus, topo II may participate in preventing replisome reassembly at the end of S phase. Overall, topo II plays a nonessential role in DNA strand synthesis but seems to nonetheless facilitate this process in problematic regions of the genome.

References

1. Meselson M., Stahl F. The relication of DNA in E. coli. Proc. Natl. Acad. Sci. USA. 1958;44:671–682. doi: 10.1073/pnas.44.7.671. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

2. Postow L., Crisona N.J., Peter B.J., Hardy C.D., Cozzarelli N.R. Topological challenges to DNA replication: Conformations at the fork. Proc. Natl. Acad. Sci. USA. 2001;98:8219–8226. doi: 10.1073/pnas.111006998. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

3. Peter B.J., Ullsperger C., Hiasa H., Marians K.J., Cozzarelli N.R. The structure of supercoiled intermediates in DNA replication. Cell. 1998;94:819–827. doi: 10.1016/S0092-8674(00)81740-7. [PubMed] [CrossRef] [Google Scholar]

4. Vos S.M., Tretter E.M., Schmidt B.H., Berger J.M. All tangled up: How cells direct, manage and exploit topoisomerase function. Nat. Rev. Mol. Cell Biol. 2011;12:827–841. doi: 10.1038/nrm3228. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

5. Wang J.C. Interaction between DNA and an Escherichia coli protein ω J. Mol. Biol. 1971;55:523–533. doi: 10.1016/0022-2836(71)90334-2. [PubMed] [CrossRef] [Google Scholar]

6. Champoux J.J., Dulbecco R. An activity from mammalian cells that untwists superhelical DNA—A possible swivel for DNA replication (polyoma-ethidium bromide-mouse-embryo cells-dye binding assay) Proc. Natl. Acad. Sci. USA. 1972;69:143–146. doi: 10.1073/pnas.69.1.143. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

7. Gellert M., Mizuuchi K., O’dea M.H., Nasht H.A. DNA gyrase: An enzyme that introduces superhelical turns into DNA (Escherichia coli/ATP-dependent reaction/superhelix density) Proc. Natl. Acad. Sci. USA. 1976;73:3872–3876. doi: 10.1073/pnas.73.11.3872. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

8. Goto T., Wang J.C. Yeast DNA Topoisomerase II. An ATP-dependent type II topoisomerase that catalyzes the catenation, decatenation, unknotting, and relaxation of double-stranded DNA rings. J. Biol. Chem. 1982;257:5866–58720. [PubMed] [Google Scholar]

9. Kreuzer K.N., Cozzarelli N.R. Formation and resolution of DNA catenanes by DNA gyrase. Cell. 1980;20:245–254. doi: 10.1016/0092-8674(80)90252-4. [PubMed] [CrossRef] [Google Scholar]

10. Tsai-Pflugfelder M., Liu L.F., Liu A.A., Tewey K.M., Whang-Peng J., Knutsen T., Huebner K., Croce C.M., Wang J.C. Cloning and sequencing of cDNA encoding human DNA topoisomerase II and localization of the gene to chromosome region 17q21-22. Proc. Natl. Acad. Sci. USA. 1988;85:7177–7181. doi: 10.1073/pnas.85.19.7177. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

11. Tan K.B., Dorman T.E., Falls K.M., Chung T.D.Y., Mirabelli C.K. Topoisomerase II alpha and topoisomerase II beta genes: Characterization and mapping to human chromosomes 17 and 3, respectively. Cancer Res. 1992;52:231–234. [PubMed] [Google Scholar]

12. Grue P., Gräßer A., Sehested M., Jensen P.B., Uhse A., Straub T., Ness W., Boege F. Essential mitotic functions of DNA topoisomerase IIα are not adopted by topoisomerase IIβ in human H69 cells. J. Biol. Chem. 1998;273:33660–33666. doi: 10.1074/jbc.273.50.33660. [PubMed] [CrossRef] [Google Scholar]

13. Lyu Y.L., Lin C.-P., Azarova A.M., Cai L., Wang J.C., Liu L.F. Role of Topoisomerase II in the Expression of Developmentally Regulated Genes. Mol. Cell. Biol. 2006;26:7929–7941. doi: 10.1128/MCB.00617-06. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

14. Ju B.-G., Lunyak V.V., Perissi V., Garcia-Bassets I., Rose D.W., Glass C.K., Rosenfeld M.G. A Topoisomerase IIbeta-Mediated dsDNA Break Required for Regulated Transcription. Science. 2006;312:1798–1802. doi: 10.1126/science.1127196. [PubMed] [CrossRef] [Google Scholar]

15. Madabhushi R., Gao F., Pfenning A.R., Pan L., Yamakawa S., Seo J., Rueda R., Phan T.X., Yamakawa H., Pao P.C., et al. Activity-Induced DNA Breaks Govern the Expression of Neuronal Early-Response Genes. Cell. 2015;161:1592–1605. doi: 10.1016/j.cell.2015.05.032. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

16. Holm C., Goto T., Wang J.C., Botstein D. DNA topoisomerase II is required at the time of mitosis in yeast. Cell. 1985;41:553–563. doi: 10.1016/S0092-8674(85)80028-3. [PubMed] [CrossRef] [Google Scholar]

17. Uemura T., Ohkura H., Adachi Y., Morino K., Shiozaki K., Yanagida M. DNA topoisomerase II is required for condensation and separation of mitotic chromosomes in S. pombe. Cell. 1987;50:917–925. doi: 10.1016/0092-8674(87)90518-6. [PubMed] [CrossRef] [Google Scholar]

18. Heck M.M.S., Hittelman W.N., Earnshaw W.C. Differential expression of DNA topoisomerases I and II during the eukaryotic cell cycle. Proc. Natl. Acad. Sci. USA. 1988;85:1086–1090. doi: 10.1073/pnas.85.4.1086. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

19. Woessner R.D., Mattern M.R., Mirabelli C.K., Johnson R.K., Drake F.H. Proliferation- and Cell Cycle-dependent Differences in Expression of the 170 Kilodalton and 180 Kilodalton Forms of Topoisomerse II in NIH-3T3 Cells. Cell Growth Differ. 1991;2:209–214. [PubMed] [Google Scholar]

20. Baxter J., Sen N., López Martínez V., Monturus De Carandini M.E., Schvartzman J.B., Diffley J.F.X., Aragón L. Positive supercoiling of mitotic DNA drives decatenation by topoisomerase II in eukaryotes. Science. 2011;331:1328–1332. doi: 10.1126/science.1201538. [PubMed] [CrossRef] [Google Scholar]

21. Kroll D.J. Homologous and heterologous protein-protein interactions of human DNA topoisomerase. Arch. Biochem. Biophys. 1997;345:175–184. doi: 10.1006/abbi.1997.0267. [PubMed] [CrossRef] [Google Scholar]

22. Porter A.C.G., Farr C.J. Topoisomerase II: Untangling its contribution at the centromere. Chromosom. Res. 2004;12:569–583. doi: 10.1023/B:CHRO.0000036608.91085.d1. [PubMed] [CrossRef] [Google Scholar]

23. Downes C.S., Clarke D.J., Mullinger A.M., Giménez-Abián J.F., Creighton A.M., Johnson R.T. A topoisomerase II-dependent G2 cycle checkpoint in mammalian cells. Nature. 1994;372:467–470. doi: 10.1038/372467a0. [PubMed] [CrossRef] [Google Scholar]

24. Chen T., Sun Y., Ji P., Kopetz S., Zhang W. Topoisomerase IIα in chromosome instability and personalized cancer therapy. Oncogene. 2015;34:4019–4031. doi: 10.1038/onc.2014.332. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

25. Kimura K., Saijo M., Ui M., Enomoto T. Growth state- and cell cycle-dependent fluctuation in the expression of two forms of DNA topoisomerase II and possible specific modification of the higher molecular weight form in the M phase. J. Biol. Chem. 1994;269:1173–1176. [PubMed] [Google Scholar]

26. Adachi N., Nomoto M., Kohno K., Koyama H. Cell-cycle regulation of the DNA topoisomerase IIα promoter is mediated by proximal CCAAT boxes: Possible involvement of acetylation. Gene. 2000;245:49–57. doi: 10.1016/S0378-1119(00)00040-8. [PubMed] [CrossRef] [Google Scholar]

27. Magan N., Szremska A.P., Isaacs R.J., Stowell K.M. Modulation of DNA topoisomerase IIα promoter activity by members of the Sp (specificity protein) and NF-Y (nuclear factor Y) families of transcription factors. Biochem. J. 2003;374:723–729. doi: 10.1042/bj20030032. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

28. Hopfner R., Mousli M., Jeltsch J.M., Voulgaris A., Lutz Y., Marin C., Bellocq J.P., Oudet P., Bronner C. ICBP90, a novel human CCAAT binding protein, involved in the regulation of topoisomerase IIα expression. Cancer Res. 2000;60:121–128. [PubMed] [Google Scholar]

29. Falck J., Jensen P.B., Sehested M. Evidence for repressional role of an inverted CCAAT box in cell cycle- dependent transcription of the human DNA topoisomerase IIα gene. J. Biol. Chem. 1999;274:18753–18758. doi: 10.1074/jbc.274.26.18753. [PubMed] [CrossRef] [Google Scholar]

30. Goswami P.C., Roti Roti J.L., Hunt C.R. The cell cycle-coupled expression of topoisomerase IIalpha during S phase is regulated by mRNA stability and is disrupted by heat shock or ionizing radiation. Mol. Cell. Biol. 1996;16:1500–1508. doi: 10.1128/MCB.16.4.1500. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

31. Goswami P.C., Sheren J., Albee L.D., Parsian A., Sim J.E., Ridnour L.A., Higashikubo R., Gius D., Hunt C.R., Spitz D.R. Cell cycle-coupled variation in topoisomerase IIα mRNA is regulated by the 3′-untranslated region: Possible role of redox-sensitive protein binding in mRNA accumulation. J. Biol. Chem. 2000;275:38384–38392. doi: 10.1074/jbc.M005298200. [PubMed] [CrossRef] [Google Scholar]

32. Salmena L., Lam V., Peter McPherson J., Goldenberg G.J. Role of proteasomal degradation in the cell cycle-dependent regulation of DNA topoisomerase IIα expression. Biochem. Pharmacol. 2001;61:795–802. doi: 10.1016/S0006-2952(01)00580-9. [PubMed] [CrossRef] [Google Scholar]

33. Fielding A.B., Concannon M., Darling S., Rusilowicz-Jones E.V., Sacco J.J., Prior I.A., Clague M.J., Urbé S., Coulson J.M. The deubiquitylase USP15 regulates topoisomerase II alpha to maintain genome integrity. Oncogene. 2018;37:2326–2342. doi: 10.1038/s41388-017-0092-0. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

34. Eguren M., Álvarez-Fernández M., García F., López-Contreras A.J., Fujimitsu K., Yaguchi H., Luque-García J., Fernández-Capetillo O., Muñoz J., Yamano H., et al. A Synthetic Lethal Interaction between APC/C and Topoisomerase Poisons Uncovered by Proteomic Screens. Cell Rep. 2014;6:670–683. doi: 10.1016/j.celrep.2014.01.017. [PubMed] [CrossRef] [Google Scholar]

35. Whalen A.M., McConnell M., Fisher P.A. Developmental regulation of Drosophila DNA topoisomerase II. J. Cell Biol. 1991;112:203–213. doi: 10.1083/jcb.112.2.203. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

36. Bermejo R., Doksani Y., Capra T., Katou Y.M., Tanaka H., Shirahige K., Foiani M. Top1- and Top2-mediated topological transitions at replication forks ensure fork progression and stability and prevent DNA damage checkpoint activation. Genes Dev. 2007;21:1921–1936. doi: 10.1101/gad.432107. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

37. Gonzalez R.E., Lim C.U., Cole K., Bianchini C.H., Schools G.P., Davis B.E., Wada I., Roninson I.B., Broude E.V. Effects of conditional depletion of topoisomerase II on cell cycle progression in mammalian cells. Cell Cycle. 2011;10:3505–3514. doi: 10.4161/cc.10.20.17778. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

38. Hossain M.S., Akimitsu N., Takaki T., Hirai H., Sekimizu K. ICRF-193, a catalytic inhibitor of DNA topoisomerase II, inhibits re-entry into the cell division cycle from quiescent state in mammalian cells. Genes Cells. 2002;7:285–294. doi: 10.1046/j.1365-2443.2002.00521.x. [PubMed] [CrossRef] [Google Scholar]

39. Hossain M.S., Kurokawa K., Akimitsu N., Sekimizu K. DNA topoisomerase II is required for the G0-to-S phase transition in Drosophila Schneider cells, but not in yeast. Genes Cells. 2004;9:905–917. doi: 10.1111/j.1365-2443.2004.00783.x. [PubMed] [CrossRef] [Google Scholar]

40. Lucas I., Germe T., Chevrier-Miller M., Hyrien O. Topoisomerase II can unlink replicating DNA by precatenane removal. EMBO J. 2001;20:6509–6519. doi: 10.1093/emboj/20.22.6509. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

41. Charbin A., Bouchoux C., Uhlmann F. Condensin aids sister chromatid decatenation by topoisomerase II. Nucleic Acids Res. 2014;42:340–348. doi: 10.1093/nar/gkt882. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

42. Fachinetti D., Bermejo R., Cocito A., Minardi S., Katou Y., Kanoh Y., Shirahige K., Azvolinsky A., Zakian V.A., Foiani M. Replication Termination at Eukaryotic Chromosomes Is Mediated by Top2 and Occurs at Genomic Loci Containing Pausing Elements. Mol. Cell. 2010;39:595–605. doi: 10.1016/j.molcel.2010.07.024. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

43. Snapka R.M., Powelson M.A., Strayer J.M. Swiveling and Decatenatio of Replicating Simian Virus 40 Genomes in Vivo. Mol. Cell. Biol. 1988;8:515–521. doi: 10.1128/MCB.8.2.515. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

44. Richter A., Strausfeld U. Effects of VM26 (teniposide), a specific inhibitor of type II DNA topoisomerase, on SV40 chromatin replication in vitro. Nucleic Acids Res. 1987;15:3455–3468. doi: 10.1093/nar/15.8.3455. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

45. Dewar J.M., Budzowska M., Walter J.C. The mechanism of DNA replication termination in vertebrates. Nature. 2015;525:345–350. doi: 10.1038/nature14887. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

46. Deegan T.D., Baxter J., Ortiz Bazán M.Á., Yeeles J.T.P., Labib K.P.M. Pif1-Family Helicases Support Fork Convergence during DNA Replication Termination in Eukaryotes. Mol. Cell. 2019;74:231–244. doi: 10.1016/j.molcel.2019.01.040. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

47. Gilson E., Géli V. How telomeres are replicated. Nat. Rev. Mol. Cell Biol. 2007;8:825–838. doi: 10.1038/nrm2259. [PubMed] [CrossRef] [Google Scholar]

48. Zhang T., Zhang Z., Li F., Hu Q., Liu H., Tang M., Ma W., Huang J., Songyang Z., Rong Y., et al. Looping-out mechanism for resolution of replicative stress at telomeres. EMBO Rep. 2017;18:1412–1428. doi: 10.15252/embr.201643866. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

49. Ye J., Lenain C., Bauwens S., Rizzo A., Saint-Léger A., Poulet A., Benarroch D., Magdinier F., Morere J., Amiard S., et al. TRF2 and Apollo Cooperate with Topoisomerase 2α to Protect Human Telomeres from Replicative Damage. Cell. 2010;142:230–242. doi: 10.1016/j.cell.2010.05.032. [PubMed] [CrossRef] [Google Scholar]

50. Yekezare M., Gó mez-González B., Diffley J.F.X. Controlling DNA replication origins in response to DNA damage—Inhibit globally, activate locally. J. Cell Sci. 2013;126:1297–1306. doi: 10.1242/jcs.096701. [PubMed] [CrossRef] [Google Scholar]

51. Truong L.N., Wu X. Prevention of DNA re-replication in eukaryotic cells. J. Mol. Cell Biol. 2011;3:13–22. doi: 10.1093/jmcb/mjq052. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

52. Cuvier O., Stanojcic S., Lemaitre J.M., Mechali M. A topoisomerase II-dependent mechanism for resetting replicons at the S-M-phase transition. Genes Dev. 2008;22:860–865. doi: 10.1101/gad.445108. [PMC free article] [PubMed] [CrossRef] [Google Scholar]


Source:
Florida International University

LEAVE A REPLY

Please enter your comment!
Please enter your name here

Questo sito usa Akismet per ridurre lo spam. Scopri come i tuoi dati vengono elaborati.