Dissertations in Forestry and Natural Sciences

ANU HANGAS

TISSUE SPECIFICITY AND TOPOISOMERASE FUNCTIONS

IN MITOCHONDRIAL DNA MAINTENANCE

PUBLICATIONS OF THE UNIVERSITY OF EASTERN FINLAND

TISSUE SPECIFICITY AND TOPOISOMERASE FUNCTIONS IN MITOCHONDRIAL DNA

MAINTENANCE

Anu Hangas

TISSUE SPECIFICITY AND TOPOISOMERASE FUNCTIONS IN MITOCHONDRIAL DNA

MAINTENANCE

Publications of the University of Eastern Finland Dissertations in Forestry and Natural Sciences

No 410

University of Eastern Finland Joensuu

2020

Academic dissertation To be presented by permission of the Faculty of Science and Forestry for public

examination in the Auditorium N101 in the Natura Building at the University of Eastern Finland, Joensuu, on December, 11, 2020, at 12 o’clock noon

Grano Oy Jyväskylä, 2020

Editor: Raine Kortet Distribution: University of Eastern Finland / Sales of publications

www.uef.fi/kirjasto ISBN: 978-952-61-3668-4 (nid.) ISBN: 978-952-61-3669-1 (PDF)

ISSNL: 1798-5668 ISSN: 1798-5668

ISSN: 1798-5676 (PDF)

Author’s address: Anu Hangas University of Eastern Finland

Depart. of Environmental and Biological Sciences P.O. Box 111 80101 JOENSUU, FINLAND email: anu.hangas@uef.fi Supervisors: Steffi Goffart, Ph.D. University of Eastern Finland Depart. of Environmental and Biological Sciences P.O. Box 111 80101 JOENSUU, FINLAND email: steffi.goffart@uef.fi Jaakko Pohjoismäki, Ph.D. University of Eastern Finland Depart. of Environmental and Biological Sciences P.O. Box 111 80101 JOENSUU, FINLAND email: jaakko.pohjoismaki@uef.fi Reviewers: Assistant Professor Paulina Wanrooij, Ph.D. University of Umeå Depart. of Medical Biochemistry and Biophysics Umeå Universitet,

901 87 Umeå, Sweden email: paulina.wanrooij@umu.se

Assistant Professor Marcos Tulio de Oliveira, Ph.D. Assistant Professor Sao Paulo State University Department of Technology Jaboticabal, SP, Brazil email: marcos.t.oliveira@unesp.br Opponent: Professor Dr. Rudolf Wiesner Institut für Vegetative Physiologie Universität zu Köln 50931 Köln, Germany email: rudolf.wiesner@uni-koeln.de

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Hangas, Anu Tissue specificity and topoisomerase functions in mitochondrial DNA maintenance Joensuu: University of Eastern Finland, 2020 Publications of the University of Eastern Finland Dissertations in Forestry and Natural Sciences 2020; 410 ISBN: 978-952-61-3668-4 (print) ISSNL: 1798-5668 ISSN: 1798-5668 ISBN: 978-952-61-3669-1 (PDF) ISSN: 1798-5676 (PDF)

ABSTRACT

Mitochondria, the energy-producing organelles in nearly all eukaryotic cells, possess their own small circular genome. The maintenance of this mitochondrial DNA re-quires a considerable set of nuclear-encoded proteins. As mitochondrial mass and function are strongly influenced by the cell type, both the expression levels of mito-chondrial proteins and mitochondrial DNA copy number vary greatly between dif-ferent tissues and organs. Mitochondrial tissue specificity is also reflected through the replication mechanisms cells use depending on their energy requirement, but also on the oxidative stress level. Here is shown that the utilized replication mechanism correlates with mitochondrial DNA damage in tissues and thus is likely an adapta-tion for the differences in the oxidative environments of the tissues.

Among the proteins, whose expression level varies between different tissues, are mitochondrial topoisomerases, essential for many aspects of genome maintenance and integrity. Mitochondria contain four topoisomerases, Top1mt, Top3α, Top2α and Top2β. While Top1mt is the only mitochondria-specific topoisomerase and has been investigated to some extent, the mitochondrial function of the topoisomerases Top2 and Top3 have barely been studied. This study shows that the mitochondrial type II topoisomerase Top2 is a regulator of mitochondrial DNA topology and is essential for the relaxation of supercoils. Top3 in contrast is not only involved in resolution of freshly replicated mtDNA molecules, as shown before, but also acts as a replicative topoisomerase in close contact with the replisome, relieving torsional stress behind the replicating fork.

The potential bacterial origin of mitochondria and the homology of many mito-chondrial proteins to their bacterial analogues make mitochondria especially vulner-able to antibacterial pharmaceuticals used to treat bacterial infections. Thus, these drugs occasionally have severe side effects caused by mitochondrial dysfunction, but

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the exact mechanisms are often unclear. Here it is shown that a commonly used an-tibiotic, ciprofloxacin, interferes with mtDNA replication by inhibiting the function of mitochondrial topoisomerase 2β, explaining the rare but devastating side effects linked to ciprofloxacin therapy.

Proper maintenance of mitochondrial DNA is essential for survival of the cell and the whole organism, and this thesis aims to understand tissue-specific aspects of mtDNA replication and the role topoisomerases play in the maintenance of this tiny but important genome.

National Library of Medicine Classification: QU 137, QU 360, QV 350 Medical Subject Headings: Mitochondria; Genome, Mitochondrial; DNA, Mitochondrial; DNA Replication; DNA Topoisomerases; DNA Damage; Topoisomerase II Inhibitors; Anti-Bacterial Agents/adverse effects; Ciprofloxacin/adverse effects; Cells, Cultured; Mice Yleinen suomalainen ontologia: mitokondriot; mitokondrio-DNA; ylläpito; replikaatio; anti-biootit; soluviljely; hiiret

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ACKNOWLEDGEMENTS

I am extremely grateful to my supervisors Steffi Goffart and Jaakko Pohjoismäki for introducing me to the mysteries of mitochondria and giving me the great oppor-tunity to learn more about this tiny organelle.

I would also like to thank all the past and current lab members, Nina Kekäläinen, Elena Herbers, Riikka Tapanainen, Craig Michell, Georgios Fragkoulis, Chiara Ru-tanen, Koit Aasumets and Petra Tikkanen for their help and delightful company in the lab. I would like to address special thanks to Rubén Torregrosa, this would have been much more boring without you. I also wish to thank my master students, whose work I had the privilege to supervise, Obaidur, Kauko and Waleed. I am thankful to the whole staff of the department; you have made this a great place to work. I would also like thank our collaborators in Tartu for their important contribution to my re-search.

I would like to thank my pre-examiners Paulina Wanrooij and Marcos Oliveira for their helpful comments and suggestions for improving the manuscript. Thank you to Prof. Rudolf Wiesner for accepting to be my opponent. I’m also thankful to Rubén, Elena and Craig for taking the time to read and comment this thesis. I like to thank Jaakko, Rubén and Marcos Torregrosa for their contribution to graphical illus-trations, where my talents were limited.

I am grateful to all my friends from Joensuu and Koski Tl and my family, Miia, Unto, and Sinikka, who have been supporting me during this time. Thank you to Kristiina, my beloved goddaughter, with whom I have been sharing most of my ac-ademic life. Also big thanks to Niko, Mari, and Heli for trying to keep me sane during this stressful process.

Lastly, I wish to express my gratitude to the funding organizations for making this research possible, Academy of Finland, Jane & Aatos Erkko Foundation, Finnish Cultural Foundation (North Karelia Regional fund, A.E. Kalsta Fund), University of Eastern Finland Doctoral School, and Faculty of Science and Forestry.

Joensuu, 9th November 2020 Anu Hangas

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LIST OF ABBREVIATIONS

2D-AGE Two-dimensional agarose gel electrophoresis 7S DNA A small third strand associated with the non-coding region of mtDNA ADP Adenosine diphosphate ATP Adenosine triphosphate BER Base excision repair BioID Proximity-dependent biotin identification BLM Bloom syndrome protein BTRR Holliday junction-resolving BLM-Top3α-RMI1-RMI2 complex C2C12 A murine myoblast cell line CBS Conserved sequence block DMEM Dulbecco’s modified Eagle medium DNA Deoxyribonucleic acid DSB Double strand break DTT Dithiothreitol EDTA Ethylenediaminetetraacetic acid HEK 293 Human embryonic kidney cell line HR Homologous recombination HSP Heavy-strand promoter LSP Light-strand promoter MEF Mouse embryonic fibroblast MELAS Mitochondrial encephalopathy, lactic acidosis, stroke-like episodes MGME1 Mitochondrial genome maintenance exonuclease 1 MMR Mismatch repair MTS Mitochondrial targeting sequence mtSSB Single-strand DNA-binding protein, mitochondrial mtDNA Mitochondrial DNA mTERF Mitochondrial transcription termination factor nDNA Nuclear DNA NCR Non-coding region NER Nucleotide excision repair NHEJ Non-homologous end joining NLS Nuclear localization sequence nt Nucleotide OH Heavy (leading)-strand origin of replication OL Light (lagging)-strand origin of replication Ori-z Initiation zone of bidirectional replication OXPHOS Oxidative phosphorylation PBS Phosphate buffered saline Polγ Polymerase γ

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POLRMT DNA-directed RNA polymerase, mitochondrial RITOLS RNA introduced throughout the lagging-strand RMI RecQ-mediated genome instability protein RNA Ribonucleic acid RNaseH1 Ribonuclease H1 ROS Reactive oxygen species SDM Strand displacement model siRNA Small interfering RNA smY Slow-moving Y SOD Superoxide dismutase TAS Termination-associated sequence TFAM Mitochondrial transcription factor A TFB1M Mitochondrial transcription factor B1 TFB2M Mitochondrial transcription factor B2 TIM Translocase of the inner membrane TOM Translocase of the outer membrane Top Topoisomerase TOPcc Topoisomerase cleavage complex TWNK mtDNA helicase (formerly Twinkle)

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LIST OF ORIGINAL PUBLICATIONS

This thesis is based on data presented in the following articles, referrred to by the Roman Numerals I-IV. I Herbers E., Kekäläinen N. J., Hangas A., Pohjoismäki, J. L., Goffart S. (2019)

Tissue specific differences in mitochondrial DNA maintenance and expression. Mitochondrion, 44: 85-92.

II Hangas A., Aasumets K., Kekäläinen N. J., Paloheinä M., Pohjoismäki J. L.,

Gerhold J. M., Goffart S. (2018) Ciprofloxacin impairs mitochondrial DNA replication initiation through inhibition of Topoisomerase 2. Nucleic Acids Research, 46: 9625-9636.

III Hangas A., Kekäläinen N. J., Michell C., Aho K. J., Pohjoismäki J. L., Goffart

S. Top3α is the replicative topoisomerase in mitochondrial DNA replication. Manuscript. IV Goffart S., Hangas A., Pohjoismäki J.L.O. (2019) Twist and Turn –

Topoisomerase Functions in Mitochondrial DNA Maintenance. International Journal of Molecular Sciences, 20: 2041.

The above publications have been reproduced with the copyright holders’ permission.

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AUTHOR’S CONTRIBUTION I) The author contributed to study design, experimentation, interpretation of the

results and manuscript writing.

II) The author conducted and interpreted most of the experimental work and contributed to study design and manuscript writing.

III) The author conducted and interpreted most of the experimental work and contributed to study design and manuscript writing.

IV) The author was one of two main contributors to manuscript design and

writing.

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CONTENTS

ABSTRACT ........................................................................................................... 7 ACKNOWLEDGEMENTS ..................................................................................... 9 LIST OF ABBREVIATIONS ................................................................................11 LIST OF ORIGINAL PUBLICATIONS ................................................................13 1 INTRODUCTION ..........................................................................................17

1.1 Mitochondria .........................................................................................17 1.2 Mitochondrial DNA ................................................................................19

1.2.1 mtDNA replication .........................................................................20 1.2.2 7S DNA .........................................................................................24 1.2.3 Mitochondrial transcription ............................................................25

1.3 Mitochondrial DNA damage ..................................................................26 1.3.1 mtDNA damage and repair ...........................................................26 1.3.2 mtDNA recombination ..................................................................27

1.4 mtDNA topology....................................................................................28 1.4.1 Topoisomerases ...........................................................................29 1.4.2 Top1mt ..........................................................................................31 1.4.3 Top3α ............................................................................................31 1.4.4 Top2 ..............................................................................................32

1.5 Mitochondrial diseases .........................................................................33 1.5.1 Drug-induced mitochondrial dysfunction ......................................34 1.5.2 Inhibition of topoisomerase function .............................................35 1.5.3 Ciprofloxacin .................................................................................36

2 AIMS OF THE STUDY .................................................................................39 3 MATERIALS AND METHODS .....................................................................41

3.1 Cell lines and mice ................................................................................41 3.1.1 Mice ..............................................................................................41 3.1.2 Cell culture ....................................................................................41 3.1.3 Cloning of expression constructs..................................................41 3.1.4 Transient transfections and immunocytochemistry ......................42 3.1.5 siRNA knock-down .......................................................................42

3.2 Mitochondrial extraction ........................................................................43 3.3. DNA ......................................................................................................43

3.3.1 DNA extraction ..............................................................................43 3.3.2 mtDNA topology............................................................................43 3.3.3 7S DNA .........................................................................................44 3.3.4 Determination of mitochondrial DNA copy number ......................44 3.3.5 Two-dimensional neutral/neutral agarose gel electrophoresis .....45 3.3.6 Quantification of DNA damage .....................................................46 3.3.7 In vitro Topoisomerase assay .......................................................46 3.3.8 Nanopore sequencing ..................................................................47

3.4. Protein ..................................................................................................47 3.4.1 Protein extraction and Western blots ...........................................47 3.4.2 Top3α-BioID purification ...............................................................48 3.4.3 Flotation gradient ..........................................................................49

3.5 RNA ......................................................................................................49

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3.5.1 RNA extraction and Northern blot ................................................ 49 4 RESULTS AND DISCUSSION .................................................................... 51

4.1 Tissue specificity of mitochondrial maintenance ................................. 51 4.1.1 Mitochondrial DNA copy number and topology ........................... 51 4.1.2 Tissue specificity in replication and correlation with mtDNA

damage and replisome components ........................................... 52 4.2 Mitochondrial topoisomerases ............................................................. 55

4.2.1 Topoisomerase localization in mitochondria and tissue and proliferation status specificity ....................................................... 55

4.2.2 Topoisomerase 2α, 2β and topoisomerase 3α functions in mitochondria ................................................................................ 56

4.2.3 Division of labor of mitochondrial topoisomerases ...................... 62 4.2.4 Ciprofloxacin-induced inhibition of mitochondrial function ........... 64

5 FINAL REMARKS AND FUTURE PERSPECTIVES .................................. 67 BIBLIOGRAPHY ................................................................................................ 69

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1 INTRODUCTION

1.1 MITOCHONDRIA Mitochondria are semi-autonomous organelles present in nearly every eukaryotic cell. Their core function is the conversion of food-derived energy into ATP by oxidative phosphorylation. Additionally, mitochondria host a plethora of other metabolic pathways, store calcium and regulate many aspects of cellular growth and death.

Mitochondria have a defined structure consisting of two membranes, a smooth outer mitochondrial membrane and a folded inner membrane (Figure 1). These mem-branes separate the functionally distinct compartments of intermembrane space and mitochondrial matrix. The protein complexes participating in electron transport and energy conversion are located in the inner mitochondrial membrane and allow elec-trons from the citric acid cycle and β-oxidation to transfer gradually to oxygen, gen-erating water. This process is coupled to the pumping of protons from the matrix to the intermembrane space, generating a proton gradient used to transform ADP to ATP by addition of Pi.

Mitochondria are often depicted as bacteria-sized ellipsoids of 2-8 µm, but their shape and size may differ greatly, and through constant fusion and fission they form a mitochondrial network (Westermann 2010). This flexible network is essential for mitochondrial quality control and turnover. Mitochondria exist in multiple copies per cell, being most abundant in tissues with high energy demand, like muscle and heart. In most animals, mitochondria are considered to be maternally inherited only, although exceptional cases of paternal inheritance have been reported (Luo et al. 2018, Schwartz & Vissing 2002). Their essential role in energy production makes any disturbance of mitochondrial function possibly life-threatening, and in fact many dis-eases are connected to mitochondrial dysfunction caused by a mutation in genes en-coding mitochondrial proteins. Mitochondrial function can also be impaired by dif-ferent extrinsic factors like drugs or environmental toxins.

According to a long-standing theory, mitochondria originate from proteobacteria, that were engulfed by another cell (Sagan 1967). Among the closest still existing rel-ative to the ancient mitochondria appears to be the α-proteobacterium Rickettsia prowazekii, an obligate endocellular parasite of eukaryotes causing louse-borne ty-phus in humans (Andersson et al. 1998). A bacterial origin is further supported by the fact, that the mitochondrial inner membrane contains cardiolipins, that are not found in any other mammalian cell membranes, but are common in bacterial mem-branes (Daum 1985).

The origin of mitochondria is still under debate. Common open questions are for example the identity of the host cell (e.g. archaeon or bacterium), the initial metabolic relationship (e.g. syntrophy, predation or parasitism) and the original evolutionary

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benefit of this relationship (e.g. energy or heat production). Also, the α-proteobacte-ria origin and affinity with Rickettsiales have been questioned (Zachar & Szathmáry 2017).

Figure 1. A typical mitochondrion possessing two membranes, a smooth outer membrane and a folded inner membrane. These two membranes divide the mitochondrion into functionally distinct compartments, the intermembrane space and the matrix. The protein complexes par-ticipating in oxidative phosphorylation are situated in the inner membrane. Proton pumping from the matrix through the complexes into the intermembrane space creates a proton gradi-ent, which is used by ATP synthase to transform ADP and Pi into ATP.

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Figure 2. Human mtDNA. Human mitochondrial DNA encodes 13 proteins (blue) which are part of the oxidative phosphorylation complexes, two ribosomal RNAs (red) and 22 tRNAs (black). Mitochondrial DNA is compact, and the only longer non-coding region (NCR) contains transcription promoters for light (LSP) and heavy strand (HSP), the H-strand origin of replica-tion OH, the termination associated sequence TAS and three conserved sequence blocks (CBS1, 2, 3). The bidirectional replication origin area Ori-z is located downstream of the non-coding region. The short 7S DNA is bound to the non-coding area, creating the displacement or D-loop (modified from Torregrosa-Muñumer 2018 and Uhler & Falkenberg 2015). 1.2 MITOCHONDRIAL DNA Mitochondria have their own multicopy genome. The human mitochondrial DNA (mtDNA) is a double-stranded circular molecule 16 569 base pairs in length. This genome is arranged in protein-DNA complexes, so-called nucleoids, in the mitochon-drial matrix (Satoh & Kuroiwa 1991). The two DNA strands can be distinguished by their nucleotide composition and are commonly referred to as heavy (H-) and light (L-) strand. Mammalian mtDNA encodes 13 protein-coding genes that are subunits of the oxidative phosphorylation (OXPHOS) complexes (Figure 2). Additionally, it

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encodes two ribosomal RNAs and 22 transfer RNAs required for the autonomous translation of the mtDNA-encoded proteins. The vast majority of mitochondrial pro-teins, among them many subunits of OXPHOS complexes and all mtDNA mainte-nance proteins, are encoded in the nuclear DNA and transferred into mitochondria post-translational (Chacinska et al. 2009).

Mitochondrial DNA is extremely compact; the protein-coding genes are separated only by tRNAs, it lacks introns and some genes even overlap. The only longer area not encoding any genes is the control region of the genome, and for this reason it is also called the non-coding region (NCR) (Figure 2). Here, transcription of mtDNA initiates from three promoters, the L-strand promoter LSP and two H-strand promot-ers, HSP1 and HSP2.

1.2.1 mtDNA replication

Because of the vital role of mitochondrially encoded proteins in ATP production and many other metabolic pathways, proper mtDNA maintenance is essential for cells, and mutations or depletion of this genome may lead to mitochondrial dysfunction and a number of diseases, affecting e.g. the central nervous system, heart or skeletal muscles.

Mitochondrial DNA replication is independent of the cell cycle and happens con-tinuously during the whole lifespan of the cell, also in post-mitotic and non-prolifer-ating cells without nuclear DNA replication (Bogenhagen & Clayton 1977). The pro-cess is rather slow, estimated to take approximately one to two hours to replicate the full genome (Berk & Clayton 1974).

Across the eukaryotic kingdom, several distinct mitochondrial replication mech-anisms exist (Pohjoismäki & Goffart 2011), and even for the well-studied mammalian replication mechanism no consensus has been reached. The first replication mecha-nism proposed for mammalian mtDNA based on electron microscopy observations was the strand-displacement mechanism (SDM) (Clayton 1982, Robberson et al. 1972), also referred as a strand-asynchronous replication mechanism (Figure 3A). In the SDM model, mtDNA replication initiates from a defined origin OH in the NCR, and H-strand synthesis proceeds continuously and unidirectionally (Kang et al. 1997). L-strand synthesis starts only when approximately two thirds of the H-strand have been replicated and the replication fork passes the origin of light strand repli-cation OL. The exposure of the single-stranded conformation of OL allows its folding into a stem-loop structure, which is recognized by the mitochondrial DNA-directed RNA polymerase (POLRMT) (Fusté et al. 2010). POLRMT synthetizes an RNA pri-mer, that initiates DNA synthesis of the light or lagging strand, oriented in the oppo-site direction of H-strand synthesis. In this model, both strands are synthetized con-tinuously, and therefore no Okazaki fragments are required.

While Clayton and co-workers proposed the exposed lagging-strand to be cov-ered and stabilized by a mitochondrial single strand binding protein (mtSSB), later

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studies found partially single-stranded replication intermediates to be covered with 200-600 nucleotides long RNA fragments (Pohjoismäki et al. 2010a, Yang et al. 2002, Yasukawa et al. 2006) (Figure 3B). To acknowledge the involvement of RNA in this replication mechanism, it was named Ribonucleotide Incorporation Throughout the Lagging Strand (RITOLS) (Yasukawa et al. 2006). Later, the RNA covering the lag-ging strand was shown to consist of processed L-strand transcripts, and the process of their laydown termed “bootlace mechanism” (Reyes et al. 2013). However, this issue is not yet settled, as there are also studies supporting the notion of a mtSSB-coated displaced strand as proposed by the SDM model (Fusté et al. 2014).

Mammalian mtDNA replication was for a long time believed to proceed exclu-sively via the above described strand-asynchronous, asymmetric replication mecha-nism, until in the year 2000 this mode was found to co-exist with a more conventional synchronous replication mechanism in human and mouse (Holt et al. 2000) (Figure 3A). This mechanism was named conventional strand-coupled, Okazaki-fragment associated (COSCOFA) replication. However, evidence of a synchronous, coupled type of replication had been published already 26 years earlier (Koike & Wol-stenholme 1974). In this mechanism replication intermediates are fully double-stranded, and the leading strand is replicated continuously, but unlike in the strand-asynchronous mechanism, the lagging strand is synthetized discontinuously and in-volves the formation of Okazaki fragments (Holt et al. 2000). Replication initiates from a broader zone (Ori-z) upstream of the non-coding region and continues bidi-rectionally until the first replication fork reaches OH and is paused, after which rep-lication continues only in the other direction (Bowmaker et al. 2003).

While the debate about the precise replication mechanisms of mtDNA continues, it is generally accepted that the mitochondrial genome is replicated by mtDNA pol-ymerase γ (Polγ), a trimer consisting of the catalytic subunit PolγA (140 kDa, p140 encoded by POLG) and the homodimeric accessory subunit PolγB (110 kDa, p55 en-coded by POLG2) which increases the processivity. Polγ also possesses reverse tran-scriptase activity (Murakami et al. 2003) and proofreading capability due to its 3’->5’ exonuclease activity (Longley et al. 2001). Polγ has been shown to function in vitro together with the mitochondrial helicase TWNK and mtSSB, but replication likely requires also several other proteins, among them at least one topoisomerase and a ligase. While candidates for these functions have been proposed, there is only indi-rect evidence for their identity.

The mitochondrial replicative helicase TWNK is a helicase closely related to the T7 phage helicase/primase gp4. The protein is encoded by the TWNK gene (also known as PEO1) and works as a hexamer or heptamer, with each monomer consist-ing of three functional domains, a primase domain, a linker region and a helicase region (Fernández-Millán et al. 2015). TWNK functions as a 5’-3’ helicase, unwinding the double-helix of the DNA ahead of the replication fork. Because of its primase domain, early studies proposed that TWNK also acts as primase in mtDNA replica-tion (Spelbrink et al. 2001, Korhonen et al. 2003), but a phylogenetical analysis of

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TWNK proteins throughout the animal kingdom showed this primase function to be lost in higher animals (Shutt & Gray 2006). This hypothesis has later been supported by several experimental studies finding no evidence of primase activity (Farge et al. 2007, Oliveira M.T., personal communication, 30.10.2020). TWNK has been demon-strated to regulate mitochondrial DNA copy number, with overexpression increasing and knock-down decreasing mtDNA levels in human cells (Tyynismaa et al. 2004). The interaction with mtSSB specifically stimulates the unwinding activity of TWNK (Korhonen et al. 2003), but also the DNA synthesis rate of Polγ (Oliveira & Kaguni 2011)

POLRMT, the mitochondrial RNA polymerase involved in mtDNA transcription, was shown to also function as a primase (Wanrooij et al. 2008). Based on studies in knock-out models, POLRMT was postulated to act as regulatory switch between mi-tochondrial DNA transcription and replication (Kühl et al. 2016).

Other important proteins include the mitochondrial transcription factor A (TFAM), that despite its name also appears to have a major role as a mitochondrial histone-like DNA-binding protein. TFAM binds to DNA nonspecifically, but also se-quence-specifically upstream of HSP1 and LSP (Dairaghi et al. 1995a, Wong et al. 2009). TFAM is able to bend and unwind DNA and thus it is the main component of packaging and organizing mtDNA into nucleoids.

The amount of TFAM in mitochondria is a contested issue. While several studies have found it be present in high concentrations, able to cover basically the whole mtDNA, others found it to be present in much lower levels (Maniura-Weber et al. 2004, Cotney et al. 2007, Kukat et al. 2011, Wang et al. 2013). It has been demonstrated that the amount of TFAM correlates with mtDNA copy number with the ratio staying stable during postnatal development of cardiomyocytes (Pohjoismäki et al. 2013a).

Ribonuclease H1 (RNase H1) is an endoribonuclease capable of hydrolyzing the RNA strands in RNA-DNA hybrids (Cerritelli & Crouch 2009). RNase H1 was shown to be essential for the removal of RNA primers in mtDNA replication, and the ab-sence of the protein leads to persistent RNA primers, creating obstacles for polymer-ase γ and thus for DNA replication (Holmes et al. 2015).

Like most replication mechanisms, also mitochondrial DNA replication requires the ligation of freshly replicated stretches of DNA strands. The only known mito-chondrial ligase is Ligase III, an enzyme having both nuclear and mitochondrial isoforms arising from different start codons of the same LIG3 gene (Simsek & Jasin 2011, Lakshmipathy & Campbell 1999). Loss of ligase III leads to depletion of mtDNA in mouse embryonic fibroblasts and to increased nicking and attenuation of mtDNA replication under stress conditions (Rauhanen et al. 2011, Shokolenko et al. 2013), supporting its importance for mtDNA replication. Overexpression of DNA ligase III in mitochondria protects cells against oxidative stress and improves mitochondrial DNA base excision repair (Akbari et al. 2014), indicating its role not only in mtDNA replication, but also in repair of mtDNA.

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1.2.1.1 Phases of mitochondrial DNA replication Replication of mitochondrial DNA can be separated into three distinct phases, 1. in-itiation, 2. elongation, and 3. termination, each needing a distinct set of protein fac-tors.

The initiation phase of asynchronous replication starts from the precise origin OH, or less frequently from Ori-b 500 bp downstream from OH (Crews et al. 1979, Kang et al. 1997, Yasukawa et al. 2005). Although no specific primase has been identified, replication is most likely initiated through the abortion of POLRMT-driven L-strand transcription at the conserved sequence block II (CSB II), producing a primer used for the start of DNA synthesis by Pol.

While asynchronous mtDNA replication appears to be the default mechanism for mtDNA replication in cultured cells, synchronous replication was described to dom-inate under situations of replication stress and mtDNA depletion. Initiation of this type of replication starts from a broader zone upstream of the non-coding region, named Ori-z (Bowmaker et al. 2003). The priming events starting this type of H-strand replication as well as the lagging strand replication are not yet known, al-though in vitro POLRMT was shown to function for both (Wanrooij et al. 2008). The regulatory factors or circumstances deciding the replication mode and the place of initiation are unclear.

The elongation phase continues the synthesis of mtDNA after the initiation of rep-lication. Here, the core replisome consists of Pol, TWNK and mtSSB, that also func-tion in a minimal replisome in vitro (Korhonen et al. 2004). Within the mitochondrion, additional factors are required, among them a topoisomerase relieving the torsional stress caused by the moving replication fork. In the case of synchronous replication involving Okazaki-fragments, a primase creating frequent primers and a ligase are also required.

In the termination phase of mtDNA replication, two replication forks converge in the non-coding region. Similar to other genetic systems, termination of mtDNA rep-lication requires four actions, although the order of events might vary: firstly, forks converge until the DNA is completely unwound. Secondly, any remaining gaps are filled by a polymerase and ligated. Thirdly, proteins of the replication machinery are unloaded and lastly, catenanes are removed. In a case of a circular genome like mtDNA, both hemi- and full catenanes might arise, requiring different topoisomer-ase activities for their resolution.

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Figure 3. Mammalian mitochondrial DNA is replicated using either the asynchronous or the strand-coupled mode. (A) In the asynchronous mechanism leading strand replication origi-nates from OH and proceeds unidirectionally until replication passes the origin of light strand OL, allowing the initiation of light strand replication. The lagging strand template is covered with RNA in the RITOLS model or with mtSSB in the SDM model. (B) Initiation of replication in the strand-coupled mechanism starts from a broader zone upstream of the non-coding region. The DNA synthesis of both strands is coupled and proceeds bidirectionally. The lagging strand replication is discontinuous and requires Okazaki-fragments. When the first replication fork reaches OH, it will pause until the forks converge in the non-coding region and the daughter molecules are separated by a topoisomerase (modified from Torregrosa-Muñumer 2018).

1.2.2 7S DNA Depending on the organism and tissue, between 1% and 90% of mtDNA molecules contain a triple-stranded region called the D-loop (reviewed in Nicholls & Minczuk 2014). The third strand is a ca. 650 nt long sequence called 7S DNA, named after its sedimentation rate in Svedbergs units. 7S DNA locates at the non-coding region of mitochondrial DNA, typically between OH (5´ end) and TAS (3´ end) (Figure 2). The 5´ end can vary, giving rise to a group of molecules with variable size. Although this

A B

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peculiar structure was found 50 years ago, its precise function still remains unknown. It has been suggested that the D-loop is the result of prematurely terminated heavy-strand replication and might be involved in the control of transcription initiation (Clayton 1982). It might also serve as a binding center for proteins due to its partially relaxed conformation (He et al. 2007) or mediate replication fork arrest through the formation of a structural barrier, as replication almost never extends beyond OH (Bowmaker et al. 2003). The half-life of 7S DNA is only ca. 45-60 min in rodent cells (Bogenhagen et al. 1979, Gensler et al. 2001), raising the question of why mitochon-dria invest so much energy into maintaining this structure. 1.2.3 Mitochondrial transcription Mitochondrial transcripts are produced as polycistronic precursors and later pro-cessed into separate RNAs. Both strands of the mitochondrial DNA are transcribed, although the L-strand is the main coding strand, and the H-strand encodes only one protein and several tRNAs. Transcription initiates at defined promoters located in the non-coding region, HSP and LSP. H-strand transcription appears to initiate occa-sionally from an alternative promoter, leading to the nomenclature of HSP1 and HSP2 (Montoya et al. 1982, Montoya et al. 1983). It has been proposed that HSP1 promotes only the transcription of ribosomal genes, whereas initiation from HSP2 would produce a whole-genome-length transcript (Montoya et al. 1983). The exact regulation of choice between these two promoters is not yet fully known, although the binding of the mitochondrial termination factor mTERF1 appears to stimulate HSP1, while high concentrations of TFAM seem to repress HSP2 (Lodeiro et al. 2012, Martin et al. 2005).

RNA synthesis during mtDNA transcription is catalyzed by POLRMT, a typical DNA-dependent RNA polymerase. Two accessory proteins interacting with POLRMT have been described, the mitochondrial transcription factors A and B, or TFAM and TFBM (Falkenberg et al. 2002). Two isoforms exist for TFBM, with TFB1M likely being a methyl transferase involved in translation, while TFB2M truly interacts with POLRMT and stimulates its activity.

The role of TFAM for mitochondrial transcription is less clear. Although a homo-zygous knock-out of TFAM is embryonically lethal (Larsson et al. 1998), it has been debated whether TFAM is essential for transcription in general, as in vitro transcrip-tion is initiated at all promoters also in the absence of TFAM. Thus, TFAM might play a role in the transcription activation without being an essential component of the in-itiation complex (Shutt et al. 2010). On the other hand, in higher salt concentrations TFAM appears to be essential for transcription also in vitro (Shi et al. 2012). In addi-tion, mutation of TFAM binding sites abolishes transcription, indicating that TFAM and structural changes caused by its DNA binding are essential for transcription

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under normal conditions (Dairaghi et al. 1995a, Dairaghi et al. 1995b, Gaspari et al. 2004).

Transcription, like replication, can be divided in three steps, initiation, elongation and termination (reviewed in Shokolenko & Alexeyev 2017). Initiation of mitochon-drial transcription starts by TFAM binding upstream of the promoter and inducing a bend in mtDNA. This conformational change recruits POLRMT to the site, followed by TFB2M, which facilitates promoter melting and recruitment of the priming nucle-otide, which allows POLRMT to initiate RNA synthesis. After transcription has started, TFB2M dissociates and the mitochondrial transcription elongation factor (TEFM) increases the processivity of the polymerase on the L-strand.

In a circular genome, prevention of potentially hazardous collisions between tran-scription and replication machineries is important. TEFM has been proposed to work as a molecular switch between replication and transcription, facilitating the bypass of a G-quadruplex structure in the human conserved sequence block CSBII, which might otherwise terminate transcription and instead create a primer for mtDNA rep-lication (Agaronyan et al. 2015). It has been suggested that TFAM acts in a concen-tration-dependent manner, increasing transcription of rRNAs at low concentration, at moderate concentrations increasing mRNA transcription and mtDNA replication and at high concentration suppressing both transcription and replication (Lodeiro et al. 2012, Pohjoismäki et al. 2006, Shutt et al. 2011).

How mitochondrial transcription is terminated is still poorly known. mTERF1 is thought to mediate termination specifically at the 3’-end of the 16S rRNA gene, where its binding to the DNA induces a bend and interferes with the transcription machin-ery (Yakubovskaya et al. 2010). Martin et al. (2005) proposed a novel model, suggest-ing mTERF1 to simultaneously bind to the sites of mtDNA transcription termination and initiation, creating a loop enabling the circulation of the transcription machinery and thus increasing the expression rate of ribosomal transcripts. In fact, transcripts are unequally present in mitochondria, with HSP1 contributing to 80%, HSP2 to 9% and LSP to 11% of all transcripts, based on RNAseq analyses (Mercer et al. 2011). 1.3 MITOCHONDRIAL DNA DAMAGE 1.3.1 mtDNA damage and repair Oxidative phosphorylation is essential for ATP production, but it is also a source of reactive oxygen species (ROS). Up to 10% of electrons exit the electron transport chain prematurely and reduce oxygen incompletely to create superoxide radicals (Ja-stroch et al. 2010). Although low levels of ROS are important cellular signaling mol-ecules, in high quantities they can damage DNA, proteins and lipids by oxidation. The close proximity of mtDNA to the OXPHOS chain and the ROS produced thus makes it prone to oxidative damage (Yakes & Van Houten 1997). Additionally, the asynchronous replication mechanism of mtDNA makes the H-strand more

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vulnerable for damage during the single-stranded phase of replication. In the nu-cleus, DNA is protected by histones; and although mitochondrial DNA lacks a chro-matin structure, TFAM is at least to some extent protecting mtDNA by compacting it into nucleoids. The DNA damage caused by ROS can arrest replication and tran-scription and is therefore deleterious for both nuclear and mitochondrial DNA. The C8 atom of guanine is most frequently oxidized, generating 8-oxo-Guanosine, a com-monly used marker of oxidative damage (Ba & Boldogh 2018). On the other hand, recent studies have questioned the significance of ROS for mitochondrial DNA mu-tations, as the observed mutations appear to arise from replication errors and/or spontaneous base hydrolysis rather than oxidative damage (Kennedy et al. 2013, Ameur et al. 2011).

In addition to oxidative damage, mitochondrial DNA is also exposed to other types of DNA damages, like alkylation, hydrolytic damage, adduct formation, and DNA strand breaks (García-Lepe & Bermúdez-Cruz 2019). For a long time, it was suspected that mitochondria lack DNA repair mechanisms, and damaged molecules would be simply degraded and replaced by newly replicated ones. This hypothesis was based on the absence of a pyrimidine dimer repair mechanism in mammalian mitochondria after UV light exposure (Clayton et al. 1974). However, years later sev-eral repair mechanisms were found to function in mitochondria (LeDoux et al. 1992), the best studied being base excision repair pathway (BER) involved e.g. in the repair of oxidative lesions in mitochondrial DNA. Likewise, there is evidence to support the existence of homologous recombination (HR) (Thyagarajan et al. 1996), nonhomolo-gous end joining (NHEJ) (Coffey et al. 1999), and mismatch repair (MMR) (Mason et al. 2003). While many proteins involved in mitochondrial DNA repair pathways are shared with the nucleus, the mitochondrial systems appear to have less components. 1.3.2 mtDNA recombination Mammalian mitochondrial DNA recombination has long been a controversial sub-ject. Given the usually homogenic mtDNA pool, distinguishing recombined mole-cules is difficult. Therefore, recombination has been observed in humans only in rare cases, where paternal inheritance of mtDNA leads to a heteroplasmic situation, with more than one type of mitochondrial genomes coexisting (Kraytsberg et al. 2004). Inter-molecular heterologous mtDNA recombination was observed in a study con-ducted with human cytoplasmic hybrid cell lines, where two cell lines with different pathogenic mtDNA mutation were fused together (D’Aurelio et al. 2004). Neverthe-less, no recombination was observed in the liver or kidney of a mouse model with two different mtDNA haplotypes (Hagström et al. 2014). Conversely, very low levels of recombined mtDNA has been observed in the mitochondria of skeletal muscles of hetoroplasmic mice and humans (Sato et al. 2005, Zsurka et al. 2005). These conflict-ing observations might be explained by tissue-specific differences in the mtDNA

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replication mechanism, as high levels of cruciform, possibly recombining, molecules might be caused by a greater need for DNA repair by recombination due to oxidative damage. Fittingly Bacman et al. (2009), using the same mtDNA haplotype mouse model than Hagström et al., found that upon induction of double-strand breaks the level of recombination was increased in mtDNA. Similar results were seen in rat tes-tes, brain, kidney, and spleen, when double-strand breaks were introduced artifi-cially (Dahal et al. 2018).

As little is known about mammalian mitochondrial recombination, also the fac-tors involved in its mechanism are yet unknown. However, DNA recombination ma-chineries require several essential components. In the nucleus three alternative path-ways are known to process double-Holliday junctions formed by homologous re-combination. One of them is the BTRR complex formed by the RecQ family helicase BLM (Bloom syndrome protein), Top3α and the RecQ-mediated genome instability proteins 1 and 2 (RMI1, RMI2) (Sarbajna & West 2014). In the nucleus, Top3α has been observed to bind double-strand ends, possibly recruiting other components such as DNA2 (Daley et al. 2014). In mitochondria, Top3α seems to work inde-pendently of the BTR complex (Nicholls et al. 2018) and its involvement in recombi-nation is unclear. Interestingly, a RecQ helicase, RecQ4, has been found in mitochon-dria, along with other helicases (Croteau et al. 2012, Ding & Liu 2015).

For recombination-mediated replication initiation, the TWNK helicase might be an important component, as it has been shown to possess the annealing activity re-quired for the catalysis of strand-exchange reactions and branch migration (Sen et al. 2012, 2016). Fittingly, overexpression of TWNK in mouse increases mtDNA recom-bination (Pohjoismäki et al. 2009, 2013b).

The existence of mitochondrial recombination has been questioned by the obser-vation that typically mitochondrial nucleoids contain only one copy of mtDNA, re-ducing the chance of recombination (Kukat et al. 2011). Again, this might depend on the type and the physiological state of a cell.

Specific areas of the D-loop are observed to be hot-spots for recombination, and the region around 16 070 bp in the human mtDNA D-loop has been suggested to be prone to intra-molecular recombination and deletion formation (Krishna et al. 2008).

1.4 mtDNA TOPOLOGY Mitochondrial genomes have enormous variability in gene content and size, but they also have several different organizations, diverging from simpler linear molecules to complex interlocked multicircular kinetoplast networks present in Trypanosoma (Nosek & Tomáška 2003). Mammalian mitochondrial DNA molecules are mainly present as single one-genome-size double-stranded circle with different degrees of supercoiling, but they can also form circular dimers or catenanes of two or more in-terlinked molecules (Kolesar et al. 2013) (Figure 4). Topological forms of mtDNA can

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also vary between tissues; and for example, human brain and heart have a high level of catenated molecules with abundant replication and recombination junctions (Poh-joismäki et al. 2009, Pohjoismäki et al. 2010b). The complexity of DNA topology seems to correlate with an increase in mtDNA copy number (Pohjoismäki et al. 2010b).

Figure 4. Topological forms of mtDNA. Mammalian mitochondrial DNA can have several dif-ferent topological forms, typically being open or supercoiled circles of one-genome-length. Two or more interlinked DNA molecules form catenanes. mtDNA can also exist as head-to-tail or head-to-head dimers or in linear form. 1.4.1 Topoisomerases DNA topology cannot change freely; instead most alterations require the action of topoisomerases. These DNA-modifying enzymes are capable of catalyzing DNA strand breaks, rotating or passing one strand through another and finally religating the cut ends without creating permanent breaks in the DNA. Topoisomerases can change coiling and winding of a DNA molecule or separate interlinked molecules, and their function is needed in replication, transcription, segregation, repair, and re-combination. Topoisomerases induce transient strand breaks by transesterification between the active center tyrosyl and the phosphor group in the DNA, leading to the topoisomerase being covalently attached to the DNA. This transient bond is broken only after a change in the DNA topology is made, and the strands are resealed (Champoux 2001).

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Topoisomerases can be divided into two groups based on their mechanism (topoi-somerase functions and classifications are summarized in table 3). Type I topoiso-merases cut one DNA strand and transfer the other strand through before rejoining the ends. Typically, enzymes in this group are monomers and do not require ATP for their function. Enzymes belonging to this group can further be separated into type IA and IB, with the two groups sharing no sequence or structural similarities (Champoux 2001). Type IA enzymes can relax negative supercoils and decatenate single-stranded DNA by binding to the 5’ end of the cut DNA. Type IB topoisomer-ases catalyze relaxing of positive and negative supercoils by rotation, binding to the 3’-end of the DNA.

Type II topoisomerases cut both strands of the double-stranded DNA, enabling a second DNA double-strand to pass through the cut (Nitiss 2009). Enzymes in this group are multimeric; they relax both negative and positive supercoils or decatenate linked molecules and require ATP for their function. Type II topoisomerases are also comprised of two subgroups. The topoisomerase IIA group contains the majority of type II enzymes such as bacterial TopIV and eukaryotic Top2, while type IIB topoi-somerases are mainly archaebacterial enzymes. The basic mechanism of action is si-milar between IIA and IIB, but topoisomerases belonging into the IIA group create four base overhangs during the double strand cut (Morrison & Cozzarelli 1979, Sander & Hsieh 1983), while the overhangs formed by type IIB topoisomerases con-tain two bases (Buhler et al. 2001).

The importance of topoisomerases for DNA maintenance is well recognized, and their function in the nucleus is studied extensively, while until recently, little has been known about their mitochondrial counterparts. Like any longer DNA molecule, mitochondrial DNA is prone to topological stress, and the circular organization of mammalian mtDNA adds to the topological challenges. Until now, only four differ-ent topoisomerases have been identified in mammalian mitochondria. The only mi-tochondria-specific topoisomerase is Top1mt, encoded by its own gene (TOP1MT) distinct from nuclear Top1 (Zhang et al. 2001). Another mitochondrial type I topoi-somerase is Top3α, with the mitochondrial protein being transcribed from the same gene (TOP3A) as its nuclear counterpart but using an alternative start codon (Wang et al. 2002). The search for mitochondrial type II topoisomerases has been eventful. The need for a Top2 function is obvious, as the circular organization of mtDNA should lead to catenation, but different studies have had conflicting data of the pres-ence of Top2 in mitochondria. Based on mass spectrometry analyses, an N-terminally truncated version of nuclear Top2 enzyme was proposed to exist in mitochondria, and because of the bacterial origin of mitochondria even the presence of a bacterial type gyrase has been speculated (Low et al. 2003). However recent studies, including our own, suggest the presence of two isoforms of Top2 similar or identical to nuclear Top2α and Top2β (Zhang et al. 2014).

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1.4.2 Top1mt Top1mt is the only mitochondrial topoisomerase in vertebrates encoded by a mito-chondria-specific gene. Mitochondrial Top1 is highly similar to the nuclear version, although it lacks part of the N-terminal end containing nuclear localization signals; instead it has a mitochondrial localization sequence (Zhang et al. 2001). In compari-son to nuclear Top1, Top1mt has reduced DNA-binding affinity, and nuclear Top1 is therefore unable to replace Top1mt in mtDNA maintenance (Dalla Rosa et al. 2009).

Lack of Top1mt leads to an increased negative mtDNA supercoiling, suggesting that relaxing negative supercoils is a prominent function of Top1mt (Zhang et al. 2014). In contrast to its nuclear counterpart, it was suggested to negatively regulate mitochondrial transcription, as deficiency or deletion of Top1mt increases mitochon-drial transcripts, whereas overexpression lowers them (Sobek et al. 2013). Top1mt binds to the non-coding region and its cut sites also map to the D-loop region, espe-cially to the beginning and the end of the D-loop, as well as to OL. Inhibition of Top1mt reduces the level of 7S DNA, and it has been suggested that Top1mt plays a role in D-loop homeostasis and could serve as a topological barrier to shift from tran-scription to replication (Zhang & Pommier 2008, Dalla Rosa et al. 2014, Dalla Rosa et al. 2017). However, Top1mt is not essential for mitochondrial DNA maintenance, as Top1mt -/- knock-out mice are viable, fertile and express mtDNA-encoded proteins, even though MEF cells derived from these mice suffer from dysfunctional mitochon-drial respiration associated with decreased ATP production and increased ROS (Douarre et al. 2012). Mice lacking Top1mt were also found to suffer from impaired liver regeneration, probably due to mtDNA amounts being insufficient to support cell proliferation, suggesting that Top1mt has an important role during development (Khiati et al. 2015). This theory is supported by the observation that mitochondrial dysfunction in Top1mt -/- MEF cannot be rescued by ectopic expression of Top1mt (Sobek et al. 2013). The adverse effects of Top1mt deletion become more evident after exposure of Top1mt -/- mice to the type II topoisomerase inhibitor doxorubicin. Treated animals suffered from mtDNA depletion and mtDNA damage in heart tis-sue, among other mitochondrial defects, leading to increased cardiotoxicity and death (Khiati et al. 2014).

1.4.3 Top3α Eukaryotic cells possess two Top3 proteins, Top3α and Top3β. Top3α has a function in recombination and resolution of hemicatenanes and double-Holliday junctions. It also resolves strand separation-caused solenoids, thus being essential for transcrip-tion. Top3β is the only human topoisomerase active also on RNA, and it is suggested to be important in transcription and R-loop resolution (Wilson-Sali & Hsieh 2002, Stoll et al. 2013). Like Top1, Top3 works as monomer. As type IA enzymes, Top3α and Top3β cleave one strand of the DNA double-strand and change topology by

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passage of the other strand. For their action they require Mg2+, but not ATP. They can relax negatively, but not positively supercoiled DNA (Champoux et al. 2001).

In mitochondria, only Top3α has been found. Mitochondrial Top3α is encoded by the same gene as its nuclear counterpart, using an alternative start codon creating a mitochondrial targeting signal at the N-terminus of the protein (Wang et al. 2002). How this alternative transcription is regulated is yet unknown, and the mistargeting of Top3α has been suggested to be a possible cause of human disorders associated with mtDNA depletion (Wu et al. 2010). In Drosophila, mitochondrial Top3α defi-ciency was demonstrated to lead to mtDNA depletion, causing loss of germline stem cells and defects in fertility (Wu et al. 2010).

The function of Top3α in vertebrate mitochondria has been studied less, but re-cently Nicholls et al. (2018) proposed a role in decatenation of newly synthesized mtDNA molecules after replication termination. In their study, the silencing of Top3α expression by siRNA lead to replication stalling, mtDNA copy number de-crease and accumulation of catenated molecules. The importance of mitochondrial Top3α was also supported by the identification of patient mutations in Top3α caus-ing progressive external ophthalmoplegia (PEO), a syndrome frequently associated with mtDNA replication dysfunctions. In E. coli TopIII has been shown to function with a RecQ-like DNA helicase in the resolution of converging replication forks (Suski & Marians 2008).

1.4.4 Top2 Eukaryotic cells have two type II topoisomerase isoforms, Top2α and Top2β. While these isoforms are encoded by separate genes, they share 68% sequence identity, with the N-terminal part being even more conserved with 78% identity (Tsai-Pflugfelder et al. 1988, Austin et al. 1993, Tan et al. 1992, Jenkins et al. 1992). The protein forms a homodimer, with each subunit consisting of an N-terminal ATPase domain, a central cleavage and re-joining domain and a C-terminal domain (Austin et al. 1995). A nu-clear localization signal is located in the C-terminal domain of the protein, and this is also a site for post-translational modifications (Mirski et al. 1997).

Top2α and β have different cellular expression profiles and specific cellular func-tions in the nucleus. Top2α is predominantly expressed in proliferating cells, whereas expression of Top2β is not tied to the cell cycle and more closely related to differentiation and maturation (Watanabe et al. 1994, Turley et al. 1997). Top2α has an essential function in chromosome segregation during the cell division in prolifer-ating cells, and the knock-out of Top2α arrests murine embryonic development, lead-ing to embryonic death (Akimitsu et al. 2003). On the other hand, Top2β seems to play an important role in the development of neurons, affecting the expression of particular postmitotic genes. Lack of Top2β caused dysfunction in axon develop-ment, and Top2β-/- mice die shortly after birth due to defects in axonal growth,

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resulting in failure to move or breathe (Yang et al 2000). Additional non-canonical functions for Top2β have been suggested in the transcriptional promoters of espe-cially long genes, involving targeted double-strand break formation. Topoisomerases are also important factors in DNA repair, and topoisomerase 2β activity was shown to correlate with DNA repair capability (Popanda & Thielmann 1992, Sabourin & Osheroff 2000, Emmons et al. 2006, Mandraju et al. 2008, Mandraju et al. 2011, Kenig et al. 2016).

The mitochondrial localization of type II topoisomerases has long been controver-sial. Low et al. (2003) suggested that Top2β would localize to bovine mitochondria as a shorter version, lacking the C-terminal domain containing the nuclear targeting sequence, and its absence would allow mitochondrial localization (Low et al. 2003). As no mitochondrial targeting sequence has been identified in Top2, in contrast to Top1mt and Top3α, the idea of a mitochondrial truncated Top2β version sounds fea-sible. However, other studies have seen no evidence of this shorter version, instead full-length Top2α and Top2β proteins have been detected in mitochondrial protein lysates (Zhang et al. 2014). Nevertheless, a recent study failed to detect either type II topoisomerases in the mitochondrial compartment (Nicholls et al. 2018).

While the debate about the presence of Top2 enzymes in mitochondria continues, the function of these enzymes in mitochondria has not been studied. The potential dual localization of Top2 enzymes makes the analysis of their mitochondrial func-tions difficult. Given that expression of Top2α is tightly connected to the proliferation state of the cell, Top2β should be more important for mitochondrial function, as mtDNA replication, transcription and other maintenance is required also in non-pro-liferating cells lacking Top2α expression. Although not essential for mitochondria, Top2α was suggested to be involved in scaffolding the NCR and protecting the 7S ends from degradation, as Top2α cleavage complexes map to the ends of the D-loop region (Zhang et al. 2014). The function of Top2β in mitochondria has been virtually unstudied before the research presented in this thesis. 1.5 MITOCHONDRIAL DISEASES To date, nearly 300 diseases have been connected to mitochondrial dysfunction caused by mutations in mitochondrial DNA. They form a big heterogenic group of pathologies with varying appearances. The first syndromes linked to mitochondrial DNA mutations were Leber’s hereditary optical neuropathy and mitochondrial my-opathies in 1988 (Holt et al. 1988, Wallace et al. 1988). As most mitochondrial proteins are encoded in the nuclear genome, also mutations in these nuclear genes can affect mitochondrial function and result in a mitochondrial disease. The same disease phe-notype can be caused by different mutations, for example 20% of known mutations causing Leigh’s syndrome are of mtDNA origin, while in 80% of the cases the muta-tion is in a nuclear gene (NIH). Although mitochondrial diseases are individually

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reasonably rare, they are usually severe and often lethal, since they interfere with the cell’s fundamental requirement for mitochondrially produced energy.

A special characteristic of mitochondrial diseases caused by mtDNA mutations is the so-called threshold effect, when a certain level of mutated mtDNA has to be reached before any clinical symptoms manifest. This results from multiple copies of mtDNA existing in each mitochondrion and the organelle’s capability to function as long as sufficient mtDNA copies are unharmed (Rossignol et al. 2003). The degree of heteroplasmy often correlates with the severity of the disorder. Mitochondrial dis-eases often have distinguished tissue specificity, naturally affecting most commonly organs and tissues with high energy demand. Mitochondrial defects often affect mul-tiple tissues, leading to multi-system diseases (Nunnari & Suomalainen 2012). How-ever, it should be noted that the same mutation can cause very different presentations in different patients. While some patients with MELAS, caused by a point mutation in a mitochondrial tRNA gene, may suffer from strokes, others might develop cardi-omyopathy, myopathy or hearing loss and diabetes (Suomalainen 2011). Common symptoms of mitochondrial diseases include, but are not limited to, muscle weak-ness, neurological problems, visual and hearing problems, and gastrointestinal dis-orders.

In addition, many common diseases are connected to mitochondrial dysfunction, including neurodegenerative diseases such as Parkinson’s disease and Alzheimer’s disease, diabetes mellitus, metabolic diseases, and many cancers (Chan et al. 2006). Mitochondria have also been speculated to be a major player in aging. Even though efforts have been made, currently there is no cure for any of the above-mentioned mitochondrial or mitochondria-related diseases; and a deeper understanding of the mitochondrial physiology is needed to achieve better clinical outcomes. 1.5.1 Drug induced mitochondrial dysfunction Mitochondrial diseases can also originate from external factors, such as drugs, infec-tions, or other environmental causes. The importance of mitochondrial dysfunction has been long understated in the investigation of adverse side effects of pharmaceu-tical drugs. Typically, safety studies of pharmaceutical drugs do not assess mitochon-drial DNA damage, copy number or transcription changes, although many drugs have been shown to interfere with mitochondrial function, causing severe side effects recognized only after the substances have been in use (Fromenty 2020). Often the exact mechanism of the observed effects remains unknown. Drugs can cause mito-chondrial dysfunction through many ways, e.g. by inhibition of oxidative phosphor-ylation complexes or disruption of the membrane potential (Nadanaciva & Will 2011). Pharmaceuticals can also affect mitochondrial DNA maintenance through ROS-mediated oxidative mtDNA damage, or directly inhibit mtDNA replication, causing reduced mtDNA levels. Some pharmaceutical drugs can also impair mtDNA translation or mtDNA methylation.

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Because of the supposed bacterial origin of mitochondria, antibiotics targeting bacterial components of DNA maintenance or protein production often have the po-tential to also inhibit their mitochondrial counterparts. Many antibiotics inhibit bac-terial protein synthesis by binding to ribosomes, and some, like linezolid and tetra-cyclines, also target mitochondrial ribosomes, disturbing mitochondrial protein syn-thesis (Moullan et al. 2015, Santini et al. 2017). Many nucleoside analogue-based re-verse transcriptase inhibitors, employed as antiretrovirals, impair DNA polymer-ase γ, causing defective mtDNA replication and mtDNA depletion (Lewis & Dalakas 1995).

Drug-induced mitochondrial dysfunction can lead to variable symptoms, whose mitochondrial origin is often not recognized. Like in mitochondria-related diseases, the effects of drug-induced mitochondrial impairment are typically observed in tis-sues with high energy demand, like heart or kidney, but also in tissues exposed to high drug concentrations, such as liver (Dykens & Will 2007). The side-effects of mtDNA-affecting drugs can occur long after the beginning of the treatment, when a threshold is reached. The question why only some patients are affected, while the drug is well tolerated by most, is still partly unanswered. As animal studies employ-ing healthy animals might not give reliable results, a mouse model with reduced su-peroxide dismutase 2 levels (Sod2 +/-) has been used to study drug-induced toxic ef-fects in an animal suffering from clinically silent mitochondrial stress (Ong et al. 2006, Ong et al. 2007). Most likely, the patient’s genetic background and pre-existing mito-chondrial conditions play a major part in the appearance of mitochondrial adverse effects. For an example, some off-target effects of aminoglycosides are linked to spe-cific mutations in mtDNA (Prezant et al. 1993). Hence it would be of great benefit to study the genetic background of patients affected by a drug to develop safer and more personalized treatment. 1.5.2 Inhibition of topoisomerase function Topoisomerases are essential for the maintenance of the genome, but their action can also damage DNA, if cleaving and ligation of DNA strands are not properly regu-lated. Therefore, levels of topoisomerase are closely controlled. On one hand, an in-sufficient amount of topoisomerases leads to problems in DNA replication and chro-mosome segregation, on the other hand too much might cause permanent double-strand breaks, leading to mutations, cell apoptosis or development of cancer (Deweese & Osheroff 2008).

Many compounds have the potential to cause breaks in DNA by interfering with the cutting or rejoining reactions of topoisomerases. These substances can be divided in two main groups, topoisomerase poisons and topoisomerase inhibitors (reviewed in Deweese & Osheroff 2008). Poisons target the topoisomerase-DNA complex and increase the amount of cutting complexes, while inhibitors interfere with the cataly-tical turnover, reducing enzyme activity. Topoisomerase 2 poisons turn

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topoisomerases into a genome-damaging factor, which creates DNA breaks and pos-sibly leads to mutagenesis. Topoisomerase poisons consist of a heterogenic group of chemicals which increase cutting complex number by two different ways. Some of them inhibit the ability to rejoin cut ends, e.g. etoposide, a chemical used in cancer therapy. Other poisons increase the speed of cutting. Topoisomerase poisons can also be divided by their mechanism of action in two different groups: covalent and non-covalent poisons. Non-covalent poisons affect the protein-DNA interface, such as etoposide, genistein, doxorubicin and quinolones. Covalent poisons instead change the structure of the enzyme or DNA covalently. The best-known covalent topoiso-merase poisons are quinones. The classification of topoisomerase-interfering chemi-cals is not always straightforward, as substances can cause inhibition by several dif-ferent mechanisms and the mechanisms can be dose- or time-dependent.

Topoisomerase function is critical for the DNA maintenance and proliferation of any cell, and their activity is specially needed in rapidly dividing cancer cells. Ac-cordingly, many anticancer drugs in clinical use target the DNA cleavage/ligation reaction of topoisomerases in order to cause extensive DNA damage in cancer cells (Fortune & Osheroff 2000). Typically, Top2α is the intended target, as it is essential for chromosome separation and its inhibition leads to interrupted cell division. Due to the extensive similarity between Top2α and Top2β, a strictly isoform-specific agent has not been identified, and the unintentional inhibition of Top2β, needed also in post-mitotic cells, might be the cause for the observed side effects of Top2 inhibi-tors (Azarova et al. 2007). 1.5.3 Ciprofloxacin Quinolones are a group of antibacterial drugs widely used to treat bacterial infec-tions. Nalidixic acid was the first compound used clinically in 1962 (Lesher et al. 1962). Later on, several modifications were made to improve the biological activity, pharmacokinetic profile and tolerability, most significantly the addition of fluorine in the C-4 position, creating a group of antimicrobials called 4-fluoroquinolones (Ball 2000). This optimization has led to a second, third and fourth generation of quin-olones, with increasing spectrum of action.

Ciprofloxacin is a 4-fluoroquinolone belonging to the second generation of quin-olones. It is a broad-spectrum antibiotic commonly used to treat many bacterial in-fections, including urinary tract and respiratory tract infections. Its effect is based on the ability to inhibit the function of bacterial type II topoisomerases DNA gyrase and TopIV, which leads to high levels of double-strand breaks in the bacterial genome and ultimately to the death of the bacterial cell (Anderson et al. 1998). Despite the drug being considered to be well tolerated, it occasionally causes severe side effects, affecting e.g. gastrointestinal, hepatic and central nervous system functions.

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Lawrence et al. (1996) suggested ciprofloxacin to specifically inhibit the function of an uncharacterized mitochondrial type 2 topoisomerase, and later ciprofloxacin was indeed shown to affect mitochondria, causing increased oxidative stress and ef-fects on mtDNA copy number, oxygen consumption, mitochondrial membrane po-tential, production of ATP and activation of apoptosis. The exact mechanism of the observed effects was unknown (Aranha et al. 2000, Aranha et al. 2002, Jun et al. 2003, Gürbay et al. 2005, Kozieł et al. 2006, Lowes et al. 2009, Abdel-Zaher et al. 2012, Ghaly et al. 2014).

Only in 2018 the European medicines agency proposed the suspension of the mar-keting authorization of some quinolones and fluoroquinolones as well as the re-stricted usage of the remaining fluoroquinolone antibiotics because of their poten-tially permanent side effects. Currently, these drugs are not recommended for the treatment of infections that get better without treatment or are not severe (EMEA/H/A-31/1452). Ciprofloxacin has been described to inhibit also eukaryotic topoisomerase 2, and consequently ciprofloxacin and other fluoroquinolones have been proposed as anticancer drugs (Jamieson et al. 2014, Sharma et al. 2020).

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2 AIMS OF THE STUDY

The aim of the present study was to analyze the tissue specificity of mitochondrial DNA maintenance mechanisms, with a special focus on the functions of topoisomer-ases in mtDNA replication. The specific objectives were to:

1. Study the tissue specificity of mitochondrial DNA maintenance and expres-sion in different mouse tissues.

2. Determine the function of topoisomerase 2 and 3 in mitochondrial DNA maintenance.

3. Study the mechanism of mitochondrial involvement in drug-induced ad-

verse effects of antibiotics.

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3 MATERIALS AND METHODS

3.1 CELL LINES AND MICE 3.1.1 Mice (I) The tissue variation of mtDNA maintenance factors and mechanisms was studied in 10-13 week old female C57 BL/6OlaHSd mice. Mice were sacrificed by cervical dislo-cation and the tissues extracted. 3.1.2 Cell culture (II, III)

Cell lines used in this work were HEK293 cells, HeLa cells, Flp-In T-REx 293 cells and murine C2C12 myoblasts. As a control, mouse embryonic fibroblasts derived from a Top2 -/- mouse model were also employed (Zhang et al. 2013). All cells were cultured in Dulbecco’s modified Eagle medium supplemented with 10% FBS at 37 °C and 8.5% CO2. T-REx Flp-In cell lines were grown in this medium containing also 150 µg/mL hygromycin and 15 µg/mL blasticidin. The differentiation of confluent C2C12 my-oblasts to myotubes was induced by a growth medium switch upon confluency to DMEM containing 2% horse serum, and the cells were harvested after 6 days in dif-ferentiation medium unless mentioned otherwise.

For T-REx Flp-In cells 5 ng/mL doxycycline was added for 24 hours, if not indi-cated otherwise, to induce protein expression. 3.1.3 Cloning of expression constructs (II, III) Human Top2β with C-terminal myc-tag was equipped with the mitochondrial tar-geting sequence of cytochrome C (kind gift of Dr. Stefan Sobek) and cloned into the pcDNA5 FRT/TO vector.

The human Top3α coding sequence (accession number NM_004618) including the mitochondrial start AUG, the targeting sequence and with a flag tag fused to the C-terminal end was cloned into pcDNA5 FRT/TO. The nuclear start AUG was mutated to GCG to create the MTS-Top3α-flag construct. To destroy the nuclear localization signal in the C-terminal end of Top3α and enhance the mitochondrial localization of the resulting protein, the bases at position 3 207-3 211 were mutated from AAAAG to CGACG, resulting in the mutation K965A R966A and the MTS-Top3α-NLSmut-flag construct (hereafter mtTop3α). The catalytically inactive version MTS-Top3α-NLSmut-Catmut-flag (hereafter mtTop3α-Y362F) was created by the replacement of the catalytic tyrosine at position 362 by phenylalanine (TAT->TTT). As control the empty pcDNA 5 FRT/TO vector was employed.

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For BioID labelling the MTS-Top3α-NLS coding sequence was fused to the N-ter-minus of the BioID2 biotin ligase in the addgene vector #74224 (Kim et al. 2016), and the fusion construct containing the mitochondrially targeted mtTop3α, the BioID2 gene and a C-terminal HA tag recloned into pcDNA5 FRT/TO. As control, a construct containing only the mitochondrial targeting sequence of Top3α directly fused to the BioID2-HA coding sequence was created.

3.1.4 Transient transfections and immunocytochemistry (II, III) For immunochemical detection of the mitochondrially targeted Top2β or various Top3α constructs, HeLa cells grown on coverslips were transiently transfected with the above described pcDNA5 FRT/TO constructs. 4 µg of plasmid DNA was diluted in 140 µL of OptiMEM and 2 µL of Turbofect transfection reagent (Thermo) was added. The reaction was incubated 20 min at room temperature and pipetted drop-wise to the cells. One day after transfection the cells were fixed with 3.3% PFA for 25 min at room temperature and permeabilized in PBS + 0.5% Triton-X, 10% FBS for 15 min. The slides were incubated with primary antibodies in PBS + 0.1% Triton, 10% FBS for 1 h, washed in PBS and incubated with secondary antibodies for 1 h. After washing the cells were mounted and the fluorescent staining analysed with a Zeiss Axiovert /Axiocam fluorescent microscope.

3.1.5 siRNA knock-down (II, III) Cells on a 6-well palate were grown to 50% confluency and the medium was changed one hour prior to transfection with 25 pmol of siRNA (or 12.5 + 12.5 pmol for com-bined transfections). siRNA was diluted with 125 µL of OptiMEM and combined with 4.5 µL of Lipofectamine RNAiMAX also diluted in OptiMEM. The reaction was incubated for 10 min at room temperature and pipetted dropwise to the cells. After 3 days, the siRNA transfection was repeated, and protein and DNA samples were collected after a total of 6 days.

siRNAs used: Negative siRNA control Cat. No. AM4613 Top2α Cat. No. 4390824 ID s14309 Top2β Cat. No. 4390824 ID s108 Top3α (A) ID s14310 Top3α (B) ID s14312

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3.2 MITOCHONDRIAL EXTRACTION (I, II, III) Mitochondria were extracted from mouse tissues and cultured cells using differential centrifugation and sucrose gradient purification. Mouse tissues were rinsed and chopped in cold tissue homogenization buffer (224 mM Mannitol, 75 mM Sucrose, 20 mM Tris pH 7.4, 10 mM EDTA, 1 mg/mL BSA, 1 mM DTT) prior to homogeniza-tion with a Dounce homogenizer. Cultured cells were harvested, incubated for 30 min in growth medium containing 20 ug/ml Cytochalasin B, washed with PBS and resuspended in cell homogenization buffer (224 mM Mannitol, 75 mM Sucrose, 20 mM HEPES pH 7.4, 10 mM EDTA, 1 mg/mL BSA, 1 mM DTT) before homogeniza-tion. Homogenates of mouse tissues and cultured cells were centrifugated twice for 5 min at 800 g and 4 °C to pellet cell debris and nuclei. Mitochondria were pelleted by centrifugation for 10 min at 15 000 g and 4 °C. Mitochondria were further purified by ultracentrifugation for 1 h at 50 000 g and 4 °C, employing a two-step sucrose gradient of 1.5 M and 1 M (0.8 M for brain) sucrose in 10 mM HEPES pH 7.4, 10 mM EDTA. The mitochondrial layer was recovered from the interphase of the gradient. 3.3. DNA 3.3.1 DNA extraction (I, II, III) DNA was extracted either from purified mitochondria or as total DNA from cultured cells. For total DNA extraction, cells were washed with PBS and harvested either by trypsinization or pipetting. The cell pellet was resuspended in DNA lysis buffer (10 mM Tris pH 7.4, 10 mM EDTA, 150 mM NaCl, 0.4% SDS, 100 g/mL Proteinase K) and incubated at 37 °C for 2 hours. In order to remove proteins and lipids, an equal volume of Phenol:Chloroform:Isoamyl alcohol (25:24:1, pH 6.7) was added, the sam-ple was mixed and centrifuged and the upper phase recovered. This phenol:chloro-form extraction was repeated twice, followed by an only-chloroform step to remove phenol traces. The DNA was precipitated using two volumes of ice-cold 100% etha-nol and 0.1 volume of 3 M sodium acetate pH 5.3. The sample was incubated at -20 °C overnight and the DNA pelleted by centrifugation for 30 min at 16 000 g and 4 °C. The pellet was washed with 70% ethanol and air-dried. The DNA pellet was resus-pended in 1x Fastdigest restriction buffer containing either BglI (human mtDNA) or KpnI (mouse) to cut nuclear DNA and incubated overnight at 37 °C. 3.3.2 mtDNA topology (I, II, III) Different topological forms of mtDNA were visualized by agarose gel electrophore-sis and Southern blotting. 1.5 µg of mouse mtDNA, 0.5 µg of mtDNA or 1.5-2 µg of

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total DNA from cultured cells was separated over a 0.4% agarose gel in 1x TBE with-out ethidium bromide. The gel was blotted and the DNA was hybridized with probes corresponding to nts 35-611 of human mtDNA, nts 14 783-15 333 of mouse mtDNA or nts 3 774-45 71 for mouse tissue samples. 3.3.3 7S DNA (II, III) Mitochondrial 7S DNA levels per mtDNA were quantified by Southern blotting. 0.5-2 µg of total DNA were digested with HindIII and heated for 10 min at 65 °C prior to separation on a 0.6% agarose gel. The DNA was blotted onto a membrane and probed against nts 16 177-40 of human mtDNA or nts 15 467-16 011 of mouse mtDNA. Sig-nals were quantified by phosphor storage screens using Molecular Imager FX and QuantityOne software, and the ratio of 7S per mtDNA was calculated. 3.3.4 Determination of mitochondrial DNA copy number (I, II, III)

Mitochondrial DNA copy number per cell was measured from total DNA samples using a Taqman-based quantitative PCR method. Relative levels of mtDNA were de-termined in triplicates by TaqMan-based duplex real-time PCR using 100 ng DNA template, 300 nM nuclear NDUFV1 primers, 100 nM 16S primers and 125 nM of both probes in a final volume of 20 μl AccuTaq readymix (VWR). The reactions were de-natured at 95 °C for 3 min followed by 40 cycles of denaturation (95 °C for 20 s), hy-bridization (52 °C for 20 s) and elongation (72 °C for 20 s) on an AriaMx realtime PCR system (Agilent Technologies). Sequences of primers and probes used for mtDNA copy number determination by quantitative PCR: NDUFV1forward 5′-CTT CCC CAC TGG CCT AA-3′ NDUFV1 reverse 5′-CCA AAA CCC AGT GAT CCA GC-3′ NDUFV1 probe 5′-VIC-GAG CCT TAG GGA AGA AGA GGC AG-MGBNFQ-3′ 16S rRNA forward 5′-TGC CTG CCC AGT GAC TAA AG-3′ 16S rRNA reverse 5′-GAC CCT CGT TTA GCC GTT CA-3′ 16S rRNAprobe 5′-FAM-TGA CCG TGC AAA GGT AGC AT-MGBNFQ-3′ HSmtDNA13456 forward 5′-ACC ATT GGC AGC CTA GCA TT-3′ HSmtDNA13593 reverse 5′-TGT CAG GGA GGT AGC GAT GA-3′ HSmtDNA 13546F probe 5′-FAM-ACA AAC GCC TGA GCC CTA- MGBNFQ-3′

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3.3.5 Two-dimensional neutral/neutral agarose gel electrophoresis (I, II, III)

The detailed processes of mitochondrial DNA replication can be studied using neu-tral/neutral two-dimensional agarose gel electrophoresis (2D-AGE), which allows the separation of replication intermediates by their size and structure (Friedman & Brewer 1995, Holt et al. 2000, Yang et al. 2002, Reyes et al. 2007). When restriction-digested DNA samples are analyzed on 2D-AGE, coupled leading- and lagging-strand DNA synthesis creates typical bubble arcs, containing molecules with a repli-cation bubble indicative of replication initiation in the observed region, and Y arcs consisting of molecules with a replication fork, arising from initiation outside of the studied sequence. The asynchronous replication typical for mtDNA produces addi-tional patterns called slow-moving Y-like arcs (smY). These are longer molecules arising from incomplete digestion, as most restriction enzymes are unable to cut sin-gle-stranded DNA or RNA:DNA hybrids. Also molecules containing two approach-ing replication forks or Holliday crossover junctions can be detected on 2D-AGE, thus providing information about the resolution of fully replicated mtDNA and DNA recombination processes. For 2D-AGE 5 µg of mtDNA or 10 µg of total DNA was used. The DNA was digested with ClaI, HincII, MluI or Bcll FastDigest restriction enzymes (Thermo Sci-entific) for 5-6 hours in 37 °C and extracted with 1 Vol phenol:chloroform before sep-aration on a first dimension gel of 0.4% agarose in 1xTBE buffer for 16 h at 20-30 V. When the DNA fragments of interest had migrated for ca. 10 cm, the sample lanes were cut out from the gel and separated over a second dimension consisting of a 0.95% agarose gel with 0.5 µg/mL ethidium bromide for 16-18 h at 110V in 1xTBE with 0.5 µg/mL EtBr at 4 °C with buffer circulation. In this step, DNA fragments were separated by their size and shape. The gel was run until the 1n DNA fragments had migrated to the edge of the gel, the gels were depurinated for 2x 15 min in 0.2 M HCl, denatured for 2x 20 min in 0.5 M NaOH, 1.5 M NaCl and capillary-blotted onto Hy-bond XL-membrane. The DNA was hybridized with probes as indicated in table 1.

For single cut 2D gels the DNA was digested with the indicated single cutting restriction enzyme as previously described, but the first dimension was run on a 0.28% agarose gel and the second dimension on a 0.58% agarose gel for 2 days at 50 V in room temperature.

Table 1. Restriction enzymes and probes used for 2D-AGE. NCR and ND4 are from human mtDNA and Cytb from mouse mtDNA. Enzyme mtDNA fragment Probe

HincII 13 637-1 007 35-611 (NCR)

BclI 11 922-8 592 11 164-11 640 (ND4)

MluI Full-length 14 783-15333 (Cytb)

ClaI 12 086-635 14 783-15 333 (Cytb)

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3.3.6 DNA damage (I, III) TaqMan-based long-range real-time PCR was used to determine the levels of poly-merase-inhibiting mtDNA damage. The 20 µL PCR reactions contained 100 ng of to-tal DNA template, 500 nM mtDNA primers, 125 nM 16S probe and 1x AccuTaq re-dymix (VWR). The DNA was denatured at 94 °C for 5 min followed by 40 cycles of denaturation at 94 °C for 20 s, annealing at 64 °C for 20 s and elongation at 72 °C for 7 min using the AriaMax Real-Time PCR system (Agilent Technologies). The amount of template was quantified from the 16S probe signal and relative damage was cal-culated using the LORD-Q method (Lehle et al. 2014). Primers and probes used for the PCR:

Mouse mtDNA1978 Forward 5′-TCC GAG CAT CTT ATC CAC GC-3′ Mouse mtDNA8496 Reverse 5′-ACC ATT TCT AGG ACA ATG GGC A-3′

Mouse mtDNA3567 Forward 5′-TTC GAG CAT CTT ATC CAC GC-3′ Mouse mtDNA8496 Reverse 5′-ACC ATT TCT AGG ACA ATG GGC A-3′ 16s-rRNA probe 5′-FAM-TGA CCG TGC AAA GGT AGC AT-MGBNFQ-3′

Human mtDNA10131 Forward 5′-ACC ACA ACT CAA CGG CTA CA -3′ Human mtDNA14841 Reverse 5′-TTT CAT CAT GCG GAG ATG TTG GAT GG-3′ 3.3.7 In vitro Topoisomerase assay (II, III) The in vitro activity of human Top2 on mtDNA was tested using Top2α (Topogen, TG2000H) and Top2β (Inspiralis, HTB205) and mtDNA isolated from HEK293 cells. 0.25-1 U of the enzymes was diluted in reaction buffer (50mM Tris-HCl pH 7.5, 125 mM NaCl, 10 mM MgCl2, 5 mM DTT, 100 µg/mL albumin and 1µM ATP) without or with 50 ng/mL EtBr, 80 µg/mL ciprofloxacin or 3.4 µM doxorubicin and incubated at room temperature for 10 min. 200 ng mtDNA in 10µL of reaction buffer was added and samples were incubated 30 min at 37 °C. The reactions were stopped by addition of 5 µL DNA loading dye (10 mM Tris-HCl pH 7.6, 0.03% bromphenol blue, 0.03% xylene cyanol FF, 60% glycerol, 60 mM EDTA). The samples were separated over a 0.4% agarose gel and blotted and probed as described above.

The identity of the various topological forms of mtDNA was investigated by the treatment with T7 endonuclease (Thermo), E. coli Topo I (NEB) and E.coli TopoIV (Inspiralis, Cat. No. T4001). 850 ng total cellular DNA were incubated for 30 min at 37 °C in 20 l 1x Cutsmart buffer with 5U Topo1, or in 20 ul TopoIV buffer with 10 U T7 endo, 10 U TopoIV or both. The reaction was stopped by addition of 5 l DNA loading dye, separated over a topology gel, blotted and probed as described above.

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3.3.8 Nanopore sequencing (III) The coordinates of mtDNA break points were determined using mtDNA isolated from HEK293 T-REx cells expressing mtTop3α or mtTop3α-Y362F as well as empty vector cells. 5 µg mtDNA were treated with 20 U RNase I for 6h at 37 °C to remove RNA and linearized with NheI, which cuts human mtDNA once at position 4 581. The linearized mtDNA sequencing libraries were prepared using the PCR-free, Liga-tion Sequencing Kit (SQK-LSK109, Oxford Nanopore Technologies, UK) along with the Native Barcoding Expansion (EXP-NBD104, Oxford Nanopore Technologies, UK) following the manufacturers protocol. The barcoded libraries were then se-quenced on a MinION Flongle flowcell. Demultiplexing, basecalling and removal of barcodes and adapters was done offline using Guppy version 4.2.2 (Oxford Na-nopore Technologies, 2020). The sequencing reads were mapped to the human mtDNA sequence (NC_012920) with first base pair set as 4 581 (Nhel restriction cut site) using minimap2 with the setting -x map-ont (Li 2018) and alignments were out-put in PAF format. 3.4. PROTEIN 3.4.1 Protein extraction and Western blots (I, II, III) Protein was extracted from mitochondria or cultured cells using TotEx buffer (20 mM HEPES pH 7.9, 400 mM NaCl, 20% glycerol, 1% IGEPAL, 1 mM MgCl2, 0.5 mM EDTA, 0.1 mM EGTA, 10 mM -Glycerophosphate, 10 mM NaF, 9 mM DTT, 1xcOm-plete EDTA-free protease inhibitor cocktail (Roche)). The mitochondria or cell pellet was suspended in ca. 4 volumes of buffer, vortexed and incubated on ice for 20 min. To aid solubilization, samples were snap-frozen in -80 °C and after thawing incu-bated for 10 min on ice. DNA and cell debris ware pelleted by centrifugation for 10 min at 17 000 g and 4°C. Protein concentrations were determined by Bradford as-say.

40-150 µg of protein were separated on 8, 12 or 15% Tris-Glycine SDS-PAGE gels or 4-12% Tris-Tricine SDS-PAGE gels. Proteins were transferred to nitrocellulose membrane (ProTran, Life Technologies), and the membrane was blocked for 1 h at room temperature with 3% skim milk in TBST (50 mM TRIS pH 7.4, 150 mM NaCl, 0.1% Tween-20). The membrane was incubated over night at 4 °C with primary anti-bodies diluted in 3% BSA in TBST (Table 2). After 3x 5 min washing with TBST, the blots were incubated for 1 h at room temperature with HRP-coupled secondary an-tibodies in TBST. Proteins were visualized by chemiluminescence detection with film or Biorad Chemidoc.

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Table 2. Antibodies used for Western blots.

Antibody Dilution Source Identifier

Rabbit-anti-mtSSB 1:1 000 Sigma HPA002866

Rabbit-anti-ATAD3 1:25 000 kind gift of Dr. Cooper Rabbit-anti-TFAM 1:1 000 Aviva ARP31400

Rabbit-anti-TFB2M 1:1 500 Abcam ab66014

Rabbit-anti-PolG1 1:500/1 000 Abcam ab128899

Rabbit-anti-POLRMT 1:500 Abcam ab32988

Mouse-anti-VDAC1 1:2 000 Sigma SAB5201374

Rabbit-anti-TWNK(PEO1) 1:2 000 antibodies online ABIN2405996

Rabbit-anti-TWNK(PEO1) 1:1 000 Elabscience EAP1298

Rabbit-anti-COXI 1:2 000 Abcam ab14705

Rabbit-anti-Top2α 1:1 000 Abgent AP9248b

Rabbit-anti-Top2β 1:1 000 Abcam ab15565

Rabbit-anti- Top2β 1:1 000 Genetex GTX102640

Rabbit-anti-Top3α 1:4 000 ProteinTech 14525-1-AP

Rabbit-anti-Top1mt 1:1 000 Bioss bs-1211R

Rabbit-anti TFAM 1:2 000 Abgent AN1221

Mouse-anti-HSP60 1:20 000 Antibodies-online ABIN361784

Mouse-anti-vinculin 1:10 000 Sigma V9264

Rabbit-anti-Histone 2.1 1:2 000 Abcam ab181973

Rabbit anti-Tomm20 1:4 000/5 000 Sigma HPA011562

Goat-anti-rabbit IgG HRP 1:10 000 Antibodies-online ABIN101744

Goat-anti-mouse IgG HRP 1:15 000 Life Technologies A16104

AlexaFluor594Goat-anti.rabbit IgG 1:1 000 Invitrogen A11037

AlexaFluor488Goat-anti-mouse IgG 1:1 000 Invitrogen A11029

3.4.2 Top3α-BioID purification (III) BioID pull-down was done as previously described by Hensen et al. (2019) and Roux et al. (2018) with minor adjustments. mtTop3α-BioID2-HA expressing cells and control cells were induced with 3 ng/mL of Doxycycline. 24 hours later the growth medium was changed to medium containing 50 µM biotin (Sigma-Aldrich, B4501) and the cells were grown for another 24 h. Mitochondria were extracted as described earlier in section 3.1. The mitochondria pellet was resuspended in 1 mL of lysis buffer (50 mM Tris pH 7.5, 500 mM NaCl, 0.4 % SDS, 1 mM DTT) and 100 µL of 20% Triton-X100 was added. The samples were sonicated on ice (15 pulses, 50% amplitude, 0.5 cycle), and 1 Vol of 50 mM Tris-HCl pH 7.5 was added. The samples were

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centrifugated at 16 000 g for 10 min at 4 °C, 100 µL of Dynabeads MyOne Streptavidin C1 (Invitrogen, 654001) beads were added to the supernatant and incubated over night at 4 °C on rotation. The beads were washed four times for 5 min with 1 mL wash buffer (50 mM Tris-HCl pH 7.4, 8 M Urea). Proteins were eluted from the beads with 60 µL of 1x SDS sample buffer (50 mM Tris pH 6.8, 2% SDS, 6% glycerol, 0.1 M DTT, 0.004% Bromphenol blue) and boiling for 5 min at 98 °C. 20 µL of input, flowthrough, wash samples and eluate were separated over a Bolt 4-12% Bis-Tris Plus SDS gel (Invitrogen) and analysed by Western blot.

3.4.3 Flotation gradient (II) The co-localization of mitochondrial topoisomerases with mitochondrial nucleoids and known mtDNA interactors was analyzed by mitochondrial sub-fractionation and flotation gradient analysis as described in Rajala et al. 2014. Mitochondria iso-lated from HEK293 cells were treated with digitonin to separate mitochondrial mem-brane-associated proteins in the pellet fraction and soluble mitochondrial proteins in the supernatant fraction after centrifugations. Both fractions were separated over bot-tom-up flotation iodixanol gradients, and their protein and mtDNA content were a-nalyzed by Western blot and Southern dot blot using nts 14 837- 15 367 as probe. 3.5 RNA

3.5.1 RNA extraction and Northern blot (I, II, III) Total RNA from tissues or cultured cells was extracted using TriReagent (Sigma) and analyzed by Northern blot. 4 µg RNA were separated over a 1,2% agarose MOPS/Formaldehyde gel, equilibrated for 20 min in 20x SSC and capillary-blotted onto Hybond-XL membrane. Mitochondrial RNA levels in mouse tissues were quan-tified using the following probes and normalized against 28S and 18S probes. Mouse tissues Cultured human cells ND1 nts 3 774-4 373 ND2 nts 1 170-5 511 Cytb nts 14 783-15 333 ND5 nts 13 640-13 777 ATP6 nts 8 165-8 494 18S rRNA nts 850-1 347 bp, accession ND5 nts 13 288-13 840 number NR_0032862 28S rRNA nts 4 165-4 703, accession number NR_145822.1

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4 RESULTS AND DISCUSSION

4.1 TISSUE SPECIFICITY OF MITOCHONDRIAL MAINTENANCE Different cell types in different tissues require different levels of mitochondrial activ-ity for their proper function. Tissues with high energy demand, like muscles and heart, rely heavily on mitochondrial ATP production (Benard et al. 2006, Férnandez-Vizarra et al. 2011). However, studying the tissue-specific aspects of mitochondrial OXPHOS activity, mtDNA copy number, transcription and their correlations has been challenging, partly due to the difficulty of normalization. Still, this information is desirable, as diseases resulting from mitochondrial dysfunction often show strong tissue-specific symptoms, and some cell types are more affected than others. It has been proposed that either different mutation load or differences in energy demand and mitochondrial function would be the main reason for the tissue specificity of mitochondrial diseases, but the cause of this phenomenon is not yet fully understood (Lightowlers et al. 2015).

This thesis sought to elucidate the cell type-specific variations of mitochondrial DNA maintenance in a range of mouse tissues, to see whether mtDNA levels and integrity correlate with mtDNA replication mode, abundance of replication factors or the level of mitochondrial gene expression.

4.1.1 Mitochondrial DNA copy number and topology (Paper I) Mitochondrial function relies on the coordinated expression of both nuclear and mi-tochondrially encoded proteins. Initially, the expression of mitochondrial proteins has been thought to be regulated by mtDNA copy number, but newer studies suggest that expression might rather be controlled by protein factors, like TFB2M (Litonin et al. 2010, Hillen et al. 2017).

We found mitochondrial DNA copy number, determined as the ratio of mtDNA to nuclear DNA, to be highest in brown fat. Our results were in line with previous findings by Férnandez-Vizarra et al. (2011). This analysis of mtDNA copy number per cell does not consider that cell size might be very different between different cell types. As a central function of mitochondria is the production of ATP, it is advisable to also compare the levels of mtDNA per ATP-consuming tissue mass. Interestingly, no significant difference of mtDNA levels per total protein content was found in the investigated tissues, suggesting that the same amount of mtDNA is able to support the energy demands of a tissue volume, regardless of the cell physiology (Paper I figure 2B).

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While the copy number of mtDNA was basically identical in the studied tissues,

clear differences in mitochondrial DNA topology were observed (Paper I figure 2C). All tissues had high levels of relaxed circle molecules, while in cultured cells super-coiled mtDNA dominates. In heart, muscle, kidney, and brown fat also linear mole-cules were abundant. This might be an artefact produced during the extraction of mtDNA from these rigid tissues or an indication of extensive mtDNA damage in these cell types.

4.1.2 Tissue specificity in replication and correlation with mtDNA damage and replisome components (Paper I) After the identification of asynchronous and strand coupled replication mechanisms in mammalian mitochondria, these mechanisms were suggested to co-exist but to be utilized at varying degrees in different cell types and situations. Cultured cells mainly use the asynchronous mechanism under normal conditions, but shift to syn-chronous replication after exposure to DNA-depleting agents (Holt et al. 2000, Tor-regrosa et al. 2015). Also the majority of studies investigating mtDNA replication in solid tissues found that the asynchronous mechanism to dominate, but most of these experiments were conducted in liver and placenta (Holt et al. 2000). Consequently, the abundance of the different replication modes utilized and the reasons to use one or the other as well as the regulatory mechanisms are still mainly unknown. We aimed to find a correlation between physiological conditions and choice of replica-tion mechanism.

The mtDNA replication mode of six different mouse tissues was analyzed using 2D-AGE (Figure 5A-F). The replication intermediates observed in liver and kidney were typical for strand-asynchronous replication (RITOLS), while heart, skeletal muscle, brain, and brown fat contained replication intermediates representative of conventional strand-coupled replication (COSCOFA). In addition, these tissues also contained abundant four-way junctions, which were absent from liver and kidney.

Earlier studies described that mitochondrial DNA topology in adult human heart as well as mammalian brain is significantly different from other tissues, exhibiting high levels of complex catenated and possible recombining molecules (Pohjoismäki et al. 2009, Goffart et al. 2009). This complex organization was interpreted to arise from recombination, serving as a protective mechanism against oxidative mtDNA damage in energy-demanding tissues (Pohjoismäki et al. 2013b).

In order to study this possibility, the level of mtDNA damage was quantified us-ing a PCR-based method. Interestingly, the highest rate of mtDNA damage was ob-served in brown fat and brain, followed by heart and muscle (Figure 5G). mtDNA damage was thus most abundant in tissues utilizing strand-coupled replication and having abundant four-way junctions. These findings suggest that the mechanism

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employed to replicate mtDNA correlates with mtDNA damage, the strand-coupled synchronous replication being predominant in tissues with higher DNA damage. Both strand-coupled replication and recombination could thus be adaptations to ge-notoxic stress, matching earlier findings that cultured cells recovering from mtDNA depletion or stress conditions show both strand-coupled replication and signs of re-combination (Torregrosa et al. 2015).

To elucidate whether the differences in replication mode are caused by changes in the composition of the replicative machinery, the expression levels of mitochon-drial replisome components were analyzed in the same mouse tissues. Interestingly, protein levels varied considerably between tissues (Paper I, figure 3H). mtSSB pro-tein levels were highest in liver and kidney, tissues employing the asynchronous rep-lication mode, in line with the important role of this protein during asynchronous replication. During asynchronous replication, the lagging strand has been suggested to be covered with either mtSSB or RNA. The RNA needed for replication is thought to originate from transcription, and transcript availability has indeed been demon-strated to be important for this mechanism (Reyes et al. 2013, Cluett et al. 2008). Cluett et al. (2008) found cultured cells to change their replication mechanism after shortage of mitochondrial transcripts. In our analysis, transcript levels did not correlate with the RITOLS mechanism (Paper I, figure 4A), conflicting with the earlier observation that transcript availability dictates the balance between replication mechanisms (Cluett et al. 2008). However, the difference in transcription levels in the analyzed tissues was very small, and it is unlikely to have any limiting effect on replication.

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Figure 5. Mitochondrial DNA replication intermediates in different mouse tissues correlate with mtDNA damage. (C) Replication intermediates of mtDNA in different mouse tissues digested with ClaI and probed against the cytochrome c region (14 783-15 333 nt). (A, D) Liver and kidney replicate their mtDNA using the strand-asynchronous replication mode (RITOLS), showing the characteristic blunt bubble arc and slow moving smY intermediates. (B, E) In heart and brain tissue, mtDNA is replicated via strand-coupled replication (COSCOFA), recognizable from a sharply defined bubble arc and near absence of smY. X-spikes, representing recombi-nation intermediates, are present in both tissues. (F) Muscle and brown adipose tissue employ a mixture of asynchronous and strand-coupled mtDNA replication. In both tissues, also X- spikes are visible. (G) Quantification of mtDNA damage by real-time long-range PCR revealed that kidney and liver, tissues using RITOLS replication, had the lowest levels of damage.

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4.2 MITOCHONDRIAL TOPOISOMERASES Topoisomerases are important factors in DNA replication, transcription, and repair, but although also mtDNA maintenance requires topoisomerases in multiple aspects, the exact identity and division of labour of these proteins in mitochondria are not fully known. Until now, four topoisomerases have been identified from vertebrate mitochondria, Top1mt, Top3α, Top2α and Top2β.

The levels of these mitochondrial topoisomerases showed clear differences be-tween the analyzed cell types and mouse tissues, with Top2β and Top3α being highly expressed in mouse brain (Paper II). This expression profile might be connected to the replication mechanism employed in these tissues, as brain has the highest level of X-shaped replication intermediates, indicative of mtDNA recombination (Sánchez & Antequera 2018, Dandjinou et al. 2006). The mtDNA of brain and heart has previ-ously been shown to contain multiple junctions and topologically challenging forms (Pohjoismäki et al. 2009), which would highlight the importance of topoisomerases in the maintenance of mtDNA in these tissues. To better address the function of topoisomerases in mtDNA maintenance, both isoforms of Top2 as well as Top3α were studied in detail. 4.2.1 Topoisomerase localization in mitochondria and tissue and proliferation status specificity (Paper II) Due to the conflicting data about the presence of topoisomerase 2 in mitochondria, we strove to determine the presence of both known isoforms in mitochondria. We did this using purified mitochondria from various cultured human and mouse cell lines as well as mouse tissues. Top2β was present in all studied cell lines and also expressed in mouse tissues, with expression levels being especially high in brain (Fig-ure 6). Top2α instead was present exclusively in proliferating cells, and absent in differentiated cultured myotubes and mouse tissues. This expression profile is in agreement with the known gene expression profiles of the nuclear enzyme isoforms. Furthermore, the observed sizes of both isoforms of mitochondrial topoisomerase 2 were similar to the nuclear versions. Similar to Zhang et al. (2015), we were unable to see a shorter version of topoisomerase 2β, described previously by Low et al. (2003), suggesting that this truncated version could be an experimental artefact, since Top2β seems to be highly sensitive to degradation during the mitochondrial purifi-cation and protein extraction.

The mitochondrial localization of Top1mt and Top3α is well established by now, and in the present study they were detected in all studied cell lines.

Interestingly, all of the studied mitochondrial topoisomerases were found to have a tissue-specific expression profile (Figure 6B). The expression levels of Top2β and Top1mt correlated, being especially high in all studied brain parts, while Top3α was present mainly in cortex and midbrain, but non-detectable in cerebellum (Figure 6B).

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Figure 6. The expression of mitochondrial topoisomerases in cultured cells and mouse tissues. (A) Expression of mitochondrial topoisomerases was detected in all proliferating cell lines, Top2α was absent from differentiated C2C12 myotubes. ATAD3 used as a loading control. (B) In mouse tissues Top2β, Top1mt and Top3α were mainly expressed in brain, Top3α was mainly present in cortex and midbrain, while Top1mt and Top2β both were equally expressed in all studied brain parts. TFAM indicates mtDNA content and TOM20 serves as a loading control. 4.2.2 Topoisomerase 2α, 2β and topoisomerase 3α functions in mitochondria (Paper II, III and IV) The function of the only mitochondria-specific topoisomerase Top1mt has been in-tensively studied throughout the last decade. In contrast, the mitochondrial functions of other topoisomerases are still unknown, mainly due to the difficulty of studying proteins with dual localization in nucleus and mitochondria. To address the function of Top2 and Top3 in mitochondria, first the association with replicating nucleoids was studied by flotation gradient, then siRNA knock-down of these proteins and in-hibition of topoisomerase 2 function by ciprofloxacin were used. Top3α function was also further studied using overexpressing cell lines. Changes in mtDNA copy num-ber, topology, levels of 7S DNA, transcription and replication were evaluated. 4.2.2.1 Topoisomerase 2 (Paper II) Replicating mitochondrial DNA nucleoids are tightly associated with the inner mem-brane, while the majority of mtDNA nucleoids are freely located in the matrix (Rajala et al. 2014). Thus, replicating and non-replicating mtDNA-protein complexes can be separated and fractioned by flotation gradient. Top2α and Top2β are present in the membrane-associated fractions also containing the replicative helicase TWNK and other members of the mitochondrial replisome, indicating a function in mtDNA

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replication (Paper II, figure 1C). Top1mt instead was co-localizing mainly with POLRMT, supporting the earlier suggested role in transcription (Sobek et al. 2013). The functions of mitochondrial Top2 were further studied using siRNA knock-down of the two isoforms Top2α and Top2β. The loss of Top2α did not have any effect on mtDNA topology or mtDNA copy number, suggesting that it is not im-portant for mitochondrial DNA maintenance (Figure 7). This fits to the fact that mtDNA in post-mitotic cells is replicated without Top2α being expressed. Top2β knock-down instead caused the strong accumulation of supercoiled mtDNA and a clear reduction in mtDNA copy number (Paper II, figure 2B). A similar, but more dramatic effect was seen when the type II topoisomerase inhibitor ciprofloxacin was used, and the supercoiled molecules were shown to be positively twisted (Figure 7).

Figure 7. Topological changes in HeLa cells induced by Top2 knock-down or ciprofloxacin inhibition. (A) A 3 day knock-down of Top2β leads to accumulation of supercoiled mtDNA, while loss of Top2α alone dose not have any effect on mtDNA topology. The reduction of Top2α and Top2β protein levels after knock-down was confirmed by Western blot, using vinculin as loading control. (B) A 24 hour-treatment with 80 µg/mL ciprofloxacin caused a similar but more extreme accumulation of supercoiled molecules. The observed changes in mtDNA topology were connected to impairment of mi-tochondrial DNA replication. MtDNA copy number decreased after ciprofloxacin treatment, and the loss of 7S DNA, a marker of mtDNA replication initiation, indi-cated a nearly complete stop of mtDNA replication (Figure 3C). This was confirmed through the analysis of replication intermediates by 2D-AGE. As seen in cultured cells amplifying mtDNA after depletion stress, also ciprofloxacin shifted the replica-tion mechanism from RITOLS replication towards conventional replication in a con-centration-dependent manner, while simultaneously causing replication stalling (Figure 8A). In differentiated C2C12 cells the impairment of replication was even

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more dramatic, and ciprofloxacin induced the almost complete loss of replication in-itiation (Figure 8B, C). Taken together the results presented here suggest that Top2β has an important function controlling mitochondrial DNA topology and thus replication initiation. As replication initiation requires a specific topological environment, accumulation of supercoils can prevent replication initiation, which in turn leads to a decrease in mtDNA copy number over time.

Figure 8. Ciprofloxacin induces replication stalling and impairs replication initiation. (A) Replication intermediates of HEK293 cells treated with 80 µg/mL ciprofloxacin for 2 and 20 h. mtDNA was digested with HincII, separated by two-dimensional agarose gel electrophoresis and probed for the fragment containing the non-coding region of mtDNA. The level of RITOLS replication intermediates decreased upon ciprofloxacin treatment and after 20 h intermediates represented nearly exclusively the strand-coupled replication mode. (B) Replication intermediates of differenciated C2C12 mouse cells after 24 h treatment with 80 µg/mL ciprofloxacin digested with ClaI and probed for the non-coding region. Ciprofloxacin led to near-complete loss of replicating molecules. (C) Identical samples from ciprofloxacin-treated C2C12 cells, digested with the single-cutting enzyme MluI allowing the analysis of the whole mtDNA molecule. Ciprofloxacing inhibited the replication severely in all regions of the mtDNA.

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4.2.2.2 Topoisomerase 3α (Paper II, III) Like Top2α and Top2β, Top3α was mainly associated with replicating nucleoids in the gradient fractionation, but a proportion was present in other fractions not con-taining mtDNA, suggesting this protein to be less frequently bound to mtDNA (Pa-per II, figure 1C).

The function of Top3α was studied using siRNA knock-down, which led to a pro-nounced decrease in mtDNA copy number and 7S DNA and the accumulation of high molecular weight forms of mtDNA that were immobile on regular topology gels, as described previously (Nicholls et al. 2018). The loss of 7S DNA could indicate reduced initiation of replication, which in turn leads to a decrease in mtDNA copy number. Nicholls et al. (2018) proposed Top3α to have an important function in mtDNA segregation at the end of the replication, accumulation of high molecular forms in the absence of Top3α supports this theory. As Top3α is a type I topoisomer-ase unable to separate fully catenated molecules, this would require the replication end products to be hemicatenanes, or segregation to happen before the daughter molecules are fully double-stranded.

Figure 9. Top3α knock-down effects of mtDNA replication. HeLa cells with reduced Top3α protein levels show decreased levels of asynchronous replication and accumulation of fully double-stranded replication intermediates, indicative of replication stalling. This effect is observed not only in the non-coding region harbouring replication initiation and termination (HincII digest), but also in the area of the ND4 gene (BclI digest), suggesting that replication fork progression is impaired throughout the replication process.

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The data presented in this thesis also supports a possible role of Top3α in replica-tion progression. SiRNA knock-down of Top3α induced replication stalling all over the mitochondrial genome and not only close to the termination region, suggesting that the enzyme is required for the progression of the replication fork (Figure 9). As Top3α is a type IA topoisomerase, incapable of removing the positive supercoils formed ahead of the replication machinery, it could relax the negative supercoils ac-cumulating after the replication fork.

To elucidate whether Top3α interacts with the replication machinery, we em-ployed a BioID approach, where the protein of interest is fused to the BioID2 biotin ligase. Proteins in close proximity to the fusion proteins get biotinylated and can then be purified and identified (Roux et al. 2018). The BioID experiments with Top3α sug-gest that the topoisomerase works in close proximity of TWNK, the replicative hel-icase, which supports the hypothesis of a replicative role for Top3α (Figure 10). Also in Escherichia coli, Topoisomerase IIIα has been demonstrated to act at the replication fork together with the helicase to remove precatenanes (Lee et al. 2019). A collabora-tion with TWNK could also suggest a role in the recombination of mitochondrial DNA, as TWNK has been suggested to participate in strand exchange and recombi-nation of mtDNA (Sen et al. 2016). In the nucleus, Top3α plays an important role during recombination as a part of the BTRR complex, and it is feasible that in mito-chondria it has a similar role.

Figure 10. Analysis of Top3α interaction with mtDNA nucleoid proteins by BioID2 labeling and affinity purification. Biotinylated proteins from HEK293 T-REx mtTop3α BioID2 and control mtBioID cells were purified using streptavidin affinity beads and analyzed by Western blot. The mitochondrial helicase TWNK and polymerase γ were both detected in the elution of mtTop3α-BioID-expressing cells, but not in cells expressing only a mitochondrially targeted BioID2 pro-tein. In contrast, MRE11 and Top2β were not purified, indicating the BioID2 labeling to be specific. MGME1 and TFAM instead were detected also in the mtBioID suggesting that the purification of these proteins was unspecific. I=input, FT=flow through, W=wash, E=elution.

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The function of Top3α for mitochondrial DNA maintenance was further studied using inducible cell lines overexpressing either a mitochondrially targeted Top3α or a catalytically inactive version of the protein. Both transgenes carried the mitochon-drial targeting sequence and mutations in the nuclear start codon as well as in the C-terminal nuclear targeting sequence, excluding the possibility of nuclear accumula-tion of the proteins. Mitochondrial DNA copy number decreased in both versions of the Top3α expressing cell lines, however even high expression levels of the catalyti-cally inactive Top3α mutant had only little effect on mtDNA topology (Figure 11). In contrast. cells overexpressing mitochondrially targeted Top3α exhibited dramatic differences in mtDNA topology; the abundance of supercoiled molecules was de-creased and the amount of relaxed mtDNA increased, indicating that Top3α relaxed negative supercoiled molecules. In addition, high molecular weight forms of mtDNA decreased upon mtTop3α overexpression, supporting a decatenating function. Inter-estingly, linearized mtDNA molecules were present at much higher level than in con-trol cells. The linear mtDNA molecules were also observed on agarose gels with di-gested mtDNA, and the breakpoint was located in the non-coding region of mtDNA (Figure 11, Paper III, figure 3C).

Mapping of linear mtDNA ends by nanopore sequencing did not identify any precise breakpoint, indicating that either the breaking occurs randomly in the non-coding region or that the end of the linear mtDNA molecules is not ligatable even after an end repair step and thus escapes the sequencing. As Top3α should induce single-strand cuts in the vicinity of OH during decatenation, any pre-existing nicks in the non-coding region might lead to the linearization of the mtDNA molecule. Simi-lar breaks and linearized molecules are present in control cells, although at a lower level, indicating that this occurs also in normal situations. The mutation of the cata-lytic center of Top3α creates an enzyme that is still able to bind to DNA, but without the cleavage function, and thus no strand break is created.

The analysis of mtDNA by one-dimensional gel electrophoresis revealed an inter-esting accumulation of mtDNA fragments of ca. 1-2 kb of size in cells overexpressing functional mtTop3α, but no change in 7S to mtDNA ratios compared to control cells (Figure 11B). The observed DNA species could possibly originate from replication intermediates being broken and released during the early stage of replication. Cells overexpressing catalytically inactive Top3α instead showed a decrease in 7S DNA, suggesting that although the topology of mtDNA is unchanged, mtDNA mainte-nance is still affected.

The presented data shows, that additional to its role as a decatenase at the end of replication, Top3α also has a role in replication progression, interacting with the mi-tochondrial replisome.

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Figure 11. Effects of Top3α overexpression in HEK T-REx cells. (A) Overexpression of mito-chondrially targeted Top3α (mtTop3α) caused changes in mtDNA topology: the amount of supercoiled molecules was decreased, and the amounts of relaxed circles and linear mole-cules were increased. Furthermore, the level of higher molecular weight forms was reduced. The catalytically inactive version of Top3α (mtTop3α-Y362F) had only little effect on mtDNA topology, however, the relative amount of high molecular weight forms increased, supporting the decatenation function of Top3α. (B) When mtDNA was digested with HindIII, broken mtDNA molecules together with mtDNA signal in the range of 0.7-2 kb were detected in mtTop3α cells, possibly originating from broken strands of arrested replication. 4.2.3 Division of labor of mitochondrial topoisomerases (Paper II, III, IV) As presented in the previous chapters, mitochondrial topoisomerases, like topoiso-merases in the nucleus, have specific roles in DNA maintenance based on their mech-anism of action. Surprisingly, many functions of the mitochondrial topoisomerases differ from their nuclear counterparts. Since the mitochondrial genome is small and circular and the topological problems arising vary from nuclear ones, mitochondrial topoisomerases need to adapt to this different and challenging topological environ-ment.

As mentioned earlier, knock-down or overexpression of different Top3α versions lead to impairment of mtDNA replication process. As Top3α interacts with replica-tive helicase and polymerase, it most likely functions in replication progression by removing the accumulation of negative supercoils behind the replication fork. It was also shown to function in segregation of chromosomes at the end of mtDNA replica-tion, and the data presented here supports this hypothesis. Positive supercoils formed ahead of replication machinery could instead be removed by Top2β or Top1mt. The lack of Top1mt causes the accumulation of negative supercoils in mtDNA (Zhang et al. 2014), so this enzyme is therefore an unlikely candidate for the relaxation of positive supercoils during replication. Inhibition of topoisomerase 2β instead leads to an increase of positively supercoiled molecules, as presented earlier

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in this thesis, suggesting that it might have a function during the replication process. Top1mt instead has been reported to negatively regulate mtDNA transcription and to interact with POLRMT, suggesting that it might regulate supercoiling during tran-scription (Sobek et al. 2013), although also another topoisomerase is likely involved. In the present study, Top3α knock-down and overexpression were seen to modulate transcription, so this protein is a potential candidate to function during mtDNA tran-scription (Paper III, figure 5).

Although topoisomerases do have specific roles in mitochondrial DNA mainte-nance, they appear to be somewhat redundant. In Top1mt -/- MEFs, Top2β levels are significantly up-regulated, while Top2β -/- MEFs do not exhibit any mtDNA topolog-ical changes (Sobek et al. 2013), suggesting that Top1mt and Top2β can substitute each other to some extent. Ciprofloxacin treatment leads to more dramatic conse-quences in differentiated cells, indicating that in proliferating cells Top2α might be able to contribute to the maintenance. Based on the data presented here, Top2β acts in balance with Top1mt under normal conditions, with Top2β relaxing positive supercoils and Top1mt negative supercoils, as demonstrated before by Zhang et al. (2014). The identified and potential functions and properties of known mitochondrial topoisomerases are presented in table 3.

Table 3. Mitochondrial topoisomerases. Modified from paper IV.

Top1mt Top2α Top2β Top3α Classification Type IB Type IIA Type IIA Type IA

Shown mito-chondrial func-tion

Regulation of transcription [1,2] and replication [3]

Regulation of replication [6]

Decatenation of hemicatenates [7] and replication [8]

Potential mito-chondrial func-tions

Regulation of translation [4]

Scaffolding of the non-coding region, 7S protection [1]

Decatenation, recombination

Recombination

Encoding de-tails

Mitochondria-spe-cific gene [5]

Probably identical to nuclear protein

Probably identical to nuclear protein

Alternative start codon [9]

Mitochondrial targeting se-quence

Yes [5] Unknown Unknown Yes [9]

Enzyme struc-ture

Monomer Homodimer Homodimer Monomer

Formed bond 3’ 5’ 5’ 5’

Cofactors Stimulated by Mg2+ or Ca2+ [5]

Mg2+, ATP Mg2+, ATP Mg2+

Protein size 70 kDa 174 kDa 180 kDa 110 kDa

1=Zhang et al. 2014, 2=Sobek et al. 2013 3=Khiati et al. 2015, 4=Baechler et al. 2019, 5=Zhang et al. 2001, 6=Paper II, 7=Nicholls et al. 2018, 8=Paper III, 9=Wang et al. 2002

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4.2.4 Ciprofloxacin-induced inhibition of mitochondrial function Ciprofloxacin, a fluoroquinolone, is a medical therapeutic used as an inhibitor of bac-terial gyrase and TopIV, but fluoroquinolones have been demonstrated also to inhibit human topoisomerases (Jadhav & Karuppayil 2017). Ciprofloxacin-mediated inhibi-tion of type II topoisomerases restrains the religation of cleaved DNA, generating protein-linked double-strand breaks in the bacterial genome. Interestingly, we were unable to see linearization of mitochondrial DNA after ciprofloxacin treatment, al-though total mtDNA damage in ciprofloxacin-treated post-mitotic cells was in-creased. However, as the PCR-based quantification method does not differentiate be-tween oxidative DNA damage and ciprofloxacin-mediated DNA breaks, the ob-served increase in DNA damage might be related to oxidative stress induced by ciprofloxacin (Lowes et al. 2009). In a study conducted with five different fluoroqui-nolones, these compounds did not significantly increase DNA breakage with human Top2α and Top2β, thus questioning the proposed anticancer potential of fluoroqui-nolones (Fief et al. 2019).

In this present study, ciprofloxacin was shown to impair mitochondrial DNA maintenance by disrupting the topology and replication of mtDNA. These effects im-pacted also cell physiology, as differentiation of C2C12 myoblasts to myotubes was impaired in the presence of ciprofloxacin (Paper II, figure 4E, F). The effects of cipro-floxacin on cellular differentiation has been observed already earlier, in processes known to require functional mitochondria, such as during spermatogenesis and brain development (Khaki et al. 2008, Dogan et al. 2019).

Ciprofloxacin has been suggested to intercalate into DNA, however, the in vitro short treatment of mtDNA with ciprofloxacin did not change the electrophoretic fea-tures of DNA, whereas the known DNA intercalator doxorubicin altered mtDNA migration on an agarose gel. Therefore, the observed topology changes in the mtDNA appear to be caused by the inhibition of the enzymatic activity of Top2β.

An interesting question is why ciprofloxacin seems to inhibit the mitochondrial version of topoisomerase 2 preferentially, while the nuclear isoform is less affected. As mentioned before, some mitochondrial proteins do share high similarity with bac-terial counterparts and are therefore vulnerable to antibiotics targeting bacterial homologues. The exact sequence and structure of mitochondrial topoisomerase 2 is not yet known, e.g. the mitochondrial targeting mechanism is still unclear. Many mi-tochondrial proteins do undergo post-translational modifications in the mitochon-dria to regulate their function and activity, and also topoisomerase 2 is known to be the target of several different modifications in the nucleus (Stram & Payne 2016). Therefore, it is possible that the mitochondrial version has small structural differ-ences compared to the nuclear version, even if there is currently no evidence for this found in the sequence of the TOP2 gene.

Also, the structure of mitochondria affects the capability of a drug to enter the organelle. Lipophilic cationic drugs can move through mitochondrial membranes

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with the help of the electrochemical gradient and thus reach high concentrations in the matrix (Ross et al. 2006). The negative charge and slightly alkaline pH of the ma-trix also accumulates amphiphilic chemicals, like ethidium bromide or doxorubicin. Cytochrome P450 can metabolize and activate chemicals in mitochondria that were harmless before. The similarity of the inner mitochondrial membrane with bacterial membranes (the presence of cardiolipins) may also play a role particularly in the case of antibiotics (Meyer et al. 2013).

Many possible mechanisms have been suggested to underlie ciprofloxacin-caused adverse effects. Fief et al. (2019) demonstrated in their in vitro studies that fluoroquin-olones in fact inhibit topoisomerase 2-mediated relaxion of DNA, as seen here, but only at drug concentrations considerably higher than clinically relevant. However, elevated plasma levels of ciprofloxacin have been reported in patients with altered pharmacokinetics, e.g. because of obesity, reaching the concentrations used in the present study (Hollenstein et al. 2001).

In conclusion, the inhibition of mitochondrial topoisomerase 2β is quite likely not the only cause of the observed side effects of fluoroquinolones, but it is a mechanism that should be considered in future applications of these drugs.

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5 FINAL REMARKS AND FUTURE PERSPEC-TIVES

Over the last decades the importance of mitochondrial DNA and the maintenance of this tiny genome have been acknowledged, especially considering the increasing number of human disorders and diseases connected to mutation in mitochondrial DNA itself or protein components securing the integrity and proper upkeep of the mitochondrial genome. Mitochondrial function can also be impaired by extrinsic fac-tors, a phenomenon whose extent we are only now starting to realize. However, de-spite the intensive investigation of the secrets of mitochondria, many questions re-main unanswered, and even the exact mechanism of mitochondrial DNA replication in higher animals is still under debate. The aim of this thesis was to cast some light on to the important and complex process of mitochondrial DNA maintenance.

The tissue-specific symptoms of mitochondrial diseases are well recognized, but their cause is not yet fully understood. In the present study tissue-specific aspects of mitochondrial DNA maintenance were addressed, in order to get a better perspective on this phenomenon. When the mtDNA replication mechanisms across a range of different mouse tissues were analyzed, strand-coupled replication together with high levels of X-form replication intermediates were seen in tissues with high amounts of DNA damage, suggesting that this replication mode is an adaptation to a highly oxi-dizing environment and extensive genotoxic stress. The relative abundance of Hol-liday junctions in mtDNA intermediates suggests an active repair or recombination-mediated replication of mtDNA, but the exact identity of these molecules remains inconclusive. The regulators of the switch between different modes of replication are not yet known and the investigation of this mechanism will be important in the fu-ture.

Maintenance of any longer DNA molecules requires the function of topoisomer-ases, enzymes specialized in resolving tangles and other topological problems origi-nating from the helical structure of DNA. Among other mitochondrial proteins, also the levels of topoisomerases were observed to vary between studied tissues, raising the question of their specific functions in mitochondrial DNA maintenance. As the roles of Top3α, Top2α and Top2β in mitochondria are hardly studied, their functions were further investigated in the present study. Top2α was not present in post-mitotic cells or tissues, so Top2β was hypothesized to be more important for mitochondrial DNA maintenance, because a type II topoisomerase is much needed also in those cells. Indeed, inhibition or knock-down of Top2β lead to an increase of mtDNA supercoils, which in turn impaired replication initiation, suggesting that this protein is important for the proper upkeep of mitochondrial DNA. The function of mitochon-drial Top3α was studied using siRNA mediated knock-down and overexpression of mitochondrially targeted functional or catalytically mutated versions of the protein.

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The data presented here indicates that Top3α has a function during replication fork progression, in addition to its already established role in mtDNA segregation. Its close proximity to TWNK helicase supports this suggestion and also gives the oppor-tunity to speculate about their possible involvement in mitochondrial recombination. This controversial aspect of mitochondrial DNA maintenance is an interesting topic for future research.

While mitochondrial topoisomerases clearly have specific functions in mitochon-drial DNA maintenance, there might be a broad overlap of functions that is not re-cognized yet. Since mouse models and cell lines of topoisomerase knock-outs are vi-able, topoisomerases appear able to cover each other’s task to some extent. This level of functional redundancy needs to be investigated further in the future.

Mitochondria are sensitive organelles, and the impairment of their function has far-reaching consequences. The findings of the present thesis emphasize the im-portance to assess mitochondrial side effects of therapeutic drugs such as fluoroqui-nolones. Although valuable in the treatment of life-threatening infections, the fre-quent use of these substances increases the risk of adverse side effects, especially in patients with already existing mitochondrial impairments. The exact mechanism of ciprofloxacin-induced side effects is probably manifold and the inhibition of mito-chondrial topoisomerase 2β by ciprofloxacin might be only one contributor, but it is an important factor to consider during the development of new drug generations.

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PUBLICATIONS OF THE UNIVERSITY OF EASTERN FINLAND

Dissertations in Forestry and Natural Sciences

ISBN 978-952-61-3668-4ISSN 1798-5668

Mitochondria are energy-producing organelles and therefore essential for the function of a cell, especially in tissues with high energy demand. The maintenance of the circular

mitochondrial genome, mtDNA, requires an armory of nuclear encoded proteins, among

them topoisomerases that control the topology of mtDNA. This thesis investigates the tissue

specificity of mtDNA replication and the function of topoisomerases in the maintenance

of this tiny but important genome.

ANU HANGAS