EFFECTS OF MITOCHONDRIAL DNA REPLICATION STRESS AND DOUBLE-STRAND BREAKS ON DNA … · 2018-02-28 · ITÄ-SUOMEN YLIOPISTO Ympäristö- ja biotieteiden laitos, biologia HAIKONEN,

EFFECTS OF MITOCHONDRIAL DNA REPLICATION STRESS

AND DOUBLE-STRAND BREAKS ON DNA DAMAGE

RESPONSE PATHWAYS AND MITOCHONDRIAL GENE

EXPRESSION

JUHA HAIKONEN

Pro gradu -tutkielma

Itä-Suomen yliopisto

Ympäristö- ja biotieteiden laitos

Biologia

2018

ITÄ-SUOMEN YLIOPISTO

Ympäristö- ja biotieteiden laitos, biologia

HAIKONEN, JUHA: Effects of mitochondrial DNA replication stress and double-strand

breaks on DNA damage response pathways and mitochondrial

gene expression

Pro gradu -tutkielma (40 op), 52 s., liitteitä 0

Helmikuu 2018

avainsanat: mitokondriot, mtDNA, ddC, HEK-293, ApaLI

TIIVISTELMÄ

Mitokondrioiden DNA (mtDNA) sisältää mitokondrioille elintärkeää geneettistä informaatiota,

esimerkiksi oksidatiiviseen fosforylaatioon osallistuvien proteiinien geenejä. Jokaisessa

mitokondriossa on enemmän kuin tarpeeksi mtDNA:ta ylläpitämään normaalia transkriptiota ja

replikaatiota. Koska mtDNA:n oikoluku- ja korjausmekanismit eivät vedä vertoja tuman

DNA:lle, mtDNA on altis vaurioille ja inaktivoiville mutaatioille. Vauriot sekä kemialliset

inhibiittorit häiritsevät mtDNA:n replikaatioon osallistuvia entsyymejä, pysäyttäen

replikaatiohaarukat. Pysähtyneet replikaatiohaarukat voivat romahtaa, aiheuttaen mtDNA-

juosteen kaksoiskatkoksia, jotka lopulta johtavat mtDNA-molekyylien hajoamiseen ja

mtDNA:n kopiolukumäärän köyhtymiseen. Mitokondriot sietävät väliaikaisesti suurtakin

pudotusta mtDNA:n lukumäärässä, ja jäljelle jäänyt mtDNA replikoituu suotuisissa

olosuhteissa nopeasti normaalille tasolle.

Tässä pro gradu -tutkielmassa selvitettiin mtDNA:n replikaation inhibition vaikutuksia

solujen DNA vauriosignalointiin ja mitokondrioiden omaan geeniekspressioon.

HEK-solujen mtDNA:n replikaatio estettiin 2′-3′-dideoksisytidiini-käsittelyllä (ddC), jonka

kesto oli 48 tuntia. Altistumista seurasi 64 tuntia kestävä toipumisjakso kasvatusliuoksessa,

joka ei sisältänyt ddC:tä. Solunäytteitä kerättiin sekä altistumisen että toipumisen aikana.

Näytteet käytettiin Southern blot -analyysiin, jolla mitattiin käsittelyn vaikutus mtDNA:n

kopiolukumäärään. Lisäksi tehtiin useita Western blot -analyyseja, joilla mitattiin millaisia

muutoksia mtDNA:n replikaation inhibitio aiheutti tärkeimpien DNA vauriota signaloivien

proteiinien tasoissa. Northern blot -analyysiä käytettiin transkriptiossa mahdollisesti

tapahtuvien muutosten löytämiseen.

Replikaation inhibitio 175 µM ddC-altistuksella sai aikaan solujen mtDNA tasojen

putoamisen 30%:een alkuperäisestä. Kun ddC poistettiin elatusliuoksesta, kopiolukumäärä

palautui normaalille tasolle 32 tunnissa. Kaikkien kokeeseen valittujen yhdeksän proteiinin

tasot pysyivät muuttumattomina koko kokeen ajan. Sen sijaan mtDNA:n transkriptio lisääntyi

huomattavasti ddC-altistuksen yhteydessä.

Altistuminen ddC:lle ei vaikuttanut tässä tutkielmassa käsiteltävien, HEK-soluissa

tavallisesti vaurio- tai stressivasteista aktivoituvien proteiinien tasoihin. Transkription

lisääntyminen on luultavasti kompensaatiomekanismi, jonka avulla solu yrittää selvitä

altistuksesta hyödyntämällä tehokkaammin jäljellä olevaa mtDNA:ta. Solut näyttäisivätkin

pitävän yllä suurempaa mtDNA kopiolukumäärää kuin mitä olisi tarpeellista mitokondrioiden

toiminnalle. On mahdollista, että tämä ylimäärä toimii puskurina mahdollisia stressitilanteita

vastaan tai kompensoi mtDNA:n vaurioalttiutta.

UNIVERSITY OF EASTERN FINLAND

Department of environmental and biological sciences

HAIKONEN, JUHA: Effects of mitochondrial DNA replication stress and double-strand

breaks on DNA damage response pathways and mitochondrial

gene expression

MSc. Thesis (40 cp), 52 pp., appendices 0

February 2018

key words: mitochondria, mtDNA, ddC, HEK-293, ApaLI

ABSTRACT

Mitochondria contain circular mitochondrial DNA (mtDNA) encoding essential genetic

information, such as components of the oxidative phosphorylation pathway. Because mtDNA

lacks some of the sophisticated error-correcting processes intrinsic in nuclear DNA, it is

considerably more vulnerable to damage and following deleterious mutations. The abundance

of mtDNA might represent a quantity over quality strategy to overcome inactivation via

mutagenesis. mtDNA damage as well as chemical inhibitors can interfere with the replication

of mtDNA, ultimately leading to the depletion of mtDNA copy number. However, cells can

survive a high degree of depletion and repopulate lost mtDNA once the inhibiting factors are

removed.

The aim of this master’s thesis was to elucidate the consequences of mtDNA replication

inhibition on multiple key protein involved in DNA damage response as well as to monitor

changes in mitochondrial gene expression.

mtDNA replication in HEK cells was inhibited with 2′-3′-dideoxycytidine (ddC) and the

treatment resulted in depletion of mtDNA over 48 hours, after which the cells recovered for 64

hours in ddC-free environment. Samples taken at designated timepoints were used to perform

a Southern blot to measure mtDNA copy number depletion, a Northern blot to measure

transcription, and multiple Western blot assays to elucidate changes in proteins involved in

DNA damage responses.

Replication inhibition using 175 µM ddC resulted in mtDNA depletion to 30% of the original

over a 48-hour. Repopulation to baseline levels occurs within 32 hours in ddC-free

environment. No determinable differences were observable in the nine proteins selected for

Western blotting. Instead, depletion of mtDNA induced a marked increase in mitochondrial

transcripts.

ddC exposure did not influence the expression of any of the studied damage or stress

response proteins. The increase in mitochondrial transcription is likely a compensatory

mechanism to maintain sufficient mitochondrial functions using the remaining mtDNA.

Interestingly, the cells seem to maintain higher mtDNA copy number than would be essential

for mitochondrial function. The excess mtDNA could buffer sudden stress events or

compensate for the higher susceptibility for mtDNA damage.

TABLE OF CONTENTS

1 INTRODUCTION ................................................................................................................... 1

2 REVIEW OF THE LITERATURE ......................................................................................... 3

2.1 Mitochondria .................................................................................................................... 3

2.2 Cellular energetics ............................................................................................................ 5

2.2 Reactive oxygen species ................................................................................................... 6

2.3 mtDNA and mitochondrial biogenesis ............................................................................. 7

2.3 The replication of mtDNA ............................................................................................. 10

2.4 Mitochondrial gene expression ...................................................................................... 12

2.5 Antiretroviral drugs as inhibitors of mtDNA replication ............................................... 13

2.6 The eukaryotic cell cycle and its key regulators ............................................................ 14

2.6.1 The cell cycle .......................................................................................................... 14

2.6.2 Regulators of the cell cycle ..................................................................................... 15

2.7. Role of histones in the repair of eukaryotic DNA ......................................................... 18

2.8 Mitochondrial quality control ......................................................................................... 19

3 AIM OF THE THESIS .......................................................................................................... 20

4 MATERIALS AND METHODS .......................................................................................... 21

4.1 Exposure and recovery of HEK cells to ddC ................................................................. 21

4.2 DNA extraction, purification and Southern blot -analysis ............................................. 22

4.3 Total protein extraction and Western blot analysis ........................................................ 23

4.4 RNA extraction and Northern blot analysis ................................................................... 25

4.6 DNA double-strand break experiment with transfected HEKs expressing ApaLI ........ 26

5 RESULTS .............................................................................................................................. 27

5.1 Depletion of mitochondrial DNA copy number by ddC ................................................ 27

5.2 Quantification of proteins of ddC-treated HEK cells ..................................................... 30

5.3 Quantification of RNA levels of cells exposed to ddC .................................................. 41

5.4 Double-strand breaks in ApaLI induced HEK cells ....................................................... 43

5.5 Protein levels of HEK cells transfected with ApaLI. ..................................................... 44

6 DISCUSSION ....................................................................................................................... 45

6.1 Depletion of mitochondrial DNA ................................................................................... 45

6.2 Activation of cellular DNA repair signalling ................................................................. 46

7 CONCLUSIONS ................................................................................................................... 47

THANKS .................................................................................................................................. 48

REFERENCES ......................................................................................................................... 48

1

1 INTRODUCTION

The scope of this thesis focuses on mitochondria, and mainly on the mitochondrial DNA

(mtDNA) found in abundance within each mitochondrion (Kühlbrandt 2015). The replication,

transcription, and translation of mtDNA are reviewed alongside the damage response pathways

associated with it. Several important proteins functioning inside and outside the mitochondria

are discussed in the context of their function, ranging from the cell cycle, chromatin and

oxidative phosphorylation to DNA double-strand repair. Mitochondrial dysfunction, often

linked to defective mtDNA, is the culprit of several rare and severe diseases. Many such

diseases can likely be cured in the near-future by a targeting a selective restriction enzyme to

attack the defective mtDNA, the elimination of which allows remaining and healthy mtDNA to

repopulate and take its place (Moraes et al. 2010; Moraes et al. 2012; Chan & Mishra 2014).

The experimental section of the thesis focuses on utilizing 2´-3´-dideoxycytidine (ddC) to

induce a depletion of mtDNA copy number in HEK-293 cells, and then allowing a rest period

so the remaining mtDNA can recover back to baseline level (Magnani et al. 1999; Jazayeri et

al. 2003). Cell samples gathered at distinct intervals throughout the exposure and recovery

period are used to perform a Southern blot analysis to measure the degree of depletion. A

Northern blot analysis measures transcription on a general level and a series of Western blots

are used to measure whether ddC induces changes in a selection of proteins, the majority of

which are activated by stress or DNA damage. Finally, to study how DNA double-strand breaks

affect mtDNA, a stable cell line of transfected HEKs, containing a gene construct for

mitochondrial targeting and the inducible expression of a restriction enzyme, ApaLI, is utilized.

Expression of ApaLI is induced by doxycycline treatment (Bayona-Bafaluy et al. 2005).

The proteins of interest were selected due to their roles in DNA-damage response, cell cycle,

oxidative stress, and other essential cellular processes. Proteins of interest functioning outside

the mitochondria are Chk1, a cell cycle mediator; p21, a cyclin dependent kinase inhibitor; p53,

a tumour suppressor; H2AFX, a histone protein associated with DNA double-strand breaks;

PKB, a multi-purpose serine/threonine-specific protein kinase; and Parkin, a ligase associated

with mitophagy.

Proteins of interest functioning inside the mitochondria are MRPL11, a mitochondrial

ribosomal component useful for estimating mitoribosomal abundance; TFAM, the major

2

mitochondrial transcription factor; and UQCRC2, a subunit of Complex III of the electron

transport chain and associated with oxidative stress.

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2 REVIEW OF THE LITERATURE

2.1 Mitochondria

Mitochondria are maternally inherited, cytoplasmic organelles popularly known as the

powerhouses of the cell. Mitochondria originate from a symbiotic eubacterium acquired by a

larger eukaryotic cell in a process known as endosymbiosis, which explains the organelle’s

relatively large dimensions: 1 to 2 µm in length and 0.1-0.5 µm in width – approximately the

size of an E. coli bacterium (White et al. 2016; Van der Giezen 2011). For 1.5 billion years, co-

evolution has worked to tighten the symbiosis to a degree that was considered absolute for all

eukaryotes, right up to the recent discovery of a eukaryotic microorganism, which has both

gained and lost mitochondrial function during its evolutionary history (Karnkowska et al.

2016).

Mitochondria are essential bioenergetics and biosynthetic factories. Cells primarily rely on

breakdown of adenosine triphosphate (ATP), a molecule with large amounts of chemical energy

stored in its phosphate bonds, for their energy needs. Mitochondria are capable of synthesizing

ATP from inorganic phosphorus and adenosine diphosphate (ADP) (Klingenberg 2008). Due

to great demand for ATP, each cell contains hundreds of mitochondria (White et al. 2016).

Notably, mitochondria are not defined in shape but form dynamic, tubular networks in cells that

undergo constant change through fusion and fission (Lackner 2013). Mitochondrial population

adapts to the energy needs of the cell. Physiological and environmental conditions and the

metabolic functions of the cell ultimately dictate its specific energy requirements (Lezza &

Picca 2015).

Mitochondria have two membranes that both segregate them from their immediate

surroundings and define their three distinct, structural mitochondrial compartments. The outer

mitochondrial membrane allows ions and small, uncharged molecules to diffuse freely through

its porous surface. Large molecules, such as nuclear-encoded mitochondrial proteins, are unable

to pass through the pores and rely on active transport by translocases on the membrane surface.

The pores are formed by specific pore-forming membrane proteins, collectively known as

porins. One such protein is the Voltage-dependent Anion Channel (VDAC), which forms

general diffusion pores for small hydrophilic molecules. Due to the porosity of the outer

mitochondrial membrane, no electrochemical gradient forms across it (Kühlbrandt 2015).

4

Unlike the outer mitochondrial membrane, the inner mitochondrial membrane acts as a tight

diffusion barrier, blocking all ions most molecules, although some are transported via specific

and selective membrane transport proteins (Kühlbrandt 2015). The innermost compartment,

surrounded by the inner membrane, is the mitochondrial matrix, which can be thought as the

cytoplasm of the ancestral endosymbiotic bacterium. Within the matrix’s alkaline pH of 7.9 to

8.0, mtDNA is stored, replicated and transcripted, proteins synthesis occurs, and numerous

enzymatic reactions take place. The absence of pores and the non-permeability of the inner

membrane to small ions and molecules, combined with the alkaline environment of the matrix,

creates an electrochemical trans-membrane potential of approximately 180 mV to build up

between the matrix and the intermembrane space (Kühlbrandt 2015).

The intermembrane space between the outer membrane and the part of the inner membrane,

known as the inner boundary membrane, can be thought as the periplasm of the ancestral

bacterium. All mitochondria-targeted matrix proteins pass through both membranes and this

~20 nm gap in between them (Kühlbrandt 2015).

Cristae are the third mitochondrial compartment, formed when the inner mitochondrial

membrane at the inner boundary membrane extends into the matrix, forming distinct, tubular-

like extensions. These extensions are the main site for biological energy conversion, greatly

increasing the surface area available for electron transport chain complexes and ATP synthases.

Mitochondrial site and cristae organizing system (MICOS) anchors the cristae to the outer

membrane (Kühlbrandt 2015). Likewise, cristae are connected to the inner mitochondrial

membrane by narrow, slot-like structures of varying length, so called crista junctions (Reichert

et al. 2009).

Although ATP synthesis is their primary function, mitochondria partake in a multitude of

critical cellular processes. Biosynthesis of many small molecules include steps that involve

mitochondria, requiring precursors and products to be shuttled into and out of the mitochondria

via transport proteins. Some of the mitochondrial functions that do not involve ATP production

include the synthesis and processing of fatty acids, steroid hormones, pyrimidines, iron-sulphur

clusters, phospholipids, ubiquinone and amino acids. For example, biosynthesis of heme begins

inside mitochondria, requires cytoplasmic modification steps in between to yield a precursor,

which then transports back to mitochondria for terminal steps. Additionally, mitochondria

regulate reactive oxygen species and ion homeostasis, ammonia detoxification, thermogenesis

5

and fatty acid oxidization (Shadel et al. 2006; Soria et al. 2012; Eisenberg-Bord & Schuldiner

2017).

2.2 Cellular energetics

ATP is the universal cellular energy unit, which provides the cell with chemical energy to power

everything from osmotic work to biosynthesis, transport of molecules, etc. ATP synthesis

process is so rapid that the human body can synthesize its own weight of ATP in a single day.

The thermodynamic efficiency of ATP synthesis process has recently been given an exact

estimation of 40 to 41 %, which is significantly lower from the estimate of 55 to 60 % that

many dated textbooks frequently suggest (Nath 2016).

ATP is spent to perform work by hydrolysing its chemical bond with phosphate and coupling

this energy releasing reaction with different, energy requiring reaction. Hydrolysis cleaves the

ATP molecule into ADP and inorganic phosphate. ADP can then be hydrolysed further into

AMP to release more energy, or be transported back into the mitochondria for regeneration into

ATP. Inorganic phosphate is transported for regeneration by phosphate carriers in the inner

mitochondrial membrane gather up inorganic phosphate (Krämer 1996; Warshel & Kamerlin

2009).

The term cellular respiration encompasses the series of chemical reactions which break down

acquired nutrients into high-energy molecules that are fed into a common metabolic pathway

to produce ATP, with carbon dioxide (CO2) and water (H2O) forming as by-products.

Additionally, cellular respiration involves the passage of high-energy electrons through a series

of oxidation and reduction reaction steps, called an electron-transport chain. When combined

with the synthesis of ATP from ADP and inorganic phosphor, the process as whole is known

as oxidative phosphorylation (OXPHOS). OXPHOS occurs in the mitochondria of nearly all

eukaryotic cells. In a biological system, oxygen – a relatively strong oxidant – serves as the

terminal electron acceptor in aerobic respiration, maximizing the conversion of energy in

nutrients into ATP. Some prokaryotes, such as anaerobic archaea and bacteria, use weaker

oxidants instead of oxygen as their final electron acceptor, which allows them to thrive in

oxygen free environments. Sulphates (SO42-) and nitrates (NO3-) can substitute for oxygen as a

final electron acceptor, although at a cost of a lesser ATP yield (Lodish et al. 2016: 515).

6

Glucose is the preferred fuel for ATP synthesis. A single glucose molecule yields six

molecules of CO2 and as many as 30 molecules of ATP (Rich 2003). The conversion of glucose

to ATP begins with glycolysis in a series of enzyme-catalysed reactions in the cytosol, which

yields two molecules of pyruvate, with some of the released energy captured as two molecules

of ATP and one molecule of NADH. Pyruvate transports into the mitochondrion where it is

converted into Coenzyme A (CoA), which is fed into the citric acid cycle where it oxidizes into

CO2. The energy released during oxidation charges nicotinamide adenine dinucleotide (NAD+)

and flavin adenine dinucleotide (FAD) with high-energy electrons, reducing them to NADH

and FADH2, respectively (Nelson & Cox 2008: 528, 542, 616).

The mitochondrial respiratory chain in the inner mitochondrial membrane is composed of

four multimeric protein complexes: I, II, III, and IV. The complex III, known as cytochrome c

reductase, is assembled from 10 nuclear encoded proteins and one protein encoded by mtDNA.

Cytochrome c reductase is incorporated into a respirasome supercomplex with complexes I and

IV. This supercomplex functions like a single enzyme (Enriquez et al. 2008; Miyake et al.

2013).

The high-energy electrons carried by NADH and FADH2 are passed along the respiratory

chain all the way down to the final electron acceptor. As the electrons are passed from complex

to complex in this multi-step process, their energy is released safely and in manageable

quantities. The released energy is used to pump protons from the matrix across the inner

mitochondrial membrane into the inter-membrane space, generating the proton-motive force.

Finally, electrons flowing back down their concentration and voltage gradients powers the F1F0-

ATP synthase, which synthesizes ATP from ADP and inorganic phosphate (Nelson & Cox

2008: 712, 723-725).

2.2 Reactive oxygen species

Oxidative phosphorylation generates reactive oxygen species (ROS) as by-product. ROS are

traditionally viewed as damage-inducing and detrimental to mitochondrial function and cellular

health, but more recent research has determined that moderate ROS levels have important

physiological functions and play a role in a wide range of cellular responses, including signal

transduction, proliferation, differentiation, and other regulatory functions. ROS are generated

in mitochondria when a single electron escapes the electron transport chain and encounters

7

molecular oxygen, forming a radical and dangerous superoxide (O2-). Typically, superoxide is

rapidly converted into hydrogen peroxide (H2O2) by superoxide dismutase. Although H2O2 is

detrimental in high quantities, it is kept in check by catalase enzyme and scavenging

peroxidases. Simultaneously, thioredoxins and glutaredoxins seek out and reverse oxidative

damage caused by H2O2. The escape of electrons from iron-sulphur groups, flavin-containing

proteins, or from ubisemiquinone of the Q cycle may generate superoxide in complexes I, II

and III (Schumacker et al.2016).

Superoxide generated on the matrix side of the inner mitochondrial membrane – a major site

of ROS production – is released into the aqueous matrix environment, where it may encounter

and damage mtDNA. Compared to its nuclear counterpart, mtDNA has a high rate of mutation,

and ROS-induced damage is believed to be one contributing factor. However, mitochondria can

repair most DNA lesions via homologous recombination or non-homologous end joining, and

the presence of multiple copies of mtDNA suggests that homologous recombination is the

primary system for repairing double-strand breaks (DSB) in mitochondria. When mtDNA

suffers multiple DSBs, the recombination process often results in large deletions (Moraes &

Williams 2009).

In humans, the nuclear gene UQCRC2 encodes for one of the core protein components of

complex III. Elevated level of UQCRC2 protein is associated with oxidative stress and elevated

levels of mitochondrial ROS (Pang et al.2015). Detection of excess UQCRC2 in a cell sample

may therefore be an indicator of oxidative stress.

2.3 mtDNA and mitochondrial biogenesis

Human mtDNA is circular 16,569 base pairs long chain consisting of a cytosine-rich light strand

and a guanine-rich heavy strand (Fig. 1). It contains 37 genes encoding for two ribosomal

ribonucleic acid (rRNA), 22 transfer RNA (tRNA), and 11 messenger RNA (mRNA) species

translating to a total of 13 proteins (Anderson et al. 1981; Kaufman et al. 2012). Additionally,

mtDNA has two non-coding regions (NCR) that regulate its replication and gene expression.

The major, 900 base-pairs long NCR contains a single promoter for the light strand (LSP), and

two heavy strand promoters (HSP1 and HSP2), as well as the origin of replication for the heavy

strand (OH), making it the major site of transcription regulation. The minor – only 30 base-pairs

8

long – NCR is located between the coding sites for tRNA-Cysteine and tRNA-Asparagine, and

contains the origin of replication of the light strand (OL) (Lezza & Picca 2015).

Aside from the two rRNAs translating to 12S and 16S mitoribosomal subunits and the 13

protein components of the OXPHOS machinery, mtDNA does not contain sufficient genetic

information for functional mitochondria (Mai et al. 2017). Indeed, all other mitochondrial

components and proteins are nuclear in origin. The remaining ~80 OXPHOS constituents, along

with 1200 to 1500 additional proteins found in mitochondria are transcripted from nuclear DNA

and translated in the cytosol. Specialized targeting and translocation mechanisms import these

proteins into the mitochondria (Shadel & Bestwick 2013).

Although most of mitochondrial proteins are synthesized and imported from the nucleus,

each cell retains approximately 1000 copies – far more than necessary – of mtDNA-containing

nucleoids, where compacted mtDNA exist as spherical, supramolecular assemblies (Kühlbrandt

2015). The compaction is primarily due to mitochondrial transcription factor A (TFAM), which

binds mtDNA non-specifically, bending and wrapping it (Lezza & Picca 2015). Based on the

level of TFAM within cells, estimates on the number of mtDNA molecules per nucleoid have

been made. The estimates vary from 1 to 1.4 to up to 3 mtDNA molecules per nucleoid (St.

John 2014). In addition to segregating mtDNA, TFAM is capable of binding, unwinding and

bending mtDNA at specific sites upstream of mtDNA promoters (Shadel & Bestwick 2013).

TFAM is involved in mtDNA transcription, maintenance and replication. Furthermore, there is

some evidence TFAM might have a role in mtDNA base excision repair (Canugovi et al. 2010).

Finally, the transcriptional coactivators of the peroxisome proliferator activated receptor

gamma coactivator-1 (PGC-1) family controls the expression of mtDNA-encoded proteins by

regulating TFAM (Lezza & Picca 2015).

9

Figure 1. Map of the human mitochondrial genome. The black band stands for the heavy strand

while the grey band stands for the light strand. Ribosomal RNA components are depicted in

purple. Each blue line marks the location of a tRNA, denoted with a single letter. Genes located

on the heavy strand are labelled outside the circle and while the single protein of the light strand,

ND6, is labelled inside. The light strand contains a single promoter, abbreviated as LSP. Heavy

strand contains two promoters; HSP1 and HSP2. Both heavy and light strand contain a primary

origin of replication, denoted by OH and OL, respectively. Above the genome map, an area

very high in transcriptional activity, known as the D-loop, is presented with detailed locations

of the three promoters, TFAM binding site, binding sites at LSP and HSP1 in relation to TFAM,

and the conserved sequence blocks (CSBs I, II, and III). Adapted from Shadel & Bestwick

2013.

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2.3 The replication of mtDNA

Replication of mtDNA differs considerably from nuclear DNA. The study of mtDNA

replication has proved difficult, with exact details and regulators requiring further analysis.

Several models for mtDNA priming and replication have been proposed, and the strand

displacement model discussed below is perhaps the most credible. Curiously, up to 95 % of

mammalian mtDNA replication events initiated at the heavy-strand origin of replication are

aborted after transcription of approximately 650 nucleotides, leading to the formation of triple-

stranded region of mtDNA known as the displacement loop (D-loop). The abortive mtDNA

product, denoted 7S DNA, remains bound to the parental L-strand, displacing the H strand. It

is possible that 7S DNA is maintained by antihelicase activity that prevents the helicase

mitochondrial DNA helicase TWINKE from unwinding it, and the obstacle is only overcome

once rising TWINKLE levels exceed a certain threshold. The D-loop contains all major

regulatory sites of both strands for mtDNA replication and transcription (Taanman 1999; Kühl

et al. 2016). The 7S DNA is easily detectable in a Southern blot by briefly heating the purified

DNA samples before they are loaded onto an agarose gel for electrophoresis.

The replication of mtDNA requires the concerted effort of the mitochondrial replication

machinery. mtDNA polymerase gamma (POL γ) is the primary polymerase of mtDNA repair

and replication, possessing 3'-5' exonuclease and 5' dRP lyase activity in its catalytic subunit

(Stumpf & Copeland 2011). In addition to POL γ, replication requires the mitochondrial RNA

polymerase (POLRMT), TWINKLE, the mitochondrial single-stranded DNA binding protein

1 (SSBP1), and the primer-processing enzymes: RNA processing endonuclease (MRP) and

endonuclease G. It is worth underscoring that few elements distinguish transcription and gene

expression of mtDNA from its replication, as the two processes are intimately linked, with the

former using widely the same molecular machinery as the latter. This is characterized in

mtDNA by an abrupt switch from expression to replication, discussed below (Taanman 1999;

Lezza & Picca 2015; Kühl et al. 2016).

For mtDNA replication process to begin, a RNA primer is needed. According to the strand

displacement model, transcription of the primers begins at OH approximately 100 base pairs

downstream of LSP. Once transcription passes the primer sequence located in the D-loop, it

terminates prematurely, leaving behind the immature primer used for mtDNA replication. This

transition from transcription to replication takes place at several distinct sites, which

11

collectively constitute the OH, in a region of three short, evolutionary conserved sequence

blocks denoted CSB I, II and III. It is speculated that these sequence blocks direct precise

cleavage of primary transcripts to provide the appropriate primer species as well as allow the

RNA precursors to form stable and persistent three-stranded RNA-DNA hybrids known as R-

loops (Taanman 1999; Kühl et al. 2016).

R-loops are enzymatically processed by mitochondrial RNA processing endonuclease

(MRP) and endonuclease G to yield mature, functional primers. Furthermore, recent evidence

points out that primer synthesis and initiation of replication are prioritized over gene

transcription, with POLRMT functioning as concentration-dependent molecular switch

between the two. Low level of POLRMT favours initiation of transcription initiation at LSP,

ensuring that primer synthesis is maintained. Conversely, high level of POLRMT allows

transcription to proceed, leading to gene expression. Additionally, once POLRMT is depleted

from the cell, 7S DNA no longer forms, suggesting that stalled replication events resume to

completion (Taanman 1999; Kühl et al. 2016).

Once the primer for the H-strand is in place, replication proceeds unidirectionally from the

temporally and spatially distinct, strand-specific origins of replication, OH and OL. Replication

begins at OH with the synthesis of daughter H-strand and continues along the parental L-strand

to produce a full H-strand circle. Progression of the replication fork leaves the L-strand single-

stranded, and once the replication fork passes OL, located two-thirds of the genomic distance

away from OH, the region forms a stem-loop structure to which POLRMT binds and synthesizes

the primer necessary for initiation of mtDNA replication at the L-strand. L-strand replication

fork proceeds in the opposite direction from the H-strand replication (Taanman 1999; Kühl et

al. 2016). For replication to proceed past initiation, the catalytic subunit of POL γ must be

activated by TFAM, and supported by mtSSB (single-stranded DNA-binding protein,

mitochondrial), TWINKLE, and POLGB – the accessory subunit of POL γ (St. John 2014).

Mitochondrial biogenesis requires the coordinated expression of both nuclear DNA and

mtDNA, and regulators such as members of the PGC-1 and PGC-related coactivator (PRC)

protein families. One pathway of mitochondrial biogenesis occurs when PGC-1 proteins

activate nuclear transcription factors: nuclear respiratory factor 1 and 2 (NRF-1 and NRF-2)

and the estrogen-related receptor alpha (ERRα) that regulate the expression of mitochondrial

proteins encoded by nuclear DNA. Thus, expression of many mitochondrial proteins, such as

12

the mitochondrial transcription factor A, (TFAM) increases. The presence of TFAM is essential

for regulating mtDNA copy number in mitochondrial biogenesis (Lezza & Picca 2015).

2.4 Mitochondrial gene expression

Because mtDNA resides in the matrix, which is a highly oxidative environment, mtDNA has

gradually lost all but the most essential genes to preserve genetic integrity. The remaining genes

are highly conserved and compact. Thus, a minimal set of 22 tRNAs encoded by human mtDNA

are sufficient for de-coding the genetic information during transcription. The genetic code used

by mitochondria differs from the universal one. In addition to UAA and UAG, human

mitochondria use the arginine codons AGG and AGA for termination while UGA codes for

tryptophan instead of a stop codon (Watanabe 2010; Ott et al. 2016). Additionally, initiation of

transcription does not require specialized methionine to act as a starter codon. Instead, tRNAMet

functions both in initiation and elongation (Watanabe et al. 1994).

Human mtDNA is transcribed from three promoters, generating polycistronic (i.e.

containing multiple genes) transcripts. The LSP controls the transcription of eight tRNAs and

the ND6 protein, while also generating the RNA primer for first strand mtDNA replication at

OH. The two promoters in the heavy strand, HSP1 and HSP2, are located upstream of the LSP.

HSP1 controls the expression of two tRNAs and two rRNA, while HSP2 controls the expression

of the remaining 12 tRNAs and 12 protein coding transcripts. Transcription from HSP1

terminates immediately after the rRNA genes, but transcripts from LSP and HSP2 are nearly

full genome in length (Kaufman et al. 2012; Shadel & Bestwick 2013).

Biogenesis of mitoribosomes has several distinct stages. First, the smaller and larger

mitochondrial ribosomal subunits are synthesized from mtDNA transcripts. Second, nuclear

gene expression produces the mitoribosomal proteins of the small (MRPS) and large (MRPL)

subunits, and necessary post-translational modifications are completed. Third, various post-

transcriptional modifications are made to the tRNAs. Finally, the smaller and larger subunits

associate with their respective proteins, then assemble to form a functional mitoribosome

(Hällberg & Larsson 2014).

The mitochondrial ribosomal protein L11 (MRPL11), encodes for a protein component of

the large subunit of mitoribosomes. As is the case with most mitochondrial proteins, MRPL11

13

is initially encoded in the nucleus. Western blot detection of MRPL11 allows for quantification

of mitoribosomal function and abundance, and its possible fluctuations during the experiment

(Hällberg & Larsson 2014).

In humans, the initiation of transcription requires primary mitochondrial transcription

components, which are the human mitochondrial polymerase (POLRMT) itself, human

mitochondrial transcription factor B2 (TFB2M), and TFAM. Just like all mitochondrial RNA

polymerases, POLRMT is DNA-dependent and consists of a single subunit. POLRMT cannot

initiate promoter specific transcription on double-stranded DNA alone; instead, it generates

RNA primers for initiation of DNA replication, thereby coupling transcription of mtDNA to

replication. It is still unclear whether TFAM is required for transcription initiation, or whether

it merely acts as a transcriptional activator or repressor, leaving transcription initiation to

TFB2M (Shadel & Bestwick 2013).

Transcription by POLRMT yields two primary transcripts, one originating from LSP and the

other from HSP. Further processing by various RNases excises the primary transcripts, liberates

the tRNAs flanking the mRNAs and rRNAs in the process and releases the clear majority of

individual tRNAs, rRNAs, and mRNAs. Currently, the exact processing of mRNAs not

liberated in this way remains to be determined (Hällberg & Larsson 2014; Shadel & Bestwick

2013).

The mitochondrial primary transcripts differ from their nuclear counterparts, because they

do not contain any introns (Anderson et al. 1981; Kaufman et al. 2012). Furthermore, because

mtDNA contains only components for its own protein synthesis and the OXPHOS machinery,

there is no evidence suggesting that any RNA transcripts or proteins of mitochondrial origin

are ever transported out of the mitochondria they are produced in, eliminating the need for any

kind of nuclear targeting machinery (Lezza & Picca 2015).

2.5 Antiretroviral drugs as inhibitors of mtDNA replication

2´,3´-Dideoxycytidine (ddC; ddCyd; or Zalcitabine) is a nucleoside transcriptase inhibitor

(NRTI) that was sold under the trade name Hivid to combat acquired immunodeficiency

syndrome (AIDS) caused by human immunodeficiency virus (HIV) in the US. NRTIs are used

in the highly active antiretroviral therapy (HAART) to significantly increase the life expectancy

14

of HIV patients. Zalcitabine is a pyrimidine analogue of the naturally occurring nucleoside, 2´-

deoxycytidine. The sale of Hivid began in 1992 and it was discontinued in 2006 due to serious

adverse side effects and the development of a new generation of NRTIs with more favourable

risk/benefit profiles (Birgerson 2006). In addition to inhibiting HIV-1 reverse transcriptase

required for the replication of retroviruses, Zalcitabine is toxic to mitochondria. The toxic effect

arises from the inhibition of DNA polymerase γ, which was, prior to the discovery of a novel

polymerase PrimPol (Blanco et al, 2013), presumed to be the only polymerase mediating DNA

synthesis related to the replication and repair of mtDNA (Magnani et al. 1999, Lodi et al. 2015).

Within a biological system, ddC transforms into dideoxyCTP (ddCTP), a biologically active

and toxic form. ddCTP readily transforms further into ddCDP-choline. Indeed, ddCDP-choline

may be the preferred form for accumulation and localization into the mitochondrion. Once in

the mitochondrion, ddCDP-choline readily transforms back into toxic ddCTP due to the

reversible nature of phosphocholine cytidylyltransferase reaction. Thus, ddCDP-choline may

act as a reservoir for ddCTP, prolonging exposure time. Inhibition of polymerase γ by ddCTP

ultimately depletes mtDNA copy number, causing a delayed toxicity effect (Magnani et al. et

al. 1999).

2.6 The eukaryotic cell cycle and its key regulators

2.6.1 The cell cycle

The cell cycle consists of several and distinct checkpoint responses, which help cells to achieve

precise and error-free mitosis. It is a tightly regulated process influenced by both internal and

external signals. The cell cycle can be partitioned into S-phase (Synthesis) and M-phase

(Mitosis) with intermediate G1 and G2 (Gap) phases in between. Replication of DNA occurs in

the S-phase while division of DNA occurs in the M-phase. The cell prepares for replication and

mitosis in G1 and G2, respectively. A defective cell cycle is detrimental to the genome and may

ultimately prove fatal to the cell (Kubiak 2011: 421-422, 461).

On molecular level, the transition from G1 to S-phase is a precisely timed and concerted

effort of enzyme phosphorylations and dephosphorylations coupled to the expression of key

regulators. The cyclin-dependent kinases are the driving force of the cell cycle, but they remain

15

inactive when growth signals are absent. To activate, they require the binding of a specific

kinase, removal of inactivating phosphate groups, and addition of activating phosphate groups

(Kubiak 2011: 421).

The sophisticated molecular machinery of the cell cycle can detect lesions in the genome

and arrest the cell cycle progress for repairs. The location of the DNA lesion determines the

checkpoint where the cell cycle arrests. The molecular machine reacting to DNA lesions can be

divided into three distinct parts: 1) the sensor which detects the lesion and emits a signal, 2) the

signal transduction cascade relaying the signal to the 3) effector, which ultimately arrests the

cell cycle. When the cell cycle arrests in G1, replication of damaged genome is halted. Likewise,

if the cell cycle arrest in G2 due to chromosomal damage, entrance to mitosis is prevented,

providing time for repairs and avoiding mitotic catastrophe or irreversible loss of genetic

information. However, there exists a point within G1 which, when crossed, irreversibly commits

the cell for transition to S-phase and genome replication. The intra-S checkpoint allows for

repairs of DNA lesions in cells already committed to mitosis by transiently slowing the rate of

DNA synthesis (Kubiak 2011: 76; Smits & Gillespie 2015).

Cells facing replication stress can delay the onset of mitosis until genome replication is

completed by utilizing a distinct S-M checkpoint. Severe stress may prevent replication

completion, leading to stalled replication forks, which are thought to require either an active

process of checkpoint-mediated stabilization or rescue by converging forks to prevent a

dangerous fork collapse. During such crisis, the cells minimize the firing of additional

replication forks (Smits & Gillespie 2015, Cortez 2015).

2.6.2 Regulators of the cell cycle

Protein kinase B (PKB) is a highly conserved, critical signalling molecule. PKB belongs to the

AGC group of protein kinases, and it has over 50 proteins as putative substrates. Mammals have

three PKB isoforms: PKBα is ubiquitous, PKBβ is restricted to insulin-sensitive tissues, and

PKBγ is found in the brain and testis. Each isoform is encoded by a separate gene, yet amino

acid sequence, structure, and the three functional domains are highly similar between isoforms.

The functional domains consist of an amino terminal domain, a central catalytic domain, and a

carboxyl-terminal regulatory domain with a hydrophobic motif (Rommel et. al 2010: 32)

16

Phosphoinositide 3-kinase signalling activates PKB via phosphorylation of its catalytic

domain and PKB is stabilized by phosphorylation of the hydrophobic motif in the regulatory

domain. Active PKB phosphorylates a wide range of substrates involved in multiple cellular

processes, such as progression through the cell cycle, cell growth and differentiation, cell

survival or suppression of apoptosis, metabolism, angiogenesis, and motility (Rommel et. al

2010: 33).

Another prominent regulator, thee checkpoint kinase 1 (Chk1), controls all but the G1

checkpoint of the cell cycle. It is active in both embryonic and most proliferating somatic cells.

Although Chk1 is known best for its role in perturbed cell cycles, it is generally required for

successful cell division as well. When a cell experiences replication stress or suffers damage to

its genome, Chk1 function is amplified to trigger the activation replication and DNA damage

checkpoints, respectively. Although cell cycle checkpoints are crucial in preventing cell death

under conditions of acute genotoxic stress, each arrest must also be reversed after the event, or

the cell will be unable to resume mitosis (Smits & Gillespie 2015).

In response to DNA damage at G2, Chk1 inhibits CDC25 family phosphatases via a

combination of protein degradation and association with 14-3-3 proteins. Additionally, Chk1

stimulates the activity of Wee1 by phosphorylation. The combined effect causes a lasting

inhibition of CDK1 for as long as DNA damage persists, blocking the entry to mitosis. During

S-phase, Chk1 can mediate the degradation of CDC25A phosphatase, suppressing the activity

of CDK2. Inhibition of CDK2 slows the rate of DNA synthesis at S-M checkpoint (Smits &

Gillespie 2015).

The checkpoint functioning outside Chk1 influence is G1, which functions primarily under

p53, a protein encoded in mammals by Tp53 – a well-known and extensively researched

oncogene. Due to its role in apoptosis and tumour suppression, Tp53 has been popularly dubbed

as “Cellular Gatekeeper” and “Guardian of the Genome” (Woods & Vousden 2001). Tp53 is

the most frequently mutated gene in cancer, highlighting the crucial need of functional p53 in

DNA damage response (Speidel 2015). Roughly half of cancer incidents in humans can be

attributed to mutated, inactive Tp53 while the rest can be linked to defective Tp53-dependent

signalling pathways. Most common mutations are point mutations of a single amino acid,

leading to the production of defective p53, which lacks sequence-specific DNA binding ability

(Vazquez et al. 2008).

17

Tp53 is expressed continuously to produce p53. However, in normal, unstressed cells, p53

is quickly degraded due to efficient downregulation. Without any post-translational

modifications, p53 is checked by Mouse double minute 2 homolog (Mdm2). In addition to

forming complexes susceptible to proteasome degradation with p53, Mdm2 can directly inhibit

its translation, making it the primary negative regulator of Tp53 (Kubiak 2011: 431). However,

unmodified p53 is activated via signal transduction upon detection of multiple inner and

external signals of DNA damage and cellular stressors, such as lack of oxygen, decrease of

growth factors, ionizing radiation, UV-light, harmful chemicals, chemotherapy agents,

oncogene signalling and defective nucleotide synthesis. In response to these insults, p53 levels

inside the cell increase drastically within one hour of exposure (Speidel 2015; Chen &

Rajewsky 2007; Vazguez et al 2008). When exposure to such an event occurs, upstream sensory

kinases such as ATM and ATR phosphorylate the two serine residues of p53, stabilizing and

heightening its tetrameric structure, increasing lifespan and activity while simultaneously

decreasing affinity for Mdm2. Active p53 then binds to sequence-specific DNA and regulates

the transcription of its target genes, continuing the reaction initially triggered by a detected

stressor by a sensory kinase (Speidel 2015; Kubiak 2011: 431, Chen & Rajewsky. 2007;

Vasquez et al 2008; Gartel 2008).

One target gene targeted by active p53, CDKN1A, encodes a relatively small, 164 amino

acids long p21, which functions in the cell cycle downstream of p53. The promoter area of

CDKN1A contains two conserved binding sites with high affinity for active p53, the binding of

which accelerates p21 synthesis. In humans, transcription of CDKN1A is mainly regulated by

p53, although several factors with control over p53 indirectly regulate p21 synthesis as well

(Hayat et al. 2013: 154-155). A simple knock-out of the Tp53 encoding for p53 leads to decrease

in p21 levels and increase in cyclin-kinase-complexes, stimulating cell proliferation (Harper et

al 1993).

p21 associates with cyclins A, B, D1, and E, as well as cyclin-dependent kinases Cdk1,

Cdk2, and Cdk4/6. Additionally, p21 is well-documented to inhibit cyclin-kinase-complexes

D-Cdk4/6. Furthermore, p21 indirectly limits E-Cdk2 activity by inhibiting D-Cdk4/6 complex

activity and B-Cdk1 activity by association with 14-3-3σ-protein, which blocks B-Cdk1

complex localization to nucleus (Chen & Rajewsky 2007, Harper et. al 1993, Hayat et al. 2013:

155-156). In addition to having a binding-site for cyclins and kinases in its amino- and carboxyl-

terminal ends, respectively (Abbas & Dutta 2009), p21 contains separate binding sites for

procaspase-3 and proliferating cell nuclear antigen (PCNA). Finally, p21 contains a nuclear

18

localization sequence which allows it to move to the nucleus from the cytosol. All in all, p21 is

known to induce the expression of 55 genes, but disrupting the expression of up to 77 genes

(Hayat et al. 2013: 155-157). Aside from cell cycle arrest, p21 plays a role in multiple cellular

events, such as apoptosis, rescue from apoptosis, senescence, DNA replication and repair, and

in some cases, it may even accelerate cell cycle progression (Yousefi & al. 2016).

p21 is a potent regulator of the cell cycle, capable of arresting progression in G1/S and G2/M

checkpoints by inhibiting CDK4,6/cyclin-D and CDK2/Cyclin-E complexes, respectively.

CDK-cyclin-complexes advance the cell cycle by partially phosphorylating retinoblastoma

(Rb), and p21 prevents this interaction. Although an arrest of the cell cycle initiated by sensory

kinases in G1/S phase does occur via a p53-dependent process, p53 is not an absolute factor for

cell cycle arrest. In this case, however, p53 activates p21, which maintains the arrest by

suppressing CDK2 activity (Kubiak 2011: 431; Yousefi & al. 2016).

2.7. Role of histones in the repair of eukaryotic DNA

Eukaryotic DNA is tightly bound to small histone proteins and orderly packaged within the cell

nucleus. The histones – called H1, H2A, H2B, H3 and H4 – are rich in positively charged amino

acids arginine and lysine, which facilitates binding to negatively charged DNA. Histones form

the nucleosome core particle consisting of a histone octamer with two copies each of histone

H2A, H2B, H3, and H4. The DNA is wrapped around the core particle and sealed by histone

H1. Repeating units of nucleosomes connected by linker DNA form chromatin, which, due to

the abundant presence of various nonhistone proteins, typically has a protein-to-DNA ratio of

2:1. These nonhistone proteins are involved in a range of activities, including DNA replication

and gene expression (Zhu et al. 2016; Cooper et al. 204-207).

The degree of chromatin condensation allows for additional control over gene expression.

Loosely condensed chromatin is readily accessed by replication and gene expression

machinery, and often contains additional RNA elements associated with them. Conversely,

densely packaged chromatin blocks access to the DNA due to steric constraints. Multiple

enzymes can modify histones post-translationally to induce methylation, acetylation, or

phosphorylation, all three of which dramatically influence how DNA is accessed (Zhu et al.

2016; Cooper et al. 207 p.).

19

H2AFX (H2A histone family, member X) is a gene variant of histone H2A encoding for

protein H2AFX, which is essential to the cell due to its role in DNA DSBs. H2AFX has at least

two upstream activators depending whether the damage originates from replication stress or

ionizing radiation. Inactive H2AFX activates via phosphorylation at serine-139 to form active

γH2AFX in reaction to a DSB. Activated form contributes to error-free homologous

recombination repair and genomic stability. Experiments determined that H2AFX-deficient

mice are more vulnerable against genotoxic insults than their wild-type counterparts. Because

γH2AFX spreads over a large area around a DNA lesion, it makes for an excellent biomarker

for detection of DSBs (Zhu et al. 2016).

γH2AFX functions by recruiting chromatin remodelling complexes and promoting histone

acetylation to render the chromatin environment surrounding the DNA more accessible for

repair factors. For above reasons, γH2AFX is tightly regulated by several factors, such as ATM

and MDC1. Once the cell has completed repairs, γH2AFX must be eliminated before the cell

cycle can resume. SWR1 can replace γH2AFX with another variant of the H2, and several

phosphatases can also remove γH2AFX by reversing the phosphorylation (Zhu et al. 2016).

2.8 Mitochondrial quality control

Healthy mitochondria are essential for cellular physiology and homeostasis. Cells possess

multiple control pathways to maintain a healthy population of mitochondria. Mitochondrial

components regularly sustain damage from ROS by-products despite the presence of ROS-

scavenging enzymes with antioxidant properties. Damaged mitochondrial organelles are either

sequestered or diluted through the dynamic processes of mitochondrial fission and fusion.

Mitophagy-pathway for autophagy of mitochondria ultimately targets excessively damaged

mitochondria to lysosomes for degradation (Bingol & Sheng 2016).

Two important genes linked to mitochondrial integrity and quality control are the E3

ubiquitin-ligase Parkin – named after its association with Parkinson’s disease – and a

Serine/Threonine protein kinase PINK1, located upstream of Parkin in the genetic pathway.

PINK1, a mitochondrially localized kinase, is responsible for activating and translocating

Parkin into the damaged mitochondria. Activated Parkin then builds ubiquitin chains on

damaged mitochondria to mark them for destruction by mitophagy (Bingol & Sheng 2016).

20

Parkin is a protein of interest in mitochondrial toxicity studies because elevated levels of Parkin

may indicate damaged or stressed mitochondria.

Typically, some copies of mtDNA within the cell are defective mutants. A mixture of wild-

type and defective mtDNA in a cell is a state known as heteroplasmy. Because mitochondria

are inherited maternally, high degree of heteroplasmy can pass directly to offspring.

Additionally, because mitochondrial components are dual-genomic in origin, heteroplasmy

may be inherited indirectly via loss-of-function mutations of nuclear genes that code for

mitochondrial elements. In either case, severe heteroplasmy causes mitochondrial dysfunction.

Mutated mtDNA can expand clonally and create mosaic patterns of respiratory chain deficiency

in various tissues (Hällberg & Larsson 2014; Shadel & Bestwick 2013; White et al. 2016).

The deleterious effects of heteroplasmy manifest only when the percentage of functional

wild-type mtDNA decreases significantly. Depending on the mutation, defective mtDNA must

accumulate to >60-90 % of total mtDNA before OXPHOS activity is compromised. A host of

mitochondria-related diseases with a broad range of clinical phenotypes, collectively termed

encephalomyopathies, can arise when the heteroplasmy threshold is crossed. The exact nature

of the disease depends on the tissue affected. Deletion, depletion and damage of mtDNA can

be the primary cause of disease, as is the case with Alpers syndrome, but is often secondary,

such as in Parkinson’s disease (Moraes et al. 2010; Chan & Mishra 2014).

Targeted cleaving of mtDNA by specific restriction endonucleases may become a viable

therapy method for patients with heteroplasmic mtDNA disorders. In cases where the mutation

causes a unique binding site, there is potential for a specific restriction enzyme to recognize and

cleave subpopulations of defective mtDNA. Cleaved mtDNA degrades rapidly, allowing wild-

type mtDNA to expand clonally and cause a shift in heteroplasmy (Moraes et al 2010; Moraes

et al 2012).

3 AIM OF THE THESIS

The aim of this thesis is to examine how HEK-293 cells respond to partial and transient mtDNA

depletion using an inhibitor of mtDNA replication, 2′-3′-dideoxycytidine (ddC). Although

essential for mitochondrial function, it is not clear how much mtDNA is required by the cell.

By eliminating mtDNA by replication inhibition, it is possible to make a rough estimate of how

21

little mtDNA suffices to maintain mitochondrial function. Furthermore, replication inhibition

results in stalled replication forks and double-strand breaks, whose effects on the host cell have

been unexplored. To obtain insight into these cellular responses, the proteins involved in in

mitochondrial function and DNA damage response were assayed during and after ddC

exposure. Additionally, mitochondrial transcript levels were measured to determine whether

the decline in mtDNA influences mitochondrial gene expression. To complement the data

obtained from mtDNA depletion experiment, double-strand breaks of mtDNA were examined

separately by utilizing a mitochondrially targeted restriction enzyme, ApaLI.

4 MATERIALS AND METHODS

4.1 Exposure and recovery of HEK cells to ddC

Human embryonic kidney cells (HEK-293) were grown in order to measure ddC-induced

mtDNA copy number depletion, damage, and cellular stress. The cells were grown in Biowest

low glucose Dulbecco’s Modified Eagle Medium (DMEM) containing stable glutamine and

sodium pyruvate. Foetal bovine serum (FBS) was added to the DMEM for a FBS concentration

of 10 %. Introduction of ddC into the growth medium occurred 24 and 48 hours prior to

collection. One batch of cells was designated as control and did not receive any ddC. To study

cell recovery following ddC toxicity, two cell batches were exposed to ddC for 48 hours, after

which one of them was collected while the other was re-seeded onto three growth plates

containing ddC-free growth medium. These cells constituted the three timepoints, denoted as

+16, +32 and +64 hours, of the recovery phase. The seeding of recovering cells was done in

suitable cell densities to ensure a sufficient and consistent yield for each batch at the end of

each of each recovery time, after which the cells were collected. Thus, a total of five timepoints

constitute the experiment time window for measuring the exposure and subsequent recovery of

HEKs to ddC. The time window was replicated in quadruplicate.

Pure ddC is white, crystalline powder with a molecular weight of 211,24. A stock

concentration of 100 mM ddC was prepared by adding 21 mg of ddC in 1 ml of sterile water

and mixed on a rotator for 20 minutes. HEK cells were exposed to 175 µM ddC concentration

by adding 5,25 µl of 100 mM stock solution in 3 ml of growth medium. After exposure, the

growth medium was removed and the cells were first washed and then suspended in phosphate-

22

buffered saline (PBS), pelleted down and frozen at -20 °C to await analysis. Approximately 60

% of cell yield was allocated for DNA extraction and the remaining 40 % for protein extraction.

4.2 DNA extraction, purification and Southern blot -analysis

Frozen cell pellets were immersed in 400 µl of DNA lysis buffer (10 mM Tris, pH 7.4; 10 mM

EDTA; 150 mM NaCl; SDS 0.4 %). After thorough mixing, 20 µl of Proteinase K (10 mg/ml)

was added. The samples were incubated at 50 °C for 2 hours to allow protein digestion. RNA

digestion was accomplished by adding of10 µl of RNAse A (10 mg/ml) followed by 10-minute

incubation at 37 °C. Finally, 40 µl of 5 M NaCl was added to assist phase separation for the

next stage.

DNA purification and extraction was done by adding an equal volume of 25:24:1

phenol/chloroform/isoamyl alcohol. After manual shaking, the samples were centrifuged at

15,000 g for 5 minutes. Next, the upper water phase was collected into fresh Eppendorf tubes,

and the extraction was repeated once with phenol/chloroform/isoamyl alcohol and again with

pure chloroform to remove phenol traces. The nucleic acids were precipitated by adding 0.1

times the total sample volume of 3 M sodium acetate and 0.6 times the total sample volume of

isopropanol alcohol. After a brief mixing, the samples were placed in −20 °C for two hours and

then centrifuged at 15,000 g for 10 minutes. The supernatant was decanted and the pellet

washed with 99 % EtOH, then allowed to air-dry. The near-dry pellets were dissolved in 300

µl of TE-buffer (10mM Tris-HCl, 1mM EDTA•Na2) overnight at 4 °C. A 30 µl fraction from

each sample was digested with 3 µl of restriction enzyme BamH1 in 3.5 µl of Fast Digest buffer.

The samples were left to incubate overnight at 4 °C. DNA concentration of each sample was

measured with a Nanodrop ND-1000 spectrophotometer.

The samples were heated for 5 minutes at 60 ° C to allow the detection of 7sDNA prior to

electrophoresis, then 1 µg of DNA from each sample was loaded on 0.4 % agarose gel with 2

µl of 10x DNA loading buffer and sterile water for a total loading volume of 20 µl.

Electrophoresis was done overnight at 30 V and the gel was stained with 10 µl of EtBr in 200

mL of TBE for 30 minutes with gentle agitation. The gel was immersed in depurination solution

(0.25 M HCl) twice for 15 minutes each, and then immersed in denaturation solution (0.5 M

NaOH; 1.5M NaCl, pH 7.2) for 25 minutes. The contents of the gel were transferred on a nylon

blotting membrane in 20x SSC (3M NaCl, 0.3M Na-Citrate) buffer. After transfer, the

23

membrane was neutralized by a one minute in neutralization buffer (1 M Tris-HCl, 2M NaCl),

then baked for 2 hours at 80 °C to cross-link the DNA to the membrane.

The membrane was placed in a hybridization cylinder and pre-hybridized in Church’s

hybridization buffer (240 mM NaPi; pH 7.2, 7 % SDS, 1mM EDTA) for 20 minutes at 65 °C

in the hybridization oven. A 12S probe was prepared and added. Hybridization continued

overnight. The hybridized membrane was washed twice with 5xSSC and twice with 1xSSC for

20 minutes each time. The membrane was exposed on a Kodak SO230 storage phosphor screen

and the data quantified with Quantity One® software (Bio-Rad).

4.3 Total protein extraction and Western blot analysis

The total protein content of each sample was extracted by lysing frozen cell pellets in four times

of their volume of TOTEX buffer (20 m Hepes; at pH 7.9, 0.35 M NaCl, 20% glycerol, 1% NP-

40, 1 mM MgC12, 0.5 mM EDTA, 0.1 mM EGTA, 50 mM NaF and 0.3 mM NaV03). The lysed

samples were kept on ice and vortexed occasionally for 10 minutes. The samples were flash

frozen with liquid nitrogen to enhance extraction. Once thawed, they were centrifuged for 10

minutes at 15,000 g in 4 °C. The supernatant was collected, and a 2 µl fraction from each sample

was used for Bradford protein assay (Bradford 1976). Protein content was measured using a

FLUOstar Omega microplate reader by BMG Labtech.

SDS-PAGE electrophoresis was done by mixing 30 µg of protein with 2 µl of 5x Laemmli

Sample Buffer (60 mM Tris-Cl pH 6.8, 2% SDS, 10% glycerol, 10% dithiothreitol, 0.01%

bromophenol blue). Volume of each sample was equalized with sterile water to 20 µl. The

samples were heated in the mixing block for 10 minutes at 95 °C before loading. The acrylamide

content of the gel was 12 % for optimal separation of proteins in the 10-200 kDa range.

Electrophoresis duration was 90 minutes at 100 V.

The Western blot membrane was primed by brief submerge in Millipore water and a 5-

minute immersion in transfer buffer. Transfer membrane was done at 4 °C at 40 V overnight.

Ponceau S staining was used to visually confirm transfer. To prevent undesired antibody

binding, the membrane was incubated in Tris-buffered saline (TBST; 50 mM Tris-Cl, pH

7.6; 150 mM NaCl) containing 5 % milk powder for 60 minutes. After a brief wash with TBST,

the membrane was submerged in 10 mL of 3 % bovine serum albumin (BSA) in TBST

24

containing the primary antibody (AB) for the protein of interest. Primary AB incubation was

done overnight at 4 °C with gentle agitation.

After primary AB incubation, the membrane was washed in TBST three times for 20

minutes each, and then immersed in 10 mL of 3% BSA in TBST containing the secondary AB.

Incubation was for 60 minutes at room temperature with gentle agitation. After three washes in

TBST for 20 minutes each, the membrane was covered with Luminol solution (50 mg Na-

Luminol sodium salt by Sigma-Aldrich, 100 ml 0.1 M Tris-HCl; pH 8.5, 62 µl 30 % H2O2) and

imaged using a UVP Biospectrum 810 imaging system. The acquired data was quantified using

VisionWorksLS Acquisition and Analysis software version 8.16.

The proteins of interest for WB detection were chosen due to their roles in DDR, cell cycle,

oxidative stress, and other essential cellular processes. Proteins functioning both inside and

outside mitochondrial origin were selected. Nuclear proteins that do not translocate inside

mitochondria were Chk1, a cell cycle mediator; p21, a cyclin dependent kinase inhibitor; p53,

a tumour suppressor; H2AFX, a histone protein associated with DSBs; PKB, a multi-purpose

serine/threonine-specific protein kinase; and Parkin, a ligase of the proteasomal degradation

machinery. Proteins that function inside mitochondria were MRPL11, a mitochondrial

ribosomal component that is useful when estimating mitoribosomal abundance; TFAM, the

major mitochondrial transcription factor; and UQCRC2, a component of the complex III

associated with oxidative stress.

To quantify detected proteins, a ubiquitous and continuously expressed control protein was

needed. Vinculin is a 117 kDa focal adhesion protein and one of the main components of the

cell’s mechanosensory machine (Atherton et al. 2016). Due to its vital role in mechanosensing

and homeostasis, Vinculin level in a cell remains constant, making it useful in protein

expression studies. In this thesis, Vinculin is used as the primary control for WB quantification

of cytosolic proteins. For mitochondrial proteins, the ubiquitous, pore-forming VDAC is used

instead.

During initial quantification, p21 was observed to fluctuate considerably. The fluctuation

may be explained by active cell cycles of dividing cells instead of exposure to ddC. For this

reason, the original experiment was repeated with the exception that the HEK cells were

confluent prior to harvesting, reducing the number of mitotic cells. Three replicates of each

timepoint were used. The harvested cells were used for an additional Western Blot detection of

p21, with Vinculin as control.

25

4.4 RNA extraction and Northern blot analysis

To determine RNA content of HEK cells during ddC exposure and recovery, the ddC exposure

and recovery experiment was repeated. The cells were lysed by removing the growth medium

and adding 1 mL of TRI-reagent. The resulting homogenate was scraped into 1 mL Eppendorf

tubes and 200 µl of chloroform was added. The samples were vortexed continuously for 10

seconds and immediately centrifuged at 12,000 g for 15 minutes at 4 °C. The upper phase was

collected into fresh tubes with 500 µl of isopropanol. After a 10-minute incubation at RT, the

samples were centrifuged at 12,000 g for 10 minutes at 4 °C. Supernatant was decanted and 1

mL of 70 % EtOH was added. The samples were vortexed and centrifuged for 5 minutes at

15,000 g at 4 °C. Ethanol was removed and the pellets were dissolved in 50 µl of purified,

sterile water. The samples were heated at 60 °C for 10 minutes prior to RNA concentration

measurement on a Nanodrop-1000 spectrophotometer.

Electrophoresis equipment was prepared with RNaseZap® RNase Decontamination

Solution and a 2 % agarose gel was prepared. The running buffer used contained 37 %

formaldehyde in 1x MOPS (3-(N-morpholino) propanesulfonic acid) solution. The

electrophoresis samples were prepared by adding 5 µg of RNA into fresh Eppendorf tubes and

equalizing the sample volumes to 5.5 µl with RNase-free water, then adding 1 µl of 10x MOPS

buffer, 3.5 µl formaldehyde, and 10 µl formamide. The samples were heated at 65 °C for 15

minutes and snap cooled on ice for 5 minutes. Meanwhile, the agarose gel was pre-run for 5

minutes at 60 V. After adding 2 µl of RNA loading dye for a total loadout volume of 22 µl, the

samples were vortexed, spun down and immediately loaded. The electrophoresis was done

overnight at 50 V.

Successful electrophoresis was confirmed under UV-light. The gel was washed with sterile

water before incubation in 20x saline-sodium-citrate buffer (SSC) for 20 minutes with gentle

agitation. The gel was capillary blotted onto a nylon membrane. The RNA was crosslinked onto

the membrane by baking it at 80 °C for 2 hours.

The crosslinked membrane was pre-hybridized in Church’s hybridization buffer for 60

minutes. A probe to detect the ND2 transcript was added and hybridization continued overnight

at 65 °C. The hybridized membrane was washed once with 5x SSC containing 0.1 % SDS and

1x SSC containing 0.1 % SDS for 5 and 20 minutes, respectively. Finally, the hybridized

26

membrane was developed using a phosphor screen. A similarly prepared probe for 18S was

used to acquire a quantification control.

4.6 DNA double-strand break experiment with transfected HEKs expressing ApaLI

To study how double-strand breaks (DSB) affect mtDNA, a stable cell line of transfected HEKs

with a gene construct for the expression of a restriction enzyme, ApaLI, was utilized. Because

ApaLI is nuclear in origin and cleaves nuclear DNA in addition to mtDNA, the gene construct

contained an attached mitochondrial targeting sequence (MTS), which transports it to

mitochondria. For easy transfection confirmation with WB detection, the gene construct

contained a Human Influenza hemagglutinin (HA) protein tag (Bayona-Bafaluy et al. 2005).

The MTS-ApaLI-HA was constructed by cleaving two plasmids with a single restriction

enzyme. One plasmid contained ApaLI and the MTS, while the other contained the HA tag.

Additionally, the plasmid containing ApaLI was cut in the ´3 direction to accommodate the HA

tag. The resulting fragments were separated by agarose gel electrophoresis, extracted and

ligated together.

The expression of ApaLI is inducible with a small concentration of doxycycline, which lifts

the repressor bound to the ApaLI operator. Active ApaLI transports inside mitochondria and

cleaves mtDNA at a specific site, generating DSBs and degrading the mtDNA.

Three replicates of transfected HEKs, each containing five timepoints and an untreated

control, were cultured. The experimental timepoints were 4, 8, 16, 24, and 48 hours of

doxycycline exposure, with no timepoints dedicated for recovery. Doxycycline stock solution

is 1000 µg/ml and 10 µl of it was diluted in 99 % EtOH for 100 µg/ml concentration. Adding

5 µl of diluted doxycycline in 10 ml of growth medium yielded a final concentration of 50

ng/ml.

The cells were collected and divided into protein and DNA fractions and prepared according

to the protocol described in chapter 3.1. Preparation and analysis of harvested samples

proceeded according to the protocols described in chapters 3.2 and 3.3, with a few exceptions.

Instead of using 1 µg of DNA for SB electrophoresis, 500 ng of DNA was used and 7S detection

was forgone. Additionally, a ND6 probe was used for hybridization instead of 12S. Western

27

blot detection included p53 and p21, along with Anti-HA for transfection verification, with

Vinculin as control for all three proteins.

5 RESULTS

5.1 Depletion of mitochondrial DNA copy number by ddC

We treated HEK cells with ddC to induce depletion of mitochondrial DNA copy number via

inhibition of DNA polymerase γ, and studied the expression of proteins associated with damage

control and DNA repair. A Southern blot analysis reveals the depletion ddC exerts on mtDNA

of HEK cells (Fig. 2).

024

48

+64

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+112

0

5 0

1 0 0

1 5 0

d d C m tD N A c o p y n u m b e r d e p le tio n

E x p o s u r e a n d re c o v e r y t im e (h o u rs )

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28

-2 0 0 -1 0 0 0 1 0 0 2 0 0

0 - 2 4

0 - 4 8

0 - + 6 4

0 - + 8 0

0 - + 1 1 2

2 4 - 4 8

2 4 - + 6 4

2 4 - + 8 0

2 4 - + 1 1 2

4 8 - + 6 4

4 8 - + 8 0

4 8 - + 1 1 2

+ 6 4 - + 8 0

+ 6 4 - + 1 1 2

+ 8 0 - + 1 1 2

9 5 % C o n fid e n c e In te rv a ls (T u k e y )

D if fe r e n c e b e tw e e n g r o u p m e a n s

C o lu m n m e a n s d iff.

****

**

**

****

***

Figure 2. Above: The level of mtDNA as percentage of an untreated control group (0-hour)

during a 48-hour exposure to ddC with two timepoints (24 and 48), followed by a subsequent

64-hour recovery period in a ddC free environment with three timepoints (+64, +80, and +112).

Below: ANOVA results displayed as 95% confidence intervals between timepoints. Statistical

significance between groups, if present, represented by stars (* = P ≤ 0.05; ** = P ≤ 0.01; ***

= P ≤ 0.001). Error bars indicate standard deviation both above and below.

Additionally, the depletion effect observed in mtDNA copy number depletion is correlated

by the depletion of 7S DNA (Fig. 3).

29

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0

5 0

1 0 0

1 5 0

2 0 0

7 s m tD N A c o p y n u m b e r d e p le tio n


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-2 0 0 -1 0 0 0 1 0 0 2 0 0

0 - 2 4

0 - 4 8

0 - + 6 4

0 - + 8 0

0 - + 1 1 2

2 4 - 4 8

2 4 - + 6 4

2 4 - + 8 0

2 4 - + 1 1 2

4 8 - + 6 4

4 8 - + 8 0

4 8 - + 1 1 2

+ 6 4 - + 8 0

+ 6 4 - + 1 1 2

+ 8 0 - + 1 1 2

9 5 % C o n fid e n c e In te rv a ls (T u k e y )

D if fe r e n c e b e tw e e n g r o u p m e a n s


****

**

**

****

***

Figure 3. Above: The level of 7S DNA as percentage of an untreated control group (0-hour)






30

5.2 Quantification of proteins of ddC-treated HEK cells

Multiple nuclear genes encode for proteins functioning outside the mitochondria in cellular

stress and damage responses. Several such proteins were quantified with Western blot analysis.

The proteins of interest were p21, p53, H2AFX, Chk1, and PKB. Vinculin was used as control

for each quantification. Some fluctuation in p21 levels was observed, and an unpaired t-test

does display a significant statistical difference between 24-hour and +64-hour timepoints (P =

0.04). However, the significance is not detectable with ANOVA (P = 0.16, Fig. 4).

31

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0

5 0

1 0 0

1 5 0

2 0 0

p 2 1 /V in c u lin


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-2 0 0 -1 0 0 0 1 0 0 2 0 0

0 - 2 40 - 4 8

0 - + 6 40 - + 8 0

0 - + 1 1 22 4 - 4 8

2 4 - + 6 42 4 - + 8 0

2 4 - + 1 1 24 8 - + 6 44 8 - + 8 0

4 8 - + 1 1 2+ 6 4 - + 8 0

+ 6 4 - + 1 1 2+ 8 0 - + 1 1 2

9 5 % C o n f id e n c e In te rv a ls (T u k e y )

D iffe re n c e b e tw e e n g ro u p m e a n s


Figure 4. Above: The level of p21 as percentage of an untreated control group (0-hour) during

a 48-hour exposure to ddC with two timepoints (24 and 48), followed by a subsequent 64-hour

recovery period in a ddC free environment with three timepoints (+64, +80, and +112). Below:

ANOVA results displayed as 95% confidence intervals between timepoints. Error bars indicate

standard deviation both above and below.

The level of Chk1, PKB, H2AFX, and p53 remained consistent throughout the experiment,

with no statistical significance between the timepoints (Figs. 5-8).

32

024

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0

5 0

1 0 0

1 5 0

C h k 1 /V in c u lin


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Figure 5. The level of Chk1 as percentage of an untreated control group (0-hour) during a 48-

hour exposure to ddC with two timepoints (24 and 48), followed by a subsequent 64-hour

recovery period in a ddC free environment with three timepoints (+64, +80, and +112).

33

024

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0

5 0

1 0 0

1 5 0

p 5 3 /V in c u lin


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Figure 6. The level of p53 as percentage of an untreated control group (0-hour) during a 48-



34

024

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0

5 0

1 0 0

1 5 0

2 0 0

H 2 A F X /V in c u lin


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Figure 7. The level of H2AFX as percentage of an untreated control group (0-hour) during a

48-hour exposure to ddC with two timepoints (24 and 48), followed by a subsequent 64-hour


35

024

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0

5 0

1 0 0

1 5 0

2 0 0

P -A k t/V in c u lin


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Figure 8. The level of P-Akt as percentage of an untreated control group (0-hour) during a 48-



In addition to non-mitochondrial protein quantification, several proteins functioning inside

the mitochondria were quantified with Western blot analysis. The proteins of interest were

MRPL11, TFAM, UQCRC2, and Parkin. Although nuclear in origin, these proteins translocate

into the mitochondria after translation. For this reason, VDAC was used as a control protein

instead of Vinculin. The levels for all proteins remained consistent throughout the experiment,

and no statistically significant differences were detected (Figs. 9-12).

36

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0

5 0

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1 5 0

2 0 0

M R P L 1 1 /V D A C


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Figure 9. The level of MRPL11 as percentage of an untreated control group (0-hour) during a



37

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0

5 0

1 0 0

1 5 0

2 0 0

T F A M /V D A C


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Figure 10. The level of TFAM as percentage of an untreated control group (0-hour) during a



38

024

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0

5 0

1 0 0

1 5 0

2 0 0

U Q C R C 2 /V D A C


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Figure 11. The level of UQCRC2 as percentage of an untreated control group (0-hour) during



39

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0

5 0

1 0 0

1 5 0

P a rk in /V D A C


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Figure 12. The level of Parkin as percentage of an untreated control group (0-hour) during a



The fluctuation of p21 levels observed during exposure to ddC are not explained by cellular

stressors alone, because p21 levels naturally change in mitotic cells during their cell cycle. To

test this, the original ddC experiment was repeated in detail, but this time the cells reached

confluence prior to harvesting, reducing the number of mitotic cells. The confluent cells have

a statistically significant difference in p21 levels between 24-hour and +112-hour timepoints (P

= 0.04, Fig. 5).

40

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0

5 0

1 0 0

1 5 0

C o n tro l fo r p 2 1 /V in c u lin


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-5 0 0 5 0 1 0 0

0 - 2 40 - 4 8

0 - + 6 40 - + 8 0

0 - + 1 1 22 4 - 4 8

2 4 - + 6 42 4 - + 8 0

2 4 - + 1 1 24 8 - + 6 44 8 - + 8 0

4 8 - + 1 1 2+ 6 4 - + 8 0

+ 6 4 - + 1 1 2+ 8 0 - + 1 1 2




*

Figure 13. Above: The level of p21 as percentage of an untreated control group (0-hour) during


recovery period in a ddC free environment with three timepoints (+64, +80, and +112). This

experiment was a replication of the original ddC exposure. Below: ANOVA results displayed

as 95% confidence intervals between timepoints. Statistical significance between groups, if

present, represented by stars (* = P ≤ 0.05; ** = P ≤ 0.01; *** = P ≤ 0.001). Error bars indicate

standard deviation both above and below.

41

5.3 Quantification of RNA levels of cells exposed to ddC

To determine whether ddC influences protein expression via interactions with RNA, the ddC

exposure and recovery experiment was replicated to collect RNA for a Northern blot. The blot

was then probed with ND2 and 18S for control. The RNA levels increase visibly within 24-

hours into ddC exposure and remain elevated throughout the experiment (Fig 6). Statistically

significant difference was found with ANOVA between 0-hour and 24-hour timepoints (P =

0.04), and 0-hour and 64-hour timepoints (P = 0.0095).

42

024

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0

5 0

1 0 0

1 5 0

2 0 0

2 5 0

d d C R N A N o rth e rn N D 2 /1 8 S


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-2 0 0 -1 0 0 0 1 0 0 2 0 0

0 - 2 40 - 4 8

0 - + 6 40 - + 8 0

0 - + 1 1 22 4 - 4 8

2 4 - + 6 42 4 - + 8 0

2 4 - + 1 1 24 8 - + 6 44 8 - + 8 0

4 8 - + 1 1 2+ 6 4 - + 8 0

+ 6 4 - + 1 1 2+ 8 0 - + 1 1 2




*

**

Figure 14. Above: The level of RNA as percentage of an untreated control group (0-hour)






43

5.4 Double-strand breaks in ApaLI induced HEK cells

A stable cell line of transfected HEK cells were used to study ApaLI induced mtDNA DSBs.

The MTS-ApaLI-HA gene construct was inducible with doxycycline to express ApaLI

restriction enzyme to interact with mtDNA. The HA-protein tag was utilized for confirmation

of transfection with Western blot analysis. A Southern blot analysis of mtDNA levels revealed

no statistically significant differences between experiment timepoints (Fig 7., P = 0.10).

0 4 816

24

48

0

1 0 0

2 0 0

3 0 0

A p a L I m tD N A D S B


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-3 0 0 -2 0 0 -1 0 0 0 1 0 0 2 0 0

0 - 40 - 8

0 - 1 60 - 2 40 - 4 8

4 - 84 - 1 64 - 2 44 - 4 88 - 1 68 - 2 48 - 4 8

1 6 - 2 41 6 - 4 82 4 - 4 8




44

Figure 15. Above: The level mtDNA as percentage of an untreated control group (0-hour)

during a 48-hour ApaLI induction with doxycycline. Below: ANOVA results displayed as 95%

confidence intervals between timepoints. Error bars indicate standard deviation both above and

below.

5.5 Protein levels of HEK cells transfected with ApaLI.

Nuclear proteins p53 and p21, involved in DNA repair and damage response, were quantified

to determine whether ApaLI was successfully transported and remained sequestered within the

mitochondria. No statistically significant differences were detected either on p53 or p21 levels,

suggesting that ApaLI remained sequestered within mitochondria and did not interact with

nuclear DNA (Figs. 16 and 17).

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2 0 0

A p a L I p 5 3 /V in c u lin


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hour ApaLI induction with doxycycline. Error bars indicate standard deviation.

45

0 4 816

24

48

0

5 0

1 0 0

1 5 0

2 0 0

2 5 0

A p a L I p 2 1 /V in c u lin


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hour ApaLI induction with doxycycline. Error bars indicate standard deviation.

6 DISCUSSION

6.1 Depletion of mitochondrial DNA

The mitochondrial toxicity of ddC is apparent by the marked decrease of mtDNA in exposed

HEK-293 cells. Inhibition of DNA polymerase γ by ddC halts replication progress, but not

DNA-unwinding helicase activity or the firing of replication forks. Fired replication forks stop

advancing and, unable to resume, collapse and cause double-strand breaks in the mtDNA,

leading to its degradation and ultimately the depletion of mtDNA copy number. A ddC

concentration of 175 µM in the growth medium is enough to deplete 70% of mtDNA in 48

hours (Fig 2).

Significant loss of mtDNA has consequences to mitochondrial function and various

biosynthetic pathways of the cell (Chandel & Schumacker 1999). However, it is possible to

46

create a HEK-293 cell line that is entirely devoid of mtDNA, but at the cost of functional

OXPHOS machinery, resulting in cells that are auxotrophic for both pyruvate and uridine

(Jazayeri et al. 2003).

In the ddC exposure and recovery experiment, HEK cells exposed to ddC recovered for 64

hours in a ddC-free growth medium. The reservoir of ddCTP-choline that accumulates during

the exposure period potentially slows the repopulation of mtDNA of HEK cells in a clean

environment by gradually transforming back into toxic ddCTP, indicated by only a modest

repopulation of mtDNA after 16 hours. Within 32 hours, however, mtDNA repopulates to

baseline level (Fig. 2).

6.2 Activation of cellular DNA repair signalling

In addition to studying ddC induced mtDNA copy number depletion, Western blot analyses of

p53, p21, and H2AFX were performed to determine whether ddC triggers signal transduction

cascades associated with key DNA repair factors or the cell cycle of affected cells.

Level of p53 remained constant throughout exposure to ddC and the subsequent recovery.

This would suggest that p21 level remained constant as well. However, there is a declining

trend in p21 levels throughout the experiment and a statistically significant difference (P < 0.04)

after 24 hours of exposure and 64 hours of recovery. The ability of p21 to act – in some cases

– independently of p53 may explain the inconsistency.

Exposure to ddC does not appear to recruit H2AFX to combat DSBs caused by stalled

replication forks. No statistical significance of H2AFX levels between different timepoints

could be elucidated by Western blot analysis. In conclusion, it appears that ddC-induced

mtDNA replication inhibition does not trigger DNA repair signalling cascades.

Furthermore, the level of MRPL11, TFAM, and UQCRC2 remained relatively stable

throughout the experiment, suggesting that mtDNA depletion did not have a significant effect

on mitochondrial gene expression.

Mitophagy is a cellular mechanism for the decommission mitochondria damaged beyond

repair. Active form of Parkin builds ubiquitin chains on damaged mitochondria, tagging them

for mitophagy. Based on the data acquired by Western blot analysis of Parkin, degradation of

mtDNA does not translate to increased mitophagy events, as no statistically significant

47

difference was found between the timepoints of ddC exposure and recovery. Reason for this

could be the failure of PINK1 to activate or translocate Parkin into the mitochondria.

Alternatively, the depletion of mtDNA copy number may not be sufficient grounds for

mitophagy, in part due to functional mtDNA remaining available to some extent.

Mitochondria likely possess compensatory mechanisms which allow them to tolerate

transient albeit severe mtDNA depletion while simultaneously maintaining mitochondrial

function and gene expression. In the ddC exposure and recovery experiment, mtDNA

repopulated to baseline level in 32 hours after more than two thirds of mtDNA was observed to

be depleted (Fig 2).

One probable way mitochondria compensate for mtDNA depletion is a temporal increase in

transcription and translation of remaining mtDNA, which appears to be the case with ddC-

induced depletion, during which a marked increase of RNA transcripts was observed for both

ddC exposure and early stages of recovery (Fig. 6). However, this increase in RNA transcript

levels did not translate to a marked increase of mitochondrial OXPHOS proteins, the levels of

which remained stable throughout the entire experiment.

An alternative explanation is that mitochondrial lifespan is extended as a response to mtDNA

depletion, but the observed increase in RNA expression does not support it, as ND2 should

behave similarly to mtDNA and get divided between daughter cells during mitosis.

A successful transfection of ApaLI was confirmed via detection of the HA-tag. Stable levels

of p53 and p21 suggest that ApaLI expression, if any, remained sequestered in the

mitochondrion.

7 CONCLUSIONS

Mitochondrial DNA replication stalling or double-strand breaks did not result in any detectable

stress in host cell, such as activation of DNA repair signalling or removal of non-functional

mitochondria by mitophagy. This indicates that acute mtDNA damage by itself is, first of all,

not signalled outside and well tolerated by the cell. Any possible deleterious effects that arise

might occur only when mitochondrial OXPHOS or other central mitochondrial functions are

impaired during more chronic stress.

48

Mitochondria lacking sufficient mtDNA copy number are nonviable, because they are missing

key RNA species and proteins for functional OXPHOS machinery. However, HEK239 cells

demonstrate robust tolerance to a severe degree of mtDNA depletion, which suggests that the

clear majority of mtDNA molecules are redundant. Furthermore, as soon as conditions turn

favourable, the remaining mtDNA is replicated and mtDNA copy number returns to baseline

level.

Just as a cell needs a certain number of functional mitochondria to survive, it could be

postulated that a mitochondrion needs a certain amount of functional mtDNA to provide the

barest minimum of gene expression for maintaining the OXPHOS machinery. Based on the

data from the ddC exposure and recovery experiment, more than two thirds of mtDNA copies

are superfluous.

Why is mtDNA so abundant? One possibility is that, because mtDNA is far more vulnerable

to inactive mutations than its nuclear counterpart, mtDNA may have evolved a strategy of

quantity over quality, thus ensuring that functional mtDNA is always available for transcription

and replication.

THANKS

I wish to thank my thesis instructors, Dr. Jaakko Pohjoismäki and Dr. Steffi Goffart for the

opportunity and guidance. Additionally, I wish to thank Rubén Torregrosa-Muñumer for always

offering hands-on assistance in the laboratory whenever it was needed. I appreciate the effort

of several friends who took the time to revise my work. Finally, I wish to thank my parents,

Leevi and Maija Haikonen, for patiently supporting my pursuit for a higher education.

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EFFECTS OF MITOCHONDRIAL DNA REPLICATION STRESS AND DOUBLE-STRAND BREAKS ON DNA … · 2018-02-28 · ITÄ-SUOMEN YLIOPISTO Ympäristö- ja biotieteiden laitos, biologia HAIKONEN,