The cellular response to alkylating agents: a complex interplay between DNA repair, oxidative signalling and

energy metabolism pathways

Ali Jbbar

Submitted for the degree of Doctor of Philosophy

October 2016

Supervisors

Dr. Ruan Elliott

FACULTY OF HEALTH AND MEDICAL SCIENCESSCHOOL OF BIOSCIENCES AND MEDICINE

Dr. Lisiane Meira

Statement of originality

I affirm that this thesis and the work to which it refers are the results of my own

efforts. Any ideas, data, images or text resulting from the work of others (whether

published or unpublished) are fully identified as such within the work and attributed

to their originator in the text, bibliography or in footnotes. This thesis has not been

submitted in whole or in part for any other academic degree or professional

qualification. I agree that the University has the right to submit my work to the

plagiarism detection service TurnitinUK for originality checks. Whether or not drafts

have been so-assessed, the University reserves the right to require an electronic

version of the final document (as submitted) for assessment as above.

Ali Jbbar

October 2016

ii

ABSTRACT

It is estimated that 10,000 lesions arise in the genome of a cell every day.

Cells have therefore also evolved ways to protect the integrity of their genomes

using direct DNA repair enzymes and multi-step pathways including base excision

repair and nucleotide excision repair. Alkylating agents are reactive chemicals that

transfer alkyl groups to biological molecules, including DNA. The base excision

repair pathway mainly repairs non-bulky lesions produced by alkylation, oxidation or

deamination of bases. This pathway is initiated by alkyladenine DNA glycosylase

(Aag).

Antioxidants neutralise free radicals including reactive oxygen specied (ROS),

and have been widely reported to protect against disease. However, some studies

have also reported that anti-oxidants may instead make disease progression worse.

This thesis aims at evaluating the role of antioxidants in the cellular response to the

alkylating agent, methylmethane sulfonate.

WT and Aag-deficient mouse embryonic fibroblasts (MEFs) were pre-treated

with the antioxidant N-acetylcysteine (NAC) and exposed to MMS. NAC increased

MMS-induced cell death in both Aag-deficient and wild-type (WT) MEFs. These were

further confirmed with embryonic stem cells (ESc) being also sensitized to MMS-

induced cell death by the anti-oxidant 2-mercaptoethanol (2-ME); and with 661W

photoreceptor cells being sensitised to MMS-induced cell death by a commercial

antioxidant mixture and NAC.

MEFs exhibited ROS generation when exposed to MMS, which was

abrogated with NAC. The mitochondrial superoxide probe MitoSox proves that the

MMS-induced ROS generation did not originate from the mitochondria. The NADPH

iii

oxidase inhibitor Diphenyleneiodonium (DPI) abrogated MMS-induced ROS

generation and also sensitised cells to MMS in a similar fashion to NAC. Collectively,

we conclude that cells generate ROS as a response to MMS treatment, and that this

ROS generation is essential for cell survival.

We also show by using different glucose concentrations, ATP levels appear

to be irrelevant to MMS-induced cell death, and that higher basal NAD levels

correlates with higher amount of MMS-induced cell death.

iv

ACKNOWLEDGMENT

I would like to pay special thanks to my supervisor Dr. Ruan Elliott for his guidance

throughout this project, and his persistence to encourage me to finish on time.

I would also like to pay an equal special thanks to my co-supervisor Dr. Lisiane

Meira for giving me this opportunity to do a PhD, as well as her guidance throughout

the project especially for the first year.

I am grateful to all my friends that that have supported me in various ways. Most

notably Dr. Fahad Alhumaydhi for his support with the mouse embryonic fibroblast

cells, and Dr. Rati Mohan for her support during the write up stage.

My greatest thanks goes to my mother Sagida Gaber, who has always stood by me,

my younger sister Maha Faraj and my younger brother Ammar Faraj.

v

ABBREVIATIONS

O6MeG O6-methylguanine

2-ME 2-mercaptoethanol

5-OHU 5-hydroxyuracil

7MeG N7-methyl guanine

Aag Alkyladenine DNA glycosylase

ALKBH Alkylation repair homolog

AP apurinic/apyrimidinic

APE-1 apurinic/apyrimidinic endonuclease 1

BER Base excision repair

CEA Chloroethylamine

CPDs Cyclobutane-pyrimidine dimers

DCF-DA Dichlorfluorescein diacetate

DMS Dimethyl sulphate

DNA-PK DNA-dependent protein kinase holoenzyme

DPI Diphenyleneiodonium

dRP 5’deoxyribose phosphate

DSBs double strand breaks

ESc Embryonic stem cells

FBS Foetal bovine serum

Fpg Formamidopyrimidine-DNA glycosylase

GG-NER Global genome nucleotide excision repair

HR Homologous recombination

LIF Leukaemia inducible factor

vi

LP-BER Long-patch base excision repair

MAM Methylazoxymethanol

MEFs Mouse embryonic fibroblasts

MGMT O6-methylguanine-DNA methyltransferase

MMR Mismatch repair pathway

MMS Methyl methanesulfonate

MNNG N-methyl-N-nitro-Nnitrosoguanidine

NAC N-Acetylcysteine

NER Nucleotide excision repair

NOX NADPH Oxidase

NHEJ Nonhomologous end-joining

Nth endonuclease III

OGG1 8-oxoguanine DNA glycosylase

PBS Phosphate-buffered saline

PI Propidium iodide

RPA Replication protein A

ROS Reactive oxygen species

SN-BER Single nucleotide base excision repair

TCR Transcription-coupled repair

TC-NER Transcription-coupled nucleotide excision repair

TMRE Tetramethylrhodamine ethyl ester perchlorate

UV Ultraviolet

UV-DDB Ultraviolet radiation-DNA damage-binding protein

WT Wild-type

XPC Xeroderma pigmentosum C

vii

viii

Contents

Statement of originality ……………………………………………………………………...i

Abstract ………………………………………………….…………………………………...ii

Acknowledgement ……………………………………….……………………………… ...iii

Abbreviations …………………………………………….………………………………....iv

1. Introduction ………………..………………………………………………………........1

1.1 DNA as the blueprints of life ……………………………………………….

…………..2

1.2 Alkylation-induced DNA lesions …………………………………….…………………

4

1.3 DNA repair of alkylated lesions

……………………………………………….............7

1.3.1 Direct reversal of DNA lesions

………………………………………………...........7

1.3.2 Non-homologous end joining ………………...……………………………….

……..9

1.3.3 Nucleotide excision repair ………………...………………………………….

…….10

1.3.4 Base excision repair ………………………………………………………….

……..12

1.4 DNA glycosylases ……………………………………………………………...

……...15

1.5 Alkyladenine DNA glycosylase ……………………………………………….

……...17

ix

1.6 DNA repair in pathology ……………………………………………………….

…......20

1.7 Hypothesis and aims ……………………………...

…………………………………..24

2. Materials and Methods …………………………………………………………....

….25

2.1 Cell culture of embryonic stem cells, mouse embryonic fibroblasts, and

661W photoreceptor cells ………………………………………………………………...26

2.2 Isolation of primary wild-type MEFs…………………………………………..………

27

2.3 Cell freezing and thawing

……………………………………………………….........27

2.4 Embryonic stem cell characterisation ……………………………..

………..............29

2.5 Cell death detection with propidium iodide

………………………………………….30

2.6 Antioxidants, NOX inhibitor and cell death

…………………………………………..31

2.7 Detection of reactive oxygen

species………………………………………………...32

2.8 Detection of mitochondrial

superoxide……………………………………………….33

2.9 Detection mitochondrial depolarisation……………………………………………...34

2.10 MEFs transfection and selection……………………………………………………34

2.11 Trypan blue cell count assay………………………………………………………..36

x

2.12 Total NAD+/NADH content quantification…………………………………………

36

2.13 ATP quantification…………………………………………………………………....37

2.14 Glucose utilisation quantification……………………………………………………

38

3.0 Effects of antioxidants on the cellular response to MMS………………………

39

3.1 Introduction……………………………………………………………………………..40

3.2.1 Phenotyping embryonic stem cells ……………………………………………...…

42

3.2.2 MMS induced cell death in wild-type and Aag-deficient ESc and

MEFs………..44

3.2.3 Antioxidant increased cytotoxicity in MMS treated

cells………………………….46

3.2.4 Antioxidant effect on NAD+/NADH levels in MMS treated

MEFs………………..55

3.2.5 Antioxidant effect on ATP levels in MMS treated

MEFs………………………….57

3.3 Discussion………………………………………………………………………………

59

4.0 Characterisation of MMS-induced reactive oxygen species generation……

65

4.1 Introduction……………………………………………………………………………..66

4.2.1 NAC depletes MMS-induced ROS generation in WT MEFs……………………68

4.2.2 MMS treatment results in ROS generation in WT but not

xi

Aag-deficient

MEFs………………………………………………………………………...71

4.2.3 Mitochondiral ROS are not elevated 1 hour post-MMS

treatment…………........72

4.2.4 NAC increased mitochondrial depolarisation in MMS treated

MEFs…………....76

4.2.5 DPI abrogates MMS-induced ROS generation in MEFs…………………………

79

4.2.6 DPI increases MMS-induced cytotoxicty in WT

MEFs…………………………...81

4.2.7 MEFs can be successfully transfected but not selected with

Geneticin or

Puromycin .............................................................................................83

4.3 Discussion……………………………………………………………………………...

86

5. Effects of glucose concentration on the cellular response to

MMS…………..92

5.1 Introduction…………………………………………………………………….

…….....93

5.2.1 MEFs proliferate slower at higher glucose concentrations…………..

…………..95

5.2.2 MEFs seeded in higher glucose exhibited higher MMS-induced

cytotoxicity…………………………………………….………………………………….. .98

xii

5.2.3 MEFs seeded in higher glucose exhibited higher basal NAD levels………...…

100

5.2.4 MEFs seeded in higher glucose exhibited higher ATP levels post MMS-

treatment…………………………………………………………………………………..102

5.2.5 MEFs did not utilize detectable amounts of glucose from the medium in the first

6 hours after medium change……………………………………………………………

104

5.3 Discussion………………………………………………………………………….....106

6.

Discussion……………………………………………………………………………..110

References……………………………………………………………………………….

117

xiii

Chapter 1: Introduction

1

1.1 DNA as the blueprint of life

Cells have complex networks and pathways that allow them to metabolise.

Metabolism is catalysed by enzymes that use their structure and functional groups to

mediate reactions. These enzymes are constantly under attack from reactive internal and

external sources that will be discussed in more depth in downstream sections. Their

structure and functional groups can therefore be compromised and render them inactive.

The living cell has therefore evolved to store information of enzymes in a more stable

structure we today call Deoxyribonuclease (DNA). DNA makes up the genome of a cell and

is therefore the blueprints of living cells, allowing the stored information to be used to

constantly restore enzyme quantity and function.

Although the DNA structure is more protected and stable it is also constantly under

attack, both from exogenous and endogenous sources. It is estimated that 10,000 lesions

arise in the genome of a cell every day [1]. Some of these lesions arise from exposure to

reactive oxygen species (ROS) generated as part of oxidative respiration or through redox-

cycling events. Reactive oxygen and nitrogen species can also be produced at sites of

inflammation by some immune cells to fight infections, both of which can damage DNA [2].

Other lesions include when cytosine is spontaneously deaminated to Uracil, or oxidatively

deaminated to 5-hydroxyuracil (5-OHU); these two lesions pair with Adenine during DNA

replication resulting in GC to AT transition mutations [1]. DNA replication could also

mismatch a nucleotide for another, leading to potential mutations and change in genetic

code. Ionising radiation as a result of radioactive decay or exposure to radioisotopes

naturally or during diagnosis and treatment of cancer generates double strand breaks

(DSBs) in DNA, and many types of damage can generate single strand breaks either

directly, or indirectly as a result of an intermediate product of a repair pathway attempting to

2

repair a lesion. Single strand breaks can lead to DSBs if left unrepaired. Contaminated food

with aflatoxins or carcinogens found in tobacco products also attack DNA [2].

DNA lesions are a serious concern to the cell because they can prevent transcription,

block replication or lead to mutations and therefore a change in the genetic code. Cells have

evolved ways to protect the integrity of their genomes using DNA repair machinery, these

include several pathways each specialised in repairing a range of lesions.

In the next section, I will specifically focus on alkylating agents and the lesions they

induce, and the different DNA repair pathways designated to repair such lesions. Later, I will

focus on the Base excision repair (BER) pathway, and end this chapter by discussing how a

miscoordinated BER can lead to cell death.

3

1.2 Alkylation-induced DNA lesions

Alkylating agents are reactive chemicals that transfer alkyl groups to biological

molecules, including DNA. They are abundant in the environment, and can also arise from

endogenous intracellular reactions. Exogenous sources of alkylating agents include abiotic

plant material and pollutants, whereas endogenous sources could arise as by-products of

oxidative reactions or from cellular methyl donors such as S-adenosylmethionine [3].

DNA is rich in nucleophilic sites, and is prone to reactions with alkylating

electrophiles to produce a diverse array of DNA adducts. Monofunctional alkylating agents

contain only one reactive site, whereas bifunctional contain two reactive sites that can react

with two separate DNA bases to form interstrand crosslinks. The lesions generated from

alkylating agents can either induce mutations, change the epigenetic programme of the cell

or be directly cytotoxic by blocking RNA and DNA polymerases [4]. Although alkylating

agents pose considerate threat to human health, they continue to be a major part of

chemotherapeutic treatment against cancer [3].

The N7 atom of guanine is the most nucleophilic region in DNA, making it the most

vulnerable site for attack by alkylating electrophiles [4]. When cells are treated with the

alkylating agents methyl methanesulfonate (MMS) or N-methyl-N-nitro-Nnitrosoguanidine

(MNNG), the predominant lesion produced is the N7-methyl guanine (7MeG; 82 and 67%

respectively) [5]. This lesion is not directly mutagenic or cytotoxic, but is very prone to

depurination, resulting in a toxic apurinic/apyrimidinic (AP) site [4].

The most mutagenic lesion arising from alkylation damage is O6-methylguanine (O6MeG),

which leads to G-A transitions. This lesion is also very cytotoxic, increasing cell death of both

primary mouse embryonic fibroblasts (MEFs) and bone marrow cells [6]. The cytotoxicity

associated with this base is due to its ability in directly blocking DNA replication or its

mispairing properties that ultimately leads to a futile cyclic activation of the MMR pathway [7,

4

8]. N3-methyladenine (3MeA) lesion is not very mutagenic, but is highly cytotoxic especially

in proliferating cells due to its ability to block DNA replication [9]. Less common lesions

include 1-methyladenine and 3-methylcytosine DNA adducts, also cytotoxic by their ability to

inhibit DNA replication [4]. However these lesions can only be formed in single stranded

DNA, as these sites are protected in double stranded DNA. They are therefore more relevant

to cells that are actively dividing, where these sites can be exposed during replication [3].

As the discussed lesions are highly mutagenic or cytotoxic, cells have evolved to

protect the integrity of their DNA using DNA repair pathways. The next section aims to

discuss the relevant repair pathways employed by the cell to repair some of the discussed

alkylating lesions.

5

Fig 1.1

Figure 1.1 Sites of alkylation on DNA bases. The N7 atom of guanine is the most nucleophilic region in DNA, making it the

most vulnerable site for attack by alkylating electrophiles. The most mutagenic lesion arising from alkylation damage is on the

O6 atom of guanine. This lesion is also very cytotoxic. Lesions affecting the N3 atom of adenine are less mutagenic than those

on O6 guanine, but N3 –adenine lesions are also very cytotoxic. Figure modified from [3].

6

1.3 DNA repair of alkylated lesions

There are different mechanisms and pathways for the repair of alkylated lesions.

These can be direct DNA repair reactions by alkylation repair homolog (ALKBH) (E.coli) and

MGMT enzymes, or multi-step pathways of the BER and nucleotide excision repair (NER)

[3]. We will discuss them in the subsequent sections.

1.3.1 Direct reversal of DNA lesions.

The first enzyme discovered able to directly reverse DNA damage was Ada in E.coli

[10]. The mammalian homologue was later discovered and named O6-methylguanine-DNA

methyltransferase (MGMT) [11]. MGMT was able to demethylate both O6-methylG and O4-

methylT, by transferring the alkyl group in a suicide reaction to one of its internal cysteine

residues, thereby inactivating the enzyme [12]. A second direct reversal enzyme was also

discovered in E.coli and named AlkB, its mammalian homologue is ABH. This enzyme is

able to demethylate 1-methylA and 3-methylC through an oxidative dealkylation reaction [1].

MGMT-deficient animals develop thymic lymphomas and lung adenomas when

treated with low dose methylating agents [13], arising from mutations induced by the O6MeG

lesion; and display an increased level of cell death in rapidly proliferating tissue such as the

bone marrow, intestine, thymus, and spleen when treated with high dose of methylating

agents [14]. The expression of MGMT is high in the liver and colon, but comparatively low in

the brain [15], suggesting an important role of this enzyme in actively proliferating tissues

and in senescent tissues with regenerative capacity. The mismatch repair pathway (MMR),

although not directly involved in repairing alkylated damage, operates in a highly co-

operative fashion with MGMT.

7

Mismatch repair (MMR) works by detecting base-base mismatches, therefore

preventing mutations to become permanent in dividin g cells [16]. The O6MeG substrate of

MGMT, if unrepaired, readily pairs with thymine during DNA replication which activates MMR

and results in the removal of the newly incorporated thymine [8]. The pairing of O6MeG with

thymine again repeats the repair, resulting in a futile MMR cycling activation [17].

Fig 1.2

Figure 1.2 Direct reversal of DNA lesions. MGMT demethylates both O6-methylG and O4-methylT, by transferring the CH3

group at the lesion to one of its internal cysteine residues, thereby inactivating the enzyme. AlkB demethylates both 1-methylA

and 3-methylC through an oxidative reaction, forcing the CH3 group to leave as CHOH. Figure taken from [3].

8

1.3.2 Non-homologous end joining

There are two main repair pathways that repair DSBs, homologous recombination (HR) and

nonhomologous end-joining (NHEJ).

In NHEJ, Ku70/80 heterodimer recognises and binds to the broken DNA ends with high

affinity [18]. The Ku70/80 binding recruits the DNA-dependent protein kinase catalytic

subunit (DNA-PKCS) and stabilises the formation of a protein complex called the DNA-

dependent protein kinase holoenzyme (DNA-PK) [19]. Once formed and bound to DNA

termini, the active DNA-PK complex phosphorylates many DNA-bound proteins and DNA-PK

brings the two DNA ends together [20]. DNA-PK also autophosphorylates itself to reduce its

affinity to DNA, thereby being released from the lesion to facilitates downstream repair [21].

Nucleases and polymerases then process the lesion by either filling or removing single

stranded overhangs. The Artemis protein possesses single strand-specific 5’ to 3’

exonuclease activity, and forms a complex with DNA-PK [22]. DNA-PK phosphorylates

Artemis to activate its endonucleolytic activity on 5’ and 3’ overhangs. The ends are finally

ligated by DNA ligase IV/XRCC4 complex in the final step. XRCC4 directly interacts with

ku70/80, therefore acts as a mediator for the recruitment of DNA ligase IV to the DSB site

[23].

9

1.3.3 Nucleotide Excision Repair (NER)

NER mainly removes bulky lesions that generally disrupt the DNA-helix structure

including the major lesions induced by ultraviolet (UV) radiation, cyclobutane-pyrimidine

dimers (CPDs) and 6-4pyrimidine-pyrimidone photoproducts (6-4PPs). It consists of two

subpathways; the first subpathway is the global genome (GG) NER, which examines the

entire genome for helix distortions; the second subpathway is the transcription coupled NER

(TC-NER), which removes damage specifically from actively transcribed genes[24]. The

difference between the two pathways is believed to be the initial recognition of damage,

where the second is initiated by RNA polymerase II [25]. It has been speculated that

transcription-coupled NER is more critical for preserving non-dividing cells such as neurons

[15].

In GG-NER, helix distorting lesions result in a single stranded DNA gap. The

xeroderma pigmentosum C (XPC) is the damage sensor and binds to the single stranded

DNA opposite the lesion strand [26]. This explains the wide lesion recognition characteristic

of XPC, as many lesions would result in the single strand gap. Some lesions however,

including UV-radiation induced lesions are poor substrates for XPC due to their mild helix

distorting properties [27]. These lesions are bound by the UV radiation-DNA damage-binding

protein (UV-DDB) and destabilised further, to enhance the XPC binding properties [28].

After binding of XPC, a lesion verification step is followed before repair is continued.

The transcription initiation factor IIH (TFIIH) complex is recruited to the XPC bound site [29],

and unwinds the DNA to extend the single strand gap using its two DNA helicases, XPB and

XPD [24]. The 5’-3’ unwinding activity of XPD is essential for NER (25), as the repair may be

aborted if no damage is detected. The replication protein A (RPA) binds to and protects the

single strand DNA opposite the lesion strand [30].

10

XPA interacts with almost all NER proteins [24], including the endonuclease XPF-ERCC1

[31], which is recruited to excise the damaged strand 5’ to the lesion [32]. XPG excises the

damaged strand 3’ from the lesion resulting in a single stranded gap of 22-30 nucleotides

[33].

The proteins involved in gap filling and ligation depend on the replicative status of the cell. In

replicating cells, DNA polymerase ε and DNA ligase I are involved, whereas in non-

replicative cells, DNA polymerase δ and XRCC1-DNA ligase III are involved [24].

Transcription-coupled (TC-) NER is initiated when an RNA polymerase II is stalled at

a lesion during transcript elongation. A stalled RNA polymerase recruits ERCC6, which binds

to the stalled polymerase, and changes the DNA confirmation around the lesion [34]. ERCC6

recruits ERCC8, and other NER proteins to the site of lesion [35]. Both ERCC6 and ERCC8

are required for the assembly of TC-NER complex [36]. Subsequent steps in TC-NER are

similar to GG-NER involving TFIIH, XPF, XPG, DNA polymerase and ligase [35].

11

1.3.4 Base Excision Repair:

The BER pathway mainly repairs non-bulky lesions produced by alkylation, oxidation or

deamination of bases. BER operates at all times irrespective of the cell cycle stage, i.e. both

proliferating and non-proliferating cells [15]; and is the main repair pathway for endogenous

base lesions, and for base lesions generated by several exogenous agents. This repair

pathway is also responsible for repairing DNA single-strand breaks, as single strand breaks

are an intermediate step in the pathway [1]. The main reaction steps of BER requires only 4-

5 enzymes, including DNA glycosylase, AP endonuclease, DNA polymerase and a DNA

ligase.

The first step is the recognition of the base lesion and the excision of the N-glycosidic bond

of the damaged base by a DNA glycosylase to generate an AP site. DNA glycosylases will

be discussed in more detail in the next section. Once the BER pathway has been initiated,

intermediate products such as AP sites and single strand breaks are more toxic than

damaged bases. The AP site is processed by AP endonuclease, which was first discovered

in E.coli as exonuclease III (Xth). It’s an AP-site specific endonuclease that cleaves the DNA

strand 5’ to the AP site generating 3’OH terminus and 5’deoxyribose phosphate (dRP)

terminus [37]. It’s also a 3’ exonuclease / DNA phosphatase, and can produce 3’ OH

terminus at strand breaks with 3’ blocked ends [1]. The mammalian homologue of Xth is

apurinic/apyrimidinic endonuclease 1 (APE-1) [38], and knocking out APE-1 gene in mice

results in embryonic lethality [39].

APE-1 also has other functions not related to BER, where its N-terminal redox domain is

dispensable for DNA repair, but regulates DNA binding of transcription factors by reducing

their Cys residues [40]. It is therefore involved in the regulation of redox-regulated

transcription of certain genes.

12

DNA polymerase β cleaves the dRP moiety produced from the activity of AP endonuclease

to generate 5’ phosphate, and fills in the single nucleotide gap. It has been reported dRP

lyase activity of DNA pol β is the rate-limiting step in the BER pathway [41], and similar to

APE-1 knock-out mice, DNA Polymerase β knock out also results in embryonic lethality [42].

The resulting nick after nucleotide incorporation is ligated by either DNA ligase I or III in the

final step [43]. This has been named single nucleotide base excision repair (SN-BER).

There is another path of BER that leaves 2-8 nucleotide gap instead of just 1, this is named

the long-patch BER (LP-BER). This sub-pathway exists due to the inability of DNA Pol-β to

remove the 5’ dRP moiety if the 3’ OH group is missing [44, 45]. In such cases, the 5’-dRP

moiety and 4-6 additional nucleotides are displaced as a single stranded DNA flap, which is

then cleaved by 5’-flap endonuclease 1 (FEN-1) [46]. Downstream steps are similar to that of

DNA replication, involving DNA polymerase δ/ε and DNA ligase I [46].

X-ray cross complementation group 1 (XRCC1) and Poly(ADP-ribose) polymerase 1

(PARP1) are mediators of BER facilitating the signalling and recruitment of BER proteins to

the site of damage. XRCC1, which has no enzymatic activity [47], binds to the nick in DNA

and acts as a scaffolding protein for recruiting BER proteins to the single strand break site

[48]. It directly interacts with NEIL-2 [49], APE-1 [50], Polβ [51] and LigIIIα [52]. It also

stimulates the activity of DNA glycosylases [53, 54] and APE-1 [50].

Poly(ADP-ribose) polymerases (PARPs) are enzymes that transfer ADP ribose groups to

target proteins. PARP family members, such as PARP-1 and PARP-2, can bind to the nicks

formed from APE-1 excisions [55]. They facilitate the access of BER proteins to the site of

damage, doing so by poly(ADP)-ribosylating several histone proteins [56]. This promotes the

recruitment of several BER proteins to the site of damage, including DNA polymerase β [57].

PARP-1 and -2 interacting with XRCC1 [58, 59], and with higher affinity when ADP

ribosylated [58, 60]. It is therefore believed PARP is used to recruit XRCC1 to site of

13

damage for the formation of repair foci [61, 62]. PARP loses affinity to DNA due to its

automodification activity, and is released to allow access of other proteins [63, 64].

Figure 1.3 Base excision repair pathway. A DNA glycosylase recognises and excises the damaged base to generate an AP

site. An AP endonuclease cleaves the DNA 5’ to the AP site to generate 3’ OH and 5’ dRP moiety. In SN-BER, DNA

polymerase β cleaves the dRP moiety to generate 5’ phosphate and fills in the single nucleotide gap. DNA ligase catalyses the

final step and joins the resulting nick. In LP-BER, the 5’ dRP moiety together with 4-6 additional nucleotides are displaced as a

single stranded DNA flap. This DNA flap is cleaved by flap endonuclease. DNA polymerase δ/ε incorporates the missing

nucleotides and DNA ligase I joins the final nick.

14

1.4 DNA Glycosylases:

Eleven DNA glycosylases have been identified in mammals, each able to recognise a limited

number of damaged bases [65]. DNA glycosylases fall into two categories, monofunctional

and bifunctional. The first DNA glycosylase discovered was the bifunctional Uracil DNA

glycosylase (UDG) in E.coli [66], which excises the N-glycosylic bond as well as cleaves the

AP DNA strand at the AP site. Alkylated bases on the other hand, are repaired by

monofunctional glycosylases and MGMT. Monofunctional DNA glycosylases excise the

damaged base leaving an AP site, which is then processed by AP endonuclease. Excision of

the damages base is usually achieved by using a water molecule as a nucleophile to attack

the Sugar C1 of the nucleotide. In contrast, bifunctional DNA glycosylases are able to further

process the AP site via β- or βδ elimination reactions, often using the N-terminal proline or a

ε-NH2 of a lysine as the nucleophile [1]. The β- reaction generates the blocking residue 3’

phospho α,β-unsaturated aldehyde (3’PUA), which is processed by APE-1 to generate 3’OH

for the recognition by Polβ. The βδ elimination reaction produces 3’ phosphate moiety, which

is processed by PNK instead of APE-1 to generate 3’OH [49]. In both cases, the 5’ terminus

is a phosphate moiety. In some cases, bi-functional glycosylases leave an intact AP site, as

seen in 8-oxoguanine DNA glycosylase (OGG1), due to a weak lyase reaction [67].

Most DNA glycosylases recognise abnormal bases in DNA with the exception of MutY in

E.coli, or its mammalian homologue MYH. MutY is able to recognise normal mismatched

bases, and excises the normal base A from AG mismatches. It also can also serve to protect

against mutations caused by oxidative stress as it also removes the normal base A from A-8-

oxoG mismatches. Deficiency in MutY leads to spontaneous CG to AT transversions [68].

DNA glycosylases specific for oxidised bases were first characterised in E.coli and are

categorised into two families, Nth (endonuclease III) and Fpg (formamidopyrimidine-DNA

glycosylase) [1]. In mammals, there are four characterised DNA glycosylases specific for

oxidised bases. The first two, NTH1 and OGG1 belong to the Nth family and excise base

15

lesions only from duplex DNA [69-71], and the other two are NEIL 1 and NEIL2, belonging to

the Fpg family, preferentially excises base lesions from single stranded DNA [49, 72]. They

are all bi-functional with broad substrate specificity.

Mouse knock-out models of OGG1, NTH1, NEIL1 and MYH DNA glycosylases are viable,

these mice did however however accumulate lesions compared to wild-type (WT)

counterparts [73-76]. Although inactivation of DNA glycosylases still gives viable mice,

inactivation of key enzymes downstream of BER leads to embryonic lethality, including APE-

1 [39]. This shows that the BER pathway is indispensable for embryonic viability, whereas

single DNA glycosylase would be dispensable for embryonic viability due to the overlap in

substrate specificity between DNA glycosylases.

16

1.5 Alkyladenine DNA glycosylase:

The full length human alkyladenine DNA glycosylase (Aag) cDNA was first isolated in 1991,

and mapped to a gene on chromosome 16 [77]. This cDNA rescued the sensitivity of AlkA-

mutant E.coli (bacterial homolog) from MMS-induced cytotoxicity. The encoded protein is 32-

kDa in size and shares extensive amino acid sequence homology with homologs of other

mammalian species, but surprisingly not with the bacterial AlkA and Tag glycosylases, or the

yeast MAG glycosylases [77]. Two other sequences were derived from the full length coding

sequence (resulting from differential RNA splicing). All three proteins excised 3-methyl

adenine and 7-methylguanine [78]. Aag can also excise mutagenic hypoxanthine lesions

from DNA that results from the deamination of adenine [79], and 1-N6-ethenoadenine (εA)

that can result from ROS [80]. It therefore has the broadest substrate specificity of all known

DNA glycosylases.

The crystal structure of Aag has been reported, and the mechanism of lesion excision occurs

by flipping the base out of the double-stranded DNA into its active site pocket, where the N-

glycosylic bond is poised for a nucleophilic attack by a water molecule bound in the active

site, thereby releasing the base [81]. Using mutational analysis, it is suggested that Aag

recognises a modified base from a normal DNA base by the shape of the damaged base, its

hydrogen-bonding interactions and aromaticity [82]. The excision of DNA lesions also

depends on the genomic context, where the rate of hypoxanthine removal is decreased 20-

fold when paired opposite cytosine versus thymine. This removal was also affected by the

type of adjacent bases [83, 84].

17

Figure 1.4 Crystal structure of AAG/ε-DNA complex. (a) The εA base (black) is flipped out of the double stranded DNA into

the proteins active site pocket. A water molecule attacks the N-glycosidic bond in the active site to release the base. (b)

Schematic diagram of contacts between AAG and the εA-DNA. The flipped-out εA base (labelled εA7) participates in hydrogen-

bonding interaction and many van der Waals interactions with the active site of the protein.It is suggested that Aad recognises

the modified base from a normal DNA base from these interactions and the aromacity of the base. Figure taken from [82].

Mouse embryonic stem (ES) cells have been doubly targeted by HR to knock out the Aag

gene. Aag-deficient ES cells had no detectable 3-MeA DNA glycosylase activity using in

vitro assays, and were more sensitive to MMS induced DNA damage [85-88]. However, it is

worth noting that in Aag-deficient ES cells, 3-MeA lesions disappear from the genome

slightly faster than would be expected by spontaneous depurination alone. This suggests

there could be another glycosylase acting on 3-MeA lesion, but to a much lesser extent [87].

The same authors also showed that 7-MeG, once thought to be a specific substrate for Aag,

was equally removed from the genome in Aag KO ES cells when compared to WT ES cells.

Two separate groups have generated Aag-deficient mice [89, 90]. Aag was found to be the

major DNA glycosylase for 3-meA, hypoxanthine and 1-N6-ethenoadenine lesions; and the

only DNA glycosylase able to excise 3-MeA and hypoxanthine lesions in the liver, testes and

18

kidney [89]. Following MMS treatment, increased persistence of 7-MeG was found in liver

sections of Aag KO mice [90], confirming previous in vitro studies.

MEFs derived from Aag-deficient mice were more alkylation sensitive compared to WT mice

[89, 90]. However, overexpressing Aag in transformed MEFs also increased their sensitivity

to alkylation compared to WT [91]. This suggests that in MEFs, any imbalance in BER

renders the cells more sensitive. Knock-down of Aag in HeLa cervical carcinoma cells and

A2780-SCA ovarian carcinoma cells by at least 80-90% also resulted in a 5-10-fold increase

in sensitivity to alkylating agents [92]. Neuronal and astrocyte cell cultures derived from the

cerebellum of Aag–deficient mice were more sensitive to the 3-MeA inducing alkylating

agent Me-Lex compared to cultures derived from WT mice [93].

Contrary to the previous reports mentioned, myeloid progenitors from the bone marrow

derived from Aag-deficient mice were more resistant than those from WT to the cytotoxic

effects of several alkylating agents. This was the first study to indicate that the initiation of

the BER by Aag was more lethal to the cell than leaving the damaged base unrepaired [94].

Again, pancreatic β-cells from Aag-deficient mice were also more resistant to the selective β-

cell genotoxicant streptozotocin that normally induces b-cell necrosis and type 1 diabetes

development in WT mice [95]. Though Aag deficiency seems to rescue certain cell types

from alkylation damage, they may acquire a mutagenic phenotype. Certainly, in splenic T-

lymphocytes, MMS treatment resulted in three- to four-fold more hprt mutations in Aag-

deficient mice compared to WT mice. Most mutations were primarily AT-TA and GC-TA

transversions, likely to be caused by 3-MeA and or 3- or 7-MeG [90].

Granule cells derived from the cerebellum of Aag-deficient mice were partly more resistant to

the methylating agents methylazoxymethanol (MAM) and nitrogen mustard (HN2) compared

to WT cultures; and significantly more resistant to dimethyl sulphate (DMS) and

chloroethylamine (CEA) [96]. Moreover, alkylation induced retinal degeneration was totally

suppressed in Aag-deficient mice in both differentiating and postmitotic retinas, when

compared to WT counterparts. Transgenic expression of Aag restores the photoreceptor

19

alkylation sensitivity in Aag-deficient animals [97]. Indeed Aag dependent alkylation-induced

tissue damage was also observed in cerebellar granule cells, splenocytes, thymocytes, bone

marrow cells, pancreatic b-cells in wild type animals, exacerbated in Aag overexpressing

animals and completely suppressed in Aag-deficient animals [98]. It is clear from the latest

studies that Aag-deficiency protects certain cell types from high levels of alkylation damage.

When cells are treated with alkylating agents at levels saturating the capacity of DNA repair,

they die mainly by necrotic cell death. A study has shown that the use of PARP Inhibitors in

combination with alkylating agents, led to necrosis induction being reduced [99]. Indeed,

alkylation induced cytotoxicity seen in wild type and Aag overexpressing animals was

abrogated in the absence of PARP-1. This shows that PARP-1 plays a crucial role in Aag-

mediated tissue damage [98].

Although alkylation induced cell death in vivo is mediated by Aag, it is important to note the

role of the enzyme in preventing cancer. In a model of inflammatory bowel disease, Aag

mediated DNA repair of reactive oxygen and nitrogen species induced DNA damage

prevented colon epithelial damage and reduced the severity of dextran sulphate sodium-

induced colon tumorigenesis [100].

20

1.6 DNA repair in pathology:

Although DNA repair generally protects against disease and genome alterations, hyper-

activation of DNA repair as a result of an imbalance between DNA lesion recognition and

repair completion, could lead to cell death. In patients that have suffered stroke or

myocardial infarction, ischemia-reperfusion can generally lead to inflammation and oxidative

damage, which results in further DNA damage and PARP-1 hyperactivation. This is followed

by a depletion of intracellular NAD+ and ATP stores, causing the cell to enter necrosis [2].

Inhibition of PARP-1 has been shown to protect against traumatic brain injury, tissue

damage caused by chronic inflammation and drug induced diabetes [101, 102]. Over-

activation of the DNA repair pathways could also further develop an atherosclerotic lesion by

senescing vascular smooth muscle cells and/or killing other vascular cells [2].

Aberrant NHEJ and NER can result in neurodevelopmental defects [103-105], and the

accumulation of DNA damage leads to ageing neurological disorders, including Parkinson’s,

Alzheimer’s and Huntington’s disease [106]. The adult nervous system is mainly made up of

post-mitotic cells that are fully differentiated and unable to proliferate. There is a small

amount of neurogenesis, but this only accounts for less than 1% of neuronal tissue [107].

DNA repair is therefore very important in the maintenance of post-mitotic neurons. The

accumulation of Age-related DNA damage also affects the expression of genes involved in

learning, memory and neuronal survival, triggering brain ageing early in adult life [108]. In

contrast to proliferating early progenitors, DNA lesions don’t induce apoptosis in mature fully-

differentiated cells [107]. Instead, it is the single and DSBs resulting from imbalanced DNA

repair pathways attempting to repair the lesion that cause cell death by hyper-activating

PARP-1. Single and DSBs can also be formed spontaneously in the brain as a result of

normal neuronal activity [109].

Defects in NER result in three inborn diseases, xeroderma pigmentosum, Cockayne

syndrome and tricothiodystrophy (TTD). All three syndromes exhibit sun sensitivity [110].

21

Xeroderma pigmentosum is characterised with a mutation in one of seven genes (XPA-

XPG). Cockayne syndrome is caused by a mutation in either the CSA or CSB gene, and is a

transcription-coupled repair (TCR) specific disorder. Cockayne syndrome does not

predispose to cancer, possibly because the TCR defects makes cells more sensitive to DNA

lesion induced apoptosis, thereby protecting against tumorigenesis. This syndrome is

associated with impaired physical and neurological development, as well as premature

ageing [110].

There hasn’t been any reported human disorders for inherited BER deficiencies. Knock out

of different glycosylases does not cause any overt phenotypes apart from a high

mutagenesis rate and cancer susceptibility. This could be explained by the redundancy of

different glycosylases. Inactivation of core proteins downstream of glycosylases causes

embryonic lethality, indicating the importance of the pathway [110].

Ataxia telangiectasia mutated (ATM) protein kinase is defective in the cancer prone, X-ray

sensitive syndrome ataxia telangiectasia. ATM is involved in the phosphorylation of histone

H2AX near DSBs to provide a local chromatin state and signal for PARP-1 and other repair

proteins [110, 111].

Acquired mutations over time play a significant role in cancer. As organisms age, mutations

accumulate which activates proto-oncogenes and inactivates tumour-suppressor genes

[110]. In replicating cells, DNA damage that cannot be repaired usually triggers cell cycle

arrest followed by cell death [110]. In some cases, specialised translesion polymerases are

able to bypass damage-induced DNA lesions at replication forks. This comes at the expense

of fidelity, potentially introducing mutations that could lead to cancers [110]. Leukemias and

lymphomas are most likely formed from chromosomal translocation, where aberrant

recombination of B-cell immunoglobin or T-cell receptor are fused with oncogenes [2, 110].

This commonly results from an imbalanced NHEJ repair pathway. Defects in MMR

22

increases mutation rates, fuelling micro-satellite instability and tumour formation [112].

Cancer treatments that utilise radiotherapy and chemotherapy highly depend on the high

proliferative rate of cancer cells combined with inactivated DNA repair pathways. This is

evident in cancers that are treated with drugs that target PARP-1 [113] . Resistant cancer

cells, especially true for cancer stem cells, usually have a high DNA repair activity which

protects them from treatment [114].

The DNA repair pathways are also used by some pathogens to fuel virulence or evolve to

evade host immune responses. Avian influenza and swine-origin viruses use error prone

mutational repair and recombination to alter their genomes, while African trypanosomes use

HR to alter their surface glycoprotein coat [2]. The NHEJ pathways is used by the herpes

simplex virus to convert its linear viral double stranded DNA into circles, an essential part of

its replication [115], and retroviruses use host DNA repair machinery to integrate their

genomes into host DNA [116, 117].

23

1.7 Hypothesis:

The study was undertaken to test the hypothesis that exposing mammalian cells to

MMS produces ROS by the action of Aag, which in-turn would be detrimental to

cells.

Aims:

1- Confirm previous studies that MMS treatment produces ROS in cells.

2- Investigate if Aag activity and therefore BER initiation, is required for ROS

production.

3- Investigate which pathways or enzymes are producing ROS post MMS-

treatment.

4- Investigate the role of antioxidants in abolishing ROS, and how this effects

cell death post-MMS treatment.

5- Investigate the energy profile in terms of NAD and ATP in cells, when treated

with antioxidants, MMS, and varying glucose concentrations.

24

Chapter 2: Materials and Methods

25

2.1 Cell culture of Embryonic stem cells, mouse embryonic fibroblasts, and 661W

photoreceptor cells

Foetal bovine serum (FBS; F0804), DMEM (D6546), penicillin/streptomycin (P4333) and L-

glutamine (G7153) were purchased from Sigma-Aldrich. Emryonic stem cells (ES cells),

MEFs and STO-1 fibroblasts cells were a kind gift from Leona Samson (Massachusetts),

661W photoreceptor cells were a kind gift from Muayyad Al-Ubaidi (University of Oklahoma).

For ES culture, 60mm dishes were coated with 3ml 1% w/v gelatin for 1 hour, and mitotically

inactivated STO-1 feeder cells were either seeded overnight or no less than 4 hours prior to

ES cell seed. ES cells were cultured over feeder cells in DMEM containing 15% v/v FBS,

2mM glutamine and 100mM 2-mercaptoethanol (2-ME, Sigma-Aldrich M3148). Medium was

changed every 24 hours and cells were passaged in a 1:3 ratio every 48 hours.

MEFs and 661W photoreceptors were cultured in T75 flasks in 10ml DMEM containing 10%

v/v FBS, 100u/ml penicillin, 100µg/ml streptomycin and 2mM L-glutamine. Medium was

changed every 48 hours and cells were passaged in 1:5-1:10 ratio every 4-5 days.

26

2.2 Isolation of primary wild-type MEFs

WT mice were weaned, and checked daily for a copulation plug. Upon detection of plug, the

embryo was recorded to be 0.5 days old. Pregnant mice were euthanized by CO2 at

embryonic day 13.5, their uterine horns were dissected and placed in DMEM with 10% v/v

FBS. In a laminar hood, individual embryos were separated and amniotic sheet was

removed. Embryos were transferred to a gelatin-coated 60mm dish and homogenised in 4ml

medium. Homogenised embryos were incubated overnight at 37°C with 5% CO2. The next

day, the supernatant was transferred to a second new dish, and a fresh 4ml medium were

added to the first dish. Both dishes were incubated again overnight. The next day, the

supernatant was aspirated, dishes were washed twice with 3ml phosphate-buffered saline

(PBS) and supplemented with 3ml medium until the allowed to grow to confluence. When

confluent, dishes were washed twice with PBS, and treated with 0.5ml trypsin to detach

them from the growth surface. Cells were then centrifuged at 300xg for 5 min, the

supernatant aspirated, and the cell pellets were resuspended in FBS to commence cell

freezing (recorded as Passage 1 at this stage).

2.3 Cell freezing and thawing

For freezing, 6mm dishes or T75 flasks were washed with 1ml or 3ml PBS, respectively.

Cells were trypsinised with 0.5ml (6mm dish) or 1ml (T75 flask) trypsin-EDTA (T4049 Sigma-

Aldrich), and the trypsin then inactivated by addition of an equal amount of medium.

Harvested cells were centrifuged at 300xg for 5min, and the supernatant aspirated. Cell

pellet were re-suspended in 1ml FBS and counted using haemocytometer. The cell

suspension was adjusted to 2x106 cells per ml. An equal amount of ice-cold freezing mixture,

consisting of 20% v/v DMSO (C6164 Sigma-Aldrich) in FBS, was added dropwise to the cell

suspension with a swirling action. 10% DMSO containing freezing mixture was used for ES

cell freezing. Cells were transferred to Cryotubes and put into a Mr. Frosty box in the -80°C

27

freezer overnight. For long-term storage, frozen cells were transferred to a liquid nitrogen

facility, while the -80°C stock provided a working batch of cells stable for a period of 3

months.

Cells were thawed in 37°C water bath, and transferred to 5ml FBS. They were centrifuged at

300xg for 5min and supernatant aspirated. Cells were resuspended very gently with a 1ml

pipette tip. This process is especially relevant to ES cells to achieve good single cell

separation without damaging their fragile membranes. The suspensions were then diluted in

5-10ml of medium. They were transferred to appropriate dishes/flasks pre-coated with 1%

w/v gelatin for 1 hour.

28

2.4 Embryonic stem cell characterisation

Embryonic stem cells (ESc) were cultured in 60mm dishes until confluent. Cells were

trypsinised with 0.5ml trypsin, and the trypsin then inactivated by addition of an equal volume

of medium (containing 15% v/v FBS). Cell suspensions were centrifuged at 300xg for 5

minutes, and the supernatant aspirated. Cell pellets were resuspended with 3ml medium,

and 1ml of the cell suspension was transferred to a 2 ml microfuge tube. The cells were

centrifuged at 300xg for 5 minutes and supernatant was aspirated. Cell pellets were

resuspended with 1.75ml PBS and 250ul of 4% paraformaldehyde was added to the cell

suspension. Cells were briefly vortexed to mix with the paraformaldehyde and allowed fix at

4°C overnight. On the following day, fixed cells were centrifuged at 300xg for 5min and the

supernatant aspirated. Cells were washed with 500µl wash buffer, consisting of 2% v/v FBS

and 0.1% v/v Triton X-100 in PBS. The cell suspension was centrifuged at 300xg for 5

minutes and supernatant aspirated. The cell pellet was resuspended in 1ml of wash buffer

and incubated for 30 minutes at 4°C to permeabilise the cells. Cell suspensions were

centrifuged at 300xg for 4 minutes and resuspended with 150µl blocking buffer, consisting of

2% v/v FBS in PBS, and 1µl of mouse anti-OCT-3/4 was added and mixed by pipetting. The

samples were incubated for 2 hours in room temperature and cells were washed twice with

500µl wash buffer, resuspended with 150µl blocking buffer with 1µl anti-mouse Dylight 488

secondary antibody. Samples were incubated for 1 hour at room temperature, and cells were

washed twice before being resuspended with 500µl PBS and analysed by flow cytometry.

29

2.5 Cell death detection with propidium Iodide

MEFs and ES cells were cultured until 80-100% confluent. The medium was aspirated and

cells were washed with PBS before being trypsinised for 0.5-1min at room temperature. An

equal volume of medium was added to neutralise the trypsin, and detached cells were

transferred to a 15ml falcon tube and centrifuged at 300xg for 5 min. The supernatant was

aspirated and cell pellet resuspended with 1ml medium. A 50µl portion of the cell suspension

was mixed with 50µl Trypan blue (T8154 Sigma Aldrich) and 10µl of this mixture was

transferred to a haematocytometer where the cells were counted under a light microscope.

The cell suspension was adjusted to 1x105/ml and 3ml of each suspension was added to

60mm tissue-culture dishes. Cells in dishes were incubated overnight in a humidified

incubator. A 100mM stock MMS solution was prepared in serum-free medium on the day of

the experiment. MMS working concentrations of 0.5mM, 1.5mM and 2.5mM were generated

by diluting the stock solution in serum-free medium. Medium in the cell dishes was aspirated,

and 3ml MMS containing medium was added for 1 hour. At the end of the 1 hour incubation,

the MMS containing medium was aspirated and replaced with 3ml of normal medium and

cells were incubated for an additional 24 hours.

For glucose experiments, MEFs were seeded at 1.5x105 or 3.0x105 cells per 60mm dish, in

low, normal and high glucose medium overnight; and treated with 2.5mM MMS in serum free

medium the next day for an hour. At the end of the 1 hour exposure, the MMS containing

medium was aspirated and cells were incubated with low, normal and high glucose for an

additional 24 hours.

On the day of analysis, the supernatant in the dishes was transferred to a Falcon tube, and

the cells were washed with 0.5ml PBS, which was also transferred to the same Falcon. Cells

were then trypsinised with 0.5ml trypsin for 0.5min and the trypsin then neutralised by

addition of the same volume of medium. The Cell suspensions were also transferred to the

corresponding falcon tubes containing the original medium and PBS wash. The cell

suspensions were then centrifuged at 300xg for 5min. The supernatant was aspirated and

30

cells were washed twice more each time with 1ml PBS. Each cell pellet was finally

resuspended with 0.5ml PBS, and 5µl of propidium iodide (P4864, Sigma-Aldrich, working

concentration 10µg/ml) was added to the sample and vortexed briefly. Samples were

analysed with flow cytometer using the PE channel.

2.6 Antioxidants, NOX inhibitor and cell death

ES cells were cultured under normal conditions as described above until confluent. Cells

were then trypsinised and harvested as previously, and counted using a haematocytometer.

A total of 3x105 cells were seeded into each 60mm dish in 3ml of DMEM (containing 15% v/v

FBS and 2mM glutamine) with or without 100mM 2-ME and incubated overnight. The next

day, cells were treated with 0.5mM, 1.5mM or 2.5mM MMS in serum-free medium or just

serum-free medium as control for 1 hour. MMS medium was replaced with medium

containing with or without 100mM 2-ME and cells were incubated for an additional 24 hours.

Cells were trypsinised and harvested as previously, and cell death analysed by flow

cytometry as described in section 2.5.

MEFs and 661W cells were cultured as normal until confluent. The cells were then

trypsinised and harvested as described above, and counted using a haematocytometer. A

total of 3x105 cells was seeded into each 60mm dish in 3ml DMEM (containing 10% v/v FBS

and 2mM glutamine) and incubated overnight. The next day, fresh NAC (NAC; A7250

Sigma-Aldrich) stock solution of 50mM was made by adding 163.19mg NAC to 2ml

autoclaved H2O and 18ml normal medium. This solution was sterilised by passing through a

0.2µm filter and a working solution of 15mM NAC was produced when required. Medium in

dishes was aspirated and replaced with medium containing 15mM NAC or control medium

without NAC, and incubated for 30 minutes. Medium was aspirated and cells were treated

with MMS in serum-free medium or just serum-free medium as control for 1 hour. MMS was

aspirated and cells were returned into medium containing 15mM NAC or control medium

31

without NAC, and incubated for 24 hours. Cells were then trypsinised and harvested as

previously, and cell death analysed as described in section 2.5. The same protocol was

employed for the Sigma antioxidant mix (Sigma AO; A1345 Sigma-Aldrich) at concentrations

of 1x, 10x, 50x and 100x, and for the NOX inhibitor diphenyleneiodonium chloride (D2926

Sigma-Aldrich) at a final concentration of 2µM.

2.7 Detection of reactive oxygen species

Confluent MEFs were trypsinised and harvested as described above. Cells were centrifuged

at 300xg for 5 minutes. The supernatant was aspirated and the cell pellets were

resuspended in 1ml medium. Cells were counted as previously and 3x106 cells were added

to each 60mm dish in 3ml medium and incubated overnight. The next day, fresh 100mM

MMS and 15mM NAC solution were prepared. Samples were treated on a time-line, starting

with the last time point first, and moving down until the first time point, thereby having all the

time points on the same plate at the end of the experiment for analysis. For experiments

involving NAC or DPI, medium was aspirated and 15mM NAC, 2µM DPI or normal medium

were added to cells for 30min. NAC or DPI containing medium was aspirated and cells were

treated with MMS for 1 hour in serum-free medium. MMS was aspirated and cells were

returned back into medium containing NAC, DPI or normal medium until the final time-point

was reached. Control cells at time-point 0 hours were also pre-incubated with NAC, DPI or

normal medium for control for 30 minutes. The medium was aspirated, cells were washed

with 0.5ml PBS, and trypsinised with 250µl trypsin until the cells detached. The trypsin was

neutralised by addition of 250µl of medium and the cell suspension were centrifuged at

300xg for 5min. The cells were washed once with 0.5ml PBS and resuspended in 0.5ml

PBS. 2’, 7’-dichlorofluorescein diacetate (DCF; 35845 Sigma-Aldrich) was made fresh at

1mM concentration just before use, and 10µl of this solution was added to each 0.5ml

sample to give a final concentration of 20µM DCF. Samples were incubated at 37°C for 30

32

minutes, and subjected to flow cytometric analysis using the FITC channel for detection of

fluorescence.

2.8 Detection of mitochondrial superoxide

Confluent MEFs were trypsinised and harvested as described above. Cells were centrifuged

at 300xg for 5min, supernatant aspirated and the cell pellets were resuspended in 1ml

medium. Cells were seeded at 3x106 into 60mm dishes in 3ml medium and incubated

overnight. The next day, fresh 100mM MMS and 15mM NAC solutions were prepared.

Samples were treated on a time-line starting with the latest time first, and moving down until

control samples at 0 hours. To start, the medium was aspirated and 15mM NAC or normal

medium was added to cells for 30 minutes. The medium was then aspirated and the cells

were treated with MMS for 1 hour in serum-free medium. The MMS-containing serum free

medium was then aspirated and cells were returned to medium containing NAC or normal

medium until the final time-point was reached. Control cells at time-point 0 hours were also

pre-incubated with NAC-containing normal medium for control for 30 minutes.

Following the treatments, the medium in the dishes was transferred to Falcon tubes. The

cells were washed with 0.5ml PBS and the PBS wash solutions were also transferred to the

same respective Falcon tubes. The cells were then trypsinised with 0.5ml trypsin for 0.5

minutes and the trypsin neutralised by addition of 0.5ml medium. The detached cells were

transferred to the Falcon tube containing the corresponding original medium and PBS wash.

The cell suspensions were centrifuged at 300xg for 5min. The supernatants were aspirated

and cells were transferred to FACS tubes and further washed twice with 1ml PBS. Each cell

pellet was finally resuspended with 0.5ml PBS, and 5µl of MitoSOX (M36008 Molecular

Probes; Stock concentration 0.5mM) was added to each tube to give a final MitoSOX

concentration of 5µM. Tubes were incubated at 37°C for 30 minutes, and analysed with a

flow cytometer using the PE channel.

33

2.9 Detection of mitochondrial depolarisation

Confluent MEFs were trypsinised and harvested as described above. Cells were centrifuged

at 300xg for 5min, the supernatant aspirated and the cell pellet was resuspended in 1ml

medium. Cells were seeded at 3x106 to 60mm dishes in 3ml medium and incubated

overnight. The next day, fresh 100mM MMS and 15mM NAC solution were prepared.

Samples were treated on a time-line starting with the latest time first, and moving down until

control samples at 0 hours. To start the treatments, the medium was aspirated and medium

containing 15mM NAC or normal medium was added and the cells incubated for 30 minutes.

The medium was then aspirated and cells were treated with MMS for 1 hour in serum-free

medium. After the 1 hour treatment, the MMS-containing medium was aspirated and cells

were returned to medium containing NAC or normal medium until the final time-point was

reached. Control cells at time-point 0 hours were also pre-incubated with NAC containing

normal medium for 30 minutes.

A 200nM working solution of tetramethylrhodamine ethyl ester perchlorate (TMRE; 87917

Sigma-Aldrich) in medium was prepared from a 1mM stock solution. A volume of 1ml of the

working solution was added to the 3ml medium already in the dishes containing the cell and

the cells were incubated for 20 minutes. The cell medium was transferred to tubes, and cells

were trypsinised and added to the cell medium already in in tubes so that both adherent and

any floating cells would be combined in the analysis. The cell suspensions were centrifuged

at 300xg for 5min and the cells washed with 0.5ml PBS. Each cell pellet was resuspended in

0.5ml PBS and samples analysed by flow cytometry using the PE channel.

2.10 MEFs transfection and selection

Transfection conditions were first optimised using a pcDNA3-EGFP plasmid (13031

Addgene). WT MEFs were cultured until confluent and harvested as described above. Cells

were seeded in a black tissue-culture 96-well plate (CLS3603-48EA Sigma-Aldrich) at two

34

different cell densities; 1x103 and 2x103 cells per well. Isolated GFP plasmid was quantified

using a Nanodrop spectrophotometer and master mixes were made for 0.2µg and 0.4µg of

plasmid in 50 µl of serum-free medium for each well. Master mixes of lipofectamine reagent

(11668027 ThermoFisher Scientific) were made in a separate tubes for 0.25µl, 0.5µl and 1µl

of reagent in 50µl of normal medium excluding antibiotics for each well. The plasmid and

lipofectamine master mixes were mixed in different combinations to produce the following

ratios of plasmid to lipofectamine in a final volume of 100µl of medium excluding antibiotics

(Plasmid: lipofectamine per well; 0.2µg: 0.25µl; 0.2µg: 0.5µl; 0.4µg: 0.25µl; 0.4µg: 0.5µl;

0.4µg: 1µl). The different mixtures were incubated for 5 minutes at room temperature.

Control wells either had no GFP plasmid or no lipofectamine reagent. After the 5 minutes

incubation, the medium in wells containing the cells was aspirated and 100µl of various

combinations of transfection medium was added to wells for 48 hours. Transfection

efficiency based on the detection of GFP expression measured using a plate reader with

excitation/emission at 475/509nm.

Puromycin selection concentration was optimised using a kill curve. For this, 5x103 cells

were seeded per well in a 96-well plate and incubated overnight. Cells were then variously

treated with 0.1 µg/ml, 0.5 µg/ml, 1 µg/ml, 5 µg/ml and 10 µg/ml puromycin normal medium.

Cells were analysed under the microscope 3 days and 7 days post-treatment for viability.

For NOX shRNA (TRC ShRNA, ThermoFisher Scientific) transfection, MEFs were seeded at

2x103 cells per well in a 96-well plate and incubated overnight. The following day, 0.4µg of

purified plasmid was mixed with 1µl of lipofectamine reagent in 100µl medium (containing

5% v/v/ FBS). The medium in the wells was aspirated and the transfection medium was

added for 48 hours. At the end of the 48 hours, the transfection medium was replaced with

normal medium containing 1µg/ml puromycin for selection for 1 week.

35

2.11 Trypan blue cell count assay:

MEFs were cultured until 80-100% confluent. Cells were trypsinised and harvested as

described above. Cells were counted with a haematocytometer under the light microscope

and seeded at 1.5x105 or 3x105 cells per 60mm dish. Cells were harvested 24 hours or 48

hours after seeding by trypsinisation and total cell population was counted three independent

times to achieve three experimental replicates with trypan blue using the haemocytometer.

2.12 Total NAD+/NADH quantification

MEFs were cultured until 80-100% confluent. The medium was aspirated and cells were

harvested by trypsinisation as described above. The cell suspensions were centrifuged at

300xg for 5 min and counted using a haematocytometer. A total of 1x104 cells in 100µl

medium was seeded into each well of 96-well plates and the cells incubated overnight.

For MMS and NAC experiments, the medium was aspirated from the wells the next day and

the cells were pre-incubated with normal medium or medium containing 15mM NAC for 30

minutes. At the end of the 30 minutes, the medium was aspirated and 2.5mM MMS in serum

free medium was added for 1 hour. The MMS-containing medium was then was aspirated

and cells were returned to normal or NAC-containing medium until the final time point.

For glucose experiments, MEFs were seeded in low (1mM), normal (5.5mM) and high

(30mM) glucose medium and incubated overnight. The following day, the medium was

aspirated and the cells treated with 2.5mM MMS in serum free medium for 1 hour. The

MMS-containing medium was then aspirated and cells were returned to low, normal or high

glucose medium until the final time point.

Upon reaching final time-point, medium was aspirated, cells were lysed and total

NAD+/NADH content was measured using NAD/NADH Cell Based Assay Kit (Cayman

36

600480) according to manufacturer’s protocol. The plate was read with a plate reader at

450nm.

2.13 ATP quantification

MEFs were cultured until 80-100% confluent. The medium was aspirated and cells were

harvested by trypsinisation as described above. Cells were centrifuged at 300xg for 5 min

and counted using a haemocytometer. A total of 1x104 cells in 100µl normal medium was

seeded into each well of black 96-well plates and the cells incubated overnight.

For MMS and NAC experiments, the medium was aspirated the next day and cells were pre-

incubated with normal medium or medium containing 15mM NAC for 30 minutes. After the

30 minutes the medium was aspirated and 2.5mM MMS in serum free medium was added

for 1 hour. The MMS-containing medium was ten aspirated and cells were returned to the

normal or NAC-containing medium until the final time point.

For glucose experiments, MEFs were seeded in low, normal and high glucose medium and

incubated overnight. They were then treated with 2.5mM MMS in serum-free medium for 1

hour. The MMS-containing medium was then was aspirated and cells were returned to low,

normal and high glucose until the final time point.

Upon reaching final time-point, the medium was aspirated, and wells were washed with

100µl PBS. Cells were lysed in wells with 100µl of boiling water in an oven at 95°C for 10

minutes. A 10µl portion of each cell lysate was taken for ATP detection using the ATP Assay

Kit (Abcam ab83355) using the fluorometric detection method performed according to the

manufacturer’s instructions. The plate was read using a plate reader with Ex/Em 535/587

nm.

37

2.14 Glucose utilisation quantification

MEFs were cultured in low, medium and high glucose until 80-100% confluent. The medium

was aspirated and cells were harvested by trypsinisation as described above. Cells were

centrifuged at 300xg for 5 min and counted using a haemocytometer. MEFs were seeded at

3x105 cells in 3ml of their respective low, medium or high glucose culture medium in 60mm

dishes overnight. The next day the medium was aspirated, and fresh medium was added.

Samples (100µl) of the culture media were taken from the dishes at several time points and

used for glucose quantification using a glucose assay kit (Abcam ab65333) according to

manufacturer protocol.

38

Chapter 3: Effects of antioxidants on the cellular response to

MMS

39

3.1 Introduction:

Antioxidants neutralise free radicals including ROS, and have been widely reported to

protect against disease [118-120]. However, some studies have also reported that anti-

oxidants may instead make disease progression worse. Precancerous regions in mice

progressed quicker when mice were supplemented with anti-oxidants vitamin E and N-

Acetylcysteine (NAC) in their diet [121]; and daily supplementation with antioxidant in healthy

individuals correlated with higher incidence of lung cancer [122].

The molecular explanation for the apparent adverse effects of antioxidants has focused on

nuclear factor-erythroid 2-related factor 2 (NRF2), a transcription factor that induces the

expression of antioxidant and detoxification genes in response to redox stress [123]. This

transcription factor was shown to be mutated in 10-25 % of cancer (lung,and head and neck)

leading to its constant activation, which correlated with poor prognosis [124]. Its negative

regulator Kelch-like ECH-associated protein 1 (KEAP1) was mutated in 19% of non-small

cell lung cancers [125]. The correlation of permanent activation of NRF2 with poor prognosis

has therefore added to the speculation that enhanced antioxidant activity may in some

circumstances be pro-mutagenic.

How the intracellular redox state affects the DNA damage response is less clear. The DNA

damage response is nomally inactivated in cancer cells allowing accumulation of more

mutations and faster disease progression [126], and whether it is disrupted by a change in

cellular redox remains to be fully elucidated. One study used a high throughout genotoxicity

assay to identify several antioxidants as inducers of the DNA damage response and cell

death [127]. Another study investigated the role of pro-oxidant on cellular response to the

alkylating agent MMS. The authors reported pro-oxidants rather than anti-oxidants increased

survival of yeast cells upon moderate MMS treatment [128]. The level of damage also plays

a role, as the authors further showed that on exposure to extremely high MMS levels, pro-

oxidants were detrimental to cells. These studies suggest that that an oxidative environment

is more favourable than a reducing environment in surviving genetic insults. Two studies

40

have shown treatment with pro-oxidant prior to alkylation results in higher APE-1 activity,

which further enhanced DNA BER, strengthening the hypothesis that intracellular redox state

does play a role in determining cellular survival to alkylating damage [129, 130]. XRCC1,

another protein also involved in BER, has been shown to be redox regulated [131], and the

oxidised form binds to polymerase β with higher affinity through its redox-sensitive binding

interface in the N-terminal domain. A constant reduced form of XRCC1 fails to fully rescue

mouse fibroblasts upon alkylation, in comparison to WT XRCC1 [132].

There is therefore a complex relationship between intracellular redox state and the DNA

damage response, which potentially explains the role of antioxidants in aberrant NRF2

activation, disproportionate APE-1 and XRCC1 activation, in both cancer progression and

cellular response to DNA insults.

The studies described in this chapter evaluates the role of antioxidants in the cellular

response to the alkylating agent, MMS, and how this associates with the cellular energy

levels NAD+/NADH and ATP.

41

3.2.1 Phenotyping embryonic stem cells

ESc express markers that keeps them in an undifferentiated state [133]. We cultured ESc on

top of mitotically inactivated STO-1 feeder cells. Feeder cells provide growth factors

including leukaemia inducible factor (LIF), necessary to keep ESc in their undifferentiated

state [134]. Mitotic inactivation was achieved either by irradiation or treatment with

mitomycin-c. ESc are smaller in size and we were able to phenotype them using Flow

Cytometry, using gating based on forward scatter versus side scatter plots to separate them

from the larger more granular feeder cells (ESc gated red in figure 3.1).

An antibody to OCT3/4 was used to confirm the undifferentiated state of the ESc. Both WT

and Aag-deficient genotypes were greater than 95% positive for OCT3/4 (Fig 3.1),

confirming they are pluripotent and in an undifferentiated state.

42

Fig 3.1

WT control Aag-deficient control

WT OCT3/4 Aag-deficient OCT3/4

Figure 3.1. Flow cytometric analysis of WT and Aag deficient ESc stained for OCT3/4 pluripotency marker. WT and Aag

deficient cells were grown on top of STO-1 feeder cells. Cells were harvested by trypsinisation, fixed with 2%

paraformaldehyde, and permeabilised with 0.1% triton X-100. Cells were incubated with OCT3/4 antibody (1:250) for 1 hour at

RT, and secondary FITC-conjugated antibody (1:500) for 1 hour at RT. Cells were washed with and resuspended in PBS prior

to flow cytometry analysis. Red gated cells represents ESc. WT and AAG-deficient controls are top plots. WT and AAG-

deficient cells stained with OCT 3/4 are bottom plots. FITC-A channel is used to measure FITC fluorescence. More than 95% of

both the WT cells and Aag-deficient cells stained with OCT 3/47 shifted to the right in the FITC-A channel, indicating staining

positive for OCT3/4. Plots are a representative of a screening experiment performed to confirm ESc phenotype.

43

3.2.2 MMS induced cell death in wild-type and Aag-deficient ESc and MEFs

ESc were seeded at 0.3x 106 per 6cm tissue culture dish, and treated with 1 hour MMS in

serum free medium. MMS concentrations used were 0.5mM, 1.5mM and 2.5mM. After the 1

hour exposure, the MMS was removed and replaced with normal medium. The next day, the

cells were harvested and incubated with Propidium iodide (PI) prior to analysis. PI is a

fluorescent molecule that intercalates with DNA. The dye is impermeable to the cell

membrane and is therefore excluded from viable cells. Therefore only dead or dying cells

with perturbed cellular membranes take up the dye and stain positive.

The control population showed minimal amount of cell death, and both WT and Aag-deficient

cells treated with 0.5mM MMS showed no apparent increase in cell death compared to

control (Fig 3.2 A). At the 1.5mM MMS concentration, there were a clear increase in cell

death in both genotypes compared to control untreated cells (p<0.0005), however there

were no difference in cell death between the Aag-deficient And WT cells. The difference

between Aag-deficient cells and WT cells was only evident when 2.5mM MMS was used

(p<0.0005). Aag-deficient cells were more sensitive to MMS than WT at this concentration,

which is in agreement with published literature [85].

MEFs were also used, seeded at 0.3x 106 cells per 6cm dish and treated with 1 hour of the

same MMS concentrations the next day. As with the ESc, Aag-deficient MEFs were also

more sensitive compared to WT. This difference was evident at a lower MMS dosage

(1.5mM; p<0.05) compared to ESc (Fig 3.2 B). At the 0.5mM MMS dosage, there were no

significant differences between the two genotypes or when comparing to untreated control.

These results indicate 0.5mM MMS is not enough to induce cell death in the cell types used.

Based on these observations, the 2.5mM concentration was selected for use in most

subsequent experiments. Having confirmed that the Aag knock-out genotype was more

sensitive to MMS compared to WT, we next wanted to test how this cellular sensitivity to

MMS would change under antioxidant treatment.

44

Figure 3.2

A

Control

0.5mM M

MS

1.5mM M

MS

2.5mM M

MS0

10

20

30

40

50

% C

ell D

eath

WT EScAag deficient ESc

****

B

Figure 3.2. Aag deficient mouse ESc and MEFs are more sensitive to MMS than Wild-type. Cells were seeded overnight

and treated with different concentrations of MMS for 1 hour. The next day they were harvested and cell death was detected

with PI using flow cytometry. (A) Wild type and Aag-deficient ESc exposed to control medium or medium containing 0.5mM

MMS, 1.5mM MMS, and 2.5mM MMS. Aag-deficient ESc were more sensitive to MMS than WT counterparts when exposed to

2.5mM MMS. No difference in cell toxicity was observed in MMS concentration below 2.5mM. (B) Wild type and Aag-deficient

MEFs exposed to control medium or medium containing 0.5mM MMS, 1.5mM MMS, and 2.5mM MMS. Aag-deficient cells were

more sensitive to MMS treatment than WT MEFs when exposed to 1.5 mM and 2.5mM MMS concentration. Data shown are

the result of three independent treatments (Two-way ANOVA Tukey’s post hoc test, mean ± SEM; ** p<0.05, **** p<0.0005).

45

Control

0.5mM M

MS

1.5mM M

MS

2.5mM M

MS0

10

20

30

40

Cel

l Dea

th %

WT MEFsAag-deficient MEFs****

**

3.2.3 Antioxidant increased cytotoxicity in MMS treated cells

We next investigated how antioxidant treatment would affect the cellular response to MMS

treatment. We achieved this by the addition of antioxidants in the medium the day before the

cells were treated. WT MEFs were seeded overnight in normal medium, and the next day

the medium was changed with either normal medium or 15mM NAC-containing medium 30

minutes prior to treatment. Cells were then exposed for 1 hour to MMS in serum free

medium without NAC, after which medium was again replaced with either normal medium or

medium containing NAC for another 24 hours. Cells were harvested and analysed for cell

cytotoxicity using PI.

Treatment with 15mM NAC alone had no detectable cytotoxic effects (Fig 3.3 A). There was

also no difference in cytotoxicity in cells treated with 0.5mM MMS compared to controls,

agreeing with the previous results described in section 3.2 above, and NAC did not change

cellular response at this MMS concentration. For the WT MEFs in normal medium, treatment

with 1.5mM MMS did not significantly affect cell viability compared to controls, in line with

previous results. However, there was a significant increase in cell death in the WT MEFs

treated with NAC and exposed to 1.5mM MMS. This difference was even more marked at

2.5mM MMS, where MMS treatment caused 15% cell death in WT cells pre-treated with

normal medium, compared to 80% cell death in cells pre-treated with the antioxidant

(p<0.0005). Aag-deficient MEFs followed the same pattern, where cells pre-treated with

NAC exhibited increased cell death following MMS treatment, with the effect being

statistically significant for MMS concentrations as low as 0.5mM (p<0.5, Fig 3.3 B). These

striking observations, indicate that an antioxidant rich environment correlated with higher

MMS-induced cell death. Before attempting to determine how NAC treatment caused the

increased cell death in MMS-treated cells, we wanted to first confirm these results with other

cell lines and antioxidants.

46

Figure 3.3

A

Control

0.5mM M

MS

1.5mM M

MS

2.5mM M

MS0

20

40

60

80

100

Cel

l Dea

th %

WT MEFsWT MEFs + NAC

****

**

B

Figure 3.3. NAC exacerbates

MMS-induced cytotoxicity in

both WT and Aag-deficient MEFs. Cells were seeded overnight and pre-treated either with normal medium or NAC-containing

medium before being exposed to different concentrations of MMS for 1 hour. The next day they were harvested and cell death

was detected with PI using flow cytometry. (A) Wild type MEFs exposed to control media, 0.5mM MMS, 1.5mM MMS, and

2.5mM MMS with and without NAC. WT MEFs were more sensitive to MMS when seeded in NAC medium, compared to normal

medium. This was observed in 1.5mM MMS and up. (B) Aag-deficient MEFs control media, 0.5mM MMS, 1.5mM MMS, and

2.5mM MMS with and without NAC. Aag-deficient cells showed same response, cells seeded in NAC medium showed MMS

sensitivity in concentrations as low as 0.5mM MMS, while such concentration had no cytotoxicity in cells seeded in normal

medium. Data shown are the result of three independent treatments (Two-way ANOVA Tukey post hoc test, mean ± SEM; *

p<0.5, **** p<0.0005).

47

0

20

40

60

80

100

Cel

l Dea

th %

Aag-deficient MEFsAag-deficient MEFs + NAC

*

****

****

To verify that the increase in MMS-induced cytotoxicity by NAC was not only specific to the

antioxidant and/or cell type used, we examined the role of the antioxidant 2-ME on ESc

MMS-induced cell death.

ESc are normally grown in vitro in the presence of 0.1mM 2-ME. This reducing environment

mimics the partial hypoxic conditions of the embryo by stabilising HIF-1α, responsible for

maintaining the pluripotency ESc [135, 136]. As ESc proliferate rapidly, they also produce

high level of ROS, and an appropriate concentration of antioxidants to mimic physiological

level of ROS have reported to reduce unwanted genomic alterations when these cells are

cultured outside their native environment [137]. To investigate how antioxidants affect ESc

response to MMS, 2-ME was removed from the culture medium. It is worth emphasising that

this experimental strategy differed from the experiments described above in that in this case

an anti-oxidant normally included in the standard growth medium, based on observed

beneficial effects on ESc normal growth, instead of adding an additional antioxidant to the

medium as described in the experiments above.

The removal of 2-ME from the medium, increased intracellular ROS levels in both WT and

Aag-deficient ESc (Fig 3.4).

48

Fig 3.4

ESc WT + 2-ME ESc WT - 2-ME ESc Aag-deficient + 2-ME ESc Aag-deficient - 2-ME

Figure 3.4. Flow cytometric analysis of intracellular ROS in WT and Aag KO ESc with or without 2-ME. WT and Aag

deficient ESc were cultured on top of STO-1 feeder cells. Cells were harvested by trypsinisation, and incubated with DCF for 30

minutes at 37°C. Cells were washed with and resuspended in PBS prior to flow cytometry analysis. Red gated cells represents

ESc. The removal of 2-ME from the medium increased intracellular ROS levels in both WT and Aag-deficient ESc, as seen by a

right shift in histograms in FITC channel. Plots are a representative of a screening experiment performed.

49

WT ESs seeded in the presence of 2-ME, and treated with MMS, did not show any

difference in cell death compared to those seeded without 2-ME. This could be explained by

the low 2-ME concentration used (Fig 3.5 A). The Aag-deficient genotype, followed a similar

pattern where cells seeded with 2-ME were more sensitive to MMS than those seeded

without 2-ME, with the difference achieving statistical significance at 2.5mM MMS (Fig 3.5 B,

P < 0.005). Thus in line with the MEF results, the antioxidant-driven increase in MMS

cytotoxicity is more evident in the Aag-deficient genotype.

50

Figure 3.5

A

Control

0.5mM M

MS

1.5mM M

MS

2.5mM M

MS0

10

20

30

% C

ell D

eath

WT ESs -2MEWT ESs +2ME

B

Figure 3.5. 2-ME increases Aag-deficient embryonic stem cell sensitivity to MMS. ESc were seeded overnight either with

or without 2-ME before being exposed to different concentrations of MMS for 1 hour. The next day they were harvested and cell

death was detected with PI using flow cytometry.(A) Wild type ESc exposed to control medium, 0.5mM, 1.5mM, and 2.5mM

MMS. There were no statistical significance between WT ESc seeded with or without 2-ME and treated with MMS (B) Aag-

deficient ESc exposed to control medium, 0.5mM, 1.5mM, and 2.5mM MMS. Aag-deficient cells were more sensitive to when

treated with 2.5mM MMS and seeded in 2-ME as compared to cells seeded without 2-ME. No difference was seen inlower

MMS concentrations. Data shown are the result of three independent treatments (Two-way ANOVA, Tukey’s post hoc test,

mean ± SEM, *** p<0.005).

51

Control

0.5mM M

MS

1.5mM M

MS

2.5mM M

MS0

10

20

30

40

50

% C

ell D

eath

Aag deficient ES -2MEAag deficient ES + 2ME

***

Antioxidant mediated increases in MMS cytotoxicity were further evaluated using a third cell

type, the murine photoreceptor cell line 661W pre-treated with either NAC or a commercial

mixture of antioxidants. MMS-induced cell cytotoxicity increased following pre-treatment with

15mM NAC, and also increased in a concentration-dependent manner with the commercial

antioxidant mix (Fig 3.6). It is worth noting that even at the lowest concentration of the

commercial antioxidant mix used here (1x), the antioxidants did not rescue the cells from

MMS cytotoxicity.

52

Fig 3.6

Figure 3.6. Commercial antioxidant mix increases cell sensitivity to MMS in a concentration dependent manner. WT

MEFs were seeded overnight and pre-treated with a commercial mixture of antioxidants before being exposed to 2.5mM MMS

for 1 hour. The next day cells were harvested and cell death was detected with PI using flow cytometry. There were no

difference in cell death between control cells pre-treated with normal medium, different concentration of commercial antioxidant

mixture or NAC. Cell pre-treated with commercial antioxidant mixture exhibited an antioxidant concentration-dependent

increase in sensitivity to MMS. Data shown are a representative experiment of three independent experiments.

53

Having confirmed the consistent effects of antioxidants on MMS-induced cytotoxicity in a

selection of different cell lines, we next wanted to investigate if these effects were due

antioxidant change of cellular energy levels. MMS treatment have shown to reduce both total

NAD+/NADH and ATP levels in cells, and this reduction in energy availability forces the cell

into cell death. We wanted to investigate whether pre-treatment with the antioxidant NAC

had any effect on total NAD+/NADH and ATP levels, either pre-exposure or post-exposure

to MMS. From here on, we chose the MEF cells to continue our work with, because they are

easier to culture than ESc, and potentially provide a better model than the 661W cells as

they are embryonic and arguably could better represent most cell types in the adult mouse.

54

3.2.4 Antioxidant effect on total NAD+/NADH levels in MMS treated MEFs

Single strand breaks formed spontaneously or as intermediate products of DNA repair leads

to PARP activation [138, 139]. When single strand breaks overwhelm the repair capacity of

the cell, PARP is hyper-activated resulting in a sharp decrease in cellular total NAD+/NADH

and ATP levels [139]. This perturbation in energy promotes necrotic cell death [140].

Therefore, we investigated whether the effects of antioxidants on sensitivity to MMS-induced

cell death were linked to cellular depletion of total NAD+/NADH and ATP. WT MEFs were

seeded in 96-well plates in normal medium overnight. The medium was changed for medium

either containing or not containing 15mM NAC 30 minutes prior to treatment. Cells were then

exposed to MMS for 1 hour and cellular NAD+/NADH levels were measured at various time

points following MMS treatment.

Cells seeded in normal medium had their total NAD+/NADH levels significantly reduced

(more than 50% reduction) after 1 hour of treatment compared to controls not exposed to

MMS (p<0.05; Fig. 3.7). This is in line with previous published results [141]. Cellular

NAD+/NADH levels continued to drop over time following MMS treatment reaching their

lowest concentrations at 2 hours and remaining at this level up to 4 hours after treatment.

The total NAD+/NADH levels in cells pre-treated with NAC also decreased less than 50% of

control levels only one hour after treatment (P>0.5), and remained at this low level up to 4

hours after treatment. There were no difference in the rate of reduction of NAD+/NADH

levels between cells seeded in NAC or normal medium across the 4 hours monitored period,

suggesting that NAD+/NADH level had no direct role in the antioxidant-driven increase in

MMS cytotoxicity. Although most papers have detected the strongest PAR signal, the

product of PARP-1 activity 1 hour after MMS treatment; some published work has shown

PAR detected as soon as 5 minutes after treatment [142]. Therefore, although we did not

detect any difference between the cells seeded in normal and NAC medium 1 hour after

MMS treatment, an earlier time-point at 5-15 minutes post-treatment may have been more

informative.

55

56

Figure 3.7

0

50

100

150

200

NA

D/N

AD

H (n

M)

WT MEFsWT MEFs + NAC**

*

Figure 3.7. NAC had no effect on the MMS induced NAD+/NADH content reduction. WT MEFs were seeded overnight,

pre-treated with 15mM NAC or normal medium before being exposed to 2.5mM MMS for 1 hour. Cells were lysed and analysed

for total NAD+/NADH levels using a commercial kit and plate reader. Total NAD+/NADH levels were measured up to 4 hours

after the MMS treatment. At one hour, there was a reduction in total NAD+/NADH content levels in cells seeded in both normal

and NAC medium. NAD+/NADH levels continued to drop or remained at low levels throughout the 4hours. However there were

no significant differences between WT MEFs pre-treated with normal medium and those pre-treated with 15mM NAC.

Data shown are the result of three independent treatments (Two-way ANOVA, Tukey’s post hoc test, mean ± SEM, * p<0.5, ***

p<0.005).

57

3.2.5 Antioxidant effect on ATP levels in MMS treated MEFs

We determined cellular ATP levels at the same time points and experimental conditions as

for the NAD+/NADH analysis described above. In line with published literature, ATP levels

were significantly reduced in cells seeded in normal medium 1 hour post MMS treatment

(P>0.5; Fig 3.8). Strikingly, although published work that has examined ATP levels post

alkylation treatment report that ATP levels do not recover [143], in our experiments the ATP

levels recovered 2 hours post treatment to a level almost double the baseline ATP

(P>0.005). There were no sign of cell toxicity when looking at the cells under a microscope

up to 4 hours of the experiment. One determining factor for recovery of ATP levels is the

type and amount of damage. MNNG is a more powerful DNA damage response inducer than

MMS due to the O6MeG lesions induced by MNNG, resulting in a stronger PARP-1 hyper

activated state. Another factor to keep in mind is the cell type and time of treatment. For a

one hour 2.5mM MMS treatment, we are confident that in MEFs, ATP recovered two hours

post treatment to a level above baseline. This indicates that cell death was not a result of

ATP-depletion mediated by PARP-hyperactivation.

Another striking observation was that basal ATP levels were increased by pre-treatment with

NAC. Cells seeded in NAC medium had basal ATP approximately double those of cells

seeded in normal medium (P>0.05). Similar to cells seeded in normal medium, the

concentration of ATP in cells pre-treated with NAC reduced 1 hour post-treatment, before

recovering to levels above baseline 2 hours post-treatment.

58

Figure 3.8

Figure 3.8. NAC had no effect on the MMS induced ATP reduction. WT MEFs were seeded overnight, pre-treated with

15mM NAC or normal medium before being exposed to 2.5 mM MMS for 1 hour. They were lysed and analysed for ATP levels

using a commercial kit and plate reader. At time 0, Cells pre-treated with NAC medium had a significant increase in ATP levels

relative to normal medium. At one hour, there was a reduction in ATP levels in cells pre-treated with both normal and NAC

medium. 2 hours post treatment, ATP levels have recovered and surpassed the basal levels, in both normal and NAC medium.

4 hours post treatment, ATP levels in cells in normal medium remains high, whereas in NAC medium ATP levels dropped back

to basal level albeit still significantly higher than the basal level of cells pre-treated with normal medium. Data shown are the

result of three independent treatments (Two-way ANOVA, Tukey’s post hoc test, mean ± SEM, * p<0.5, ** p<0.05, *** p<0.005,

**** p<0.0005).

59

3.3 Discussion:

In this chapter, we have investigated the effect of antioxidants on alkylation-treated cells.

Antioxidants have been widely accepted to help protect against disease [118-120], but little

is known about whether they can also be harmful. New studies are emerging suggesting an

association between dietary antioxidant supplementation and increased incidence of cancer

[122], as well as faster progression of precancerous regions [121].

The cells used to investigate our hypothesis were mouse EScs, MEFs, and photoreceptor

cell line 661W. We had two genotypes of ESc and MEFs, WT and Aag-deficient genotypes.

ESc were used to determine whether the effects observed were general, or specific only to

differentiated cells. ESc were first characterised to confirm their pluripotent state (Fig 3.1),

and both WT and Aag-deficient cells stained greater than 95% positive for pluripotent marker

OCT3/4.

Having confirmed the ESc were pluripotent, we next wanted to see MMS-induced

cytotoxicity on the two genotypes available, WT and Aag-deficient, in both cell lines ESc and

MEFs. This has already been investigated in the literature, reporting Aag-deficient ESc to be

more sensitive than WT counterparts [85]. The published reports for MEFs on the other

hand have been inconsistent, with data showing Aag-deficient MEFs being no different in

sensitivity to MMS treatment than WT [144].

We aimed to first investigate the effect of MMS treatment on ESc. Briefly, ESc were seeded

and treated the next day with 0.5, 1.5 and 2.5mM MMS concentration for 1 hour in serum

free medium. The cells were harvested after 24 hours and analysed for cell death using PI

and Flow Cytometry. The PI method was selected as it’s a robust, reliable and rapid method

to analyse cell death. It relies on the compromised integrity of the cell membrane of dead

and dying cells to permit the take up of the PI dye which intercalates with DNA to give

enhanced fluorescence. Little or no cell death was detected in the control populations (Fig

3.2A), and Aag-deficient ESc were found to be more sensitive to MMS than WT as evident

60

by the greater degree of cell death caused by the 2.5mM MMS. This was in agreement with

previous published results [85], and indicates that the initiation of the BER pathway by Aag

plays an important role for the survival of ESc.

MEFs were also seeded and treated the next day with same MMS concentrations for 1 hour,

and analysed for cell death 24 hours later. We confirm that Aag-deficient MEFs were more

sensitive than WT (Fig 3.2 B), and this difference was observed consistently in cell cultures

of low passage (P5) up to high passage (P30). It was observed that after P30 the cells would

look notably smaller in size, proliferate more rapidly, and eventually have their MMS

sensitivity changed (data not shown). Therefore experiments were performed in lower

passages (below P20). The published reports that report different to our findings, where WT

MEFs had no different sensitivity to MMS than their Aag-deficient counterparts could relate

to the different cell death or viability assays used.

The results described in this chapter suggests that in the cell types evaluated, and at the

MMS concentration used, the initiation of BER by Aag has a favourable outcome for cell

survival. It could be that the downstream enzymes of Aag are in abundant amount, allowing

very efficient coordination between the initiation and completion of the repair pathway.

We next investigated how antioxidant treatment would affect the cellular response to MMS

treatment. The antioxidant NAC used in our experiment, is an aminothiol and a synthetic

precursor of intracellular GSH. GSH is the most abundant non-protein thiol in cells that

defends against oxidative stress, and NAC acts as an indirect antioxidant by increasing

intracellular stores of GSH [145]. However, NAC also possesses a reducing property through

its thiol-disulfide exchange activity [146]. Thiol-disulfide exchange reactions involve direct

interaction with target proteins through the cysteine residue or thiol group. NAC was shown

to be a powerful scavenger of hypochlorous acid, hydroxyl radical, and hydrogen peroxide in

cell-free experiments [147].

61

Briefly, WT MEFs were pre-treated with normal and 15mM NAC containing medium before

being exposed to MMS for 1 hour. Cells were harvested and analysed for cell cytotoxicity

using PI. On its own, incubation with 15mM NAC did not appear to be cytotoxic to cells (Fig

3.3 A). This is in line with previously published studies in which concentrations of up to

30mM NAC have been used with live cells [148]. However, NAC increased MMS-induced

cytotoxicity in both WT and Aag-deficient MEFs (Fig 3.3 A and B). This effect was more

evident in the Aag-deficient genotype where a statistically significant effect was observed at

lowest MMS concentration used (0.5mM MMS, p<0.5), whereas the WT cells the effect was

only significant at concentrations of 1.5mM or above, (p<0.05). In both genotypes, cells

seeded in NAC and treated with 2.5mM MMS showed almost no cell viability (>80% PI

positive, p<0.0005).

These results were very striking, and similar results were obtained with a different cell type

(ESc) and different antioxidant (2-ME). As is the case for NAC, 2-ME is also a thiol

containing antioxidant that is routinely used for the culture of ESc, to mimic the partial

hypoxic conditions of the embryo and maintain their pluripotency [135, 136].

2-ME therefore gave us an additional control to investigate how reducing conditions affect

MMS-cytotoxicity, as it was taken away from the medium rather than being added to it. ESc

that were seeded without 2-ME exhibited higher intracellular ROS levels than those that

were seeded with 2-ME (Fig 3.4). WT ESc seeded with 2-ME and treated with MMS showed

a pattern of apparently increased MMS induced cell death compared to ESc that were

seeded without 2-ME (Fig 3.5 A), although the differences were not statistically significant.

This might be due to a low concentration of 2-ME used. For the Aag-deficient ESc however,

the results were statistically significant and showed 0.1mM 2-ME enhanced MMS-

cytotoxicity (2.5mM MMS, p<0.005; Fig 3.5 B). To further test the generalisability of these

observations, the effects of antioxidants on a third cell line were also evaluated. The 661W

photoreceptor cell line was selected for this purpose. These are cone photoreceptors [149],

a type of cells that shows evident particular sensitivity to MMS treatment in mice [97]. Pre-

62

treatment with NAC enhanced MMS-cytotoxicity in the 661W cell line to a similar extent to

that seen with the MEFs (Fig 3.6). Moreover, pre-treatment with a commercial antioxidant

mixture also enhanced MMS-cytotoxicity in a concentration dependent manner.

Having confirmed that antioxidant loading of various cell types increased their sensitivity to

MMS treatment, subsequent experiments were designed to investigate whether this effect

might be associated with the effects of DNA damaging agents on cellular energy

metabolism. MMS treatment results in the reduction of intracellular concentrations of

NAD+/NADH and ATP [2]. This results, at least in part, via hyperactivation PARP-1 enzyme.

PARP-1 is normally is essential in BER and DNA single strand break repair. However, when

the production of DNA lesions is sufficient to overwhelm the DNA repair capacity, an

imbalanced coordination of repair processes leads to a hyperactivation of PARP-1 which

produces PAR polymer at the expense of NAD+ [2]. Initial earliest research in this area

suggested that the resulting decrease in cellular NAD+/NADH levels led to in a reduction of

ATP levels, and this perturbation of energy levels promotes mitochondrial dysfunction and

the release of the apoptotic inducible factor (AIF), resulting in necrotic cell death [140].

However, recent research has suggested the PAR polymer is itself a messenger signal for

the release of AIF regardless of NAD+/NADH levels [150].

MMS treatment reduced total NAD+/NADH levels 1 hour after treatment (Fig 3.7), the

earliest time point analysed post treatment. There was no evidence that NAD+/NADH levels

recovered at all throughout the time course up until the last time point at 4 hours. This is in

agreement with published literature [141]. Seeding cells in medium containing NAC had no

detectable effects on baseline cellular concentrations of NAD+/NADH. Nor did it have any

detectable effect on cellular NAD+/NADH depletion following treatment with MMS.

The effects of antioxidants on cellular NAD+/NADH levels have not been investigated

extensively. One paper suggests there is an association between antioxidant

supplementation and serum NAD+/NADH levels after exercise [151]. In our cells, antioxidant

63

did not change basal total NAD+/NADH levels prior to treatment, and most interestingly, did

not change the rate that total NAD+/NADH levels dropped post-treatment. The earliest time

point post-treatment that we analysed was 1 hour. Although most papers have detected the

strongest PAR signal, the product of PARP-1 activity that requires NAD+ as substrate for its

production, 1 hour after MMS treatment, some published work has shown PAR can be

detected as soon as 5 minutes after treatment [142]. Therefore, although we did not detect

any difference between the cells seeded in normal and NAC medium 1 hour after MMS

treatment, an earlier time-point at 5-15 minutes post-treatment may have been more

informative.

As was the case with NAD+/NADH, very little is available in the literature on how

antioxidants affect ATP levels in cells, or how ATP levels are affected by antioxidants post

MMS-treatment. We therefore wanted to investigate how antioxidants affect MMS-treated

MEFs at the same time-points as NAD+/NADH experiments. MMS treatment significantly

reduced ATP levels at the first time point post treatment in MEFs seeded without NAC (1

hour, Fig 3.8). However, unexpectedly, ATP levels had recovered by 2 hours post-treatment.

This observation does not match the limited published evidence that is available, which

suggest ATP levels remain below basal levels [143]. We also used boiling MilliQ autoclaved

water to lyse the cells, and incubated the plates further at 95C° for complete lysis. This lysis

method is different from the conventional method using trichloroacetic acid (TCA), and has

been reported to stabilise ATP more effectively. Another important factor that is likely to

influence the rate and extent of recovery of ATP levels is the type and amount of damage

produced. The alkylating agent MNNG is a more powerful DNA damage response inducer

than MMS due to the O6MeG lesions induced by MNNG. This is expected to result in a

greater PARP-1 hyper activation and this could also contribute to the apparent discrepancy

in findings between the study described here and that published previously [152].

64

Similarly to the cells seeded in normal medium, cells seeded in NAC-containing medium had

their ATP concentration reduced 1 hour post-treatment with MMS, before recovering to

levels above baseline 2 hours post-treatment (Fig 3.8). However, surprisingly basal ATP

levels in cells seeded with NAC was higher (approximately doubled) compared to those

seeded in normal medium (P>0.05). It is unlikely that antioxidants interfered with the assay,

as NAC medium was discarded, and cells were washed twice with PBS, before lysing cells

with water. The cell lysates were also transferred to a new plate, eliminating the possibility

that antioxidants could have adhered to the plastic of the 96-well plate. To our knowledge,

this effect has not been described in previous published literature that show or explain this

effect, and it will be difficult to draw conclusions without further results.

The studies described in this chapter evaluated the role of antioxidants in the cellular

response to the alkylating agent MMS, and whether the effects observed can be linked with

effects on energy metabolism. We have shown that the antioxidants NAC, 2-ME and Sigma

AO exacerbated the MMS-induced cytotoxicity in MEFs, ESc and the 661W photoreceptor

cell line. NAC did not alter the effects of MMS on cellular NAD or ATP levels NAC did

increase basal ATP levels in MEFs, but this would unlikely explain the NAC-driven increase

in MMS-cytotoxicity, as ATP levels recovered to above basal level 2 hours post-treatment in

both cells seeded with or without NAC. These findings suggest that the observed

enhancement of MMS-cytotoxicity by antioxidants was not due to changes alterations in the

effects of MMS on cellular energy metabolism.

65

Chapter 4: Characterisation of MMS-induced reactive oxygen

species generation

66

4.1 Introduction:

It is widely held that ROS are only detrimental to cells. More recently however, the

importance of intracellular ROS acting as signalling molecules has begun to be recognised

[153]. Whether ROS signalling plays a role in the DNA damage response is not yet entirely

clear.

There have been reports that DNA damage can result in an intracellular increase in ROS

levels [154]. One study showed that this increase in ROS was not directly linked to cell death

[155]. The authors hypothesized the ROS produced were involved in signalling the stress

response. In MEFs, treatment with MNNG caused ROS production and c-jun N-terminal

kinase (JNK) activation [156], a kinase that can be activated by ROS in response to stress

stimulus. Kang and coworkers found that ROS induced by DNA damage using the

radiomimetic neocarzinostatin (NCS) is detrimental to cells [148]. They reported that the

ROS generation was due to an increase in H2AX and found DNA damage related induction

of ROS was also caused by hydroxyurea and deoxyrubicin in different cell lines. Treating

661W photoreceptor cells with 1-methyl-1-nitrosourea (MNU) also resulted in ROS

production, and this was confirmed in vivo using mice models [157]. Contradictory to the

reports that state ROS to be detrimental, it has been shown that ascorbate, the mineral salt

for ascorbic acid (vitamin C), increases strand breakage in DNA if administered at 0.5mM

prior to MNU treatment in leukemia and pancreatic cells [158].

Overall, it is currently unclear exactly how the intracellular redox state affects the DNA

damage response after DNA insults or vice versa, with different studies reporting apparently

conflicting findings.

Intracellular ROS can be generated from several sources inside a cell. In the mitochondria,

the flow of electrons across the respiratory chain ends up reducing molecular oxygen to

water or to a lesser extent partially reducing molecular oxygen to generate the superoxide

anion [159]. As this suggests, ROS generation through the mitochondria happens as a by-

67

product of oxidative phosphorylation, and there are other enzymes that produce ROS as a

by-product such as xanthine oxidase, nitric oxide synthase, cyclooxygenases, cytochrome

p450 enzymes and lipoxygenases [153]. The family of NADPH oxidases (Noxes), first

discovered in neutrophils, are the only known enzymes whose primary role is to produce

superoxide ions [153]. The superoxide produced by Noxes in neutrophils was thought to be

only relevant in the oxidative burst, a mechanism used by neutrophils to inactivate foreign

microorganisms. However, it was found later that a wide variety of ligands could activate

Noxes in non-immune cells, and that the ROS produced served various signalling purposes

[160].

The results described in chapter 3 demonstrate the use of antioxidants can exacerbate the

cytotoxic effects of alkylation damage. This, therefore, leads naturally to the questions of

whether alkylation lesions induced by MMS affect intracellular ROS and, if so, how

antioxidant supplementation influences this production of ROS. Therefore, the experiments

described in this chapter were desgined to investigate how the effects of MMS and NAC,

separately and in combination, affect intracellular ROS levels, both in the cytosol and in the

mitochondria. If the results are in agreement with the literature about ROS generation post

genomic insults, we will try and pinpoint the responsible enzymes or pathways that results in

ROS generation post MMS treatment.

68

4.2.1 NAC depletes MMS-induced ROS generation in WT MEFs

The effects of NAC and MMS treatment on intracellular ROS were initially investigated by

making use of 2’-7’ dichlorfluorescein diacetate (DCF-DA). DCF is an acetylated redox-

sensitive probe that can freely cross the plasma membrane. Once inside the cell, it is

deacetylated by the action of cellular esterases to the reduced form DCF-H2. This conversion

traps the DCF-H2 within the cell [161]. DCF-H2 inside the cell is senstive to oxdiation by ROS

and this oxidation converts it into a fluorescent form.

MEFs were seeded overnight with normal medium and had their medium changed for normal

or NAC containing medium 30 minutes prior to MMS exposure. They were treated with

2.5mM MMS for 1 hour and the treatment medium was then changed back to normal

medium either containing NAC or not. After 24 hours cells were harvested and incubated

with the redox sensitive probe H2DCF-DA for 30min at 37°C. Cells were washed and

analysed by flow cytometry.

At time 0, immediately prior to MMS treatment, the ROS levels, as detected by the DCF

probe, in cells pretreated with 15mM NAC were not significantly different to those in control

cells (Figure 4.1). This indicates that treatment with 15 mM NAC was well tolerated by the

cells, with no evidence that it directly affected cell viability (Fig 3.3) or cellular redox (Fig 4.1).

In the absence of NAC pre-treatment, ROS levels increased in MEFs following MMS

exposure (Fig 4.1). The precise time-dependent pattern of MMS-induced ROS induction and

the absolute fold-increases compared to baseline levels varied from one experiment to

another. The earliest induction observed occured as soon as 30 minutes post-treatment,

while in most experiments ROS generation peaked at 60 minutes (p<0.005). In all

experiments, pre-treatment with NAC completely depleted the ROS generation post-MMS

treatment (Fig 4.1). These results suggest that since antioxidants deplete MMS-induced

ROS generation and increase MMS-induced cell death, the ROS generated post-MMS

treatment potntially increases the survival of cells. We will investigate this in subsequent

69

experiment to pin-point where the ROS originates from, and whether inhibiting the

production pathway also has the same effect as the antioxidants.

70

Fig 4.1

010

min30

min60

min

120m

in

240m

in0

2

4

6

8

10

Fold

cha

nge

RO

S

WTMEFs

WT MEFs + NAC***

Figure 4.1. NAC completely abrogates MMS-induced ROS production in MEFs. WT MEFs were seeded overnight, pre-

treated with 15mM NAC or normal medium before being exposed to 2.5 mM MMS for 1 hour. They were harvested at different

time-points and analysed for intracellular ROS levels using flow cytometry. At time 0, cells pre-treated with NAC did not exhibit

any different ROS levels when compared to controls. MMS exposure induced ROS generation in MEFs 1 hour after treatment

start. This was ROS generation was absent in MEFs pre-treated with NAC. The generated ROS in MEFs seemed to decrease

over time until reaching near basal levels at 4 hours post-MMS exposure. Data shown are the result of five independent

treatments (Two-way ANOVA, Tukey’s post hoc test, mean ± SEM, *** p<0.005).

71

4.2.2 MMS treatment results in ROS generation in WT but not Aag-deficient MEFs

We confirmed that MMS treatment leads to ROS generation, and that the antioxidant NAC

abrogates ROS generation post MMS-treatment, thereby increasing MMS-toxicity.

Subsequent experiments were performed in an attempt to pinpoint the enzymes or pathways

were responsible for the ROS generation.

We were interested to know whether the damage to DNA, RNA and proteins by MMS

treatment was enough to elicit ROS production, or whether it was the initiation of the BER

pathway. To explore this, WT and Aag-deficient MEFs were seeded overnight and treated

with MMS the following day for 1 hour, and analysed for ROS production with H2DCF-DA

and flow cytometry.

Strikingly, Aag-deficient MEFs did not display ROS induction post-MMS treatment,

compared to WT cells (Fig 4.2). Across all time points, up to 4 hours after MMS treatment,

there was no evdience of any increase in fluorescence indiciative of ROS production in Aag-

deficient cells. In total, this experiment was performed on 9 separate occasions using cells at

passages ranging from P7-P20, with consistent results in each experiment.

This suggests that the DNA damage MMS produces is not, on its own, sufficent to induce

ROS generation. Instead, this observation suggest that the ROS generation is induced only

after the BER pathway is initiated by Aag following the production of DNA alkylation damage

by MMS.

72

Fig 4.2

010

min30

min60

min

120m

in

240m

in0

2

4

6

8

10

Fold

cha

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RO

S

WT MEFs

Aag-deficient MEFs****

***

Figure 4.2. Aag-deficient cells did not exhibit MMS-induced ROS generation. WT and Aag-deficient MEFs were seeded

overnight, and exposed to 2.5 mM MMS for 1 hour. They were harvested at different time-points and analysed for intracellular

ROS levels using flow cytometry. At time 0, the different genotypes did not have any significant difference in intracellular ROS

levels. At 60 minutes post-MMS exposure, there was a significant increase in ROS generated by WT MEFs compared to Aag-

deficient counterparts. Aag-deficient MEFs did not display any MMS-induced ROS generation throughout the 4 hour

experiment. MMS-induced ROS detected in WT cells normalised to near basal levels 4 hours post MMS exposure. Data shown

are the result of nine independent treatments (Two-way ANOVA, Tukey’s post hoc test, mean ± SEM, *** p<0.005, ****

p<0.0005).

73

4.2.3 Mitochondrial ROS are not elevated 1 hour post-MMS treatment

Using WT MEFs, we wanted to pinpoint the responsible pathways or enzymes involved in

the ROS generation. The mitochondria is a potential site for ROS production, as some

electrons could partially reduce molecular oxygen [159]. We predict that during insult, the

flow of electrons across the respiratory chain could be less regulated, and therefore leakage

of electrons would cause more partial reduction of molecular oxygen at the site, leading to a

spike in ROS levels. We will investigate the mitochondrial ROS levels using a redox probe

called Mitosox Red, a cell permeable probe for the detection of superoxide in mitochondria in

live cells [162]. Mitosox has a lipophilic, positively charged triphenylphosphonium moiety

that targets the mitochondria, and another superoxide-sensitive dihydroethidium conjugate

that is used to detect superoxide[163]. The oxidation of dihydroethidium converts it to a red

fluorescent form that can be detected and quantified by microscopy and flow cytometry. WT

MEFs were seeded overnight and pre-treated with normal medium or NAC containing

medium for 30 minutes prior to MMS exposure.

Neither populations of MEFs seeded with or without NAC showed any evidence of elevated

mitochondrial ROS at 1 hour post MMS-treatment. This indicates that the cellular ROS spike

seen at 1 hour post treatment in WT MEFs detected with the probe DCF (Fig 4.1) was

unlikely to originate from the mitochondria. At the last time point tested (4 hours post-MMS

treatment) both populations started to show increased mitochondrial ROS production. MEFs

seeded with NAC exhibited significantly higher mitochondrial ROS compared to those

seeded without NAC (p<0.05; Fig 4.3).

Since the earlier time points (15 min to 2 hours post-MMS treatment) did not show any

superoxide level elevation, it can be concluded that mitochondrial DNA and protein damage

caused by 2.5mM MMS was not sufficient to immediately destabilise the electron transport

chain to a degree where excessive electron leakage and partial reduction of molecular

oxygen occurred. The superoxide elevation at 4 hours post-MMS treatment, however,

indicates mitochondrial dysfunction possibly as a result of dying cells. To further support this

74

hypothesis, the cells seeded with NAC exhibited higher mitochondrial superoxide formation

at 4 hours post-MMS treatment (Fig 4.3) and exhibited higher cell death detected with PI

staining 24 hours post treatment (Fig 3.3) compared to MEFs seeded without NAC.

75

Fig 4.3

010

min30

min60

min

120m

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240m

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1

2

3

Fold

Mito

chon

dria

l Sup

erox

ide

WT MEFs

WT MEFs + NAC

**

Figure 4.3. WT MEFs pre-incubated with NAC and exposed to MMS show a late phase mitochondrial superoxide

generation. WT MEFs were seeded overnight, pre-treated with normal or NAC containing medium and exposed to 2.5 mM

MMS for 1 hour. They were harvested at different time-points and analysed for mitochondrial superoxide formation. There were

no significant difference in superoxide levels between MEFs pre-treated with or without NAC at time 0. This was also true up to

2 hours post MMS-exposure. At the last time-point 4 hours post MMS exposure, cells pre-treated with NAC exhibited higher

levels of mitochondrial superoxide generation as opposed to cells pre-treated with normal medium. Data shown are the result of

three independent treatments (Two-way ANOVA, Tukey’s post hoc test, mean ± SEM, ** p<0.05)

76

4.2.4 NAC increased mitochondrial depolarisation in MMS treated MEFs

To confirm that mitochondrial superoxide formation observed in MEFs 4 hours after MMS

treatment was due to mitochondrial dysfunction associated with cell death, the extent of

mitochondrial polarisation up to 4 hours after MMS treatment was analysed using the probe

TMRE. TMRE is a lipophilic fluorescent cation probe that is cell permeable and accumulates

within the mitochondria when they are polarised (i.e. when the proton gradient across the

mitochondrial membrane is maintained appropriately to permit ATP generation) [164]. This

can be used qualitatively to compare polarisation of mitochondria between samples or

treatments.

WT MEFs were seeded in normal medium overnight, and pre-treated with normal medium or

NAC containing medium before being exposed to MMS. The population of control cells that

were not treated with MMS but had been seeded in medium containing 15mM NAC

appeared to include a slightly higher proportion of cells with low mitochondrial polarisation

compared to cells seeded without NAC. However, this apparent difference was not

statistically significant (Fig 4.4).

At 2 hours after MMS treatment, there was a significant difference (p<0.5) between the

proportion of cells having low mitochondrial polarisation in the cells pre-treated with NAC

compared to those seeded with normal medium. This difference was even more marked at 4

hours, by which time approximately 55% of cells seeded in NAC had low mitochondrial

polarisation compared with only 20% (a value close to baseline proportion at time-point 0) of

cells seeded in normal medium (p<0.0005). This shows that cells seeded with NAC undergo

mitochondrial depolarisation more readily after MMS treatment than control cells, and that

the extent of depolarisation continued to increase until at least the last time point analysed (4

hours post MMS treatment). This pattern of change mirrored the increased in mitochondrial

superoxide production described above. In summary, these data suggest that the NAC pre-

treatment increased mitochondrial susceptibility to depolarisation, mitochondrial superoxide

formation and cell death in response to MMS exposure. It not clear, however, whether the

77

mitochondrial superoxide formation is responsible for the observed cell death, or whether it

is a secondary effect of the cell death process.

On the other hand, the observed patterns of mitochondrial ROS production following MMS

exposure do not explain the peak in cellular ROS detected with DCF at 1 hour post-MMS

treatment suggesting that other sources must be responsible for production of these ROS.

Therefore, a range of further experiments were performed to evaluate other possible sources

of the ROS production elicited by MMS treatment.

78

Fig 4.4

Control

60min

120m

n

240m

in0

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40

60

80

% C

ell w

ith lo

w m

itoch

odria

l pol

aris

atio

n WT MEFsWT MEFs + NAC

*

****

Figure 4.4. WT MEFs pre-incubated with NAC and exposed to MMS show a late phase mitochondrial depolarisation.

WT MEFs were seeded overnight, pre-treated with normal or NAC containing medium and exposed to MMS for 1 hour. They

were harvested at different time-points and analysed for mitochondrial depolarisation. There were no significant difference in

mitochondrial polarisation between MEFs pre-treated with or without NAC at time 0. However, at 2 hours post-MMS exposure,

cells pre-treated with NAC exhibited higher mitochondrial depolarisation than did cells pre-treated with normal medium, this

effect was further enhanced at 4 hours. Data shown are the result of three independent treatments (Two-way ANOVA, Tukey’s

post hoc test, mean ± SEM, * p<0.5, **** p<0.0005).

79

4.2.5 DPI abrogates MMS-induced ROS generation in MEFs

The experiments described in chapter 3 demonstrated that MMS treatment of WT MEFs led

to substantial increases in cellular ROS generation and that the DNA repair enzyme Aag is

required for the production of these ROS since no equivalent increase in ROS was observed

following exposure of Aag knockout MEFs to MMS. This MMS-induced increase in ROS

generation reached a maximum between 15 minutes to 2 hours post-MMS treatment, before

declining back to basal ROS levels. Analysis of ROS production in mitochondria in MEFs

following MMS exposure suggested that these are unlikely to be the primary source of the

cellular ROS. We therefore decided to investigate ROS generation by NOXes, the only

enzymes known to produce ROS not as a by-product. Diphenyleneiodonium (DPI) was used

as an irreversible flavoprotein inhibitor that has been used previously to inhibit NOXes [165].

WT MEFs were seeded in normal medium and incubated overnight. The medium was

replaced with either fresh normal medium or medium containing 2µM DPI the next day and

the cells incubated for 30 minutes prior to MMS treatment. Cells were treated in serum-free

medium containing 2.5mM MMS for 1 hour before having this medium replaced back either

to normal medium or medium containing DPI. Cells were then harvested and incubated with

DCF prior to flow cytometry analysis.

MEFs exhibited ROS generation 1 hour post-MMS treatment (Fig 4.5), in line with previous

results (Fig 4.2). Cells that were incubated with DPI did not show any significant elevation in

ROS levels post-MMS treatment. At the 1 hour time-point, there was a significant difference

in ROS levels detected in WT MEFs seeded without DPI compared to cells seeded with DPI

(p<0.5; Fig 4.5). This suggests that NOXes could be responsible for the MMS-induced ROS

generation observed 1 hour post treatment.

We next attempted to inhibit NOXes with DPI, treat with MMS and see if that brings an

increase in cell death detected by PI as it did with antioxidant NAC.

80

Fig 4.5

010

min30

min60

min

120m

in

240m

in0

1

2

3

Fold

RO

S

WT MEFs

WT MEFs + DPI*

Figure 4.5. DPI completely abrogates MMS-induced ROS production in MEFs. WT MEFs were seeded overnight, pre-

treated with 2µM DPI or normal medium before being exposed to 2.5 mM MMS for 1 hour. They were harvested at different

time-points and analysed for intracellular ROS levels using flow cytometry. Control cells did not show any difference in

intracellular ROS levels between cells pre-treated with or without DPI at time 0. Cells pre-treated with normal medium and

exposed to MMS exhibited a significant increase in intracellular ROS 1 hour post-MMS exposure, compared to cells pre-treated

with DPI. Data shown are the result of three independent treatments (Two-way ANOVA, Tukey’s post hoc test, mean ± SEM, *

p<0.5).

81

4.2.6 DPI increases MMS-induced cytotoxicty in WT MEFs

Having confirmed that both NAC and DPI prevent the MMS-induced ROS generation, the

effects of DPI on MMS-induced cytotoxicity was investigated to determine whether or not this

was also similar to the effects observed with NAC.

WT MEFs were seeded and incubated overnight. The medium then replaced with either

fresh normal medium or medium containing 2µM DPI and the cells incubated for 30 minutes

prior to MMS treatment. Cells were treated in serum-free medium containing 2.5mM MMS

for 1 hour before having medium replaced back to normal medium or medium containing

DPI. Cells were harvested and analysed for cell death by PI using flow cytometry.

Addition of 2µM DPI alone did not lead to any detectable additional cell death compared with

control untreated cells (Fig 4.6). This same DPI concentration did however increase cell

death following MMS treatment, compared to cells in normal medium without DPI. The

extent of increase in MMS-cytotoxicity in cells seeded with DPI was not as large as that

observed in cells seeded with 15mM NAC (~20% cell death with DPI as opposed to ~80%

with NAC).

Taken together, these data strongly suggest that NOXes are responsible for, or substantially

contribute to, the rapid production of ROS observed after-MMS treatment in wild type cells.

However, since DPI is not highly specific for NOX inhibition it is not possible to absolutely

rule out other mechanisms. Therefore, NOX knock-down experiments, using shRNA, were

attempted to obtain more definitive proof.

82

Fig 4.6

Control 2.5mM MMS0

10

20

30

% C

ell D

eath

WT MEFsWT MEFs + DPI

***

Figure 4.6. DPI increased MMS-induced cell death in WT MEFs. WT MEFs were pre-treated with 2µM DPI or normal

medium before being exposed to MMS for 1 hour. Cells were analysed for cell death 24 hours after 2.5 mM MMS exposure.

Pre-treatment with DPI did not increase cell death in control cells. When cells were exposed to MMS, pre-treatment with DPI

increased cell death compared to pre-treatment with normal medium. Data shown are the result of four independent treatments

(Two-way ANOVA, Tukey’s post hoc test, mean ± SEM, *** p<0.005).

83

4.2.7 MEFs can be successfully transfected but not selected with Geneticin or

Puromycin

As discussed in section 4.3, the evidence obtained using the inhibitor DPI strongly

suggested that NOX is either responsible for or contributes substantially to, the rapid

transient increase in cellular ROS levels observed in WT MEFs following MMS treatment.

We wanted to further confirm this with NOX knock-down using small hairpin RNA (shRNA).

These are short hairpin precursors approximately 70 nucleotides in length that are

processed by Dicer to form active siRNAs of around 20 nucleotides in length [166]. siRNA

are incorporated into RNA-induced silencing complex (RSC) and target complementary

mRNA where they are cleaved by RISC [167].

Initial experiments were performed, using a green fluorescent protein (GFP) reporter plasmid

construct, to determine optimise transfection conditions. WT MEFs were seeded at two

different cell densities (1000 and 2000 cells per well) in black 96 well plates. The next day,

they were transfected with different ratios of plasmid DNA concentrations to transfection

reagent for 24 hours (0.2 µg and 0.4 µg of plasmid DNA; 0.25 µl, 0.5 µl and 1 µl transfection

reagent per well). Transfection medium was aspirated and cells were allowed to recover in

normal medium for another 48 hours before being analysed for GFP fluorescence.

This experiment demonstrated that seeding cells at a density of 2000 per well and using 0.2-

0.4 µg/1µl of plasmid DNA/transfection reagent ratio gave the best transfection results (Fig

4.7). All the wells that treated with less than 1 µl transfection reagent exhibited very poor

transfection. A seeding density of 2000 cell per well and treatment with 0.2 µg/1 µl ratio was,

therefore selected as the conditions to perform with the shRNA experiments.

Optimal concentration of the selection drug puromycin to use after transfection was also

determined. For this MEFs were seeded and incubated variously with 0, 0.2, 0.4, 0.6, 0.8

and 1 µg/ml. A puromycin concentration of 0.8 µg/ml killed approximately 50% of the

untransfected cells after 3 days and almost all the cells after 7 days. The 0.8 µg/ml was

84

chosen for selection purposes and a lower concentration of 0.4µg/ml was chosen for the

maintenance concentration.

MEFs were seeded in 96-well plates, transfected the next day with one of four different

NOX1 shRNAs for 48 hours, then selected with 0.8 µg/ml puromycin for 7 days, before

switching to the maintenance concentration to allow wells to become confluent. In all cases

the MEFs exhibited slow growth rate after selection until they senesced. The transfection

was attempted on passage 4 cells and passage 15 cell. In both cases, the cells stopped

proliferating shortly after selection. In further attempts to transfect the cells, the maintenance

concentration of puromycin was reduced to 0.2 µg/ml but this still did not improve the

outcome. Similar problems were encountered with a separate set of experiments in which

MEFs were transfected with plasmids that express ATP and NAD sensitive probes, where

transfectants were selected using geneticin. Consequently, further attempts to perform the

NOX knockdown experiments by shRNA were abandoned.

85

Fig 4.7

Figure 4.7. Transfection efficiency of MEFs using GFP. WT MEFs were seeded at different seeding densities in 96-well

plate, and transfected with different transfection reagent amount and ratio to GFP plasmid DNA. Cells seeded at 2000 per well,

with 0.2-0.4µg/1ul of plasmid DNA/transfection reagent ratio gave the best transfection results. Data shown are a

representative of a screening experiment performed.

86

4.3 Discussion:

The work described in the previous chapter demonstrated that antioxidants sensitise cells to

MMS-induced cell death. This was shown to be independent of the effects of MMS on

cellular energy status, as evaluated by cellular ATP and total NAD+/NADH levels.

There has been several reports that indicate DNA damage production results in an

intracellular increase in ROS levels [148, 154-157]. Therefore, the experiments described in

this chapter were designed to test the hypothesis that the mechanisms underlying the

sensitising effects of antioxidants on MMS-induced cell death involve changes in cellular

ROS production and/or signalling. Two different probes were used to investigate the effects

of antioxidants and MMS of cellular ROS: one that specifically measures mitochondrial

superoxide and another that measures cellular ROS. The probe DCFDA-H2 has been widely

used for the detection of intracellular ROS [161]. Using this probe we demostrated that MMS

treatment resulted in a substantial increase in cellular ROS levels (Fig 4.1). The increase in

ROS was detectable as early as 30 minutes post-MMS treatment in some experiments, with

peak levels typically being reached after 60 minutes. At later time points cellular ROS

declined, returning back to basal levels between 2 and 4 hours following MMS treatment.

The significance of ROS generation following DNA insults is not clearly understood. The

experiments described here demonstrated that preincubation of the cells with NAC

abrogated the MMS-induced increase in cellular ROS (Fig 4.1). This effects appears to be

specfic for the conditions produced by MMS treatment that trigger additional ROS production

since. on its own (i.e. in the absence of MMS treatment) NAC treatment did not cause any

detecable change in cellular ROS levels (zero time point in Fig 4.1). In conjunction with the

data described in chapter 3, this further confirms 15mM NAC was well tolerated by the

MEFS and that the senstisiting effects of NAC observed following MMS treatment are not

obviously the result of mild adverse effects of NAC on the MEFS being amplified following

the challenge with the alkylating agent.

87

There appears to be a mixed opinon about whether the increase in ROS observed following

DNA damage is protective or detrimental to cell survival. Kang and co-workers reported that

there are two ROS production phases: one that happens early approximately 1 hour after the

DNA insult, and another that occurs approximately 5 hours after insult, after the first phase of

ROS production had been normalised back to basal levels. This group showed that

abolishing the first ROS production phase at 1 hour post treatment is detrimental to cell

survival, although they were not able to determine why. In contrast, abolishing the second

ROS phase by adding NAC 2 hours after treatment was beneficial to cells [148].

Contradictory to the reports that state ROS to be detrimental, it has also been shown that

ascorbate, the mineral salt for the antioxidant ascorbic acid (vitamin C), increases strand

breakage in DNA if administered at 0.5mM prior to MNU treatment in leukemia and

pancreatic cells [158]. Splenocytes from superoxide dismutase (SOD1) gene knock out mice

exhibited more DNA damage when treated with monomethylarsonous acid (MMA3) or

dimethylarsinous acid (DMA3) than their WT counterparts [168]. SOD1 is an enzyme that

converts superoxide ions into hydrogen peroxide, which is less reactive with a longer half-life

making it more suitable as signalling molecule.

The observation that NAC pretreatment blocked the early phase induction of ROS following

MMS treatment suggested that this might somehow be involved in the mechanism

responsible for the senstising effect of NAC on MMS-induced cell death. Therefore, we used

a range of experimental strategies to investigated candidate pathways and enzymes that

might be responsible for this ROS generation.

The two predominant lesions produced by MMS are the 7meG and 3meA (82% and 11%,

respectively), both substrates for Aag [87]. Since the experiments described in chapter 3 had

demonstrated that both Aag-deficient ESc and MEFs were more senstive to MMS treatment

than either WT equivalent, we investigated the effect of Aag-deficiency on MMS-induced

cellular ROS production.

88

These experiments demonstrated that in stark contrast to WT cells, Aag-deficient MEFs did

not display ROS generation, monitored using DCFA-H2, following-MMS treatmentat any time

point up to 4 hours post-treatment (Fig 4.2). This strongly suggests that the MMS-induced

ROS production requires the initiation of BER by Aag at the sites of DNA alkylation produced

by the MMS treatment. It is not clear how the process of BER is linked with this ROS

production or which step of the BER process is key to this link. APE-1 and XRCC1, both

components of the BER machinery, have been shown to be redox regulated, having greater

activity when in oxidised forms [129-132]. ROS production post-MMS treatment could

therefore be involved in post-translational modifications of key proteins involved in BER to

further enhance repair activity. It has been previously shown that DNA insults could partly

increase ROS production through H2AX accumulation [148]. H2AX becomes quickly

phosphorylated when exposed during DSBs and acts as DNA foci for the recruitment of

repair proteins [169].

We next investigated whether the mitochondria could be the origin of the rapid ROS

production observed following MMS treatment. The mitochondria is a major site for cellular

ROS production, as electrons from the electron transport chain can partially reduce

molecular oxygen [159]. For this, Mitosox Red was used, a probe that is sequestered into

the mitochondria and is selective for superoxide detection.

WT MEFs treated with MMS did not show any elevated mitochondrial superoxide levels 1

hour post-treatment (Fig. 4.3). This suggests that the intracellular ROS production detected

by the probe DCFDA-H2 at 1 hour after MMS treatment is unlikely to originate from the

mitochondria. At the last time point analysed, 4 hours after MMS treatment, elevated

mitochondrial superoxide production was detected, and the increase was greater in the cells

that had been incubated with NAC (Fig 4.3). The timing of this increase is generally

consistent with the second phase of ROS production following DNA damage induction

reported in the study by Kang and co-workers (add the relevant citation here).This is

89

believed to be ROS involved in cell death as the cells incubated with NAC also showed

higher MMS-cytotoxicity in the previous chapter Fig 3.3).

During cell death, the mitochondrial membrane permeability (MMP) increases leading to the

release of pro-apoptotic and –necrotic factors as well as ROS production [170, 171]. These

is a key hallmark of mitochondrial dysfunction. Therefore, we investigated mitochondrial

depolarisation up to 4 hours post-MMS treatment. In control cells, it appeared that NAC

alone might promote some mitochondrial depolarisation although the apparent change was

not statistically significant (Fig 4.4). One possibility is that a reducing environment primes the

mitochondria for an easier mitochondrial membrane permeability increase following genetic

insults. There has been one study that shows glutathione reduction stress mediates

mitochondrial oxidation and cytotoxicity [172]. We did not find any mitochondrial oxidation or

superoxide formation in previous results in NAC treated control cells, neither did we detect

any cytotoxicity. However, this mitochondrial probing effect could possibly explain why

during insult, NAC treatment cells facilitates higher depolarisation and higher cytotoxicity.

Indeed as we treated MEFs with MMS, the number of cells with low polarised mitochondria

increased to a greater extent in the NAC treated cells and in the control cells (Fig 4.4).

These results further confirm that the mitochondrial superoxide detection 4 hours post-MMS

treatment, an effect observed greater in MEFs pre-treated with NAC, is due to the

mitochondria depolarising. This pool of results suggest that NAC reduces mitochondrial

depolarisation, followed by mitochondrial superoxide formation and cells death. It is unclear

whether the mitochondrial superoxide formation is responsible for cell death, or whether it is

a secondary effect of the cell death process.

Having confirmed that mitochondria are not the source of the early phase of intracellular

ROS production observed following MMS treatment, we next investigated whether ROS

generation by Noxes could be responsible. Treatment of the cells with the NOX inhibitor DPI

completely abrogated the cellular ROS production spike that peaked at 1 hour after -MMS

treatment in control WT MEFs. The effect of DPI pre-treatment, therefore, mirrored that of

90

pre-treatment with the antioxidant NAC (Fig 4.5). This suggests that Noxes are a strong

candidate for the source of the MMS-induced ROS generation 1 hour post treatment. DPI is,

however, not a highly specific inhibitor of NOX. Although it has been widely used in the

literature as a NOX inhibitor, it is actually general flavoprotein inhibitor. Other flavoproteins

that also produce ROS include cytochrome P450s [173]. It is unlikely that cytochrome P450s

were responsible for generating the intracellular ROS observed 1 hour after MMS treatment

because it is located in the mitochondria and Mitosox Red failed to detect ROS level change

at this time-point. DPI also increased MMS-cytotoxicity (Fig4.6).

It will be appropriate in the future to generate concentration response curve of DPI toxicity to

MEFs, to pick the highest non-toxic concentration to perform experiments with. On its own,

2µM DPI was not toxic to MEFs in control samples. Although we do not have any data to

show that within the 30 mins DPI pre-treatment, that NOXes were sufficiently inhibited, the

findings from the experiments using DPI align well with those from experiments using the

antioxidant NAC. The extent of the increase in MMS-cytotoxicity in cells seeded with DPI

was not as great as for cells seeded with 15mM NAC. The MMS-induced cytotoxicity was

further increased with higher concentrations of DPI, but at these higher concentrations,

some cytotoxicity was observed due to the DPI treatment alone. This may well be the result

of the non-specific inhibitory action of DPI on all flavoproteins. Overall, these findings were

entirely consistent with NOX being responsible for the increase in cellular ROS 1 hour

following-MMS treatment, but the poor specificity of DPI meant that this conclusion could not

be firmly drawn based on these experiments alone. Therefore, we attempted to obtain

confirmation by NOX knock-down using small hairpin RNA (shRNA). Although we were able

to transfect WT MEFs successfully with a plasmid carrying a GFP reporter construct,

attempts to obtain transfectants expressing shRNAs to knock-down NOX enzyme activities

were unsuccessful. MEFs proliferation slowed down when incubated with the selection drug,

entering a senescent state. Reducing the selection concentration from 0.8 µg/µl did prevent

MEFs entering senescence. The transfection experiments therefore had to be abandoned.

91

Overall, this chapter shows that the exposure of cells to MMS induces ROS generation,

which is abrogated by NAC. Aag-deficient cells did not exhibit this ROS generation post-

MMS exposure, suggesting that the initiation of BER was critical. MMS-induced ROS is

generated by NOXes and not the mitochondria, and inhibiting NOXes yields similar pattern

of MMS cell death results as was seen with the antioxidant results seen in previous

chapters.

92

Chapter 5: Effects of glucose concentration on the cellular

response to MMS

93

5.1 Introduction

The effect of glucose on MMS-induced cell death has been a relatively unexplored area. In

physiological conditions, cells produce their ATP through both glycolysis and oxidative

phosphorylation. In cell culture, low glucose medium makes cells consume a higher amount

of glutamine, which pushes the ATP generation more through TCA cycle and oxidative

phosphorylation compared to cells in high glucose that rely more on glycolysis [174]. Little is

known about how the different glucose levels supplemented to cells in medium directly affect

total NAD+/NADH and ATP levels. The glucose concentration in the medium used in our and

many other laboratories is 30mM, which is high compared to the physiological 5.5mM. 1mM

is below the physiological levels and is considered as low concentration.

It is generally believed that NAD depletion results in energy failure due to a drop in ATP. Our

results in the previous chapter challenge this scenario as we show ATP levels recover post-

MMS treatment even when total NAD+/NADH levels remain low. A recent paper has shown

that PARP-1 hyperactivation in cells cultured in the absence of glucose exhibit NAD

depletion but also an unexpected PARP-1 activity-dependent ATP increase [175]. The

authors also report that in the absence of glucose, AIF release from the mitochondria was

reduced. Another study has shown that treating yeast with MMS on a glucose rich medium is

accompanied by a strong suppression of glycolysis and increased toxicity [176]. Cells that

are supplemented with more nutrients, including glucose, tend to proliferate faster as nutrient

availability is not a limiting factor. The 3-MeA lesion generated by MMS blocks replication,

and triggers cell death if left unrepaired [85]. Cells with higher proliferation rates complete

the cell cycle more often in a given time, therefore have higher chance of replication fork

collapse and cell death due to the 3-MeA lesion [177]. It would be interesting to see how

glucose levels affect cell proliferation rate, and how this correlates with MMS-induced cell

death.

The aim of this chapter is to culture MEFs in different glucose concentrations, low (1mM),

normal (5.5mM) and High (30mM), over several passages. Cells will be treated with MMS to

94

see how different glucose concentrations affect MMS induced cell death. ATP and NAD

levels will also be measured to try and correlate findings with energy levels. We will also

detect cell proliferation rates of cells in difference glucose concentrations to see whether

different glucose has changed cell proliferation and how that potentially affects cell death

data.

95

5.2.1 MEFs proliferate slower at higher glucose concentrations.

Cells that are cultured in different glucose concentration may have different proliferation

rates. The experiment described here investigated whether culturing MEFs through five

passages in media with different glucose concentration altered their proliferation rate. Two

seeding densities were used: 150,000 (150k) and 300,000 (300k) total cells to be seeded in

6cm dishes. The reason for using two seeding densities was to account for the possibility

that the higher seeding density would reach confluency after 48 hours of culture, in which

case the lower seeding density would be more informative. Briefly MEFs were seeded at the

two seeding densities in low (1mM), normal (5.5mM) and high (30mM) glucose. Cell

numbers were determined at 24 and 48 after seeding using trypan blue.

At the 150k seeding density, there were no significant differences in proliferation between

MEFs seeded in media with different glucose concentration, either at 24 hours or 48 hours

after seeding (Fig 5.1 A). At the 300k cell seeding density, however, there was a significant

difference in proliferation between cells in high glucose compared to low and normal glucose

(Fig 5.1B). Cells in high glucose proliferated more slowly although this was only observed in

the 48h time point and not the 24h time point. Since the MEFs used are primary cells

(Passage 8-10), it is anticipated that too much glucose may cause the cells to halt

proliferation, until they are able to adapt to the high glucose. The cells had been cultured in

these concentrations for at least 3 passages before these experiments, with the only

exception of the cells cultured in 1mM glucose. MEFs that were to be cultured in 1mM

glucose were seeded from the cells that had been cultured in 5.5mM glucose previously.

This was because preliminary experiments had suggested that cells cultured in 1mM

glucose for several passages exhibited higher amount of cell death during culture. We

therefore routinely cultured them in 5.5mM and 30mM glucose concentrations, and then

seeded them in the three different glucose concentrations when performing the experiments.

Although this experiment provided an overview of MEFs proliferation rates in different

glucose concentrations, it was difficult to choose the most appropriate cell seeding density to

96

investigate MMS-induced cell death based on these data. Therefore, for the subsequent

experiment both cell seeding densities were used for the investigation of the effects of MMS

on MEFs cultured at different glucose concentrations.

97

Fig. 5.1

(A) 150k seeding density

(B) 300k seeding density

Figure 5.1. WT MEFs cell proliferation. Cells were seeded at (A) 150,000 cells and (B) 300,000 cells in 6cm dishes, in low

(1mM), medium (5.5mM) and high (30mM) glucose medium. Cell count was performed 24h and 48h after seeding. In both cell

seeding densities, cells did not have significant differences in cell counts after 24 hours between the different glucose

concentrations. After 48 hours, cells seeded in high glucose had lower cell counts when seeded in 300k seeding density,

compared to normal and low glucose concentrations. Cells seeded in the 150k cell density did not have any significant

difference in cell count after 48 hours between the different glucose concentration media. Data shown are the result of three

independent treatments (Two-way ANOVA Tukey post hoc test, mean ± SEM; **** p<0.0005).

98

1 mM

5.5 m

M

30 m

M

1 mM

5.5 m

M

30 m

M 0

100000

200000

300000

400000

500000

Glucose Medium

Cel

l cou

nts

per d

ish

24 hours48 hours

0

200000

400000

600000

800000

Glucose Medium

Cell

coun

ts p

er d

ish

24 hours48 hours

********

5.2.2 MEFs seeded in higher glucose exhibited higher MMS-induced cytotoxicity

The role of glucose concentration in MMS-induced cytotoxicity was investigated. The

previous results indicated that MEFs divide at equal proliferation rates when seeded at 150k

cells. In the cells seeded at 300k, MEFs proliferated more slowly in the 30mM glucose

concentration. To fully investigate the MMS-induced cytotoxicity of MEFs in different glucose

concentrations, MEFs were seeded at 150k and 300k cell densities in different media

containing different concentrations of glucose, treated with 2.5mM MMS the next day for one

hour in serum-free glucose-free medium, before changing medium back to media containing

the original glucose concentration. Cell death was determined with PI staining in conjunction

with flow cytometry.

At the 150K cell seeding density, there were no differences in cell death in control samples

(i.e. without MMS exposure) between the different glucose concentrations (Fig 5.2A). The

same was for true for cell seeded at the 300K cell density (Fig 5.2B). In the MMS treated

cells seeded at 150K cell density, there were no differences in cell death levels between

MEFs in low and high glucose media (Fig 5.2A). The same was true for the MEFs seeded at

300K cell density (Fig 5.2B). MEFs seeded at the 150k cell seeding density in high glucose

exhibited higher cell death compared to those in low glucose (Fig 5.2A), and cells seeded at

the 300k cell seeding density in high glucose exhibited higher cell death levels compared to

those cultured both in low and medium glucose (Fig 5.2B).

These results clearly demonstrated that cells seeded in high glucose levels were more

susceptible to MMS treatment, compared to medium or low glucose levels. These

observations contrasted with the previous cell proliferation data that demonstrated MEFs in

high glucose proliferated more slowly than MEFs in physiological or low glucose. To

investigate the associations between glucose levels, cell proliferation and sensitivity of MMS

exposure in more detail, the effects of different glucose concentration on cellular energy

levels, specifically NAD and ATP, were investigated.

99

Fig 5.2

(A)

Control

2.5mM M

MS0

20

40

60

80

Cel

l Dea

th %

1mM Glucose5.5mM Glucose30mM Glucose

150k Cell Density

*

(B)

Figure 5.2. Cell death of MEFs in

different glucose concentrations

when exposed to MMS. Cells were seeded overnight at (A) 150,000 cells and (B) 300,000 cells in 6cm dishes, in low (1mM),

medium (5.5mM) and high (30mM) glucose medium. Cells were exposed to 2.5 mM MMS for 1 hour. The next day they were

harvested and cell death was detected with PI using flow cytometry. There was no difference in cell death in control samples

between all media with different glucose concentration, at both the 150k and 300k cell densities. MEFs seeded at 150k in high

glucose exhibited higher cell death compared to cells seeded in low glucose. MEFs seeded at 300,000 in high glucose had

100

Control

2.5mM M

MS0

20

40

60

300k Cell Density

Cel

l Dea

th %

1mM Glucose5.5mM Glucose30mM Glucose

***

higher cell death than cells seeded in low and medium glucose. Data shown are the result of three independent treatments

(Two-way ANOVA Tukey post hoc test, mean ± SEM; * p<0.5, ** p<0.05).

5.2.3 MEFs seeded in higher glucose exhibited higher basal NAD levels.

To further investigate what could possibly cause the effect of MEFs cultured in high glucose

to be more sensitive to MMS-induced cell death, despite proliferating slower, total

NAD+/NADH levels pre and post MMS treatment in cell grown in the different glucose

concentrations were measured.

For this experiment only the higher cell seeding density (300k) was used as this was the

seeding density that had exhibited the most marked difference between glucose

concentrations in terms of cell proliferation and sensitivity to MMS. In control cells (not

exposed to MMS), MEFs cultured in high glucose had significantly higher total NAD/NADH

levels than did MEFs in normal and low glucose media (Fig 5.3). Upon MMS treatment, NAD

levels dropped in all three glucose concentrations 1 hour post treatment, with no difference

between samples. Intracellular NAD levels remained low for up to 4 hours following MMS

treatment. These results are striking because they again indicate the NAD levels post MMS-

treatment do not necessarily correlate with cell survival levels, as all MEFs cultured in

different glucose concentrations exhibited the same level of decrease in NAD levels

regardless of their sensitivity to MMS. The higher levels of total NAD+/NADH in MEFs

cultured in high glucose concentration could explain how these cells were more sensitive to

MMS-induced cell death than did the cells cultured in lower glucose concentrations; as the

higher NAD levels would theoretically translate into more PAR synthesis by PARP-1.

Sufficient PAR production alone, regardless of cellular energy levels, has been shown to be

able to induce necrotic cell death.

101

Fig 5.3

Control

1hr

2hr

4hr

0

100

200

300 1mM Glucose5.5mM Glucose30mM Glucose

NA

D/N

AD

H (n

M)

****

Figure 5.3. NAD+/NADH levels of MEFs in different glucose concentrations when exposed to MMS. Cells were seeded

overnight at 300,000 cells in 6cm dishes, in low (1mM), medium (5.5mM) and high (30mM) glucose medium. Cells were

exposed to 2.5 mM MMS for 1 hour and total intracellular NAD+/NADH levels were determined using a commercial kit. In

control cells, MEFs cultured in high glucose had significantly higher NAD/NADH levels than did MEFs in normal and low

glucose media. Upon exposure to MMS, MEFs in all different glucose concentrations had their NAD+/NADH levels decreased

with no significant differences between them. The NAD+/NADH levels remained low up until 4 hours post MMS-exposure. Data

shown are the result of three independent treatments (Two-way ANOVA Tukey post hoc test, mean ± SEM; ** p<0.05).

102

5.2.4 MEFs seeded in higher glucose exhibited higher ATP levels post MMS-treatment.

Having observed that MEFs cultured in high glucose had higher total intracellular

NAD+/NADH levels, intracellular ATP levels were analysed to determine whether or not

these followed a similar trend, and how the levels change following exposure to MMS. The

300k cell seeding density was used again for this experiment, the same seeding density

used in the NAD/NADH experiment. MEFs were seeded at 300k cell density in low, normal

and high glucose, and treated the next day with 2.5mM MMS for one hour. The media was

then changed back to media containing the original glucose concentration. Cells were

harvested at different time points following MMS treatment for the determination of

intracellular ATP levels.

In control cells, not exposed to MMS, there were no differences in intracellular ATP levels

between cells seeded in different glucose concentrations (Fig 5.4). There was no clear

reduction in intracellular ATP levels following MMS treatment in cells cultured in low or

normal glucose. In contrast, MEFs cultured in high glucose had ATP levels that were

significantly higher than MEFs cultured in low or normal glucose at 1 hour post-MMS

treatment, and the concentrations remained elevated until the last time-point analysed at 4

hours post-treatment. In the previous chapters, MEFs were cultured in 25mM glucose and

had a reduction in ATP levels at 1 hour post MMS-treatment before rising above base line 24

hours post treatment (Fig. 3.8). Although the results don’t agree at the 1 hour post-treatment

time point, possibly due to the difference in the high glucose concentration, they do agree in

the later time points 2-4 hours. This phenomenon of ATP levels increasing above basal

levels after MMS treatment further indicates that cell death is not caused by a drop in ATP

levels.

103

Fig 5.4

Control

1hr

2hr

4hr

0

2000

4000

6000

80001mM Glucose5.5mM Glucose30mM Glucose

ATP

Rel

ativ

e Lu

min

esce

nce

*****

Figure 5.4. ATP levels of MEFs in different glucose concentrations when exposed to MMS. Cells were seeded overnight

at 300,000 cells in 6cm dishes, in low (1mM), medium (5.5mM) and high (30mM) glucose medium. MEFs were exposed to 2.5

mM MMS for 1 hour and intracellular ATP levels were determined using a commercial kit. There were no difference in ATP

levels in control cells in MEFs seeded in different glucose concentrations. There were also no difference in ATP levels in MEFs

seeded in low and medium glucose concentrations throughout the experiment up to 4 hours post MMS-exposure. MEFs

cultured in high glucose had ATP levels higher than MEFs in low and medium glucose, after being exposed to MMS. Data

shown are the result of three independent treatments (Two-way ANOVA Tukey post hoc test, mean ± SEM; ** p<0.05).

104

5.2.5 MEFs did not utilize detectable amounts of glucose from the medium in the first

6 hours after medium change.

The previous results show that MEFs seeded in high glucose exhibited higher ATP and total

NAD+/NADH levels than their MEFs seeded in low and medium glucose levels. In regards to

NAD, the MEFs in high glucose had their NAD significantly higher than MEFs in lower

glucose before MMS treatment, compared to ATP which showed MEFs in high glucose only

had their ATP significantly elevated after MMS treatment. To try and relate this to relevant

glucose concentration, we measured glucose utilization from the culture medium by MEFs

seeded in different glucose concentrations. Briefly, MEFs were seeded in 3ml low, normal

and high glucose in 6-cm dishes. The medium was pipetted up and down in wells before 100

µl samples was taken for glucose analysis. Surprisingly, MEFs seeded in all glucose

concentrations did not show any detectable utilization of glucose over a 6 hour time course

after addition of fresh medium (Fig 5.5). It is evident that the assay was working as expected

since the apparent glucose concentrations in each of the different media were at or close to

the concentrations expected. The most likely explanation, therefore, is that within 6 hours

MEFs did not utilise sufficient glucose to deplete the medium to any detectable degree.

105

106

Fig 5.5

0 hr 1 hr 6 hr0

10

20

30

40

50

Glu

cose

con

cent

ratio

n m

mol

/L

1mM Glucose5.5mM Glucose30mM Glucose

Figure 5.5. Glucose utilisation by MEFs in different glucose concentrations. Cells were seeded overnight at 300,000 cells

in 6cm dishes, in low (1mM), medium (5.5mM) and high (30mM) glucose medium. The next day the medium was changed and

analysed for glucose levels over a 6 hour period. There were no significant change in glucose levels throughout the 6 hours in

the medium of MEFs seeded in all three glucose concentrations. Data shown are the result of three independent treatments

(Two-way ANOVA Tukey post hoc test, mean ± SEM).

107

5.3 Discussion:

This chapter describes studied performed to investigate the effects of cellular energy

supplies and intracellular energy levels on MMS-induced cell death. MEFs were cultured

initially in two different glucose concentrations; normal (5.5mM) and high (30mM) glucose.

The cells were then seeded for experiments in using three different glucose concentrations,

low (1mM), normal (5.5mM) and high (30mM). It was anticipated that cells grown in higher

glucose concentrations will proliferate faster due to the excess energy available. It was

hypothesized that cells that proliferate faster would be more prone to alkylation-induced cell

death owing to the production of the replication blocking lesion 3MeA by MMS. The first

experiment therefore aimed at investigating growth rates of MEFs in media with different

glucose concentration. Cells were counted over time using the trypan blue method, to

determine the proliferative rates of. Cells were also seeded at two different cell seeding

densities at 150k and 300k cells, to gain a more informative data in case the higher seeding

density might achieve confluence in the wells within the 48 hours of culture. MEFs were

counted 24 hours and 48 hours after seeding. Cells seeded at the 150k cell density exhibited

no significant differences in proliferation in the different glucose concentrations either at 24

hours and 48 hours post seeding (Fig 5.1a). The 300k cell density did show a difference in

proliferation rate. The cells seeded in normal glucose proliferated faster than the cells in high

glucose (Fig 5.1b, p>0.0005). The same was observed between low and high glucose.

MEFs seeded in low and normal glucose showed had no difference in proliferative rates.

This last observation might be explained by the fact that both samples had originated from

cells cultured in normal glucose levels. We were unable to continuously culture cells in low

glucose as they started to look unhealthy after a few passages, and decided to culture them

in normal and high glucose only for routine culture. Therefore, when performing the

experiments described the cells seeded in the low glucose had to that point been

conditioned to normal glucose levels. MEFs seeded in high glucose were observed to be

dividing slower over the 48hour time point course than MEFs in low and normal glucose.

108

This is believed to be because cells cultured in high glucose reached confluency before

seeding, and when seeded in lower density it took the cells some time to start diving again;

Whereas cells cultured in normal glucose did not reach full confluency prior to the seeding,

therefore cells were always in their log phase of proliferation. The next experiments

continued to be done with both cell densities to continue having a more informative outlook

on the correlation between cell density and the results of experiments.

The role of glucose concentration on MMS-induced cytotoxicity was next investigated. There

were no difference in cell death in control samples of MEFs seeded in different glucose

concentrations (Fig. 5.2A and B). MEFs seeded in high glucose however did show elevated

cell death compared to MEFs in low glucose concertation when seeded at 150k (Fig 5.2A),

and elevated cell death compared to both normal and low glucose when seeded at 300k (Fig

5.2B). It is not possible to associate this to proliferation state, as MEFs in high glucose were

proliferating more slowly than MEFs in the low and normal glucose after being seeded. Cells

cultured in high glucose over several passages rely more on glycolysis than oxidative

phosphorylation, and some reports have suggested that a block in glycolysis drives PARP-1

mediated cell death. It is possibly that MEFs cultured in high glucose relied more on

glycolysis, and when PARP-1 activity caused a block in glycolysis the cells entered a ‘shock’

state which resulted in more cell death. This could be investigated further using specific

probes that detect oxidative phosphorylation and glycolysis activity. It would also have been

informative to investigate cell cycle analysis using flow cytometry to count amount of cells in

G0/1 phase (growth phase), S phase (DNA replication) and M phase (mitosis); and to detect

difference in cyclin levels using antibodies to see how active in the cell cycle the cells were.

Total NAD+/NADH and ATP levels of cells were also measured, in an attempt to try and

correlate them with the above results. NAD and ATP experiments were performed only with

the 300k cell density as the two densities gave very similar pattern of results. In control cells,

MEFs cultured in high glucose exhibited significantly elevated levels of NAD+/NADH than

did MEFs in low and normal glucose levels (Fig 5.3). After MMS treatment, total NAD/NADH

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levels dropped to a similar level between all glucose concentrations. NAD+/NADH levels

post treatment could therefore not explain why MEFs in higher glucose exhibited higher cell

death post-MMS treatment. However, it is interesting that MEFs in high glucose had

significantly higher total NAD+/NADH levels. The PAR molecule (product of PARP-1), was

able to initiate PARP-1 mediated cell death by itself regardless of NAD levels [150]. This

mechanism could explain our observations since NAD levels dropped to the same levels

post MMS treatment, and because MEFs in higher glucose started with higher levels of total

NAD+/NADH, this could be taken as an indication that more PAR molecules were produced

as a result of DNA damage (more substrate more product). To further investigate this it

would be appropriate to quantify PAR levels using Western blot, to determine whether cells

in high glucose did indeed produce higher PAR levels post-MMS treatment than did MEFs in

low and normal glucose. Intracellular ATP concentrations were followed in MEFs after

exposure to MMS. This experiment showed there was no significant decrease in ATP levels

following MMS treatment in MEFs seeded in low or normal glucose (Fig 5.4). However, in

high glucose medium, ATP levels appeared to increase following MMS treatment. This

observation was consistent with result described in the previous chapter where MEFs

cultured in 25mM glucose also exhibited an increase in intracellular ATP levels following

exposure to the alkylating agent. This indicates clearly that ATP levels do not necessarily

mirror changes in intracellular NAD levels and, more importantly, it indicates that a decrease

in intracellular ATP levels is not the driving force for necrotic cell death following MMS

exposure. The observed increase in ATP levels following MMS treatment was unexpected

but confirmed in two independent sets of experiments and a further detailed investigation

would be required to elucidate the mechanism involved. One possible approach would be to

determine the activities of enzymes involved in the production of ATP, so at to pinpoint any

changes in this process on a molecular level.

Finally, we wanted to investigate glucose uptake from medium by MEFs in different glucose

concentrations. Glucose uptake would indirectly mean glucose utilisation by cells, therefore

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giving a further confirmation of different glucose conditioning of MEFs. Cells were seeded in

low, normal and high glucose in wells, as before and the supernatants were sampled at

several time points over a period of 6 hours post seeding. There were no detectable

changes over the time frame analysed in the concentrations of glucose in the media samples

suggesting that glucose utilization was at a level too low to detect. We are confident that the

apparent lack of change in medium glucose concentration was not due to any technical

problems with the assay itself. The standard curves produced were as expected and the

apparent concentrations of glucose determined in the different media matched expected

levels at time 0. It would therefore be a good idea to repeat this experiment for future work

using a longer time-frame. This glucose uptake experiment was also performed without

MMS treatment, and it would also be a good idea to repeat it with MMS treatment to see how

the treatment itself affects glucose utilization. Finally, analysis of intracellular glucose

concentrations, would be highly informative; indicating how glucose is utilized by the cells in

the presence of varying medium glucose concentrations.

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Chapter 6: Discussion

112

Antioxidants have been widely accepted to help protect against disease [118-120], but little

is known about whether they can also be harmful. The aim of this work was to investigate

how ROS and antioxidant afffect the cellular repsonse to MMS. The cells used in this project

were WT and Aag deficient Esc; WT and Aag-deficient MEFs; and WT photoreceptor cell

line 661W. We confirmed the pluripotency of ESc (Fig 3.1) prior to the experiments

conducted. This gave us assurance that experiments conducted on these cells were indeed

on exmbryonic stem cells in their undifferentiated state. The MMS cytotoxicity of WT and

Aag-deficient ESc were similar to published literature [85], in that Aag-deficient ESc were

more sensitive compared to their WT counterpart (Fig 3.2A).

MEFs were initially chosen as a model for differentiated cells, and we hoped to mimic the

response of differentiated cells in the adult mouse to MMS. In the newborn and adult mice,

Aag-deficiency rescued cells from the cytotoxic effects of MMS as compared to the WT mice

[97]. In our hands, Aag-deficient MEFs followed the same suit as ESc and were more

sensitive to MMS compared to their WT counterparts (Fig 3.2 B). We speculate that MEFs

behave the same as ESc to MMS due to the fact that MEFs originate from the embryo,

where ESc also originate from. This raises the possibility that though MEFs are differentiated

cells, their embryonic origin makes them behave different than fully differentiated cells in the

adult mouse.

We believe that using PI is more efficient to measure cell death than other cell viability

assays availbe in the market including the MTT assay. As cells die, their cell membrane

become permeable, allowing the PI to enter the cell and intercalate with DNA. PI is

otherwise impermeable to viable cells.

We also tested the effect of antioxidants on MMS-induced cytotoxicity. The main antioxidant

used in this project was NAC at the concentration of 15mM. NAC was supplemented to cells

in the medium 30 minutes prior to MMS treatment, and re-supplemented immediately after

treatment has finished. No NAC was given to cells during the 1 hour MMS treatment, to keep

the treatment consistent. Strikingly, NAC increased MMS-induced cytotoxicity in both WT

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and Aag-deficient MEFs (Fig 3.3 A and B). Similar results were also obtained with different

cell types (ESc) and different antioxidant (2-ME) (Fig 3.5 A and B). The concentration of 2-

ME used in this experiment was 0.1mM, and is the same concentration normally used to

culture ESc. The reducing environment achieved with 2-ME best mimics the partial hypoxic

environment of the embryo. We therefore measured the ROS levels in ESc with or without 2-

ME, and found ESc cultured without 2-ME had a higher baseline intracellular ROS levels in

comparison to those cultured with 2-ME (Fig 3.4). This proves that 0.1mM 2-ME, same

concentration used in the MMS experiment, reduced intracellular ROS levels in ESc. The

antioxidant accelerated MMS-induced cell death was also tested on photoreceptor cell line

661W, where NAC and a second mixture of antioxidants both increased sensitivity of cells to

MMS (Fig 3.6).

To try and pinpoint why antioxidants increase MMS-induced cell death, we measured

NAD+/NADH and ATP levels at baseline and post MMS treatment, with and without NAC.

MMS treatment reduced NAD+/NADH levels 1 hour post treatment (Fig 3.7), the earliest

time point analysed post treatment, and levels remained low up to the last time-point 4 hours

post treatment. This is in agreement with published literature [141]. NAC however, had no

effect in baseline NAD+/NADH levels pre-treatment, and up to 4 hours after treatment. As

NAD+/NADH levels did not change in the presence of NAC pre-treatment and post-

treatment, it was unlikely the driving force for the increased cell death observed in cells

supplemented with NAC and treated with MMS. We also investigated how antioxidants affect

ATP of MMS-treated cells at the same time-points as NAD+/NADH experiments. Cellular

ATP levels were significantly reduced 1 hour post MMS treatment (Fig 3.8), and

unexpectedly recovered 2 hours post-treatment. This is contradictory to some published

results which show ATP levels remain down below basal levels [143]. We anticipate that the

concentration, type of damage and length of treatment all play a role in determining whether

ATP levels recover or not. MEFs supplemented with NAC had higher basal ATP levels,

however MMS treatment significantly dropped ATP levels, following similar trend to cells

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without NAC. It is therefore unlikely that ATP levels to be the drive behind antioxidant

increase MMS cytotoxicity.

As antioxidants did not change the MMS-induced drop of ATP and NAD levels in cells, the

next step was to investigate if they changed intracellular levels of ROS. To test for ROS,

DCFDA-H2 was used to measure ROS and the results were analysed by flow cytometer. We

found that MMS treatment increased intracellular ROS levels as early as 30 minutes into

treatment, and peaking at 60 minutes (Fig 4.1). As expected, the antioxidant NAC abrogated

MMS-induced increase in ROS (Fig 4.1). ROS production following an insult is normal in

dying cells, where the produced ROS is involved in apoptosis. However, ROS production

involved in apoptosis is termed ‘bad’ ROS and if eliminated could reduce apoptosis. In our

casde, eliminating ROS with NAC increased MMS-induced cell death. This pushes us into

thinking the produced ROS must be beneficial to cells, acting as a messeneger. Strikingly,

Aag-deficient MEFs did not show any MMS-induced ROS generation, as monitored by

DCFA-H2, up to 4 hours post-treatment (Fig 4.2). This shows that the initiation of BER by

Aag must play a key role in the induction of ROS generation post MMS treatment, further

imposing that ROS role here is a messenger coordinating repair once it has been initiated. It

is not clear how the process of BER is linked with this ROS production or which step of the

BER process is key to this link.

We went on to investigate the origin of MMS-induced ROS, first focusing on mitochondrial

ROS. We used the MitoSox dye that specifically gets sequestered in the mitochndria and

measures mitochondrial superoxide ions. WT MEFs treated with MMS did not show any

elevated mitochondrial superoxide levels 1 hour post-treatment (Fig. 4.3). This suggests that

the intracellular ROS production detected as a result of MMS treatment was unlikely to be

originating from the mitochondria. However, we did detect an increase in mitochondrial

superoxide ion 4 hours post treatment, and this was further enhanced by NAC (Fig 4.3). We

believe this increase of mitochondrial ROS is late enough to be linked with dying cells. To

further confirm whether this was the case, we decided to measure the mitochondrial

115

membrane permeability which is reported to increase during mitochondrial dysfunction

leading to the release of pro-apoptotic and –necrotic factors as well as ROS production [170,

171]. Indeed as we treated MEFs with MMS, the number of cells with low polarised

mitochondria increased to a greater extent, peaking at the 4 hour time-point and further

intensified in cells supplemented with NAC (Fig. 4.4). This was the same time point of which

we observed had an increase of mitochondrial ROS. Low polarisation would be directly

linked to increased mitochondrial membrane permeability. This effectively shows that the

detected superoxide 4 hours post treatment is due to mitochondrial dysfunction.

Having confirmed that mitochondria are not the source of early phase ROS production

detected following MMS treatment, we next wanted to investigate whether NOXes were

involved. NOXes are flavoproteins specialised in superoxide production [160]. We used the

NOX inhibitor DPI to inhibit NOX activity, and measured cell death as well as ROS

production with and without MMS treatment. Adding DPI to culture medium completely

abrogated the intracellular ROS produced post-MMS treatment (Fig 4.5). This strongly

indicates that a NOX is behind the ROS production post-MMS treatment. Moreover, adding

DPI to the culture medium also increased MMS-induced cell death (Fig 4.6). Collectively DPI

gave the same trend in results as the antioxidant NAC did. There must be some sort of

cross-talk between Aag activity and NOXes, where the initiation of BER signals to the NOX

units to produce ROS. ROS has been shown to be important for the activity of many parts of

the repair pathway, including APE-1 [129, 130] and XRCC1 [131] [132]. We did try and

pinpoint which NOX was responsible for the ROS production, as there are several isoforms

of NOX. To test this, we transfected MEFs with different NOX shRNA’s to specifically knock-

down individual isoforms. Although transfecting MEFs was a success with GFP, we had a

difficult time during selection phase, as the cells went into senescence when stressed with

the selection drug. We could have pursued using spontaneously transformed MEFs to

increase transfection chances as these won’t enter senescence like primary MEFs during

116

selection. However, spontaneous transformed cells harbour excessive mutations at unknown

locations, making them possibly unfit for experimental use due to the chances of obtaining

incorrect data. MEFs also have a tendency to change behaviour very quickly if allowed to be

cultured more than Passage 20, and so we did not pursue this path.

The MMS-induced ROS production, as well as the drop in NAD and ATP levels encouraged

us to look into how intracellular energy levels can affect cell survival post MMS treatment.

We decided to culture primary MEFs in medium and high glucose media (5.5mM and 30mM

respectively); and perform experiments using low, medium and high glucose media (1mM,

5.5mM and 30mM respectively).

We first investigated the proliferative state of MEFs when seeded in two densities, 150,000

and 300,000. There were no difference in proliferation of MEFs in the different glucose

concentrations up to 48 hours when seeded at 150,000 (Fig. 4.1a); However in the 300,000

seeding density, cells seeded in high glucose proliferated slower than cells in medium or low

glucose (Fig 4.1b). There is a possibility that MEFs cultured in higher glucose proliferated

slower because they were confluent prior to seeding phase. This means the proliferation rate

would have been slower due to contact inhibition as compared to the cells cultured in

medium glucose that were in an exponential proliferation state already prior to seeing stage.

Cells in high glucose also showed higher MMS-induced cell death than cells in medium and

low glucose (Fig 4.2A) at both seeding densities of 150,000 and 300,000. It was interesting

to observe slower dividing cells undergoing higher MMS-induced death, as this indicates that

MMS-induced cell death doesn’t rely much on the proliferative state of cells. It would have

been interesting to obtain cell cycle data on cells cultured in the different glucose

concentrations.

We also measured the NAD+/NADH levels as well as ATP levels in MEFs seeded in low,

medium and high glucose. High glucose MEFs showed significantly higher baseline NAD-

117

levels than did medium and low glucose MEFs (Fig 4.3). All cells had their NAD levels

dropped to the same level after MMS treatment. This could explain why MEFs in higher

glucose exhibited higher cell death, as PAR molecule itself is able to induce cell death [150].

PAR is the product of PARP-1 activity, using NAD as a substrate. Our results indicate that

more NAD could have been used to produce PAR in high glucose MEFs than in low and

medium glucose MEFs, therefore eliciting a stronger death response. It would be a good

future step to investigate PAR levels in different glucose MEFs, and linking those with the

NAD data. There were no significant drop in ATP levels in MEFs seeded in low and normal

glucose (Fig 4.4), and in high glucose, ATP levels increased post-MMS treatment similar to

results in Fig. 3.8. We also investigated glucose uptake in MEFs cultured in different glucose

concentrations. There were no significant uptake of glucose within each glucose

concentration across the time points (Fig 4.5). It is unclear to why MEFs did not take up any

glucose in the first 6 hours of medium change. We think that measuring intracellular glucose

would be a better way for future experiments and will be more informative than measuring

glucose in medium.

In conclusion, our results have shown that MMS treatment can lead to ROS production that

is necessary for cell survival. Abrogating ROS using antioxidants would lead to higher MMS

cytotoxicity. We have shown that this ROS is not produced in the mitochondria, but rather

through a NOX isoform. We were unfortunate not being able to pinpoint which NOX isoform

that was producing ROS, but we did confirm that this ROS production required the activity of

Aag and therefore the initiation of BER. We also highlight that for the MMS concentration

used in this study, energy and metabolism dictate MMS cytotoxicity more than proliferation

state.

118

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