UBC Theses and Dissertations

UBC Theses Logo

UBC Theses and Dissertations

Measuring apparent oxidative damage to mitochondrial DNA by HIV antiretroviral therapy Sohi, Gurmeet Kaur 2012

Your browser doesn't seem to have a PDF viewer, please download the PDF to view this item.

Item Metadata

Download

Media
24-ubc_2012_fall_sohi_gurmeetkaur.pdf [ 2.72MB ]
Metadata
JSON: 24-1.0072779.json
JSON-LD: 24-1.0072779-ld.json
RDF/XML (Pretty): 24-1.0072779-rdf.xml
RDF/JSON: 24-1.0072779-rdf.json
Turtle: 24-1.0072779-turtle.txt
N-Triples: 24-1.0072779-rdf-ntriples.txt
Original Record: 24-1.0072779-source.json
Full Text
24-1.0072779-fulltext.txt
Citation
24-1.0072779.ris

Full Text

 Measuring Apparent Oxidative Damage to Mitochondrial DNA by HIV Antiretroviral Therapy  by  Gurmeet Kaur Sohi  B.Sc., Simon Fraser University, 2010  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF SCIENCE in The Faculty of Graduate Studies  (Pathology and Laboratory Medicine)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  April 2012  © Gurmeet Kaur Sohi, 2012 ii  Abstract Background/objectives: HIV antiretroviral therapy, specifically nucleoside reverse transcriptase inhibitors (NRTIs) have been associated with mitochondrial DNA (mtDNA) alterations, possibly through mtDNA oxidative damage leading to mitochondrial dysfunction, which is associated with degenerative diseases and aging. A published assay exploits that oxidative damage can slow down/inhibit DNA polymerase progression, such that the PCR amplification of damaged mtDNA template yields less product compared to undamaged mtDNA. I sought to optimize this assay using in-house tools and to quantify apparent mtDNA oxidative damage in cultured cells exposed to NRTIs. Methods: Three DNA quantification methods were compared: PicoGreen fluorescence quantification, UV spectrophotometry, and qPCR mtDNA copy number. Human hepatocellular carcinoma cells (HepG2) were exposed to hydrogen peroxide (H2O2) followed by recovery time to allow mtDNA repair. To determine whether NRTI exposure induces mtDNA damage, human coronary artery endothelial cells (hCAE) cells and human colorectal adenocarcinoma cells (HT29) cells that had been exposed to various NRTIs were subjected to the assay. To assess the assay’s future applicability to clinical samples, human skeletal muscle DNA samples were also assayed. Results: Quantification of long PCR mtDNA product by UV (CV=6.5%) and qPCR (CV=7.0%) showed lowest variability while PicoGreen quantification was noticeably higher (CV=21%). DNA from H2O2-exposed cells showed decreased amplification of long PCR product that increased with repair time. MtDNA depletion occurred in both cultures treated with stavudine. While there was no apparent mtDNA oxidative damage in HT29 cells with any NRTI, both tenofovir iii  and stavudine yielded increased mtDNA oxidative damage in hCAE cells. A wide degree of apparent mtDNA oxidative damage was observed in clinical samples. Conclusions: The preferred method for DNA quantification is qPCR mtDNA copy number. The observed mtDNA depletion indicated that NRTIs were active in both cell lines. The primary hCAE cells incurred greater mtDNA oxidative damage than cancer-derived HT29 cells. Cancer cells may have enhanced anti-oxidant mechanisms, suggesting that primary cells may be better model for studying mtDNA damage. The broad range of mtDNA damage detected in clinical samples bodes well for the assay’s use with diverse samples.              iv  Table of Contents  Abstract ......................................................................................................................................................... ii Table of Contents ......................................................................................................................................... iv List of Tables ................................................................................................................................................ vi List of Figures .............................................................................................................................................. vii List of Abbreviations .................................................................................................................................. viii Acknowledgements ...................................................................................................................................... xi Dedications ................................................................................................................................................. xii 1. Introduction .......................................................................................................................................... 1 1.1 Introduction to Thesis ......................................................................................................................... 1 1.2 HIV/AIDS .............................................................................................................................................. 1 1.3 HIV Pathophysiology ........................................................................................................................... 2 1.3.1 Structure ...................................................................................................................................... 2 1.3.2 Replication/Life Cycle of HIV ........................................................................................................ 3 1.3.3 Transmission ................................................................................................................................ 6 1.3.4 Infection ....................................................................................................................................... 7 1.4 HAART ................................................................................................................................................. 8 1.4.1 NRTIs ............................................................................................................................................ 9 1.4.2 Other Classes of HAART Drugs ................................................................................................... 12 1.4.3 Combinations of Drugs ............................................................................................................... 14 1.5 Mitochondria .................................................................................................................................... 15 1.5.1 Overview .................................................................................................................................... 15 1.5.2 Mitochondrial DNA .................................................................................................................... 17 1.5.2.4.1 Mitochondrial Disorders Mechanisms ................................................................................. 21 1.5.2.4.2 Oxidative Stress .................................................................................................................... 22 1.6 NRTIs and Mitochondrial Damage .................................................................................................... 24 1.6.1 The DNA POLG Hypothesis ......................................................................................................... 25 1.6.2 NRTI-Induced Oxidative Stress ................................................................................................... 26 1.6.3 MtDNA Variants ......................................................................................................................... 27 1.7 Oxidative Damage Assays ................................................................................................................. 29 1.7.1 High Performance Liquid Chromatography (HPLC) .................................................................... 30 v  1.7.2 Comet Assay ............................................................................................................................... 30 1.7.3 Dichlorofluoroscein (DCF) Based Assay ..................................................................................... 31 1.7.4 Gas Chromatography (GC) - Mass Spectrophotometry (MS) Assay .......................................... 32 1.7.5 Quantitative PCR (qPCR) Based Assay ........................................................................................ 32 2. MtDNA Oxidative Damage Assay ............................................................................................................ 35 2.1 Introduction ...................................................................................................................................... 35 2.2 Materials and Methods ..................................................................................................................... 38 2.2.1 H2O2 Experiments ....................................................................................................................... 38 2.2.2 MtDNA Oxidative Damage Assay Optimization ......................................................................... 40 2.3 Results ............................................................................................................................................... 47 2.3.1 Initial Quantification .................................................................................................................. 47 2.3.2 Long Template PCR Conditions .................................................................................................. 48 2.3.3 H2O2-Treated Cells ...................................................................................................................... 49 3. Assay Applications............................................................................................................................... 53 3.1 Clinical Setting ............................................................................................................................. 53 3.1.1 Samples ............................................................................................................................... 53 3.1.2 MtDNA Oxidative Damage Assay ........................................................................................ 53 3.2 Cell Culture Settings .................................................................................................................... 54 3.2.1 Human Coronary Artery Endothelial (hCAE) Cell Culture ................................................... 55 3.2.2 HT 29 Cell Culture ............................................................................................................... 57 3.3 Results ......................................................................................................................................... 59 3.3.1 Clinical Samples .......................................................................................................................... 59 3.3.2 Cultured Cells ............................................................................................................................. 60 4 Discussion ............................................................................................................................................ 66 4.1 Assay Optimization ..................................................................................................................... 66 4.2 Clinical Samples ........................................................................................................................... 70 4.3 Cultured Cell Applications ........................................................................................................... 71 4.4 Future Directions ........................................................................................................................ 74 4.5 Conclusions ................................................................................................................................. 76 References .................................................................................................................................................. 78  vi  List of Tables Table 1 NRTIs with the Nucleosides they Mimic ........................................................................................... 9 Table 2 Pros and Cons of Various Oxidative Damage Assays ..................................................................... 34 Table 3: Nucleic Acid Quantification Using the LightCycler 480 ................................................................. 42 Table 4: PicoGreen Fluorescence Quantification Standard Curve Dilutions ............................................... 43 Table 5: Primers Used in Apparent MtDNA Oxidative Damage Assay ........................................................ 45 Table 6: NRTI Settings with Primary Cells ................................................................................................... 55 Table 7: Primers Used to Determine MtDNA Content ................................................................................ 57 Table 8: NRTI Settings with Immortalized Cells .......................................................................................... 58           vii  List of Figures Figure 1: HIV Replication. .............................................................................................................................. 5 Figure 2: Factors Affecting HIV Transmission ............................................................................................... 6 Figure 3: Structural Properties of NRTIs ..................................................................................................... 10 Figure 4: Overview of Mitochondria ........................................................................................................... 10 Figure 5 Electron Transport Chain. ............................................................................................................. 16 Figure 6: Mitochondrial DNA. ..................................................................................................................... 18 Figure 7: Mechanisms of NRTI-Induced MtDNA Damage. .......................................................................... 28 Figure 8: PCR Amplification Curve. ............................................................................................................. 35 Figure 9: Overview of MtDNA Oxidative Published Method ...................................................................... 37 Figure 10: Experiment to Determine Ideal Quantification Method ........................................................... 44 Figure 11: Primers for Apparent MtDNA Oxidative Damage Assay. ........................................................... 45 Figure 12: Sample Standard Curve for PicoGreen Fluorescence Quantification. ....................................... 47 Figure 13: Comparison of Three DNA Quantification Methods. ................................................................. 48 Figure 14: Cycle Testing with Control HepG2 DNA ..................................................................................... 49 Figure 15: Results of H2O2 Trials ................................................................................................................. 50 Figure 16: Changes in Relative Amplification Over Repair Time................................................................. 50 Figure 17 Apparent MtDNA Oxidative Damage Assay Revised. ................................................................. 52 Figure 18: Representative MtDNA Long PCR Amplification of Human Muscle DNA.. ................................ 59 Figure 19: Longitudinal Apparent MtDNA Oxidative Damage Following NRTI Exposure ........................... 61 Figure 20: Longitudinal MtDNA Content in Cells Exposed to NRTIs. .......................................................... 62 Figure 21: Longitudinal Changes in Relative MtDNA Content (left) and Apparent MtDNA Oxidative Damage (right) in HT29 Cells Exposed to Various Concentrations of NRTIs over 30 Days ......................... 65 viii  List of Abbreviations Abbreviation Full Name 3TC Lamivudine 8OHdG 8-hydroxyguanosine A Adenosine (nucleoside)/Adenine (base) ABC Abacavir AIDS Acquired Immune Deficiency Syndrome AP Apurine/Apyrimidine ART Antiretroviral Therapy ARV AntiRetroViral ATP Adenosine TriPhosphate AZT Zidovudine/formerly called azidothymidine BC British Columbia BER Base Excision Repair bp Base Pair C Cytidine (nucleoside)/Cytosine (base) CD4+ Cluster Designation 4 positive lymphocytes CV Coefficient of Variation d4T Stavudine DCF Dichlorofluoroscein ddC Zalcitabine/formerly called dideoxycitidine ix  ddI Didanosine/formerly called dideoxyionosine DNA DeoxyriboNucleic Acid dNTP DeoxyriboNucleoside TriPhosphate ETC Electron Transport Chain FTC Emtricitabine (thymidine analogue with a stabilising fluorine atom) G Guanosine (nucleoside)/Guanine (base) GC-MS Gas Chromatography-Mass Spectrophotometry gp GlycoProtein HAART Highly Active AntiRetroviral Therapy HIV Human Immunodeficiency Virus Type-1 HPLC High Performance Liquid Chromatography II Integrase Inhibitor kb kilobases MT mtDNA nucleotide position mtDNA Mitochondrial DNA mRNA Messenger RNA NNRTI Non Nucleoside Analogue Reverse Transcriptase Inhibitor NRTI Nucleoside Analogue Reverse Transcriptase Inhibitor OXPHOS Oxidative Phosphorylation PHAC Public Health Agency of Canada PCR Polymerase Chain Reaction PI Protease Inhibitor x  POLG DNA Polymerase γ qPCR Quantitative Polymerase Chain Reaction RNA RiboNucleic Acid ROS Reactive Oxygen Species rRNA ribosomal RNA RT Reverse Transcriptase S.D. Standard Deviation T Thymidine (nucleoside)/Thymine (base) TDF Tenofovir/disoproxil fumarate tRNA transfer RNA UBC University of British Columbia UNAIDS Joint United Nations Programme on HIV/AIDS           xi  Acknowledgements Firstly, I would like to thank my supervisor, Dr. Hélène Côté for all her guidance, support and feedback throughout my research.  I have come a long way with my abilities to think like a scientist and realize my potential because of her mentorship and guidance. I would like to thank my supervisory committee, Drs. Haydn Pritchard, Jacqueline Quandt, Angela Devlin, and Neora Pick for their time, feedback, and involvement with my project. My lab members past and present have also helped me through this experience of graduate school and for that, I am very grateful for their support and friendship. Izabelle and Beheroze can be credited with teaching me the ways of the laboratory, answering all my questions, and helping me troubleshoot through the various phases of my project. Marissa Jitratkosol, Rachel Wade, Hayley Spencer, Tuhina Imam, Deanna Zanet and Sara Saberi – I thank them all for their words of encouragement, their comforting words and advice during stressful times. I would like to thank Tuhina and Hayley in particular for being there during the most trying times of my research and for keeping my spirits high. Working with everyone in the Côté lab, including summer students, Pinky Hapsari, Janet Lee, Mehul Sharma, and Adam Ziada, has been a pleasure. Last but not least, I would like to thank my family and friends for their constant support. They always gave me the encouragement and motivation I needed to excel and for their patience and understanding, I am extremely grateful. My parents and my siblings are the reason I have come this far and I would like to take this opportunity to thank them for all their contributions and support.   xii  Dedication     To my family and friends                1  1. Introduction 1.1 Introduction to Thesis Human Immunodeficiency Virus (HIV) antiretroviral therapy (ART) has been associated with toxicity and damage to mitochondrial DNA (mtDNA) [1-3].  One possible mechanism for this increased damage is via oxidative damage directly to the mtDNA genome or to the machinery responsible for mtDNA replication. The aims of my MSc research are to: 1) Optimize and validate an assay to quantify apparent mtDNA oxidative damage 2) Determine if certain drugs of the current antiretroviral therapy induce apparent mtDNA oxidative damage and if so, to quantify the extent of the damage. Cultured cells treated with varying concentrations of antiretroviral drugs were studied. We hypothesized that cells treated with some antiretrovirals would show an increase in apparent mtDNA oxidative damage. 1.2 HIV/AIDS The Joint United Nations Programme on HIV/AIDS (UNAIDS) reported that by the end of 2010, 34 million people were living with HIV [4]. Between 1997 and 2010, the number of new infections has dropped by 21% while the number of people gaining access to ART is reaching new highs [4]. There are 6.6 million people now receiving treatment in low- and middle-income countries[4]. Despite this success, there are more people than ever living with HIV and among them, the proportion of women globally has been and continues to be 50%, although in greater proportions in sub-Saharan Africa and the Caribbean [5]. 2  The introduction of ART to low- and middle-income countries in 1995 has since averted an estimated 2.5 million deaths but it is the rapid scaling up of highly active antiretroviral therapy (HAART) in the last few years, which has had the greatest impact [6]. In 2010 alone, an estimated 700,000 AIDS-related deaths were averted [4, 5]. Despite wider access to ART, education, and awareness about HIV infection, the HIV epidemic in North America has remained relatively stable since 2004 [4]. The number of people living with HIV in North America reached 1.3 million in 2010, an increase from the 980,000 in 2001; an increase which can be attributed to ART averting AIDS-related deaths and increased incidence since then [4, 5]. In Canada, an estimated 67,000 people were living with HIV in 2009, with 2300 to 4300 new infections occurring every year [7]. 1.3 HIV Pathophysiology 1.3.1 Structure Human Immunodeficiency Virus belongs to the genus Lentivirus of the Retroviridae, or retrovirus family, a class of viruses that contain RNA as their genetic material. The HIV genome is approximately 10kb in length and encodes for nine genes. Three genes encode proteins for viral structure (gag, pol, env) [8]. The structure of HIV is heavily comprised of three Gag proteins: viral p24, p17, and p7. These viral Gag proteins are formed when the HIVp55 Gag precursor polyprotein is processed by viral protease. 3  HIV is surrounded by an envelope which consists of host cell-derived lipid bilayer and virus- encoded glycoproteins (gp): gp120 external surface envelope protein and gp41 transmembrane protein [9]. These two glycoproteins are made from the gp160 precursor protein [10] and play an important role in the infection stage of the HIV life cycle. 1.3.2 Replication/Life Cycle of HIV HIV follows a lysogenic viral life cycle targeting cells of the immune system including dendritic cells, macrophages, and helper T-cells (Figure 1)[10].  The virus enters immune cells via adsorption of its surface glycoproteins to receptors on the surface of immune cells. Specifically, viral gp120 adheres to the CD4 glycoprotein receptor and one of the two co-chemokine receptors (either CCR5 or CXCR4) on the cellular surface of CD4+ T-lymphocytes. Once adsorbed, the viral envelope fuses with the cell membrane of the immune cells [10], and the HIV capsid, including the RNA genome, is released into the cell. Once inside the cell, an HIV- encoded enzyme, reverse transcriptase (RT), detaches the single-stranded RNA from its coinciding viral proteins and reverse transcribes the RNA into complementary DNA. RT then creates a sense DNA from the antisense DNA resulting in double stranded viral DNA which is then transported in to the cell nucleus and integrated into the host cell’s genome via a viral integrase[11].  The integrated HIV DNA within the host cell’s DNA is referred to as the provirus state. It can remain inactive with no HIV replication for a latency period of up to several years. When certain transcription factors are present, the T-cell becomes activated [12]. The integrated DNA provirus is then transcribed into messenger RNA (mRNA) by the host cell’s enzyme RNA polymerase, spliced, and transported out of the nucleus into the cytoplasm where translation of viral proteins occurs[11]. Viral structural proteins such as Gag and Env are then 4  produced from the mRNA and packaging of new viral particles can occur [13]. HIV is then assembled at the plasma membrane of the host cell; upon maturation, viral proteases cleave polyproteins into individual structures that are then utilized to produce mature virions. The newly assembled virus then undergoes budding, incorporating the host cell’s plasma membrane to act as its own outer envelope. This envelope also contains the glycoproteins necessary to bind to the CD4 receptors, readying the virus to infect new cells [13]. Various viral enzymes responsible in the HIV life cycle have been targeted for therapeutic intervention sites as described in Section 1.4       5   Figure 1: HIV Replication. (1) HIV binds to CD4 receptor and CCR5/CXCR4 co-receptor (2) The virus fuses with the membrane of the host cell (3) Viral RNA is released into the cytoplasm of the host cell (4) Viral RNA is reverse transcribed to DNA by the viral enzyme, reverse transcriptase, and then replicated into double-stranded DNA (dsDNA) (5) The viral dsDNA is transported into the nucleus (6) Viral DNA is integrated into the host cell genome by viral integrase and is transcribed into messenger RNA (mRNA) (7) Ribosomes translate mRNA, producing viral proteins (8) Viral proteases cleave viral polyproteins, forming mature viral proteins (9) Packaged viral particles undergo budding to be released from the cell (10). The released mature virions can further infect other cells. 6  1.3.3 Transmission Virus transmission and consequent infection is dependent on three factors: 1) characteristics of the agent such as its virulence and infectiousness, 2) host-related factors such as susceptibility, immune response and how contagious it is, and 3) environmental factors such as social, cultural, and political (Figure 2) [14].        Figure 2: Factors Affecting HIV Transmission. Factors affecting transmission of HIV are diverse, depending on viral characteristics, environmental influences and human behaviours. The transmission of the virus in the population occurs mostly via exchange of body fluid from sexual contact and contaminated blood products [15]. The four main ways HIV transmission occurs is by unsafe sex practices, contaminated needles, breast milk, and perinatal transmission when an infected mother passes it to her child [14]. The greatest risk of HIV transmission may be during acute HIV infection or the symptomatic period [16].           Pathogen: HIV • Virulence • Infectivity • Replication Rate  Host: Human • Susceptibility • Risk Behaviours • Immune response  Environment • Social • Cultural • Political 7  1.3.4 Infection HIV infection disease progression occurs in stages: 1) primary or acute HIV infection, 2) clinically asymptomatic, 3) symptomatic HIV infection and 4) progression to Acquired Immune Deficiency Syndrome (AIDS) [17]. The initial acute period lasts a period of 2-4 weeks which results in influenza type symptoms in 50-90% of those infected [18], and these are often overlooked. During this acute phase, the virus replicates at a very rapid rate, resulting in a dramatic increase in viral load, reaching possibly several million viral particles per millilitre of blood [19, 20]. This sharp increase in viral load is accompanied with a severe decrease in the number of circulating immune cells, which are the primary targets of HIV [19]. The immune cells affected predominately are CD4+ T- lymphocytes, macrophages, and dendritic cells [21-23]. The virus destroys CD4+ T cells whilst proliferating and therefore HIV infection is associated with a progressive decrease in the CD4+ T cell count with an accompanying increase in viral load. HIV carries out this role by directly killing infected cells, increasing apoptosis in infected cells, and indirectly via CD8 cytotoxic lymphocytes which recognize infected CD4+ T cells and attack them [24]. During this stage, there are high levels of HIV RNA, but not HIV antibody. Asymptomatic HIV infection lasts for a variable period of time, averaging 6-8 years but can vary from two weeks up to twenty years [25]. HIV RNA levels, a surrogate measure for viral replication, stay relatively stable during this time, while CD4 cells gradually decline over time [26]. During asymptomatic infection, the immune cells fight the virus continuously resulting in the decrease and following stability in the plasma HIV viral load levels. However, HIV is still 8  active in the resting T-cells [27]. If ART is administered earlier in the asymptomatic phase rather than later, survival is significantly improved [28]. The third stage, symptomatic HIV infection is indicated by the presence of clinical symptoms due to opportunistic infections  [29]. A person is diagnosed as living with AIDS when the CD4 cell count drops below 200 cells per cubic millimetre (cells/mm 3 ), a CD4 cell percentage from total lymphocytes of less than 14% or suffering from an AIDS-related opportunistic infection or malignancy [30]. Pneumonia, meningitis, tuberculosis and non-Hodgkin’s lymphoma are several conditions afflicting AIDS patients. Tuberculosis co-infection is the most common cause of AIDS morbidity [4, 31]. 1.4 HAART HAART is a treatment regimen consisting of various types of drugs and has been very successful in significantly reducing morbidity and mortality associated with HIV and AIDS, the first positive change since the recognition of the HIV epidemic [32]. HAART normally consists of three agents, two of which are nucleoside reverse transcriptase inhibitors (NRTIs) in combination with either a protease inhibitor (PI) or non-nucleoside reverse transcriptase inhibitor (NNRTI) [33] . Other drugs used within HAART regimen can include newer antiretroviral drugs that interfere at other points of the HIV life cycle such as fusion and entry antagonists, integrase inhibitors, and maturation inhibitors [34, 35]. By using a combination of drugs, the development of HIV resistance to drugs can be delayed or avoided altogether [36]. Success of HAART as an effective therapeutic regimen is heavily dependent on the compliance of patients to the drugs as the therapy is life-long. Adherence is 9  one of the most important patient-enabled predictors of therapeutic success and mortality [37]. However, the effectiveness of HAART is often hindered by low compliance of patients and drug-related toxicity [37, 38]. 1.4.1 NRTIs NRTIs act by mimicking the naturally occurring nucleosides, adenine (A), thymine (T), cytidine (C), gunanine (G), but they lack the 3’ hydroxyl group (Table 1). NRTIs currently used in the clinics include zidovudine (AZT), stavudine (d4T), lamivudine (3TC), abacavir (ABC), tenofovir disoproxil fumarate, a nucleotide analogue (TDF), emtricitabine (FTC), and didanosine (ddI). Zalcitabine (ddC) is no longer used as it showed excessive toxicity [39]. Table 1: NRTIs with the Nucleosides they Mimic. Nucleosides NRTI Drugs Thymidine AZT, d4T Cytidine ddC, FTC, 3TC Adenosine ddI, TDF Guanosine ABC  NRTIs competitively inhibit the conversion of HIV RNA to DNA by RT. The lack of the 3’hydroxyl end causes chain termination during replication [40]. This translates into decreased amount of viral RNA reverse transcribed into DNA. The overall effect is important as replication of the viral genome is hindered and proliferation and multiplication of the virus in the human body halted [40, 41]. Figure 3 shows the similarities in structure of NRTIs to natural nucleosides in the body. 10  Figure 3: Structural Properties of NRTIs. Structural properties of NRTIs strongly resemble naturally occurring nucleosides Thymidine ddC 3TC FTC ddI TDF AZT d4T Cytidine Guanosine Adenosine Nucleosides Analogs ABC 11  1.4.1.1 Cellular Activation and NRTIs In order for NRTIs to inhibit RT, they need to be first shuttled into the cell, and phosphorylated into their active triphosphate form [42] . NRTIs enter the cell either by passive diffusion or carrier-mediated transport [43]. ddC, 3TC and FTC, aka the cytidine analogues, are transported into the cell via carrier-mediated transport [44]. Anabolism of NRTIs to the triphosphate form, which competes with the naturally occurring dNTPs, occurs by the action of various nucleoside kinases and phosphotransferases [44]. A number of cellular factors affect NRTI phosphorylation. These factors can be divided into intrinsic that include cell type, cell cycle, and intracellular phosphorylated NRTI ratio, and extrinsic factors such as activation state of cell cyle and infection status [41]. In certain cells such as stimulated lymphocytes, the kinetics of phosphorylation are regulated as kinases in these cells are specific to a certain phase of the cell cycle [41, 45]. However, in other cells such as macrophages and monocytes, the mitochondrial kinase  displays constitutive expression and cell cycle is not an influential factor [41]. Cell studies suggest the most important factor in generation of the active intracellular NRTI phosphate species is the cellular activation state, as in whether the cells are resting or actively replicating [40]. A higher activation state of cells generally results in upregulation of kinases responsible for NRTI phosphorylation and nucleic acid synthesis [46]. HIV infection is also associated with cellular activation markers such as proinflammatory cytokinase, interferon and tumour necrosis factor [47]. This may suggest that in HIV-infected individuals, the rate of NRTI 12  phosphorylation is elevated inside the cell, increasing the levels of intracellular active drug, which can have implications in NRTI toxicity discussed in Section 1.6 [48]. There are various mechanisms for the breakdown and excretion of NRTIs including oxidation, conjugation, and transport [44]. Certain NRTIs (ddC, 3TC, FTC, and TDF)  do not undergo many changes in their organic structure and are readily excreted [44]. However, other NRTIs such as AZT, ABC, d4T, and ddI undergo structural changes during their metabolism and excretion processes, something that can affect their apparent concentration when administered with other agents[44]. 1.4.2 Other Classes of HAART Drugs 1.4.2.1 Protease inhibitors PIs target the viral protease thereby inhibiting the cleavage of polypeptide precursors necessary for the final assembly of the virions, disrupting formation of new viruses [49]. PIs currently used in HAART include atazanavir boosted with ritonavir (RTV), lopinavir (LPV), darunavir, saquinavir, indinavir, fosamprenavir [50]. 1.4.2.2 Non-NRTIs NNRTIs, like NRTIs, also target HIV RT but their mode of action is different. Instead of competitively inhibiting the RT by interacting with the active site of the enzyme like NRTIs, NNRTIs non-competitively inhibit the enzyme by binding to an allosteric site on the enzyme[51, 52]. This causes conformational changes to the enzyme, preventing it from catalysing the reverse transcription of viral RNA to DNA [51].  NNRTIs used in HAART treatment include efavirenz (EFV), nevirapine (NVP), delavirdine, etravirine, and rilpivrine [53, 54]. 13  1.4.2.3 Fusion and Entry Inhibitors Fusion or entry inhibitors disrupt the binding and entry of the virus into the host cell by inhibiting the various receptors responsible in the fusion process. Targets of this therapy theoretically can include CD4, gp120, CCR5, CXCR4, and gp41 [55]. Maraviroc is a CCR5 receptor antagonist [56]. Its mode of action involves binding to the CCR5 chemokine receptor on the surface of the CD4 T-lymphocytes, which as mentioned previously, is a co-receptor necessary for most strains of HIV to enter target immune cells. By blocking entry of the virus in to the cell, the virus is unable to begin its life cycle. Another drug in this class is enfuvirtide (also known as Fuzeon and T-20) [57-61]. Enfuvirtide binds to gp41 on the viral surface and disrupts its ability to bind. Other potential agents targeting the other receptors responsible in this step of the HIV life cycle have been or are currently undergoing investigation [62-64]. 1.4.2.4 Integrase Inhibitors Integrase inhibitors target the viral enzyme integrase responsible for incorporating viral DNA into the target cell’s genome [65]. Raltegravir is an II often used during salvage therapy at which stage the virus has mutated and developed resistance to the other classes of drugs. [66, 67] 1.4.2.5 Maturation Inhibitors Lastly, maturation inhibitors interfere with the virus’ gag protein processing, preventing generation of the mature capsid protein, p24 [68, 69]. The defective core resulting from immature protein structures causes the virus to exhibit decreased infectivity, slowing their rate 14  of replication. Although α-interferon is the only current maturation inhibitor used, Bevirimat is a potential maturation inhibitor currently undergoing clinical trials [70-73]. 1.4.3 Combinations of Drugs As described in section 1.4.2, HAART is a combination of antiretroviral drugs. The backbone drugs of HAART are the NRTIs, typically combined with one or two drugs from other classes. Although the combination of drugs decreases the chances of resistance, it still can develop in many people [74]. Recommended guidelines for first-line and second-line treatments are provided by the World Health Organization (WHO), and are influenced by factors such as pregnancy, accessibility, and affordability of drugs [38]. In the current state of HIV treatment, WHO-recommended first line regimen of drugs consists of the NRTIs TDF and FTC (or 3TC) with a NNRTI (either EFV or NVP)[67]. Until recently, the first line treatment regimen in resource- limited settings consisted of d4T and 3TC with a NNRTI (EFV or NVP) [67]. Due to toxicity of d4T [75, 76], it is no longer recommended [67], but in practice is still used  in many resource-limited countries. If resistance by the virus is developed against these drugs, second-line treatment usually consisting of AZT and 3TC and a PI in resource limited settings [67] or AZT, 3TC and an NNRTI in the developed world [50] is introduced. However, if this therapy plan also fails, salvage therapy consisting of newer antiretroviral drug classes such as integrase or entry inhibitors may be recommended [50, 67]. HAART is initiated when certain conditions are met. These guidelines are also set out by the WHO recommending that CD4 counts below 350 cells/mm 3  warrant initiating HAART[67].  15  1.5 Mitochondria 1.5.1 Overview Mitochondria are double membrane organelles found in eukaryotic cells (Figure 4). Their main function is the production of energy for cells, in the form of adenosine triphosphate (ATP). Energy production occurs via oxidative phosphorylation (OXPHOS) at the inner mitochondrial membrane by the electron transport chain (ETC). The proteins of the ETC act as electron acceptors, harnessing the transfer of electrons and subsequent crossing of oxygen across the membrane, to produce ATP (Figure 5) [77, 78]. In addition to energy production, mitochondria play significant roles in many other areas of normal and abnormal cell function, including cell signalling, cell growth, cell differentiation, and apoptosis [79, 80]. Cellular mitochondrial content varies depending on factors such as the type of cell, the cell’s metabolic and energy requirements, and cellular functions [81]. For example, cells of more metabolically active tissues such as skeletal muscle and liver have greater mitochondrial content and density compared to less metabolically active cells such as adipose tissue [81]. Further, mitochondria are dynamic organelles, with ongoing fusion and fission occurring within cells [82].         ETC mtDNA Outer mitochondrial membrane Inner mitochondrial membrane Mitochondrial matrix Intermembrane space Figure 4: Overview of Mitochondria. The mitochondria are double membrane organelles with and outer and inner mitochondrial membrane. The mitochondrial matrix houses mtDNA and is the site of greatest energy production. 16   Figure 5: Electron Transport Chain. The electron transport chain (ETC) at the inner mitochondrial membrane is the site of oxidative phosphorylation where energy is produced, in the form of ATP. Electrons are transferred from Complex I through V through electron carriers such as ubiquinone (Uq) and Cytochrome C (Cyt c). Reaction oxygen species (ROS) are formed as a by-product of the ETC and are released into the mitochondrial matrix where mtDNA is located. 17  1.5.2 Mitochondrial DNA 1.5.2.1 Overview An independent genome exclusive to the mitochondria is believed to be the evolutionary result of the endosymbiosis of prokaryotes some 1.5 billion years ago [83]. The double membrane described in the mitochondrial structure alludes to this theory as well, with the prokaryotic similarities also present in mitochondrial ribosomal structure [84]. MtDNA, as well as the mitochondrial organelle itself have lost most of their independence and are now dependent on nuclear control [83]. MtDNA is an independently-replicating circular genome, 16,569 base pairs (bp) in length (Figure 6) [85]. It consists of 37 genes, encoding 13 ETC proteins, 22 tRNAs and 2rRNAs [83, 85]. Although the majority of the approximately 2000 distinct proteins in the mitochondria are encoded by nuclear genes, the 13 proteins encoded by the mtDNA are involve in the ETC and are therefore essential for energy production and cell survival [83, 86]. Within a cell, there are many copies of mtDNA which were initially believed to be all identical in nucleotide sequence, a concept referred to as homoplasmy [84]. However, it is now well established that mtDNA variations can affect some of the many copies of mtDNA within a mitochondrion or within a cell. This results in a mixture of wild-type and mutated mtDNA genomes, a phenomenon referred to as heteroplasmy [83]. It is the relative ratio of wild-type to variant mtDNA copies that determines whether pathogenic or clinical phenotype is incurred. A certain threshold of mutant mtDNA copies needs to be reached based on the energy requirements of a certain cell, 18  where the number of mutant mtDNA copies hinders the efficient energy production resulting in pathology [83, 87]. During cellular replication and division, mtDNA is distributed asymmetrically to the new daughter cells [83, 88]. It is possible during this normal cellular process that the proportion of variant mtDNA copies may increase or decrease in daughter cells, depending on patterns of segregation. This phenomenon is referred to as mitotic segregation and is responsible for the emergence of clinically-relevant phenotypes later in life in patients with hereditary mtDNA- associated conditions [83]. The inheritance pattern of mtDNA differs from that of the Mendelian inheritance pattern inherent in nuclear DNA (nDNA) [88]. The mtDNA follows a mostly maternal inheritance pattern, as during fertilization, the fertilized oocyte destroys sperm mitochondria. However, there have been cases where paternal mtDNA blockage has not been absolute and paternal mtDNA inherited [89].       Figure 6: Mitochondrial DNA. The mtDNA is a circular genome 16 kb in length. The D-loop region is a regulatory region where initiation of mtDNA replication occurs. The mtDNA contains 13 genes that encode for proteins of the electron transport chain. 19  1.5.2.2 MtDNA Replication/Transcription Unlike nDNA, the turnover and replication of mtDNA is a continuous process, occurring at any or all cell cycle phases [86, 90]. Replication of mtDNA is carried out by a protein-enzyme complex, DNA polymerase-γ (POLG) consisting of two subunits, a large 125-140 kDA containing the polymerase and exonucleases catalytic activity and a smaller 41-55 kDA accessory subunit which allows for tighter binding of the complex with DNA [89, 91]. Both POLG subunits are encoded by the nuclear genome [91]. MtDNA replication is initiated at the non-coding regulatory D-loop region of the mtDNA, which also contains the promoters necessary for mtDNA transcription [89, 90]. MtDNA transcription is carried out by a single subunit RNA polymerase which is encoded by the nuclear POLRMT gene [89]. Transcription and translation of the 13 structural genes encoded by the mtDNA, all of which encode  ETC proteins, is dependent on mtDNA-encoded rRNAs and tRNAs as well, illustrating the importance of the mtDNA genome in maintaining OXPHOS and energy production function [85]. 1.5.2.3 MtDNA Repair Mechanisms The mtDNA replication machinery has very limited and basic repair mechanisms for correcting mistakes or adducts compared to nDNA [92]. The repair pathways are limited to base excision repair (BER), double-strand break repair, direct repair and possibly mismatch repair [86, 93, 94]. Nuclear DNA repair mechanisms have been better studied and understood [95] and in addition 20  to those seen for mtDNA, include nucleotide excision repair and recombination repair occurs. These have not yet been seen in mtDNA [94]. 1.5.2.3.1 Base Excision Repair BER  is the most significant factor in removing oxidative lesions or DNA adducts [95]. In nDNA, BER involves a class of enzymes, the glycosylases, followed by endonuclease or lyase activity on apurinic or apyrimidine (AP) sites. The repair then continues with one of the short patch, long patch, and single strand break repair pathways [95, 96]. In mtDNA, although less is known about the specific pathways of BER [93] , it is understood that BER is mostly involved in the repair of single strand breaks to mtDNA as wells as smaller lesions such as  8-hydroxyguanosine (8OHdG) and hydrogen peroxide (H2O2)-induced mtDNA damage [96, 97] [95]. Four enzymes are required for a complete mitochondrial BER pathway consisting of DNA glycosylase, AP- endonuclease, POLG, and DNA ligase [86]. In BER, the DNA glycosylases first remove the damaged base, resulting in an abasic site [86]. The AP-endonuclease then recognizes these sites and produces a 3’OH and 5’phosphate strand break continuing with the short patch pathway in mtDNA [86]. BER activity in mtDNA has been shown to be upregulated with increasing age, which coincides with 8OHdG accumulation seen with aging [95]. 1.5.2.3.2 Mismatch Repair Mismatch repair is responsible for correcting mispaired bases and small loop structures [98]. Although no mismatch repair proteins have yet been singled out in mammalian mtDNA, 21  evidence of mtDNA mismatch repair has been demonstrated in S. cerevisiae [98] as well as in rat liver mitochondrial extracts [99]. 1.5.2.4 MtDNA Abnormalities 1.5.2.4.1 Mitochondrial Disorders Mechanisms Genetic variants or other environmental factors that illicit changes in mtDNA, or nDNA coding for mitochondrial components, can result in the development of mitochondrial disorders [83, 100, 101]. The mitotic segregation, described in Section 1.5.2.1, can result in the abundance of mutated or defective mtDNA copies becoming localized in an area or organ of the body [83]. The “threshold expression” where the number of dysfunctional mtDNA copies results in clinical symptoms, can be specific to a small group of cells or just an organ. Therefore, mitochondrial disorders may appear to affect only some parts of the body [83]. Examples of mitochondrial disorders include chronic progressive external ophthalmoplegia, Kearns-Sayre syndrome, pigmentary neuropathy and maternally inherited Leigh syndrome [102]. The frequency of mtDNA mutations is high because of the limited BER mechanisms of POLG, which misses errors identified by more extensive mechanisms of proofreading such as those of the nuclear polymerases [103]. This accumulation of mutations over time, along  with clonal expansion of mtDNA over the lifetime, can result in the spontaneous appearance of mitochondrial disorders later in life [87]. Likewise, any damage or defect to POLG or the mitochondrial components responsible for mtDNA maintenance [103] can induce changes into mtDNA which can then manifest clinically as mitochondrial disorders if the dysfunction threshold is reached [87]. 22  1.5.2.4.2 Oxidative Stress Oxidative stress is a state where there is an imbalance in the production of reactive oxygen species (ROS) and antioxidants which fight against ROS to prevent damage, where the former exceeds the latter [104]. The mitochondrial ETC is responsible for the abundant production of ROS; as much as 95% of superoxide anions are generated as by-product of the ETC in normal metabolism [105]. This production occurs mostly at the levels of NADH dehydrogenase or coenzyme Q [106]. Approximately 1-2% of oxygen consumed by the mitochondria is converted into superoxide [107]. Other ROS include H2O2, lipid peroxides, hydroxyl radicals (OH), and peroxynitrite [1]. ROS interact readily with proteins, lipids and DNA. It can induce modifications to mtDNA by oxidative damage including strand breaks, protein-DNA cross-links, damage to deoxyribose, and base modifications [105, 108] . Common oxidation products of mtDNA include 8OHdG, a mutagenic lesion, and thymine/thymidine glycols (TGs) [109]. The close physical proximity of the ETC to mtDNA enclosed within the inner mitochondrial membrane makes mtDNA especially vulnerable to ROS-induced damage [108, 110, 111]. ROS are able to release iron (Fe) (II) from aconitase, which binds to mtDNA localizing oxidant production to the mtDNA [1]. Oxidation to the mtDNA template via such mechanisms can interfere with mtDNA replication and transcription. Another way for oxidative damage to interfere with mtDNA quality is by directly oxidizing the POLG protein, which is very sensitive, decreasing significantly its DNA binding efficiency. [112]. Because of the limited repair mechanisms of mtDNA replication compared to the extensive proofreading of nDNA, mtDNA bases accumulate 2-3x more oxidation than nDNA [113]. 23  As the superoxide generation is a physiological process, mitochondria have defence mechanisms to prevent injury from ROS in form of antioxidant enzymes [106]. These enzymes include manganese superoxide dismutase (MnSOD), glutathione peroxidase, glutathione reductase, glutathione, catalase, NAD(P) transhydrogenase, NADPH, vitamins E and C [1, 104, 106, 114]. Mitochondrial membrane proteins make up approximately 65% of the membranes in a cell [106]. Therefore, it is likely that along with mtDNA, mitochondrial proteins including those that make up the ETC, are also targets of ROS interactions[111]. For example, hydroxyl radicals can oxidize cysteine and methionine residues [106]. Oxidized mitochondrial proteins can disrupt normal physiological conditions of the mitochondria, including membrane permeabilization which plays an important role in  apoptosis [106], especially in cells under oxidative stress conditions [115]. Since mtDNA is responsible for encoding vital proteins of the ETC, damage to mtDNA can disrupt in the formation or function of these proteins [116]. The combination of ROS concurrently oxidizing mtDNA and mitochondrial proteins further fuels oxidative stress within the mitochondria, potentially causing even greater insult to mtDNA and other mitochondrial components [117]. This is highly relevant to human health as oxidative mtDNA damage has been associated with the aging process as well as many other conditions including Alzheimer’s disease [118-120], diabetes [121], cardiovascular disease [122],Huntington’s disease [123, 124], Parkinson’s disease[125], and multiple sclerosis [126]. 24  1.6 NRTIs and Mitochondrial Damage Although treatment with HAART has extended the life span of HIV-infected individuals, NRTI- related mitochondrial toxicity has resulted in serious side effects, negatively influencing treatment effectiveness [127, 128]. NRTI toxicity is considered  a key factor in the development of lipoatrophy and hyperlactatemia in HIV patients taking HAART [129]. HIV patients also suffer from age-related conditions earlier in life such as diabetes, cardiovascular events, and neurological disorders which may be attributed to mitochondrial toxicities [128]. Although some of the mechanisms behind these toxicities have been identified, further research is still needed to fully understand their effect in various human tissues and develop anti-HIV drugs with less mitochondrial toxicity. Before the NRTIs induce damage to mtDNA, a critical mass of NRTIs needs to reached intracellularly in order for NRTIs to compete with the nucleotide pool within the mitochondria. To achieve this, NRTIs must successfully be internalized and transported as well as phosphorylated [1]. NRTIs in their phosphorylated active form have been mostly implicated in the toxicities by various possible mechanisms, but some NRTIs have shown to deplete mtDNA by chain termination without being anabolized to the triphosphate form [130].  NRTI activation, as mentioned earlier in Section 1.4.1, is influenced by the transfer of nucleoside analogues in the cytoplasm and mitochondria[44]. The mitochondrial NRTI levels influence the antiviral activity and subsequent toxicity they may exert at the mitochondria [131]. 25  There are several considerations for the mechanisms of NRTI toxicity to mtDNA. It is likely that not one mechanism is solely responsible for mitochondrial toxicity, but rather the combination of several mechanisms. 1.6.1 The DNA POLG Hypothesis POLG is part of a protein-enzyme complex responsible for the replication of the mitochondrial genome [110]. While they have little effect on nuclear polymerases [132], NRTIs have been shown to strongly inhibit POLG [110, 133]. While this can clearly play a role in NRTI-induced mitochondrial toxicity, there may be an array of possible mechanisms. Nucleoside analogues may affect mtDNA quantity or quality as they [110]: - become incorporated into mtDNA and cause chain termination - inhibit POLG without being incorporated into the mtDNA and block replication directly - remain incorporated due to inefficient excision repair and alter the fidelity of POLG in mtDNA replication or through a combination of the above mechanisms. Disrupted function of POLG exonucleases activity, which serves as a repair mechanism for mtDNA, may also be a contributor to mitochondrial toxicity, as AZT has been shown to inhibit repair activity [134]. Whatever the mechanism, the resulting mtDNA depletion or mtDNA mutations can affect levels of transcription of ETC proteins transcribed by mtDNA, causing dysfunction of the ETC and OXPHOS [1]. Defects in mtDNA acquired by NRTI-induced dysfunction in mtDNA replication may yield phenotypic effects that mirror mitochondrial genetic disease caused by mtDNA point mutations and mtDNA deletions [87, 91] . It has been 26  suggested that the main consequence of mtDNA depletion is the subsequent decrease in energy production from impaired OXPHOS[91]. As seen in heteroplasmy and presentation of genetic mitochondrial diseases, the accumulation of deletion mutants may occur until a threshold of energy shortage is reached and clinical symptoms displayed [83, 87].  In genetic diseases, this phenomenon is referred to as the OXPHOS paradigm, and highlights that development of symptoms depends on the tissue requirements for OXPHOS and dysfunction threshold effects [91]. 1.6.2 NRTI-Induced Oxidative Stress Oxidative stress, as mentioned earlier, is the imbalance between the production of reactive oxygen species and countering antioxidant efforts. Evidence has been rising that NRTIs may be involved in the accumulation of oxidative damage to mtDNA [105, 135, 136]. Mice treated with AZT show increased molecular and ultrastructural oxidative damage in skeletal muscle and liver mtDNA [135, 137]. In HIV transgenic animal models, NRTIs demonstrated increased oxidation to mtDNA as well as oxidative stress levels [138]. In vitro studies also demonstrated increased oxidation of mtDNA [139]. Furthermore, levels of 8OHdG and other oxidative stress markers have been shown to be markedly increased in the brains and cerebrospinal fluid of HIV infected patients [140]. It is proposed that dysfunctional ETC and OXPHOS contributes to an increased production in ROS, tipping the balance to an environment of oxidative stress in the mitochondria [135]. These ROS are then available at higher levels to oxidize mitochondrial components such as proteins and mtDNA. This oxidation can further damage and disrupt the functioning of the already 27  disrupted ETC proteins and disturbed transcription of ETC proteins from mtDNA, amplifying the NRTI-mediated toxicity [135]. Further insults to mtDNA replication can occur by the induction of oxidative lesions to mtDNA itself as well as possibly oxidatively inactivating POLG [1, 135]. 1.6.3 MtDNA Variants The original mitochondrial oxidative stress theory of aging assumed that a “vicious cycle” of apparent mtDNA oxidative damage exists whereby  apparent mtDNA oxidative damage results in further ETC dysfunction, and more apparent mtDNA oxidative damage [141]. NRTI-related mitochondrial toxicity would also feed this vicious cycle. Newer studies have suggested that mitochondrial aging may be driven by the clonal amplification of mtDNA mutation to a point of becoming pathogenic, rather than the disseminated oxidative stress somatic mtDNA mutations themselves [87]. Similarly, it has been suggested that upon NRTI-induced mtDNA depletion (such as with d4T), a compensatory mechanism increases mitochondria biogenesis resulting in a rapid rate of mtDNA turnover [142-144]. This rapid turnover may lead to the clonal expansion of mtDNA point mutations incurred from NRTI toxicity and oxidative damage, which then become more prevalent and evident in cells [142, 143]. The increase in mutated mtDNA copies then is more likely to reach the threshold of dysfunctional energy production, contributing further to mitochondrial toxicity [91, 142]. An overview of the mechanisms described is highlighted in Figure 8.    28                      Figure 7: Suggested Mechanisms of NRTI-Induced MtDNA Damage. Mechanisms of NRTI Toxicity  Inhibit POLG exonuclease activity, preventing mtDNA repair  Competitively inhibit with naturally occurring nucleotide pools . • mtDNA depletion and dysfunction . • Decreased transcription of electron transport chain proteins . • Impaired oxidative phosphorylation ↓energy production  ↑Reactive Oxygen Species    mtDNA oxidative lesions mtDNA mutations and dysfunction Clonal expansion of mtDNA mutations Inhibition of mtDNA replication by chain termination  29  1.7 Oxidative Damage Assays There is currently no gold standard for measuring oxidative stress and oxidative damage in cells, let alone in mtDNA [145, 146]. Measurement of ROS directly is problematic, although common, because ROS react readily with other macromolecules in the cellular environment and with antioxidant systems [135]. Most methods used to quantify oxidative damage focus on “ROS” footprints resulting from the interactions of ROS with macromolecules such as DNA, proteins, carbohydrates and lipids [135]. For example, DNA altered by oxidation from ROS results in the generation of a wide variety of base and sugar modification products [147]. These products can be measured by various techniques, of which the following will be discussed here: high performance liquid chromatography (HPLC), comet assay, gas chromatography-mass spectrophotometry (GC-MS), and dichlorofluoroscein (DCF) assay [145]. A newer, quantitative PCR (qPCR)-based assay that can quantify apparent mtDNA oxidative damage directly as opposed to measuring levels of oxidation by-products, will also be described [148]. It must be noted that initial products of ROS-attack undergo transformations and therefore measuring a single reaction product as an indicator of oxidative DNA damage level is likely not the most reliable approach [145]. For example, measurement of 8OHdG, produced by oxidation of dG bases is widely used as an indicator of oxidative damage but only measures one type of oxidative damage. There exists a need for assays of greater practicality [145]. Furthermore, agreement on basal levels of 8OHdG in DNA is not established, and issues of 8OHdG formation during DNA isolation and preparation may in itself introduce artifacts muddying accurate measurement [145, 149]. Techniques that address this important limitation will make the detection and quantification of oxidative damage to DNA more robust, reliable, and universal. 30  1.7.1 High Performance Liquid Chromatography (HPLC) HPLC is a chromatography-based technique which separates compounds, and when coupled with electrochemical detection, can be used to measure 8OHdG [150] in both nuclear and mtDNA [151]. However, as mentioned, measuring a single product such as 8OHdG as an indicator of oxidative damage may not reflect the overall occurrence of oxidative damage to DNA [150]. Secondly, sample preparation for HPLC may incur oxidation and therefore overestimate 8OHdG levels [145, 150, 152]. Finally, as it is often impractical or even not feasible to physically separate mtDNA from nuclear DNA, the assay does not distinguish between the two DNA genomes. 1.7.2 Comet Assay A popular method in genetic toxicology, the comet assay, also referred to as single-cell microgel electrophoresis, can detect and quantify single strand breaks in DNA of individual cells [153]. The principle of the comet assay is that faster migration of smaller DNA fragments occurs in an electric field, compared to larger DNA fragments [154]. Treated cells are firstly captured in gel, lysed by alkali solution where DNA is denatured. Electrophoresis then allows DNA to migrate, with fragmented, smaller and damaged DNA moving faster than undamaged, larger DNA. This migration pattern results in the appearance of a comet with the undamaged DNA being the “head” of the comet and the fragmented, smaller DNA as the “tail.” Assessing oxidative damage with this assay exploits the fact that DNA breaks will result in fragmented DNA when oxidized bases with specific lesion sites are removed by repair 31  endonucleases [155]. Therefore, this method can be used to measure the amount of bases oxidized by oxidative stress. As mentioned in the introduction of Section 1.7, introduction of oxidation artifacts may occur with some methods. In a comparison of  HPLC described above and the comet assay, the latter showed 10 times lower endogenous oxidative damage rate, further indicating artifact introduction [156]. Although the comet assay can be used on many different cell types and intact cells [145, 154], it only addresses nDNA and not mtDNA, because as lysis starts, mtDNA disperses from around the nucleus. By the time electrophoresis begins, there is insufficient mtDNA remaining to detect it [155]. In nDNA, it is also difficult to distinguish whether the measured strand breaks are the result of oxidative damage or of other cellular events causing strand breakage, such as apoptosis [157]. 1.7.3 Dichlorofluoroscein (DCF) Based Assay Dichlorofluorescein (DCFH) is a popular probe used to measure intracellular ROS formation by flow cytometry [158]. The principle of this assay is that non-fluorescent fluorescin derivatives become proportionally fluorescent upon being oxidized with H2O2 [158]. DCFH precursors are applied to intact cells and once they cross cell membranes, enzymatic hydrolysis by cellular enzymes converts the molecule to the ready non-fluorescent DCFH. When ROS are present, the DCFH becomes oxidized to the fluorescent form, dichlorofluorescein (DCF). The overall intracellular fluorescence of DCF reflects the level of oxidative stress in the cells and can be measured using a fluorescence microplate reader. 32  Benefits of this assay are that it is relatively easy and a quick method to determine overall levels of oxidative stress in intact cells. ROS including H2O2, nitric oxide, superoxide and peroxynitrate have shown to oxidize DCFH to the fluorescent DCF [158], which is beneficial in assessing overall intracellular oxidative stress levels. However, this assay is limited in that damage induced to cellular components, such as DNA and proteins, is not quantified let alone which regions of either nuclear or mitochondrial genomes are oxidized. The translation of oxidative stress in a cell to the levels of damage incurred to DNA is an area of greater importance for an oxidative damage assay. 1.7.4 Gas Chromatography (GC) - Mass Spectrophotometry (MS) Assay GC-MS can be used to characterize various types of DNA damage ROS. In this method, DNA hydrolysis and conversions of products to volatile derivatives occurs. These are then separated by gas chromatography and mass spectrometry is used to identify the structure [150]. This method is useful as it allows for various oxidized base products to be recognized and therefore somewhat indicate which ROS was responsible for the oxidative damage [150]. Like HPLC, preparation of samples for the GC-MS procedure is prone to oxidation, in particular during DNA extraction and the hydrolysis steps [156]. In addition, again the level of DNA oxidative damage cannot be extracted by this method rendering it a qualitative rather than quantitative method. 1.7.5 Quantitative PCR (qPCR) Based Assay This assay utilizes the principles that damage to mtDNA can slow down or block the progression of PCR polymerase, resulting in reduced PCR product formation with greater apparent mtDNA 33  oxidative damage [148]. To observe this decrease in PCR product formation, emphasis on amplification to the exponential phase of the PCR is important. The assay addresses various issues compared to previously used assays. For one, it does not require isolation of mitochondria, which in itself is thought to increase the apparent detected oxidative damage by artifacts introduced in the process, as seen with HPLC [148, 150]. Secondly, the assay focuses on detecting changes to mtDNA which affect normal mitochondrial processes such as replication, instead of focusing on detecting oxidized bases or macromolecules [148, 150]. Limitations addressed in other assays such as the comet and DCF assays about identifying where oxidative DNA damage occurs is addressed, as primers chosen for the PCR reaction amplify specific areas of interest. Other practical benefits include the applicability of the assay to a wide range of research samples, requiring only a small amount of DNA for analysis, and being able to compare levels from one region to another[148]. With its benefits though are also limitations. For instance, the assay is limited to detecting damage brought on by oxidants that would inhibit or slow down the progression of the polymerase. Therefore, comparing results obtained from this assay to previous studies which have historically used 8OHdG as an oxidative stress indicator would be difficult as 8OHdG  does not slow down or inhibit PCR polymerase progression [148]. Additionally, the damage that is detected cannot be distinguished with respect to its type or origin [159]. In addition to oxidative lesions, it would likely also detect other damage such as single strand break, double strand break, etc…. A comparison of the benefits and limitations of the described assays is summarized in Table 2. 34  Table 2: Pros and Cons of Various Oxidative Damage Assays. Assay Pros Cons HPLC Can be used to measure both mtDNA and nDNA  Procedures may introduce artifactual oxidation, overestimating damage Specific regions affected cannot be identified Measures only one oxidation product (8OHdG) Comet  -Can be used on a variety of cell types -Only uses a small amount of DNA -Allows detection from intact cells, preventing detection of artifactual DNA oxidation during DNA isolation/analysis Can only be used on nDNA and not on mtDNA May measure strand breaks from sources other than oxidative damage, such as apoptosis  DCF  Can measure overall levels of oxidative stress in cells Detects a variety of important ROS Relatively easy and quick Can achieve lots of data in a single run Cannot identify/measure damage to specific cell components and regions Cannot distinguish which ROS most responsible GC-MS Can identify various oxidation products Procedures may introduce oxidation artifacts Specific regions of genes cannot be identified qPCR Wide range of samples can be studied (clinical, cell culture models) Specific regions/genes can be targeted for detection Small quantity of DNA needed (a few nanograms) Mitochondrial isolation not needed or mtDNA purification Damage between regions can be compared Oxidative lesions that do not stall PCR polymerase are not detected DNA damage that stalls PCR polymerase cannot be differentiated 35  2. MtDNA Oxidative Damage Assay 2.1 Introduction Santos et al. [159] published an assay describing a qPCR-based approach to quantify oxidative lesions to mtDNA. The principle of the assay exploits the fact that mtDNA oxidative damage or other DNA alterations will slow down or inhibit the progression of PCR polymerase, resulting in lower amplification of long PCR products. In order to capture differences in amplification of long template PCR product, however, the PCR reaction must be measured only during the exponential phase of the reaction [148].           Cycle Number P ro d u ct  A m o u n t     Undamaged mtDNA Damaged mtDNA Exponential Phase Amplification of damaged mtDNA relative to undamaged can be detected Figure 8: PCR Amplification Curve. Quantification of PCR product that has amplified to the exponential phase of the PCR allows for the calculation of relative apparent mtDNA oxidative damage. The inhibition of the PCR polymerase by oxidative lesions results in decreased mtDNA amplification, which can be detected in this phase and compared to an undamaged DNA template. 36  The published original assay involves quantifying total DNA via PicoGreen fluorescence, adjusting the total DNA concentration to 3ng/µL, amplifying the mitochondrial genome by long template PCR, and quantification of the long template PCR product by PicoGreen fluorescence. The analysis calculates the amplification of the long template PCR products in the sample of interest relative to undamaged control template also run through the assay and uses a Poisson calculation to express results as oxidative lesions per 10kb of DNA. However, it is important to note that the damage detected by this assay may encompass alterations other than oxidative ones and therefore the assay’s measure may be better described as the apparent mtDNA oxidative damage. In my research, I sought to adapt and optimize this assay. 2.1.1 Objectives The objectives for assay optimization were to: 1. Adapt the assay to use primers, DNA quantification assays, and instruments already available in our laboratory. 2. Optimize the different steps of the assay, including the quantification of initial DNA and long template PCR product, as well as parameters of the long template PCR reaction. 3. Adapt the analysis method 4. Determine whether cells subjected to oxidative stress showed increased and quantifiable apparent mtDNA oxidative damage. 5. Assess whether apparent mtDNA oxidative damage in NRTI-exposure cell culture models can be quantified using the optimized assay. Figure 9 illustrates the various step of the assay that were adapted, studied, and optimized. 37   Figure 9: Overview of Published MtDNA Oxidative Damage Method. First, cells are exposed to some agent that may induce oxidative damage. Total genomic DNA is extracted and quantified by PicoGreen fluorescence. The DNA of interest and control undamaged DNA concentration is adjusted to 3ng/µL and a 15kb mtDNA segment is amplified by long template PCR. The long PCR products are quantified again by PicoGreen fluorescence and the quantity of amplified damaged sample relative to undamaged product is calculated. A PCR is conducted on a short mtDNA fragment to normalize the results to mtDNA copy number. Steps highlighted by red squares indicate targets of assay adaptation or optimization. 38  2.2 Materials and Methods 2.2.1 H2O2 Experiments H2O2 is a prevalent ROS that has been shown to damage mtDNA as well as nDNA in various studies [154, 160-163]. In order to validate and optimize this qPCR-based apparent mtDNA oxidative damage assay, cultured cells were exposed to H2O2 at high and low concentrations to generate study samples with various degrees of oxidative damage. 2.2.1.1 Cell Culture An immortalized human hepatocarcinoma (HepG2) cell line was used to generate oxidized DNA through H2O2 exposure. These cells,  acquired from the American Type Culture Collection (CRL- 10741, ATCC, Manassas, VA, USA), were cultured at 37 o C, 5% CO2, in Dulbecco’s Modified Eagle Medium - High Glucose (D5648, Sigma-Aldrich, St. Louis, MO, USA) containing 10% fetal bovine serum (FBS, ATCC, Manassas, VA, USA), unless otherwise indicated. An immortalized human placental cell line (JEG-3) was also used to prepare various controls. These cells were also from ATCC (HTB-36, ATCC, Manassas, VA, USA) and were cultured at 37 o C, 5% CO2,  in Eagle’s Minimum Essential Media (30-2003, ATCC, Manassas, VA, USA) containing 10% FBS, unless otherwise indicated. 2.2.1.2 High H2O2 Experiment In this initial experiment, HepG2 cells were grown in 25cm 2 -t-flasks in 5mL of growth medium that was changed twice a week. Subculturing of the cells was done as instructed in [164]. Cells approximately at 80% confluency were treated with 1.8 M H2O2 (5 mL of stock 17M H2O2 added 39  directly to 45 mL of culture medium) for a period of 2 hours, after which time the cells were trypsinized, washed with PBS, centrifuged briefly and frozen at -20 o C for extraction. Control cells were left untreated and frozen similarly. This concentration was chosen to elicit a large amount of oxidative stress and oxidative damage. 2.2.1.3 Low H2O2 Repair Experiment In this second H2O2 experiment, HepG2 cells were cultured as above, and treated with a final concentration of 0.2M H2O2 for 1 hour. The cells were then given periods of 0, 15, 30, and 60 min to recover and undergo DNA repair with regular growth medium. After the 1 hour of H2O2 treatment, the medium was removed from all cultures. The repair time =0 and the control untreated cells were immediately trypsinized and frozen as above. For the other repair times (15, 30 and 60 min), the H2O2-containing medium was removed, the cells were rinsed once with PBS and replaced with regular medium for the indicated time before they were trypsinized and frozen. 2.2.1.4 DNA Extraction Total genomic DNA was extracted from the frozen cell pellets as per the manufacturer’s protocol using QIAamp® DNA Mini Kit (QIAGEN, Mississauga, ON, Canada) on a QIACube (QIAGEN, Mississauga, ON, Canada). 40  2.2.2 MtDNA Oxidative Damage Assay Optimization 2.2.2.1 DNA Quantification The published method uses PicoGreen fluorescence with a fluorescence plate reader to quantify total DNA in the sample to be assayed [148]. Quantification is done in a two-step process: Pre-quantification to 10ng/µL followed by adjustment of the DNA concentration to 3ng/µL, which is the amount of template DNA for long template PCR. In order to quantify total DNA, I compared the three following methods: UV spectrophotometry, mtDNA copy number by qPCR and PicoGreen fluorescence quantification. 2.2.2.1.1 UV Spectrophotometry DNA concentration was determined by measuring the sample’s absorbance at 260 nm using a NanoDrop spectrophotometer (2000, Thermo Scientific, Nepean, Ontario, Canada)[165]. 2.2.2.1.2 MtDNA Copy Number by qPCR MtDNA copy number quantification was done using a real-time quantitative PCR (qPCR) assay commonly used in our laboratory [166-168]. Briefly, a short fragment of the mitochondrial regulatory region, the D-loop, introduced in Section 1.5 was amplified with 1 µM of each of the primers MT325F and MT474R (Table 5), 1x of LightCycler 480 SYBR Green I Master (Roche, Laval, Quebec, Canada) and 2µL of the total DNA sample, in a final volume of 10µL. The PCR reaction was carried out in the LightCycler® 480 thermocycler (Roche, Laval, Quebec, Canada) with conditions of 95 ° C for 10 min followed by 45 cycles of 95 ° C/5s, 60 ° C/10s and 72 ° C/5s. The standard curve for quantification was generated from a serial dilution of a plasmid containing 41  this fragment. Each sample was quantified in duplicate. Two internal controls and a negative control were included in each qPCR run. The result was expressed as copies of mtDNA/µL. This assay was used to quantify mtDNA copy number in various steps of the apparent mtDNA oxidative damage assay described in Figure 17. 2.2.2.1.3 PicoGreen Fluorescence Quantification The Quant-iT PicoGreen dsDNA reagent is an ultrasensitive fluorescent nucleic acid stain which quantifies double stranded DNA (dsDNA). Quantification by this method requires a spectrofluorometer which performs fluorescein excitation (at 480nm) and measure emission (at 520nm). The published apparent mtDNA oxidative damage assay utilizes Quant-iT PicoGreen® Kit (Invitrogen, Carlsbad, CA, USA) and a fluorescence plate reader to quantify dsDNA. Since our lab did not have access to a fluorescence plate reader, we used our LightCycler® 480 (Roche); the instrument was programmed to perform the same excitation and emission as those necessary for quantification with PicoGreen as recommended by Roche in Table 3.     42  Table 3: Nucleic Acid Quantification Using the LightCycler 480. Programs Program Name Cycle Analysis Mode Equilibration 1 None Fluorescence Measuring 20 Quantification Ending 1 None Temperature Target Target (°C) Acquisition Mode Hold (hh:mm:ss) Ramp Rate (°C/s) Acquisitions (per °C) Equilibration 37 None 00:01:00 4.4 - Fluorescence Measuring 37 Single 00:00:02 4.4 - Ending 37 None 00:00:10 4.4 - Detection Format = SYBR Green I (483/533)  The PicoGreen dye was diluted 1:200 to its working concentration in kit-provided TE buffer. A LightCycler 480 96-well plate (Roche) was then loaded with 5µL of diluted PicoGreen dye and 5µL DNA sample for a final working volume of 10µL. Once a full plate was loaded, it was sealed, vortexed, centrifuged, and incubated at room temperature (protected from light) for 2-5 minutes before each run. A standard curve was initially prepared by serial dilution of the standard DNA provided in the kit, according to protocol. However, as the resulting standard curve did not follow linearity, adjustments were made in the dilutions and the range of the standard curve. The following 43  1:1.25 dilution series (Table 4) was used to generate a linear standard curve for PicoGreen fluorescence quantification with the LightCycler® 480 (Roche). Table 4: PicoGreen Fluorescence Quantification Standard Curve Dilutions. Initial Concentration (ng/mL) Dilution Factor Final Concentration (ng/mL) Stock (100,000) 1:50 2000 2000 1:62.5 1600 1600 1:78.1 1280 1280 1:97.6 1024 1024 1:122.0 819.2 819.2 1:152.5 655.4 655.4 1:190.7 524.3 524.3 1:238.3 419.4 419.4 1:297.9 335.5 335.5 1:372.4 268.4 268.4 1:465.5 214.75 214.8 1:558.6 171.8  2.2.2.2 Comparing DNA Quantification Methods To determine which of the three quantification methods of UV spectrophotometry, PicoGreen fluorescence, and qPCR mtDNA copy number would be most appropriate for DNA quantification, I determined the DNA concentration by subjecting DNA from the control HepG2 cells (section 2.2.1.2 ) through the first steps of the apparent mtDNA oxidative damage assay (Figure 10). Control undamaged HepG2 DNA was diluted randomly to generate eight DNA samples at varying concentrations. These 8 samples were then quantified by each of the three methods and all samples were diluted accordingly to a concentration of 3ng/µL as recommended by 44  Santos et al. [159]. The samples underwent long template PCR at the long template PCR conditions optimized as described in Section 2.2.2.2 below. The long PCR products were then quantified by qPCR mtDNA copy number and the coefficient of variation (CV) was calculated.               Total DNA DNA Quantification via: Adjust all samples to the same [DNA] Long PCR Agarose Gel Long PCR Product Quantification             PicoGreen               qPCR mtDNA copy number Analysis                          1             2              3             4              5              6             7             8  Dilutions PicoGreen            NanoDrop A 260     qPCR mtDNA copy number Figure 10: Experiment to Determine Ideal Quantification Method. Control HepG2 DNA was diluted in an 8-part series to generate samples. The eight DNA samples were quantified by the three methods of quantification (PicoGreen Fluorescence, UV Spectrophotometry, qPCR mtDNA copy number) and then adjusted to 3ng/µL or equivalent. The samples were all amplified with long template PCR and the long PCR products were quantified. The CV of the 8 long PCR products per initial quantification method was calculated. 45  2.2.2.2 Long Template PCR Conditions MtDNA Long template PCR was carried out using the Expand Long Range dNTPack Kit (Roche). Each reaction contained final concentrations of: 1x Long Range PCR buffer, 2.75mM MgCl2, 0.5mM dNTP, 0.3µM of each primer MT16535F and MT8388R or MT7988F or MT708R (Table 5), 3% DMSO, 1.25 units of enzyme (U/µL) and 6 µL of template DNA, in a total PCR reaction volume of 50µL. The PCR conditions were 93°C/2:00, 26 cycles of 93°C/0:10, 58°C/0:30, 68°C/8:00, 68°C/7:00, 4°C and the reaction was conducted on a MyCycler thermocycler (Bio- Rad, Hercules, CA, USA). Table 5: Primers Used in MtDNA Oxidative Damage Assay. he PCR primers used were obtained from IDTDNA (IDTDNA, Coralville, IA, USA). The lyophilized primers are dissolved in 10mM Tris-HCl to make stock, and working primers are made in DNAse/RNAse free water.     Primer Name mtDNA region Primer Sequence Long Template PCR MT16535F Fragment 1 5’- GCCCACACGTTCCCCTTAAATAAGA-3’ MT 8388R Fragment 1 5’- CGGTAGTATTTAGTTGGGGCATTTCAC-3’ MT7988F Fragment 2 5’- CTCCTTGACGTTGACAATCGAGT-3’ MT708R Fragment 2 5’- GGGGATGCTTGCATGTGTAATCTTAC-3’ qPCR on short fragment MT325F D-loop region 5’-CACAGCACTTAAACACATCTCTGC-3’ MT474R D-loop region 5’- AGTATGGGAGTGRGAGGGRAAAA -3’ Long PCR fragment 1   Short qPCR Long PCR fragment 2 Figure 11: Primers for Apparent MtDNA Oxidative Damage Assay. Two large fragments  (8.4 and 9.4 kb) are amplified during long template PCR (indicated in orange and green). A short 149-bp fragment (purple) in the D-loop region of the mtDNA is used to quantify mtDNA copy number. 46  Our long template PCR reaction, designed by the Côté lab amplifies the mtDNA genome in two large fragments (8.4 and 9.4 kb each) whereas the initially published assay amplifies a 15kb segment [159] . As our long template PCR primers and conditions varied from the published assay, control genomic DNA from HepG2 cells was used to test how many amplification cycles were needed to ensure the long template PCR product was amplifying to the exponential phase. The control HepG2 DNA was adjusted to 3ng/µL and this template amount was amplified by long template PCR with 24 to 35 PCR cycles. The long PCR products were then visualized on agarose gel to determine at which PCR cycle number there were sufficient products. 2.2.2.3 Analysis The published assay expressed mtDNA oxidative damage by first calculating the relative amplification of damaged long PCR products to control undamaged PCR product in the same run. The relative amplification was then converted to lesion frequency by applying a Poisson distribution [148].  For our analysis, we chose to express our results as the amplification of damaged long PCR product relative to undamaged control PCR product. 2.2.2.4 Reproducibility To assess the assay’s variability between runs, internal controls consisting of moderately damaged JEG-3 DNA and  H202-damaged HepG2 DNA were included in each long template PCR batch, in addition to the control undamaged JEG-3 DNA. Amplification of these internal controls relative to undamaged JEG-3 was calculated. Two plasmid DNA samples containing the 149bp fragment (a high and a low) were included as internal controls for qPCR mtDNA copy number. 47  2.3 Results 2.3.1 Initial Quantification The standard curve that was generated using PicoGreen fluorescence quantification with the LightCycler 480 technology ultimately showed a linear relationship over an effective concentration range for quantification.  Figure 12: Sample Standard Curve for PicoGreen Fluorescence Quantification. The standard curve generated is effective and linear for quantifying total DNA concentrations between171- 1600ng/mL.  The three different DNA quantification approaches were compared using the eight DNA dilutions (Figure 10). The qPCR copy number and NanoDrop Spectrophotometer methods both showed CVs of 7.0 and 6.5% respectively while the PicoGreen method showed a higher CV at 19%. These trends were visualized on an agarose gel (Figure 13). R² = 0.9985 0 10 20 30 40 50 60 0 500 1000 1500 2000 P ic o G re e n  F lu o re sc e n ce DNA Concentration (ng/mL) 48   Figure 13: Comparison of Three DNA Quantification Methods. Eight different dilutions of the same DNA were quantified by each of the three methods (PicoGreen fluorescence, UV spectrophotometry, and qPCR mtDNA copy number), their concentrations were then adjusted to 3ng/µL or equivalent, and each was then subjected to long PCR amplification. The long PCR products were visualized on this agarose gel and quantified by qPCR mtDNA copy number. The CV indicates the variation in the resulting mtDNA copy number.  With this experiment, it was determined that, for undamaged HepG2 DNA, 24,000 mtDNA copies as quantified by qPCR corresponds approximately to the total DNA concentration of 3ng/µL used by Santos et al. [169]. 2.3.2 Long Template PCR Conditions The cycle test performed determined how many PCR cycles would yield sufficient product, within the exponential phase, and well before the plateau phase of the reaction. As visualized in Figure 14, 26 cycles fit these criteria.   PicoGreen CV=19% UV Absorbance CV=6.5% qPCR CV=6.5%    +             - 1 kb 49         2.3.3 H2O2-Treated Cells DNA from HepG2 cells treated with 1.8M H2O2 demonstrated a dramatic decrease in long PCR product amplification compared to the control (Figure 14, lanes C and D). However, when the cells were treated with 0.2M H2O2 and given time to repair, the assay readily detected the smaller differences in apparent mtDNA oxidative damage. With increasing repair time, amplification of long PCR product became apparently stronger as visualized in Figure 15, 16. The apparent mtDNA oxidative damage was expressed as the amplification of long PCR product relative to the untreated, undamaged control.     +         -        24     25      26     27     28     29      30      32     35     1kb Cycles Figure 14: Cycle Testing with Control HepG2 DNA. DNA from control HepG2 was amplified with long template PCR from cycles 24 through 35 to determine at which cycle the DNA template amplifies to the exponential phase of the PCR. The 26 cycles fit the criteria and yielded sufficient long PCR product as visualized on this agarose gel. 50   Figure 15: Results of H2O2 Trials: DNA template from cells exposed to high H2O2 (left) and low H2O2 (right) amplified less compared to the unexposed controls (C). Decreased amplification is reflective of an increase in apparent mtDNA oxidative damage following H2O2 exposure. A PCR positive control (+) of megapool DNA and a negative control (-), water were also amplified.  Figure 16: Changes in Relative Amplification Over Repair Time. Long PCR product of DNA from 1.8M and 0.2M H2O2-treated cells amplified to less than 5% of the untreated control. An increase in relative amplification of H2O2 treated long PCR products occurs with repair time over 60 min.  0.00 0.05 0.10 0.15 0.20 0.25 0.30 Damaged (1.8M            treated) 0min repair 15min repair 30min repair 60min repair Amplification of sample relative to undamaged control          +       -     C       D        C        0       15      30     60     1kb   Repair Times (min) C = control D = damaged Damaged with varying repair times 0 .2 M  H 2 O 2  t re a te d  a t va ri o u s re p a ir  t im e  p o in ts  H2O2 High H2O2 Low H2O2 51  2.3.4 Reproducibility For both long fragments that were amplified by long template PCR, the PCR products of the internal controls (moderately damaged JEG-3 DNA and damaged HepG2 DNA) were quantified by qPCR mtDNA copy number and relative amplification of these relative to the undamaged JEG-3 DNA was calculated. Over several long template PCR reactions, there was a high variation in relative amplifications. For HepG2 internal control, (Fragment 1: n=8, CV=29.8%; Fragment 2: n=11, CV=40.5%). For the JEG-3 internal control, (Fragment 1: n=8, CV=46.6%; Fragment 2: n=11, CV=36.1%). Inter-run variability was much lower for the qPCR mtDNA copy number internal controls of the assay yielded lower variation between runs (IC-1: n=9, CV=6.2%; IC-2: n=9, CV=9.2%). 52   Figure 17: Apparent MtDNA Oxidative Damage Assay Revised. The newly optimized assay includes quantifying DNA by qPCR mtDNA copy number, amplifying the DNA in two large fragments by long template PCR, and an additional step of quantifying the adjusted DNA template also by qPCR mtDNA copy number. 53  3. Assay Applications 3.1 Clinical Setting The second of aim of my thesis was to determine if apparent mtDNA oxidative damage in human clinical samples could be detected using the optimized assay. A previous study in our laboratory was conducted to assess muscle mtDNA quantity and mtDNA deletion in patients with statin-induced myopathy (SIM), compared to comparators [170]. If the apparent mtDNA oxidative damage assay was successful in detecting variations in apparent mtDNA oxidative damage in these clinical samples, I hoped to determine whether a difference in apparent mtDNA oxidative damage existed between the two groups. I did not expect the apparent mtDNA oxidative damage to be as severe as that of my H2O2 cell culture experiments. However, despite antioxidant mechanisms and repair, we would expect that with the prolonged nature of ROS exposure over the lifetime, apparent mtDNA oxidative damage may be detectable. 3.1.1 Samples 3.1.2 MtDNA Oxidative Damage Assay As part of a previous study in our laboratory, DNA was extracted from retrospective muscle biopsies obtained from SIM subjects and comparators. Comparators were used as controls even though the patients were referred for a muscle biopsy for a reason. They were only used as controls if biopsies indicated normal or subclinical muscle pathology. Details about the study subjects and the method of DNA extraction can be found in [170]. Dilutions (1:100) of the total genomic DNA extracted from the muscle biopsy samples were the only DNA samples available for this project. These samples were first quantified by qPCR 54  mtDNA copy number as previously described. From the 44 samples, only 22 were of sufficient concentration for the assay. The 24,000 mtDNA copies of sample required for long template PCR is usually obtained by diluting samples to 4000 mtDNA copies/µL as 6µL of DNA template is added to the PCR. These dilutions could be made for the sufficiently concentrated 22 samples. For the remaining 22 samples that were too dilute, the exact volume required to achieve 24,000 mtDNA copies was calculated (under 10µL for all) and added to the long template PCR. To maintain similar long PCR conditions for all samples, 4µL of DNA extract elution buffer (AE buffer, Qiagen, Mississauga, Ontario) was added to the 22 samples of sufficient concentration. AE buffer was added to each sample accordingly to ensure each long template PCR reaction had 10µL of total DNA/AE buffer. Following the long template PCR, the long PCR products were quantified by qPCR mtDNA copy number and their amplification relative to the undamaged control DNA was calculated. Muscle apparent mtDNA oxidative damage was compared between the comparators and SIM group using the Mann-Whitney statistical test in Microsoft XLSTAT (Microsoft, Redmond, WA, USA). A p value of less than 0.05 would be considered significant. 3.2 Cell Culture Settings For this part of the project, we aimed to utilize the optimized assay to address our main study objective: determine whether NRTI exposure is associated with increased apparent mtDNA oxidative damage. To do so, we took advantage of resources from collaborators and received cell pellet samples from two different cell cultures treated with NRTIs and previously used for other analyses. Based on these collaborators’ results, we already had confirmation that NRTIs 55  had indeed entered the cells, been phosphorylated and were in their active form in the cultures [136, 171]. 3.2.1 Human Coronary Artery Endothelial (hCAE) Cell Culture  3.2.1.1 Hypothesis We hypothesized that following exposure to NRTIs, human coronary artery endothelial (hCAE) cells will show detectable increases in apparent mtDNA oxidative damage. 3.2.1.2 Materials and Methods The hCAE primary cells (C-1222, PromoCell GmbH, Heidelberg, Germany) were exposed to the following NRTIs: AZT, d4T, 3TC, and TDF for 30 days [172]. The following table (Table 6) presents the concentrations used in the experiments along with the maximum concentration these drugs are found in a physiological setting, referred to as the Cmax. The samples, collected at days 0, 8, 18, and 28, were from two independent cultures and the assays were done in duplicate. The cell  culture experiments were done in a similar fashion to [172].  Table 6: NRTI Settings with Primary Cells. NRTI Concentration (µM) Cmax (µM) Dissolved in AZT 1 7.1 DMSO 3TC 10 8.3 PBS D4T 10 4 PBS TDF 1 1.2 PBS    56  3.2.1.3 MtDNA Oxidative Damage Assay The DNA extracted from frozen cell pellets was quantified with qPCR mtDNA copy number. However, some samples showed mtDNA copy numbers as low as 500 mtDNA copies/µL. This concentration of mtDNA was too low to achieve 24,000 copies in 6µL so we decided to further adapt the apparent mtDNA oxidative damage assay to address this issue of low DNA concentration. Internal control samples (both damaged and non-damaged HepG2 DNA) were diluted to 500 mtDNA copies/µL and used in a long template PCR cycle test ranging from 27 to 32 cycles, to determine which conditions would allow samples to amplify to the exponential phase, starting with the lower mtDNA copy number. Once this cycle test was complete, all samples were diluted to 500 mtDNA copies/µL and long template PCR reaction was carried out as described in Section 2.2 but with 30 PCR cycles instead of 26. An additional step was included in the assay where the DNA templates diluted to 500 mtDNA copies/µL were quantified by qPCR mtDNA copy number. The long PCR products were then quantified by qPCR mtDNA copy number and analysis carried out as described in Section 2.2. 3.2.1.4 MtDNA Content Assay The cells’ mtDNA content was determined by calculating the ratio of a mitochondrial gene copy number to a nuclear gene copy number. Typically, the mitochondrial gene amplified is for the cytochrome c oxidase subunit I, CCOI. The DNA polymerase ɣ accessory subunit (ASPG), a single copy nuclear gene previously described [166] was also quantified. The qPCR copy number of 57  these two genes was carried out with the same PCR conditions as the mtDNA copy number for the short D-loop region described in Section 2.2.2.1.2, with the exception of different primers for these regions (Table 7). Prior to the usage of the hCAE cells in my project, both CCOI and ASPG were amplified and mtDNA content was calculated by a laboratory technician. Table 7: Primers Used to Determine MtDNA Content Primer Name Primer Sequence Gene: CCOI CCOI1F 5’-TTCGCCGACCGTTGACTATT-3 CCOI2R 5’-AAGATTATTACAAATGCATGGGC-3’ Gene: ASPG ASPG3F 5’-GAGCTGTTGACGGAAAGGAG-3’ ASPG4R  5’- CAGAAGAGAATCCCGGCTAAG-3’ he PCR primers used were obtained from IDTDNA. 3.2.2 HT 29 Cell Culture 3.2.2.1 Hypothesis We hypothesized that with increased exposure to NRTIs, human colorectal adenocarcinoma (HT29) cells will show detectable increases in apparent mtDNA oxidative damage over time. 3.2.2.2 Materials and Methods HT29  (ATCC, Manassas, VA) cells were exposed to the NRTIs: AZT, ddI, ABC, 3TC, d4T [171]. Detailed methods of this cell culture experiment can be found in [171]. Cells were exposed for a duration of 30 days and were collected periodically.   58  Table 8: NRTI Settings with Immortalized Cells.  3.2.2.3 MtDNA Oxidative Damage Assay All samples were assayed together to minimize variability with the revised apparent mtDNA oxidative damage assay as described in Figure 18. The long PCR reaction was done with 24,000 mtDNA copies for 26 cycles for all samples. 3.2.2.4 mtDNA Content The mtDNA content was quantified as described in Section 3.2.1.4. However, the mitochondrial D-loop region was amplified instead of the mitochondrial gene CCOI.       NRTI Concentration (µM) Cmax (µM) Dissolved in D4T 5,10, 20, 80, 160 4 PBS AZT 62.5, 125 7.1 DMSO TDF 50, 100 1.2 DMSO ABC 5, 10, 20 11.7 PBS 3TC 80 8.3 ꎯ ꎸ  ddI 30, 60, 120 11 PBS 59  3.3 Results 3.3.1 Clinical Samples Among the 44 muscle DNA samples studied, there was variation (range: 0.03-0.47) in relative amplification or apparent mtDNA oxidative damage (Figure 17). No samples showed greater amplification than the control, suggesting that all samples had incurred some apparent mtDNA oxidative damage. Theoretically, no clinical sample should amplify greater than the HepG2 culture non-damaged control as over time, human subjects would be expected to accumulate oxidative damage. While a wide range of  apparent mtDNA oxidative damage was observed overall, there was no difference in relative apparent mtDNA oxidative damage between comparators and SIM patients (n=22 per group, 0.26 ± 0.15 for SIM vs. 0.25 ± 0.22 for comparators, p>0.5, Mann-Whitney). There was a significant correlation between the two long PCR fragments (n=44, Pearson’s r=0.622, p<0.0001).     Figure X: Representative mtDNA long PCR amplification of human muscle samples. All clinical     C= control +      -      C                       representative set of clinical samples                            1kb Figure 18: Representative MtDNA Long PCR Amplification of Human Muscle DNA. Long PCR products of random samples visualized on agarose show varying levels of amplification. All samples amplify to a fraction of the control undamaged HepG2 DNA (C), indicating some level of apparent mtDNA oxidative damage in each sample. 60  3.3.2 Cultured Cells 3.3.2.1 HCAE Cells 3.3.2.1.1 Apparent MtDNA Oxidative Damage The apparent mtDNA oxidative damage is expressed as the amplification of sample mtDNA relative to an undamaged control (JEG-3 control DNA). Additionally, for each time point, the sample’s relative amplification was normalized to that of the corresponding control PBS culture. The relative amplification of the two independent cultures was averaged and is displayed in Figure 19. The two cultures showed significant correlation (n=38, Pearson’s, R 2 =0.452, p=0.002; Spearman’s, coefficient of determination=0.564, p=0.0003). Over the period of 30 days, the greatest decreases in relative amplification, and therefore most apparent mtDNA oxidative damage, occurred with d4T, followed by TDF. Both AZT and 3TC remained relatively stable in apparent mtDNA oxidative damage levels over this time course.    61  A  B  Figure 19: Longitudinal Apparent MtDNA Oxidative Damage Following NRTI Exposure in HCAE Cells. A: Following exposure to various NRTIs, the amplification of sample mtDNA was calculated relative to an undamaged JEG-3 control and then normalized to corresponding PBS culture time points (panel B). B: The untreated PBS culture showed an increased in apparent mtDNA oxidative damage over the course of the culture.    0.0 0.2 0.4 0.6 0.8 1.0 1.2 1.4 0 5 10 15 20 25 30 a m p li fi ca ti o n  o f sa m p le  m tD N A re la ti v e  t o  u n d a m a g e d   c o n tr o l Days in culture AZT (1 µM) 3TC (10 µM) d4T (10 µM) TDF (1 µM) 0.00 0.10 0.20 0.30 0.40 0.50 0.60 0 5 10 15 20 25 30 A m p li fi ca ti o n  o f sa m p le  r e la ti v e  t o  u n d a m a g e d  c o n tr o l Days in culture 62  3.3.2.1.2 MtDNA Content With d4T treatment, over time mtDNA depletion was strongly evident. A relative increase in mtDNA content over the 30 day period was observed for all other drugs, similar to that seen in the control cultures. A  B  Figure 20: Longitudinal MtDNA Content in HCAE Cells Exposed to NRTIs. A: MtDNA content normalized to that of the untreated control culture. MtDNA depletion is observed over time with d4T treatment. B: In untreated control cells, mtDNA content increases over the course of the culture. 0.0 0.2 0.4 0.6 0.8 1.0 1.2 0 5 10 15 20 25 30 R e la ti v e  m tD N A  c o n te n t Days in culture AZT 3TC d4T TDF 0 100 200 300 400 500 600 0 5 10 15 20 25 30 m tD N A  c o n te n t (m tD N A /n D N A ) Days in culture 63   3.3.2.2 HT 29 Cells Results Results of both mtDNA content and apparent mtDNA oxidative damage of the HT29 cells exposed with NRTIs are included in Figure 21. Some important observations include the dose- dependent mtDNA depletion seen with both ddI and d4T exposure. No NRTI treatments showed any relative difference in apparent mtDNA oxidative damage over time.   64   65   Figure: L Figure 21: Longitudinal Changes in Relative MtDNA Content (left) and Apparent MtDNA Oxidative Damage (right) in HT29 cells Exposed to Various Concentrations of NRTI over 30 Days. Lower relative amplification of sample long PCR product implies greater apparent mtDNA oxidative damage 66  4. Discussion  4.1 Assay Optimization The benefit of this qPCR-based apparent mtDNA oxidative damage assay is that it addresses the direct insult on the mitochondrial genome by elevated ROS and oxidative stress. Up until this assay was developed, the most popular oxidative biomarkers and measures of oxidative damage have concentrated on measuring either the levels of known ROS or their oxidation by- products, instead of assessing whether these actually resulted into mtDNA injury. Although, the nature of this assay prevents it from quantifying mtDNA oxidative lesions that do not slow down polymerase progression, the gene specificity achieved and direct measurement of mtDNA damage provides additional information with regard to oxidative damage. A limitation of the assay is that any mtDNA damage that would slow down the polymerase progression, including oxidative damage but also other types of DNA damage, could affect measurements. Therefore, it cannot be confirmed that the damage is solely ROS-related as other alterations may have similar effects. In the future, it may be interesting to study the correlation between apparent mtDNA oxidative damage measured by this qPCR-based method in relation to those other measures of oxidative stress to determine the strength of the correlation between ROS and the damage detected with this assay. Adapting the initial quantification method from total DNA to mtDNA copy number by qPCR is more useful since mtDNA is the target of the long PCR amplification. Furthermore, depending on the type of tissue (e.g. fat vs. muscle), the actual amount of mtDNA may vary greatly for a given total DNA concentration. PicoGreen fluorescence and A260, do not distinguish between 67  mtDNA and nDNA and the A260 method measures ssDNA and RNA along with dsDNA. Since this assay relies on a specific dsDNA input into the long template PCR reaction, and since A260 does not lend itself to quantifying unpurified PCR products because of dNTP/primer interference, this method was not preferred.  Although Santos et al. quantify total DNA and normalize to mtDNA copy number afterward [169], large fluctuations in initial mtDNA copy number could influence the long template PCR amplification which would not be accounted for completely. Therefore, it is beneficial to use qPCR for mtDNA copy number. The short 149-bp segment from the D-loop region used for the qPCR reaction was chosen for several reasons. Because the D-loop is a regulatory, non-coding region, it does not have pseudogenes and would therefore truly amplify only mtDNA. This area is also not prone to deletion or mutation so chances of underestimating mtDNA copies would be avoided [173]. By quantifying mtDNA instead of total DNA, the variation in mtDNA copies in different cell types is accounted for. For example, muscle and more metabolically active cells tend to have an increased density of mitochondria within cells and consequently mtDNA, compared to less metabolically active tissue such as adipose tissue [81]. In metabolically less active tissues then, using total DNA to quantify the sample would result in the input of fewer mtDNA copies into the long template PCR reaction, something that could increase the risk of measuring long PCR product outside of the exponential range of the PCR amplification. 68  Furthermore, the additional step of quantifying the template DNA after its concentration adjustment for long template PCR introduced in our optimization allows for more accurate analysis, because the exact amount of DNA template amplified is accounted for. In choosing an optimal cycle number with our long template PCR tools, amplifying the sample sufficiently to the exponential phase of the long template PCR amplification reaction was an important aim in optimization. As shown in Figure 14, 26 cycles fit the criteria and was chosen. Although lower PCR cycles (24 and 25 cycles) fit the criteria of amplifying to the exponential phase, to ensure amplification would be sufficient with each run, we had aimed for the cycle of amplification to reflect more so the middle of the exponential phase (Figure 8). Choosing 27 or 28 cycles would risk the sample amplifying to the plateau phase and therefore render it non- applicable for relative amplification analysis.. The analysis method was changed from a Poisson analysis expressing mtDNA oxidative lesions as lesion frequency, per the equation lesion frequency = -ln (relative amplification) to simply expressing apparent mtDNA oxidative damage as the amplification of damaged samples relative to an undamaged control. As the nature of the lesions cannot be differentiated by the assay, we felt that results would be better expressed as the “apparent mtDNA oxidative damage” thereby acknowledging that oxidative stress may not be the only factor involved. With the introduction of the additional template DNA quantification step (Step 4a in Figure 17), normalizing long PCR product amount to the initial template added to long template PCR (Step 3 in Figure 17) strengthened the data. This introduction allowed small variations in initial template, which 69  could influence greatly the amplification over 26 cycles, to be addressed and incorporated into the analysis. The H2O2 experiments were paramount in establishing and confirming the functionality of the assay. The main findings of the cell culture experiments gave indication that the assay was able to detect apparent mtDNA oxidative damage both with obviously high ROS exposure (1.8M H2O2) as well as with smaller changes in apparent mtDNA oxidative damage as observed with the 0.2M H2O2 exposure followed by repair times. The repair trial experiment could be repeated with longer repair times in the future to determine if amplification eventually returns to baseline, but this was not a primary goal in establishing the assay’s functionality here. These initial experiments although conducted only once mirrored the results of Santos et al. suggesting that this assay could be a robust and effective tool in quantifying apparent mtDNA oxidative damage. The 1.8M H2O2-treated culture showed remarkable cell death and a visibly low ratio of cell adherence to the culture flask at the end of the 2-hour exposure. This cell death was anticipated as the high concentration of H2O2 was used particularly to ensure sufficient damage to cells and mtDNA. With the 0.2M H2O2 treatment, cell death was minimal and adherence to culture flask was unchanged, providing confirmation that this concentration was more appropriate in giving cells the opportunity to tolerate the oxidative environment and respond to apparent mtDNA oxidative damage.   70  4.1.1 Limitations of Assay Considering the various adaptations brought upon the assay, some considerations need to be highlighted. Run-to-run variability was an issue as long PCR enzyme efficiency was variable between batches of enzyme, affecting reproducibility. To address this issue, it was important that experiment sets be amplified with the same batch of long PCR Taq polymerase enzyme. Certain measures were also introduced to improve quality control and reproducibility of the assay, including the use of internal controls. These internal controls included moderately and undamaged DNA from JEG-3 cells and damaged DNA from HepG2 cells. While these were used to evaluate the assay’s variability, they also allowed me to visualize that expected trends were seen with each long PCR amplification reaction. Due to the high run-to-run variability reported, practices such as pooling of enzyme before experiments, running all samples for a given experiment together, and utilizing internal controls will be essential in moving forward with the assay. 4.2 Clinical Samples The main rationale and finding observed from assaying the mtDNA of muscle biopsy clinical samples was the indication that the assay could detect a broad range of apparent mtDNA oxidative damage with diverse samples. This set of samples also presented the challenge of having lower DNA concentration and therefore fewer mtDNA copies than required to follow the optimized assay procedure. By adjusting template DNA volume used in the long template PCR reaction, we were able to include all samples and carry out the apparent mtDNA oxidative damage assay. Retrospectively 71  though, the optimization later conducted with the hCAE cells, namely using 30 PCR cycles with 3000mtDNA copies/reaction instead of 26 cycles with 24000mtDNA copies/reaction was preferable. However, due to sample shortage, we could not repeat the assays on these clinical samples with the new conditions. There was no difference found between the comparators and SIM patients, however, there was no clear expectation that a difference should be seen between the groups. These experiments expanded the potential relevance and applicability of the assay to study a broader spectrum within the laboratory.  In the future, a greater variety of tissue samples need to be used to determine how the assay fares in detection of apparent mtDNA oxidative damage with various cell types. 4.3 Cultured Cell Applications The application of the apparent mtDNA oxidative damage assay to two different cell lines treated with NRTIs provided us with insight on the behaviour of NRTIs in different types of cells as well as the apparent mtDNA oxidative damage they may elicit. Firstly, the benefit of assaying cultures treated with NRTIs that had already been tested for other changes was very important in the context of my project. We had initially planned to carry out cell culture experiments with NRTI treatment in another immortalized cell line. However, with the opportunity of having access to these cell culture samples, we realized it would be more informative to experiment with these given the previous work done by others. It also had the distinct advantage of freeing some time that was directed toward other experiments not initially planned. Kyle Hukezalie had shown changes in telomere length 72  following exposure of HT29 cells to NRTIs, indicating that the NRTIs were indeed transported into the appropriate components of the cell and phosphorylated to their active form [171]. The hCAE cell experiments had also demonstrated changes in mtDNA content with some NRTI treatment, also indicating the action of active NRTIs. At least for the cell samples with reduced mtDNA content or shortened telomere, we could be confident that active NRTI were present within the cell. By carrying out these experiments, we were able to learn which cell types firstly respond to NRTI treatment and secondly how they respond to oxidative stress. The mtDNA content was also measured in both cell cultures as this measure has been useful in assessing mitochondrial toxicities associated with NRTI exposure [174]. Both hCAE and HT29 cells showed concentration-dependent depletion with exposure to d4T mirroring observations in other studies [175-177], and to ddI in HT29 cells. In hCAE cells, following exposure to AZT, 3TC, and TDF, concentration-dependent increases in mtDNA content were observed. In HT29 cells, there was no relative movement in mtDNA content with AZT, 3TC, or ABC.  The increase in mtDNA content with exposure to some of the NRTIs could be due to a possible compensatory mechanism of mtDNA proliferation occurring in response either to mild mtDNA depletion induced by NRTI toxicity [142] or to mitochondrial dysfunction through any mechanism. It is well known that d4T is one of the more toxic drugs to the mitochondria [175, 177-179] and therefore, the depletion of mtDNA is too strong to allow for compensatory mtDNA proliferation. The mtDNA depletion following d4T treatment however can be utilized as an indicator that a specific cell type allows for NRTI activation and is responsive to NRTI toxicity, therefore allowing these drugs to act in a way as a sort of control for in vitro studies with NRTI treatment. 73  The apparent mtDNA oxidative damage in NRTI-treated cells however showed very different results between the two cell lines studied. This difference could be related to the cell type with further discussion below on the difference of primary and immortalized cells. In the hCAE cells, one interesting observation was that despite no change in mtDNA content, apparent mtDNA oxidative damage was observed in the cells following TDF exposure. TDF is considered to a less toxic drug and has been recently recommended in the first-line HAART regimens [178, 180, 181]. Furthermore, in HT29 cells, TDF was cytotoxic before day 10, which further alludes to its potential adverse effects on some cells. In addition to mtDNA depletion in the hCAE cells following d4T treatment, greater apparent mtDNA oxidative damage was observed with this NRTI. This result suggests that d4T-related dysfunction brought on by the decrease in mtDNA may increase ROS generation by the ETC, resulting in the increased apparent mtDNA oxidative damage over time. In HT29 cells, although there was no change seen in apparent mtDNA oxidative damage with d4T treatment, the lack of change may be attributed to the cell type as discussed below. Although other studies have shown an increase in oxidative stress in cells following exposure to AZT [137, 182, 183], this observation was not observed in either the hCAE or HT29 cell lines. 3TC has not been historically implicated in any mitochondrial toxicity[184]; in both hCAE cells and HT29 cells, this trend was again observed with apparent mtDNA oxidative damage. 4.3.1 HCAE vs. HT29 cells: Primary vs. Immortalized Cell Lines Assaying DNA from both primary (hCAE) and immortalized (HT29) NRTI-exposed cell cultures allowed for the benefit of examining how these two types of cultures react to drug exposure 74  but also introduced some limitations in assessing oxidative damage. Although it is inherently expected that cell culture models do not entirely reflect what may be happening in vivo, some cell culture types may undergo adaptations that influence their response to various environmental stresses. Adaptations to an environment of oxidative stress can be one such example.  For instance, the HT29 cell line, being established from a cancerous cell type (colorectal adenocarcinoma), may have increased resistance to oxidative stress by means of elevated antioxidant expression and inhibition against pro-oxidation signals, as seen in other cancer cell lines such as breast [185] and epithelial [186] lines . Some cell lines have further even utilized ROS as a means of survival [187] and therefore an immortalized cell line such as HT29 may not reflect oxidative damage by NRTIs at the tissue or organism level. Referring back to the observations made in the HT29 experiment, none of the NRTIs showed any change in apparent mtDNA oxidative damage despite observing mtDNA depletion in d4T and ddI- treated cells. In the hCAE primary cell line, the d4T concentration was lower (insert actual concentrations) and the subsequent apparent mtDNA oxidative damage observed was greater. This provides indication that primary cell lines may be more appropriate in assessing factors such as oxidative stress and apparent mtDNA oxidative damage. 4.4 Future Directions With the assay being newly established with our laboratory’s instruments, ensuring reproducibility over time will be important in establishing a robust assay. 75  Given the differences seen in apparent mtDNA oxidative damage in the two cell lines, it will be important to establish whether other primary and transformed cells respond in a similar fashion or not. For future studies of HAART-related apparent mtDNA oxidative damage, the use of primary cell lines may offer a better model for changes that take place with HAART in humans. The cell cultures studied have given some insight into the patterns of apparent mtDNA oxidative damage associated with NRTI exposure. To further determine how these patterns hold, concentration-dependent studies with NRTIs should be conducted in primary cells to establish if dose dependence factors into apparent mtDNA oxidative damage. Furthermore, as HAART is administered in the form of combination therapy, such patterns need to be assessed in vitro to determine whether drug combinations exacerbate apparent mtDNA oxidative damage. If the patterns of apparent mtDNA oxidative damage are influenced by synergistic or additive mechanisms, it would be crucial to determine which combinations are related to the greatest changes in mtDNA quality. It would be important for mtDNA content analyses to be conducted concurrently in these future experiments to determine whether clonal expansion of mtDNA following periods of stress are involved in potentially altering the detectable levels of apparent mtDNA oxidative damage. Functional assays such as mitochondrial enzyme activity and antioxidant enzyme expression in relation to apparent mtDNA oxidative damage will provide further information on which mechanisms are in play in the cell. However, these assays do require larger quantities of fresh sample, which is not necessary with the mtDNA oxidative damage assay described herein. 76  Also, increased oxidative stress in cells has recently been reported following exposure to PIs [136] so utilizing this assay to measure apparent mtDNA oxidative damage with PI exposure in in vitro studies would be of paramount importance. Furthermore, since PIs are usually administered with two NRTIs in HAART, assessing apparent mtDNA oxidative damage with NRTI and PI concurrent drug exposure would also be of great interest. Since in vitro studies have their limitations in reflecting what is happening in vivo, in the future it will be relevant to analyse apparent mtDNA oxidative damage in clinical samples from patients treated with HAART. Although it can be  difficult to pinpoint whether the source of cellular oxidative stress is from HIV infection or NRTI drug exposure [110], such studies may offer insight if comparing long periods of NRTI exposure compared to shorter periods to assess NRTI toxicity versus HIV infection. 4.5 Conclusions HAART-induced mitochondrial toxicity has disrupted quality of life and limited treatment options for many HIV patients. The goal of this thesis was to determine the relationship between NRTIs and mtDNA quality and apparent mtDNA oxidative damage. This was achieved by firstly the optimization of an assay that would detect and quantify this apparent mtDNA oxidative damage. Two different cell culture models treated with NRTIs in vitro were then subjected to the assay to measure levels of apparent mtDNA oxidative damage. Clinical DNA samples from previously studied muscle biopsies for mtDNA quantity were also tested with the assay to determine the assay’s detection of apparent mtDNA oxidative damage in clinical samples. 77  The optimization and development of this qPCR based assay to measure oxidative damage in mtDNA is crucial in furthering our knowledge on the effects of redox imbalance in disease and drug toxicity. Its versatility in sample types whether the source of DNA is from cells or clinical samples, fresh or frozen, allows large-scale comparison in levels of apparent mtDNA oxidative damage in the context of various diseases. Unlike methods that measure levels of ROS and the overall oxidative stress environment, utilizing an assay that measures direct damage to mtDNA as a result of oxidative stress is of benefit. With this assay optimization, the translation of oxidative stress to oxidative damage is addressed and apparent mtDNA oxidative damage quantified more effectively. Assessing mtDNA quality and quantity from cells treated with NRTIs has provided us with grounds to further explore the relationship between oxidative stress, oxidative damage and NRTI-induced mitochondrial toxicity. The realization that primary cells are a more useful tool for studying oxidative stress will be very important in designing and developing further studies. There is still much research that needs to be conducted in this field. With greater understanding of the various mechanisms that mediate drug toxicity, safer therapy for HIV patients can be identified.     78  References 1. Kohler, J.J. and W. Lewis, A brief overview of mechanisms of mitochondrial toxicity from NRTIs. Environmental and Molecular Mutagenesis, 2007. 48(3-4): p. 166-172. 2. White, A.J., Mitochondrial toxicity and HIV therapy. Sexually Transmitted Infections, 2001. 77:: p. 158-173. 3. Foster, C. and H. Lyall, HIV and mitochondrial toxicity in children. J. Antimicrob. Chemother., 2008. 61(1): p. 8-12. 4. Joint United Nations Programme on HIV/AIDS (UNAIDS), Global Report: UNAIDS Report on the Global AIDS Epidemic 2010. 2010: Geneva. 5. World Health Organization, UNAIDS, UNICEF., Global HIV/AIDS Response: Epidemic update and health sector progress towards Universal Access Progress Report 2011. 2011: Geneva. 6. Joint United Nations Programme on HIV/AIDS (UNAIDS), UNAIDS World Aids Day Report 2011. 2011, Joint United Nations Programme on HIV/AIDS: Geneva. 7. Public Health Agency of Canada, HIV/AIDS Epi Updates 2010, Public Health Agency of Canada: Ottawa. 8. Zheng, Y.-H., N. Lovsin, and B.M. Peterlin, Newly identified host factors modulate HIV replication. Immunology Letters, 2005. 97(2): p. 225-234. 9. Jonathan Allen, C., Virology, immunology, and natural history of HIV infection. Journal of Nurse- Midwifery, 1989. 34(5): p. 242-252. 10. Cooley, L.A. and S.R. Lewin, HIV-1 cell entry and advances in viral entry inhibitor therapy. Journal of Clinical Virology, 2003. 26(2): p. 121-132. 11. Andrew Ml, L., HIV: the virus. Medicine, 2005. 33(6): p. 1-3. 12. Hiscott, J., H. Kwon, and P. Génin, Hostile takeovers: viral appropriation of the NF-kB pathway. The Journal of Clinical Investigation, 2001. 107(2): p. 143-151. 79  13. Marsh, M., K. Theusner, and A. Pelchen-Matthews, HIV assembly and budding in macrophages. Biochemical Society transactions, 2009. 37(Pt 1): p. 185-9. 14. Levy, J.A., HIV and the Pathogenesis of AIDS. Vol. 2nd. edition. 1998, Washington: Library of Congress Cataloging-in-Publication Data. 588. 15. Jaffe, H.W., D.J. Bregman, and R.M. Selik, Acquired Immune Deficiency Syndrome in the United States: The First 1,000 Cases. Journal of Infectious Diseases, 1983. 148(2): p. 339-345. 16. Hollingsworth, T.D., R.M. Anderson, and C. Fraser, HIV-1 Transmission, by Stage of Infection. Journal of Infectious Diseases, 2008. 198(5): p. 687-693. 17. Touloumi, G. and A. Hatzakis, Natural history of HIV-1 infection. Clinics in Dermatology, 2000. 18(4): p. 389-399. 18. Kahn, J.O. and B.D. Walker, Acute Human Immunodeficiency Virus Type 1 Infection. New England Journal of Medicine, 1998. 339(1): p. 33-39. 19. Piatak, M., et al., High levels of HIV-1 in plasma during all stages of infection determined by competitive PCR. Science, 1993. 259(5102): p. 1749-1754. 20. Clark, S.J., et al., High Titers of Cytopathic Virus in Plasma of Patients with Symptomatic Primary HIV-1 Infection. New England Journal of Medicine, 1991. 324(14): p. 954-960. 21. Gendelrnan, H.E., et al., The macrophage in the persistence and pathogenesis of HIV infection. AIDS, 1989. 3(8): p. 475-496. 22. Wu, L. and V.N. KewalRamani, Dendritic-cell interactions with HIV: infection and viral dissemination. Nat Rev Immunol, 2006. 6(11): p. 859-868. 23. Grossman, Z., et al., CD4+ T-cell depletion in HIV infection: Are we closer to understanding the cause? Nat Med, 2002. 8(4): p. 319-323. 24. Rosenberg, E.S., et al., Immune control of HIV-1 after early treatment of acute infection. Nature, 2000. 407(6803): p. 523. 80  25. Lewthwaite, P. and E. Wilkins, Natural history of HIV/AIDS. Medicine, 2005. 33(6): p. 10-13. 26. Lang, W., et al., Patterns of T Lymphocyte Changes with Human Immunodeficiency Virus Infection: From Seroconversion to the Development of AIDS. JAIDS Journal of Acquired Immune Deficiency Syndromes, 1989. 2(1): p. 63-69. 27. Finzi, D., et al., Latent infection of CD4+ T cells provides a mechanism for lifelong persistence of HIV-1, even in patients on effective combination therapy. Nat Med, 1999. 5(5): p. 512-517. 28. Kitahata, M.M., et al., Effect of Early versus Deferred Antiretroviral Therapy for HIV on Survival. New England Journal of Medicine, 2009. 360(18): p. 1815-1826. 29. Cunha, B.A., Antibiotic Essentials. 10th ed. 2011, Sudbury: Jones & Bartlett Learning. 30. S.D, L., AIDS in Africa: the impact of coinfections on the pathogenesis of HIV-1 infection. Journal of Infection, 2004. 48(1): p. 1-12. 31. World Health Organization, WHO policy on collaborative TB/HIV activities: Guidelines for national programmes and other stakeholders. 2012, World Health Organization: Geneva. 32. Sepkowitz, K.A., Effect of HAART on natural history of AIDS-related opportunistic disorders. The Lancet, 1998. 351(9098): p. 228. 33. Murphy, E.L., et al., Highly Active Antiretroviral Therapy Decreases Mortality and Morbidity in Patients with Advanced HIV Disease. Annals of Internal Medicine, 2001. 135(1): p. 17-26. 34. Damond, F., et al., Virological and immunological response to HAART regimen containing integrase inhibitors in HIV-2-infected patients. AIDS, 2008. 22(5): p. 665-666 10.1097/QAD.0b013e3282f51203. 35. Greenberg, M.L. and N. Cammack, Resistance to enfuvirtide, the first HIV fusion inhibitor. Journal of Antimicrobial Chemotherapy, 2004. 54(2): p. 333-340. 81  36. Barreca, M.L., et al., Design, Synthesis, Structure−Ac:vity Rela:onships, and Molecular Modeling Studies of 2,3-Diaryl-1,3-thiazolidin-4-ones as Potent Anti-HIV Agents. Journal of Medicinal Chemistry, 2002. 45(24): p. 5410-5413. 37. Mills, E.J., et al., Adherence to HAART: A Systematic Review of Developed and Developing Nation Patient-Reported Barriers and Facilitators. PLoS Med, 2006. 3(11): p. e438. 38. Monforte, A.d.A., et al., Insights into the reasons for discontinuation of the first highly active antiretroviral therapy (HAART) regimen in a cohort of antiretroviral naive patients. AIDS, 2000. 14(5): p. 499-507. 39. Grant, M., et al., Antiretroviral therapy 2010 update: Current practices and controversies. Archives of pharmacal research, 2011. 34(7): p. 1045-1053. 40. Anderson, P.L., T.N. Kakuda, and K.A. Lichtenstein, The Cellular Pharmacology of Nucleoside- and Nucleotide-Analogue Reverse-Transcriptase Inhibitors and Its Relationship to Clinical Toxicities. Clinical Infectious Diseases, 2004. 38(5): p. 743-753. 41. Stein, D.S. and K.H.P. Moore, Phosphorylation of Nucleoside Analog Antiretrovirals: A Review for Clinicians. Pharmacotherapy, 2001. 21(1): p. 11-34. 42. Basavapathruni, A., C.M. Bailey, and K.S. Anderson, Defining a Molecular Mechanism of Synergy between Nucleoside and Nonnucleoside AIDS Drugs. Journal of Biological Chemistry, 2004. 279(8): p. 6221-6224. 43. Baldwin, S.A., et al., Nucleoside transporters: molecular biology and implications for therapeutic development. Molecular Medicine Today, 1999. 5(5): p. 216-224. 44. Ray, A.S., Intracellular interactions between nucleos(t)ide inhibitors of HIV reverse transcriptase. AIDS reviews, 2005. 7(2): p. 113-25. 82  45. Munch-Petersen, B., et al., Diverging substrate specificity of pure human thymidine kinases 1 and 2 against antiviral dideoxynucleosides. Journal of Biological Chemistry, 1991. 266(14): p. 9032-9038. 46. Gao, W.Y., et al., Differential phosphorylation of azidothymidine, dideoxycytidine, and dideoxyinosine in resting and activated peripheral blood mononuclear cells. The Journal of Clinical Investigation, 1993. 91(5): p. 2326-2333. 47. Godfried, M.H., et al., Soluble Receptors for Tumor Necrosis Factor as Predictors of Progression to AIDS in Asymptomatic Human Immunodeficiency Virus Type 1 Infection. Journal of Infectious Diseases, 1994. 169(4): p. 739-745. 48. Cherry, C.L. and S.L. Wesselingh, Nucleoside analogues and HIV: the combined cost to mitochondria. Journal of Antimicrobial Chemotherapy, 2003. 51(5): p. 1091-1093. 49. Deeks, S.G., et al., HIV-1 Protease Inhibitors: A Review for Clinicians. JAMA, 1997. 277(2): p. 145- 153. 50. Thompson, M.A., et al., Antiretroviral Treatment of Adult HIV Infection. JAMA: The Journal of the American Medical Association, 2010. 304(3): p. 321-333. 51. De Clercq, E., The role of non-nucleoside reverse transcriptase inhibitors (NNRTIs) in the therapy of HIV-1 infection. Antiviral Research, 1998. 38(3): p. 153-179. 52. De Clercq, E., Non-nucleoside reverse transcriptase inhibitors (NNRTIs) for the treatment of human immunodeficiency virus type 1 (HIV-1) infections: Strategies to overcome drug resistance development. Medicinal Research Reviews, 1996. 16(2): p. 125-157. 53. de Béthune, M.-P., Non-nucleoside reverse transcriptase inhibitors (NNRTIs), their discovery, development, and use in the treatment of HIV-1 infection: A review of the last 20 years (1989– 2009). Antiviral Research, 2010. 85(1): p. 75-90. 83  54. De Clercq, E., Non-Nucleoside Reverse Transcriptase Inhibitors (NNRTIs): Past, Present, and Future. Chemistry & Biodiversity, 2004. 1(1): p. 44-64. 55. Baldwin, C.E., R.W. Sanders, and B. Berkhout, Inhibiting HIV-1 Entry with Fusion Inhibitors. Current Medicinal Chemistry, 2003. 10(17): p. 1633-1642. 56. Asmuth, D.M., et al., CD4+ T-Cell Restoration After 48 Weeks in the Maraviroc Treatment- Experienced Trials MOTIVATE 1 and 2. JAIDS Journal of Acquired Immune Deficiency Syndromes, 2010. 54(4): p. 394-397 10.1097/QAI.0b013e3181c5c83b. 57. Duffalo, M.L. and C.W. James, Enfuvirtide: A Novel Agent for the Treatment of HIV-1 Infection. The Annals of Pharmacotherapy, 2003. 37(10): p. 1448-1456. 58. Jing, S., Q. Zhao, and A.K. Debnath, Peptide and Non-peptide HIV Fusion Inhibitors. Current Pharmaceutical Design, 2002. 8(8): p. 563-580. 59. Singh, I.P. and S.K. Chauthe, Small molecule HIV entry inhibitors: Part II. Attachment and fusion inhibitors: 2004 – 2010. Expert Opinion on Therapeutic Patents, 2011. 21(3): p. 399-416. 60. Schneider, S.E., et al., Development of HIV fusion inhibitors. Journal of Peptide Science, 2005. 11(11): p. 744-753. 61. Fatkenheuer, G., et al., Efficacy of short-term monotherapy with maraviroc, a new CCR5 antagonist, in patients infected with HIV-1. Nat Med, 2005. 11(11): p. 1170-1172. 62. Whitby, L.R., et al., Discovery of HIV fusion inhibitors targeting gp41 using a comprehensive α- helix mimetic library. Bioorganic &amp; Medicinal Chemistry Letters, (0). 63. Wang, Y., et al., Structure-based design, synthesis and biological evaluation of new N- carboxyphenylpyrrole derivatives as HIV fusion inhibitors targeting gp41. Bioorganic &amp; Medicinal Chemistry Letters, 2010. 20(1): p. 189-192. 84  64. Viard, J.-P., et al., Immunological success is predicted by enfuvirtide but not interleukin-2 therapy in immunodepressed patients. AIDS, 2009. 23(11): p. 1383-1388 10.1097/QAD.0b013e32832cdc26. 65. Pommier, Y., A.A. Johnson, and C. Marchand, Integrase inhibitors to treat HIV/Aids. Nat Rev Drug Discov, 2005. 4(3): p. 236-248. 66. Steigbigel, R.T., et al., Raltegravir with Optimized Background Therapy for Resistant HIV-1 Infection. New England Journal of Medicine, 2008. 359(4): p. 339-354. 67. World Health Organization, Antiretroviral therapy for HIV infection in adults and adolescents: recommendations for a public health approach. - 2010 rev., in HIV/AIDS Programme. 2010, World Health Organization: Geneva. 68. Yu, D., et al., The discovery of a class of novel HIV-1 maturation inhibitors and their potential in the therapy of HIV. Expert Opinion on Investigational Drugs, 2005. 14(6): p. 681-693. 69. Zhou, J., et al., Inhibition of HIV-1 Maturation via Drug Association with the Viral Gag Protein in Immature HIV-1 Particles. Journal of Biological Chemistry, 2005. 280(51): p. 42149-42155. 70. Neil, S.J.D., et al., An Interferon-α-Induced Tethering Mechanism Inhibits HIV-1 and Ebola Virus Particle Release but Is Counteracted by the HIV-1 Vpu Protein. Cell Host &amp; Microbe, 2007. 2(3): p. 193-203. 71. Adamson, C.S., et al., In Vitro Resistance to the Human Immunodeficiency Virus Type 1 Maturation Inhibitor PA-457 (Bevirimat). Journal of Virology, 2006. 80(22): p. 10957-10971. 72. Arts, E.J. and D.J. Hazuda, HIV-1 Antiretroviral Drug Therapy. Cold Spring Harbor Perspectives in Medicine, 2012. 73. Wainberg, M.A. and J. Albert, Can the further clinical development of bevirimat be justified? AIDS, 2010. 24(5): p. 773-774 10.1097/QAD.0b013e328331c83b. 85  74. Clavel, F. and A.J. Hance, HIV Drug Resistance. New England Journal of Medicine, 2004. 350(10): p. 1023-1035. 75. Pujari, S.N., et al., Lipodystrophy and Dyslipidemia Among Patients Taking First-Line, World Health Organization-Recommended Highly Active Antiretroviral Therapy Regimens in Western India. JAIDS Journal of Acquired Immune Deficiency Syndromes, 2005. 39(2): p. 199-202. 76. Renaud-Théry, F., et al., Use of antiretroviral therapy in resource-limited countries in 2006: distribution and uptake of first- and second-line regimens. AIDS, 2007. 21: p. S89-S95 10.1097/01.aids.0000279711.54922.f0. 77. Erecińska, M. and D.F. Wilson, Regulation of cellular energy metabolism. Journal of Membrane Biology, 1982. 70(1): p. 1-14. 78. Chance, B. and G.R. Williams, The Respiratory Chain and Oxidative Phosphorylation, in Advances in Enzymology and Related Areas of Molecular Biology. 2006, John Wiley & Sons, Inc. p. 65-134. 79. Petit, P.X., et al., Mitochondria and programmed cell death: back to the future. FEBS Letters, 1996. 396(1): p. 7-13. 80. Duchen, M.R., Mitochondria and calcium: from cell signalling to cell death. The Journal of Physiology, 2000. 529(1): p. 57-68. 81. Robin, E.D. and R. Wong, Mitochondrial DNA molecules and virtual number of mitochondria per cell in mammalian cells. Journal of Cellular Physiology, 1988. 136(3): p. 507-513. 82. Terman, A., et al., Mitochondrial turnover and aging of long-lived postmitotic cells: the mitochondrial–lysosomal axis theory of aging. Antioxidants & redox signaling, 2010. 12(4): p. 503-535. 83. DiMauro, S., Mitochondrial DNA Medicine. Bioscience Reports, 2007. 27(1): p. 5-9. 84. Piet, B., Structure and function of mitochondrial DNA. Trends in Biochemical Sciences, 1977. 2(2): p. 31-34. 86  85. Anderson, S., et al., Sequence and organization of the human mitochondrial genome. Nature, 1981. 290(5806): p. 457-465. 86. Stuart, J.A. and M.F. Brown, Mitochondrial DNA maintenance and bioenergetics. Biochimica et Biophysica Acta (BBA) - Bioenergetics, 2006. 1757(2): p. 79-89. 87. Greaves, L.C., et al., Mitochondrial DNA and disease. The Journal of Pathology, 2012. 226(2): p. 274-286. 88. Lightowlers, R.N., et al., Mammalian mitochondrial genetics: heredity, heteroplasmy and disease. Trends in Genetics, 1997. 13(11): p. 450. 89. Falkenberg, M., N.-G. Larsson, and C.M. Gustafsson, DNA Replication and Transcription in Mammalian Mitochondria. Annual Review of Biochemistry, 2007. 76(1): p. 679-699. 90. Clayton, D.A., Replication of animal mitochondrial DNA. Cell, 1982. 28(4): p. 693-705. 91. Lewis, W., W.C. Copeland, and B.J. Day, Mitochondrial DNA Depletion, Oxidative Stress, and Mutation: Mechanisms 0f Dysfunction from Nucleoside Reverse Transcriptase Inhibitors. Lab Invest, 2001. 81(6): p. 777-790. 92. Ledoux, S.P. and G.L. Wilson, Base excision repair of mitochondrial DNA damage in mammalian cells, in Progress in Nucleic Acid Research and Molecular Biology, S.M.A.K.M.R.S.L.S.H.W. Kivie Moldave, Editor. 2001, Academic Press. p. 273-284. 93. Vilhelm A, B., Repair of oxidative DNA damage in nuclear and mitochondrial DNA, and some changes with aging in mammalian cells. Free Radical Biology and Medicine, 2002. 32(9): p. 804- 812. 94. Boesch, P., et al., DNA repair in organelles: Pathways, organization, regulation, relevance in disease and aging. Biochimica et Biophysica Acta (BBA) - Molecular Cell Research, 2011. 1813(1): p. 186-200. 87  95. Bohr, V.A., T. Stevnsner, and N.C. de Souza-Pinto, Mitochondrial DNA repair of oxidative damage in mammalian cells. Gene, 2002. 286(1): p. 127-134. 96. Sykora, P., D.M. Wilson Iii, and V.A. Bohr, Repair of persistent strand breaks in the mitochondrial genome. Mechanisms of Ageing and Development, (0). 97. Arroyo-Torres⁎, Y., et al., Repair of H2O2-induced mtDNA damage requires the BER enzyme Apn1. Mitochondrion, 2011. 11(4): p. 649. 98. de Souza-Pinto, N.C., et al., Novel DNA mismatch-repair activity involving YB-1 in human mitochondria. DNA Repair, 2009. 8(6): p. 704-719. 99. Mason, P.A., et al., Mismatch repair activity in mammalian mitochondria. Nucleic Acids Research, 2003. 31(3): p. 1052-1058. 100. Larsson, N. and D.A. Clayton, Molecular Genetic Aspects of Human Mitochondrial Disorders. Annual Review of Genetics, 1995. 29(1): p. 151-178. 101. Moslemi, A.-R. and N. Darin, Molecular genetic and clinical aspects of mitochondrial disorders in childhood. Mitochondrion, 2007. 7(4): p. 241-252. 102. Zeviani, M. and S. Di Donato, Mitochondrial disorders. Brain, 2004. 127(10): p. 2153-2172. 103. Cohen, B.H. and R.K. Naviaux, The clinical diagnosis of POLG disease and other mitochondrial DNA depletion disorders. Methods, 2010. 51(4): p. 364-373. 104. Barry, H., Reactive oxygen species in living systems: Source, biochemistry, and role in human disease. The American Journal of Medicine, 1991. 91(3, Supplement 3): p. S14-S22. 105. Hashiguchi, K., V.A. Bohr, and N.C. de Souza-Pinto, Oxidative stress and mitochondrial DNA repair: implications for NRTIs induced DNA damage. Mitochondrion, 2004. 4(2-3): p. 215. 106. Kowaltowski, A.J. and A.E. Vercesi, Mitochondrial damage induced by conditions of oxidative stress. Free Radical Biology and Medicine, 1999. 26(3-4): p. 463. 88  107. Cadenas, E. and K.J.A. Davies, Mitochondrial free radical generation, oxidative stress, and aging. Free Radical Biology and Medicine, 2000. 29(3–4): p. 222-230. 108. Croteau, D.L. and V.A. Bohr, Repair of Oxidative Damage to Nuclear and Mitochondrial DNA in Mammalian Cells. Journal of Biological Chemistry, 1997. 272(41): p. 25409-25412. 109. Bruce N, A., Endogenous DNA damage as related to cancer and aging. Mutation Research/Fundamental and Molecular Mechanisms of Mutagenesis, 1989. 214(1): p. 41-46. 110. Lewis, W., B.J. Day, and W.C. Copeland, Mitochondrial toxicity of NRTI antiviral drugs: an integrated cellular perspective. Nat Rev Drug Discov, 2003. 2(10): p. 812. 111. Costa, R., et al., The role of mitochondrial DNA damage in the citotoxicity of reactive oxygen species. Journal of Bioenergetics and Biomembranes, 2011. 43(1): p. 25-29. 112. Graziewicz, M.A., B.J. Day, and W.C. Copeland, The mitochondrial DNA polymerase as a target of oxidative damage. Nucleic Acids Research, 2002. 30(13): p. 2817-2824. 113. Richter, C., J.W. Park, and B.N. Ames, Normal oxidative damage to mitochondrial and nuclear DNA is extensive. Proceedings of the National Academy of Sciences, 1988. 85(17): p. 6465-6467. 114. Bai, J. and A.I. Cederbaum, Mitochondrial Catalase and Oxidative Injury. Neurosignals, 2001. 10(3-4): p. 189-199. 115. Petit, P.X., et al., Mitochondria and programmed cell death: back to the future. FEBS Letters, 1996. 396(1): p. 7-13. 116. Swalwell, H., et al., Respiratory chain complex I deficiency caused by mitochondrial DNA mutations. Eur J Hum Genet, 2011. 19(7): p. 769-775. 117. Ozawa, T., Oxidative Damage and Fragmentation of Mitochondrial DNA in Cellular Apoptosis. Bioscience Reports, 1997. 17(3): p. 237-250. 118. Mao, P. and P.H. Reddy, Aging and amyloid beta-induced oxidative DNA damage and mitochondrial dysfunction in Alzheimer's disease: Implications for early intervention and 89  therapeutics. Biochimica et Biophysica Acta (BBA) - Molecular Basis of Disease, 2011. 1812(11): p. 1359-1370. 119. Mórocz, M., et al., Elevated levels of oxidative DNA damage in lymphocytes from patients with Alzheimer’s disease. Neurobiology of Aging, 2002. 23(1): p. 47-53. 120. Santos, R.X., et al., Nuclear and mitochondrial DNA oxidation in Alzheimer's disease. Free Radical Research, 2012. 46(4): p. 565-576. 121. Suzuki, S., et al., Oxidative damage to mitochondrial DNA and its relationship to diabetic complications. Diabetes Research and Clinical Practice, 1999. 45(2–3): p. 161-168. 122. Corral-Debrinski, M., et al., Association of mitochondrial DNA damage with aging and coronary atherosclerotic heart disease. Mutation Research/DNAging, 1992. 275(3–6): p. 169-180. 123. Polidori, M.C., et al., Oxidative damage to mitochondrial DNA in Huntington's disease parietal cortex. Neuroscience Letters, 1999. 272(1): p. 53-56. 124. Chen, C.-M., et al., Increased oxidative damage and mitochondrial abnormalities in the peripheral blood of Huntington’s disease patients. Biochemical and Biophysical Research Communications, 2007. 359(2): p. 335-340. 125. Giasson, B.I., et al., The relationship between oxidative/nitrative stress and pathological inclusions in Alzheimer’s and Parkinson’s diseases. Free Radical Biology and Medicine, 2002. 32(12): p. 1264-1275. 126. Lu, F., et al., Oxidative damage to mitochondrial DNA and activity of mitochondrial enzymes in chronic active lesions of multiple sclerosis. Journal of the Neurological Sciences, 2000. 177(2): p. 95-103. 127. Monforte, A.d.A., et al., Clinical outcome and predictive factors of failure of highly active antiretroviral therapy in antiretroviral-experienced patients in advanced stages of HIV-1 infection. AIDS, 1998. 12(13): p. 1631-1637. 90  128. Blas-Garcia, A., N. Apostolova, and J. V Esplugues, Oxidative Stress and Mitochondrial Impairment After Treatment with Anti-HIV Drugs: Clinical Implications. Current Pharmaceutical Design, 2011. 17(36): p. 4076-4086. 129. Esser, S., et al., Side effects of HIV therapy. JDDG: Journal der Deutschen Dermatologischen Gesellschaft, 2007. 5(9): p. 745-754. 130. Cui, L., et al., Effect of Nucleoside Analogs on Neurite Regeneration and Mitochondrial DNA Synthesis in PC-12 Cells. Journal of Pharmacology and Experimental Therapeutics, 1997. 280(3): p. 1228-1234. 131. Kakuda, T.N., Pharmacology of nucleoside and nucleotide reverse transcriptase inhibitor-induced mitochondrial toxicity. Clinical Therapeutics, 2000. 22(6): p. 685-708. 132. Wright, G.E. and N.C. Brown, Deoxyribonucleotide analogs as inhibitors and substrates of DNA polymerases. Pharmacology &amp; Therapeutics, 1990. 47(3): p. 447-497. 133. Pinti, M., P. Salomoni, and A. Cossarizza, Anti-HIV drugs and the mitochondria. Biochimica et Biophysica Acta (BBA) - Bioenergetics, 2006. 1757(5-6): p. 700. 134. Eriksson, S., B. Xu, and D.A. Clayton, Efficient Incorporation of Anti-HIV Deoxynucleotides by Recombinant Yeast Mitochondrial DNA Polymerase. Journal of Biological Chemistry, 1995. 270(32): p. 18929-18934. 135. Day, B. and W. Lewis, Oxidative stress in NRTI-induced toxicity. Cardiovascular Toxicology, 2004. 4(3): p. 207-216. 136. Lagathu, C., et al., Some HIV antiretrovirals increase oxidative stress and alter chemokine, cytokine or adiponectin production in human adipocytes and macrophages. Antiviral therapy, 2007. 12(4): p. 489-500. 91  137. de la Asunción, J.G., et al., AZT treatment induces molecular and ultrastructural oxidative damage to muscle mitochondria. Prevention by antioxidant vitamins. The Journal of Clinical Investigation, 1998. 102(1): p. 4-9. 138. Prakash, O., et al., The Human Immunodeficiency Virus Type 1 Tat Protein Potentiates Zidovudine-Induced Cellular Toxicity In Transgenic Mice. Archives of Biochemistry and Biophysics, 1997. 343(2): p. 173-180. 139. Velsor, L.W., et al., Mitochondrial oxidative stress in human hepatoma cells exposed to stavudine. Toxicology and Applied Pharmacology, 2004. 199(1): p. 10-19. 140. Turchan, J., et al., Oxidative stress in HIV demented patients and protection ex vivo with novel antioxidants. Neurology, 2003. 60(2): p. 307-314. 141. Romano, A.D., et al., Oxidative stress and aging. Journal of nephrology, 2010. 23: p. S29. 142. Payne, B.A.I., et al., Mitochondrial aging is accelerated by anti-retroviral therapy through the clonal expansion of mtDNA mutations. Nat Genet, 2011. 43(8): p. 806-810. 143. Larsson, N.-G., Somatic Mitochondrial DNA Mutations in Mammalian Aging. Annual Review of Biochemistry, 2010. 79(1): p. 683-706. 144. Tuppen, H.A.L., et al., Mitochondrial DNA mutations and human disease. Biochimica et Biophysica Acta (BBA) - Bioenergetics, 2010. 1797(2): p. 113-128. 145. Halliwell, B. and M. Whiteman, Measuring reactive species and oxidative damage in vivo and in cell culture: how should you do it and what do the results mean? British Journal of Pharmacology, 2004. 142(2): p. 231-255. 146. Ravanat, J., J. Cadet, and T. Douki, Oxidatively Generated DNA Lesions as Potential Biomarkers of In Vivo Oxidative Stress. Current molecular medicine, 2012. 147. Dizdaroglu, M., et al., Free radical-induced damage to DNA: mechanisms and measurement. Free Radical Biology and Medicine, 2002. 32(11): p. 1102-1115. 92  148. Kovalenko, O.A. and J.H. Santos, Analysis of Oxidative Damage by Gene-Specific Quantitative PCR, in Current Protocols in Human Genetics. 2001, John Wiley & Sons, Inc. 149. Helbock, H.J., et al., DNA oxidation matters: The HPLC–electrochemical detection assay of 8-oxo- deoxyguanosine and 8-oxo-guanine. Proceedings of the National Academy of Sciences, 1998. 95(1): p. 288-293. 150. Halliwell, B. and M. Dizdaroglu, Commentary the Measurement of Oxidative Damage to DNA by HPLC and GC/MS Techniques. Free Radical Research, 1992. 16(2): p. 75-87. 151. Barja, G. and A. Herrero, Oxidative damage to mitochondrial DNA is inversely related to maximum life span in the heart and brain of mammals. The FASEB Journal, 2000. 14(2): p. 312- 318. 152. Lim, K.S., R.K. Naviaux, and R.H. Haas, Quantitative Mitochondrial DNA Mutation Analysis by Denaturing HPLC. Clinical Chemistry, 2007. 53(6): p. 1046-1052. 153. Takahashi, M., et al., Effect of oxidative stress on development and DNA damage in in-vitro cultured bovine embryos by comet assay. Theriogenology, 2000. 54(1): p. 137-145. 154. Ghazalla, M.B., et al., Genotoxic effect induced by hydrogen peroxide in human hepatoma cells using comet assay. 2010. 2010. 155. Collins, A.R., et al., The comet assay: topical issues. Mutagenesis, 2008. 23(3): p. 143-151. 156. Gedik, C.M., S.G. Wood, and A.R. Collins, Measuring oxidative damage to DNA; HPLC and the comet assay compared. Free Radical Research, 1998. 29(6): p. 609-615. 157. Choucroun, P., et al., Comet assay and early apoptosis. Mutation Research/Fundamental and Molecular Mechanisms of Mutagenesis, 2001. 478(1–2): p. 89-96. 158. Wang, H. and J.A. Joseph, Quantifying cellular oxidative stress by dichlorofluorescein assay using microplate reader. Free Radical Biology and Medicine, 1999. 27(5–6): p. 612-616. 93  159. Kovalenko, O.A. and J.H. Santos, Analysis of oxidative damage by gene-specific quantitative PCR. Curr. Protoc. Hum. Genet, 2009. 19(1). 160. Ballinger, S.W., et al., Hydrogen Peroxide- and Peroxynitrite-Induced Mitochondrial DNA Damage and Dysfunction in Vascular Endothelial and Smooth Muscle Cells. Circ Res, 2000. 86(9): p. 960- 966. 161. Ballinger, S.W., et al., Hydrogen Peroxide Causes Significant Mitochondrial DNA Damage in Human RPE Cells. Experimental Eye Research, 1999. 68(6): p. 765. 162. Wijeratne, S.S.K., S.L. Cuppett, and V. Schlegel, Hydrogen Peroxide Induced Oxidative Stress Damage and Antioxidant Enzyme Response in Caco-2 Human Colon Cells. Journal of Agricultural and Food Chemistry, 2005. 53(22): p. 8768. 163. Kleiman, N.J., R.-R. Wang, and A. Spector, Hydrogen peroxide-induced DNA damage in bovine lens epithelial cells. Mutation Research/Genetic Toxicology, 1990. 240(1): p. 35. 164. Freshney, R.I., Culture of Animal Cells: A Manual of Basic Techniques. 5th ed. 2005, New Jersey: Wiley-Liss. 165. Holden, M.J., et al., Factors Affecting Quantification of Total DNA by UV Spectroscopy and PicoGreen Fluorescence. Journal of Agricultural and Food Chemistry, 2009. 57(16): p. 7221-7226. 166. Côté, H.C.F., et al., Changes in Mitochondrial DNA as a Marker of Nucleoside Toxicity in HIV- Infected Patients. New England Journal of Medicine, 2002. 346(11): p. 811-820. 167. Côté, H.C.F., et al., Quality assessment of human mitochondrial DNA quantification: MITONAUTS, an international multicentre survey. Mitochondrion, 2011. 11(3): p. 520-527. 168. Wong, A. and G. Cortopassi, Reproducible Quantitative PCR of Mitochondrial and Nuclear DNA Copy Number Using the LightCycler™ Mitochondrial DNA, W.C. Copeland, Editor. 2002, Humana Press. p. 129-138. 94  169. Santos, J.H., et al., Quantitative PCR-Based Measurement of Nuclear and Mitochondrial DNA Damage and Repair in Mammalian Cells DNA Repair Protocols, D.S. Henderson, Editor. 2006, Humana Press. p. 183-199. 170. Stringer, H., Mitochondrial DNA Alterations and Statin-Induced Myopathy, in Pathology and Laboratory Medicine. 2009, University of British Columbia: Vancouver. p. 114. 171. Hukezalie, K.R., Characterizing the effects of N/NRTIs on human telomerase activity in vitro and telomere maintenance in a transformed human cell model, in Genetics. 2011, University of British Columbia: Vancouver. p. 148. 172. Lefèvre, C., et al., Premature Senescence of Vascular Cells Is Induced by HIV Protease Inhibitors. Arteriosclerosis, Thrombosis, and Vascular Biology, 2010. 30(12): p. 2611-2620. 173. Sbisà, E., et al., Mammalian mitochondrial D-loop region structural analysis: identification of new conserved sequences and their functional and evolutionary implications. Gene, 1997. 205(1– 2): p. 125-140. 174. Cote, H.C.F., et al., Changes in Mitochondrial DNA as a Marker of Nucleoside Toxicity in HIV- Infected Patients. N Engl J Med, 2002. 346(11): p. 811-820. 175. Gaou, I., et al., Effect of Stavudine on Mitochondrial Genome and Fatty Acid Oxidation in Lean and Obese Mice. Journal of Pharmacology and Experimental Therapeutics, 2001. 297(2): p. 516- 523. 176. McComsey, G.A., et al., Improvements in lipoatrophy, mitochondrial DNA levels and fat apoptosis after replacing stavudine with abacavir or zidovudine. AIDS, 2005. 19(1): p. 15-23. 177. Nolan, D., et al., Mitochondrial DNA depletion and morphologic changes in adipocytes associated with nucleoside reverse transcriptase inhibitor therapy. AIDS, 2003. 17(9): p. 1329-1338. 178. Rosso, R., et al., Effects of the Change From Stavudine to Tenofovir in Human Immunodeficiency Virus-Infected Children Treated With Highly Active Antiretroviral Therapy: Studies on 95  Mitochondrial Toxicity and Thymic Function. The Pediatric Infectious Disease Journal, 2008. 27(1): p. 17-21 10.1097/INF.0b013e31814689be. 179. Sutinen, J., Interventions for managing antiretroviral therapy-associated lipoatrophy. Current Opinion in Infectious Diseases, 2005. 18(1): p. 25-33. 180. Schooley, R.T., et al., Tenofovir DF in antiretroviral-experienced patients: results from a 48-week, randomized, double-blind study. AIDS, 2002. 16(9): p. 1257-1263. 181. Lok, A.S.F., Drug Therapy: Tenofovir. Hepatology, 2010. 52(2): p. 743-747. 182. García de la Asunción, J., et al., AZT induces oxidative damage to cardiac mitochondria: Protective effect of vitamins C and E. Life Sciences, 2004. 76(1): p. 47. 183. Modica-Napolitano, J.S., AZT Causes Tissue-Specific Inhibition of Mitochondrial Bioenergetic Function. Biochemical and Biophysical Research Communications, 1993. 194(1): p. 170. 184. Anderson, K.S., Perspectives on the molecular mechanism of inhibition and toxicity of nucleoside analogs that target HIV-1 reverse transcriptase. Biochimica et Biophysica Acta (BBA) - Molecular Basis of Disease, 2002. 1587(2–3): p. 296-299. 185. Bae, I., et al., BRCA1 Induces Antioxidant Gene Expression and Resistance to Oxidative Stress. Cancer Research, 2004. 64(21): p. 7893-7909. 186. Viard, I., et al., Clusterin Gene Expression Mediates Resistance to Apoptotic Cell Death Induced by Heat Shock and Oxidative Stress. 1999. 112(3): p. 290-296. 187. Pervaiz, S., et al., Superoxide anion inhibits drug-induced tumor cell death. FEBS Letters, 1999. 459(3): p. 343-348.   

Cite

Citation Scheme:

        

Citations by CSL (citeproc-js)

Usage Statistics

Share

Embed

Customize your widget with the following options, then copy and paste the code below into the HTML of your page to embed this item in your website.
                        
                            <div id="ubcOpenCollectionsWidgetDisplay">
                            <script id="ubcOpenCollectionsWidget"
                            src="{[{embed.src}]}"
                            data-item="{[{embed.item}]}"
                            data-collection="{[{embed.collection}]}"
                            data-metadata="{[{embed.showMetadata}]}"
                            data-width="{[{embed.width}]}"
                            async >
                            </script>
                            </div>
                        
                    
IIIF logo Our image viewer uses the IIIF 2.0 standard. To load this item in other compatible viewers, use this url:
https://iiif.library.ubc.ca/presentation/dsp.24.1-0072779/manifest

Comment

Related Items