Live Cell Fluorescence Imaging of Nucleotide Dynamics ...

Live Cell Fluorescence Imaging of Nucleotide

Dynamics: ATP Hydrolysis and DNA Damage

Response

Doctoral thesis for obtaining the

academic degree Doctor of Natural Sciences

(Dr.rer.nat.)

submitted by

Bhat, Anayat

at the

Faculty of Sciences

Department of Chemistry

Konstanz, 2021

Konstanzer Online-Publikations-System (KOPS) URL: http://nbn-resolving.de/urn:nbn:de:bsz:352-2-151sgk7n0qvtw7

Date of the oral examination: 26th February 2021

1. Referee: Prof. Dr. Andreas Zumbusch

2. Referee: Prof. Dr. Elisa May

Acknowledgements

I would like to take this opportunity for expressing my sincere gratitudeto my supervisor Prof. Dr. Andreas Zumbusch for providing me the oppor-tunity to work in his research group. I wish to thank you for your consistentsupport, excellent scientific advice, and the regular critical discussions. I deeplyappreciate the trust placed in me and the freedom that I relished while pursuingmy ideas during my Ph.D. I am heartily thankful for keeping your door alwaysopen and being so kind as to provide the personal support in turbulent times.

I would like to pay my special regards to Prof. Dr. Elisa May for super-vising my thesis committee. Also, I am grateful for her invaluable suggestions,discussions, and the assistance we got for our experiments from the bioimagingcenter.

I would like to extend my heartfelt acknowledgments to Prof. Dr. An-dreas Marx for agreeing to chair my examination committee. I am also indebtedfor his expert opinions and the fruitful collaborations with his group.

I am wholeheartedly thankful to Dr. Martin Winterhalder for beingavailable all the time to help with operating the microscopes. I thank FranziskaRabold, Martina Adam, and Brunhilde Kottwitz for their help with the cell cul-ture and other laboratory work. I am thankful to Tobias Loffler for being helpfuland kind while working together over the last few years. It has been a pleasant,successful, and memorable cooperation. I am thankful to Dr. Franziska Dollfor all her support during the starting of my work in the group. I am thank-ful to Daniel Hammler for his cooperation and the evaluative discussions thathave been helpful for the development of our project. I am thankful to all thestudents including Adrian Huck, Svenja Hehn, and Natalie Pach who had been

ii Acknowledgements

the part of this work during their bachelor’s theses and internships.

I especially appreciate the assistance of Dr. Peyman Zirak with theMatlab and for developing the LabView scripts used for imaging. I would liketo recognize the help of Imaiyan Ragupathy and Marcel Rudolf for their kindassistance during the compilation of this thesis.

I wish to thank all the present and the former members of the workinggroup of Prof. Zumbusch for providing conducive working environment in thelab. Their support and companionship has not only provided me with constantmotivation but has also bestowed me with cherished memories.

I would like to thank my Graduate School, Konstanz Research Schoolof Chemical Biology for financial support and for a wonderful environment fordoing science throughout my Ph.D.

I sincerely acknowledge all my friends in Konstanz for the unforgettabletimes that we have spent together. I thank them for their persistent motiva-tion and encouragement that made the life easy while my stay in Konstanz. Iam thankful to my friends who have been far from me but their unwaveringpersonal and professional support has been a source of comfort throughout thisdissertation.

Lastly, I express my profound gratitude to my family for unfailing sup-port. My family has been a source of encouragement throughout my studies. Iam forever indebted to my parents for their unconditional love and sympathywhile staying away from them. I am thankful to my siblings for consistentlyserving as morale boosters through all these years. This accomplishment wouldnot have been possible without the support of my family.

Zusammenfassung

Nukleotide sind eine Klasse von Biomolekülen, welche nicht nur als Zellbe-standteil dienen, sondern auch entscheidend für die Zellfunktion und Homöostasesind. Ein Nukleotid besteht aus einer stickstoffhaltigen Base, einem Zucker undeiner Phosphat Gruppe. In erster Linie dienen Nukleotide in allen Organismenals Grundgerüst für das Genmaterial, wie zum Beispiel DNA oder RNA. Jedochspielen Nukleotide auch eine entscheidende Rolle in regulatorischen Prozessen,einschließlich der Zellkommunikation, des Stoffwechsels und der Energieübertra-gung. Adenosintriphosphat (ATP) ist ein Mononukleotid, das als Zwischenen-ergiequelle fungiert und als der universelle Energieträger der Zelle bekannt ist.Nicotinamidadenindinukleotid (NAD+) ist ein zentrales Dinukleotid des Stof-fwechsels und ist in mehreren Redoxreaktionen am Elektronentransfer beteiligt.ADP und NAD+ sind ebenfalls Teil der posttranslationalen Proteinmodifika-tionen (PTM). PTMs sind verantwortlich für Änderungen der Proteineigen-schaften, um deren biologische Aktivität zu regulieren, indem Wechselwirkun-gen mit anderen Faktoren moduliert werden. NAD+ dient auch als Baustein(Edukt) für das Protein Poly(ADP-ribose)polymerase zur PARylierung, welcheseine posttranslationale Modifikation ist, die hauptsächlich als Antwort auf einenDNA Schaden vorkommt.

Diese Arbeit umfasst die Untersuchung zweier häufig vorkommender Nuk-leotide und deren Dynamik in lebenden Zellen mit Hilfe von Fluoreszenz Mikros-kopie. Fluoreszenz Mikroskopie ist zu einer nicht mehr wegzudenkenden Meth-ode zur Visualisierung intermolekularer Wechselwirkungen geworden, vor allemdurch neueste Entwicklungen in der Sensitivität, der Schnelligkeit und der Au-flösung. Die Fortschritte bei der Herstellung von Biosensoren und FluoreszenzProben hat zu einer Revolution der Aufnahmen von dynamischen Prozessen auf

iv Zusammenfassung

zellulärem und molekularem Niveau geführt. Basierend auf Fluoresenzlebens-dauer Mikroskopie (fluorescence lifetime imaging microscopy, FLIM), verwen-den wir den Förster Resonanz Energietransfer (FRET), um die Aktivität undKinetik der zwei Nukleotidanaloga des ATP und NAD+ zu beobachten. Neuar-tige fluoreszierende Nukleotide wurden hierfür in der Arbeitsgruppe von Prof.Andreas Marx hergestellt.

Um den Mechanismus der biologischen Funktionen zu verstehen, beiwelchen ATP eine Rolle spielt, ist es wichtig die ATP Hydrolyse Aktivitätzu beobachten. Die Aktivität der ATP Hydrolyse wurde mit Hilfe von Flu-oreszenzmikroskopie in Verbindung mit fluoreszenten ATP Analoga sichtbargemacht. Das ATP Analog Adenosin Tetraphosphat (Ap4), wurde mit dem Rho-damin Derivat Atto-488 an der terminalen Phosphatgruppe und an einer nichtfluoreszenten Quenchergruppe an der Adenosin Base modifiziert. Der Farb-stoff verhält sich wie ein Donor und der nicht fluoreszenten Quencher dient alsAkzeptor. Im intakten Molekül wird die Fluoreszenz aufgrund des FRETs we-gen der räumlichen Nähe zwischen dem Donor und dem Akzeptor gelöscht. DieEnzymspaltung führt sowohl zu einer Erhöhung der Fluoreszenzintensität, alsauch zu einer größeren Fluoreszenzhalbwertszeit des Farbstoffes. Dieses Prinzipwurde zum einen dazu verwendet die Hydrolyse der Analoga in vitro zu quan-tifizieren, zum anderen auch zur Quantifizierung in lebenden Zellen mit Hilfevon FLIM und konfokaler Mikroskopie. Unsere Experimente deckten auf, dassdie Hydrolyse des Ap4 Analogs zum größten Teil in Lysosomen stattfindet. DaLysosome eine äußerst wichtige Rolle im Prozess der zellulären Autophagie spie-len, werteten wir die Verwendung von Ap4 während der Autophagie mit fluo-reszenten Markern aus. Wir konnten beobachten, dass das Analog hydrolisiertwird und als eine potentielle Energiequelle in Lysosomen und Autolysosomenwährend der Autophagie dient. Wir konnten erfolgreich die Hydrolyse einesNukleotids in einer lebenden Zelle beobachten. Die Experimente ermöglichenes das Analoga zum Beobachten der Autophagie in lebenden Zellen zu verwen-den. Weitere Experimente zur Beobachtung und zur Ermittlung quantitativerDetails des autophagischen Flusses könnten durchgeführt werden.

v

Die Antwort auf einen DNA Schaden und dessen Reparatur ist ein grundle-gender biologischer Prozess zur Erhaltung der genetischen Intaktheit eines Or-ganismus. Die Protein PARylierung ist der elementare Schritt der DNA Reparatur.Die PARylierung führt zur Regulierung vonWechselwirkungen zwischen mehrerenFaktoren während der Initiierung, um den Prozess der DNA Reparatur zu bew-erkstelligen. NAD+ fungiert als Donor für die PARylierung der Proteine, welchebei der Schadensreparatur involviert sind. Ein neuartiges fluoreszentes NAD+

Analog, das TMR-NAD wurde mit einem Tetramethylrhodamin Farbstoff syn-thetisiert. Das Analog wurde in Verbindung mit dem EGFP Fusionsprotein ver-wendet, um die PARylierung des ARTD1 zu beobachten, welches hauptsächlichan der PARylierung beteiligt ist. Mit FLIM FRET konnte die Echtzeit Beobach-tung der PARylierung in lebenden Zellen erreicht werden. Die Bildgebungberuhte auf FRET zwischen dem TMR-NAD Analog und dem EGFP Fusion-sprotein. Der kleine Abstand zwischen den beiden Farbstoffen führt zu FRETund zur Änderung der Fluoreszenzhalbwertszeit des Donors, wie zum Beispielim Falle der PARylierung des Proteins und der nicht kovalenten Wechselwirkungzwischen einem Protein und den PAR Ketten. Die Abnahme der Fluoreszen-zlebenszeit des Donorfluoreszenzfarbstoffs wurde gemessen, um die PARylierungund die damit verbundenenWechselwirkungen zu veranschaulichen und zu quan-tifizieren. Die Kinetik der Rekrutierung und PARylierung von zwei Proteinen,wie zum Beispiel ARTD1 und mH2A, wurden bestimmt, um die komplexenProzesse der DNA Reparatur zu verstehen. Um einen Schaden mit gleichbleiben-den Bedingungen zu erzeugen, wurden ultra-kurze nah infrarote Laser Pulsezur Mikrobestrahlung des Zellkerns in einem wohl definierten Bereich verwen-det. Diese Studie wurde mit Hilfe der Arbeitsgruppe von Prof. Dr. Elisa Maydurchgeführt. Mit diesem Ansatz ist es vorstellbar, mit unglaublicher räumlicherund zeitlicher Auflösung die Dynamik molekularer Ereignisse zu beobachten,welche bei der DNA Reparatur auftreten.

Summary

Nucleotides are biomolecules that not only act as a cellular constituents but arealso crucial for cellular functioning and homeostasis. A nucleotide consists ofa nitrogenous base, a sugar, and phosphate groups. Nucleotides primarily actas the basic structural units of the genetic material, i.e. DNA and RNA in allorganisms. In addition, some nucleotides play a critical role in regulatory pro-cesses including cellular signaling, metabolism, and energy transfer. Adenosinetriphosphate (ATP) is a mono nucleotide that acts as intermediate energy sourceand is known as the universal energy carrier of the cell. Nicotinamide adeninedinucleotide (NAD+) is a dinucleotide central to metabolism and is involved inelectron transfer in various redox reactions. ATP and NAD+ are, however, alsoinvolved in posttranslational protein modifications (PTM). PTMs are responsi-ble for the alterations of the properties of a protein to regulate their biologicalactivity by modulation of their interactions with other factors. NAD+ also actsas a substrate for the protein poly (ADP-ribose)ylation (PARylation), whichis a posttranslational modification that occurs for example in response to theDNA damage.

The present work involves the study of two prominent nucleotides andtheir dynamics in living cells using fluorescence microscopy. Fluorescence mi-croscopy has become an essential tool for visualizing molecular interactionsbecause of recent advancements in its sensitivity, speed, and resolution. Theadvancement in the synthesis of biosensors and fluorescent probes has led toa revolution in imaging the dynamics at the cellular and molecular level. Wehave used the Förster resonance energy transfer (FRET) microscopy based onfluorescence lifetime imaging (FLIM) to monitor the activity and kinetics of two

vii

nucleotide analogs of ATP and NAD+. The novel fluorescent nucleotide analogsof ATP and NAD+ used in this work were synthesized in the working group ofProf. Dr. Andreas Marx.

To understand the mechanism of the biological functions involving ATP,it is important to monitor ATP hydrolysis activity. The activity of ATP hy-drolysis was visualized by using fluorescence microscopy in combination withfluorescent ATP analogs. The ATP analog adenosine tetraphosphate (Ap4) wasmodified with the rhodamine derivative Atto-488 at the terminal phosphate anda non-fluorescent quencher group at the adenosine base. The dye would act asa donor and the quencher as the acceptor. In the intact molecule, the fluo-rescence of the dye is quenched due to the Förster resonance energy transfer(FRET) because of the spatial proximity between the donor and acceptor. En-zymatic cleavage results in an increase in the fluorescence intensity as well asan increase in the fluorescence lifetime of the dye. This principle was used toquantify the hydrolysis of analogs in vitro as well as in living cells by using fluo-rescence lifetime imaging (FLIM) microscopy and confocal scanning microscopy.Our experiments revealed that the hydrolysis of Ap4 analogs largely takes placein lysosomes. Since lysosomes play a prominent role in the process of cellularautophagy, we evaluated the utilization of Ap4 in autophagy by using fluores-cent markers. It was observed that the analog is hydrolyzed and is used as apotential energy source in lysosomes and autolysosomes during the process ofautophagy. We thus have been able to successfully monitor the hydrolysis of anucleotide in living cells. The experiments invoke the possibility of using theanalog for monitoring the process of autophagy in living cells. Further experi-ments can be performed to visualize and get quantitative details of the processof autophagic flux.

The DNA damage response and repair is a fundamental biological pro-cess crucial for maintaining the genomic integrity in an organism. ProteinPoly(ADP-ribosyl)ation (PARylation) is a primary step in DNA damage re-sponse. PARylation results in the regulation of interactions among multiplefactors during initiation to accomplish the process of DNA repair. NAD+ is

viii Summary

used as a donor for the PARylation of proteins involved in the damage re-sponse. A novel fluorescent analog of NAD+, TMR-NAD, was synthesized withtetramethylrhodamine dye attached to it. The analog was used in combinationwith EGFP fusion proteins to monitor the PARylation of ARTD1 which is pri-marily involved in PARylation. With the application of FLIM-FRET imaging,the real-time visualization of PARylation in living cells using this analog wasachieved. The imaging was based on FRET between TMR-NAD analog and theEGFP fusion protein. The close proximity between the two fluorophores resultsin FRET and the change in the fluorescence lifetime of donor e.g. in case of thePARylation of the protein and the non-covalent interaction between a proteinand the PAR chains. The decrease in the fluorescence lifetime of the donorfluorophore is measured to visualize and quantify protein PARylation and theassociated interactions. The kinetics of the recruitment and PARylation of twoproteins i.e. ARTD1 and mH2A were determined to understand the complexprocess of DNA damage response. To induce the damage with the standardizedconditions, ultra-short laser pulses from the near-infrared (NIR) laser were usedfor the micro irradiation of the nucleus in a well-defined region. This study wasperformed with help from the working group of Prof. Dr. Elisa May. Usingthis analog approach, it is conceivable to monitor the dynamics of molecularevents involved the DNA damage response with incredible spatial and temporalresolution.

Contents

Acknowledgements i

Zusammenfassung iii

Summary vi

1 Introduction 1

2 Molecular Imaging in Living Cells 5

2.1 Microscopy in Biological Research . . . . . . . . . . . . . . . . . 5

2.1.1 Principles of Fluorescence Microscopy . . . . . . . . . . . 8

2.1.2 Types of Microscope Setups . . . . . . . . . . . . . . . . . 10

2.1.2.1 Widefield Microscopy . . . . . . . . . . . . . . . 10

2.1.2.2 Confocal Microscopy . . . . . . . . . . . . . . . . 11

2.2 Imaging Molecular Interactions and Dynamics . . . . . . . . . . . 13

2.2.1 Förster Resonance Energy Transfer . . . . . . . . . . . . . 14

2.2.1.1 Fluorescence Intensity based FRET . . . . . . . 16

2.2.1.2 Fluorescence Anisotropy based FRET . . . . . . 17

2.2.1.3 Fluorescence Lifetime based FRET . . . . . . . 18

2.2.2 Fluorescence Lifetime Imaging Microscopy (FLIM) . . . . 18

2.2.2.1 Time-Domain FLIM . . . . . . . . . . . . . . . . 19

2.2.2.2 Frequency-Domain FLIM . . . . . . . . . . . . . 20

x CONTENTS

2.2.3 Applications of FLIM FRET Imaging . . . . . . . . . . . 23

3 Kinetics of ATP Hydrolysis 243.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 243.2 Cellular ATP Hydrolyzing Enzymes . . . . . . . . . . . . . . . . 273.3 Methods for Studying ATP hydrolysis . . . . . . . . . . . . . . . 30

3.3.1 Assays Based on Inorganic Phosphate Detection . . . . . 303.3.2 Radioactivity Based Assays . . . . . . . . . . . . . . . . . 313.3.3 Enzyme Coupled Assyas . . . . . . . . . . . . . . . . . . . 313.3.4 Protein Based ATP Sensors . . . . . . . . . . . . . . . . . 323.3.5 Assays Based on Fluorescent ATP Analogs . . . . . . . . 33

3.4 Modified ATP Analogs . . . . . . . . . . . . . . . . . . . . . . . . 33

4 Protein PARylation in DNA Damage Response 374.1 Protein Modifications and Interactions . . . . . . . . . . . . . . . 37

4.1.1 Protein Poly-ADP-ribosylation . . . . . . . . . . . . . . . 394.2 DNA Damage and Repair . . . . . . . . . . . . . . . . . . . . . . 40

4.2.1 Types and Causes of DNA Damage . . . . . . . . . . . . . 414.2.2 Mechanisms of DNA Repair . . . . . . . . . . . . . . . . . 43

4.3 PARylation in DNA Damage Response . . . . . . . . . . . . . . . 454.3.1 PARP1: Structure and Function . . . . . . . . . . . . . . 454.3.2 Role of PARP1 in DNA Repair . . . . . . . . . . . . . . . 47

4.4 Methods for Studying DNA Damage Response . . . . . . . . . . 484.4.1 PCR Based Methods . . . . . . . . . . . . . . . . . . . . . 484.4.2 Molecular Marker Based Method . . . . . . . . . . . . . . 484.4.3 Fluorescence Based Methods . . . . . . . . . . . . . . . . 49

4.5 Modified NAD Analogs . . . . . . . . . . . . . . . . . . . . . . . 504.6 Fluorescence Microscopy for Studying Damage Response . . . . . 51

4.6.1 DNA Damage by Laser Microirradiation . . . . . . . . . . 524.6.2 Imaging of DNA Damage Response . . . . . . . . . . . . . 54

5 ATP analog hydrolysis in living cells 575.1 Stability and Enzyme Specificity of Ap4 . . . . . . . . . . . . . . 595.2 Ap4 hydrolysis In Vitro . . . . . . . . . . . . . . . . . . . . . . . 605.3 Incorporation of the Ap4 analog in living cells . . . . . . . . . . 62

CONTENTS xi

5.4 Imaging of Ap4 Hydrolysis in Living Cells . . . . . . . . . . . . . 65

5.4.1 Wide Field FLIM Imaging of the Ap4 Hydrolysis . . . . 65

5.4.2 Confocal FLIM microscopy of the Ap4 hydrolysis . . . . 66

5.5 Ap4 hydrolysis in lysosomes. . . . . . . . . . . . . . . . . . . . . 68

5.6 Utilization of Ap4 Analog in Autophagy . . . . . . . . . . . . . . 78

5.7 Summary and Outlook . . . . . . . . . . . . . . . . . . . . . . . . 81

6 Protein PARylation in DNA Damage Response 83

6.1 Incorporation of TMR-NAD Analog into the Living Cells . . . . 85

6.2 Laser induced DNA damage . . . . . . . . . . . . . . . . . . . . . 89

6.3 Nature of the DNA damage Induced . . . . . . . . . . . . . . . . 91

6.4 Kinetics of ARTD1 PARylation . . . . . . . . . . . . . . . . . . 93

6.5 Kinetics of Macro H2A . . . . . . . . . . . . . . . . . . . . . . . . 100

6.6 Summary and Outlook . . . . . . . . . . . . . . . . . . . . . . . . 106

7 Materials and Methods 108

7.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 108

7.1.1 Buffers and Solutions . . . . . . . . . . . . . . . . . . . . 108

7.1.2 Mammalian Cell Lines and Bacteria . . . . . . . . . . . . 109

7.1.3 Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109

7.1.4 Plasmids . . . . . . . . . . . . . . . . . . . . . . . . . . . 110

7.1.5 Reagents and Chemicals . . . . . . . . . . . . . . . . . . . 110

7.1.6 Kits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111

7.1.7 Equipments . . . . . . . . . . . . . . . . . . . . . . . . . . 111

7.1.8 Consumables . . . . . . . . . . . . . . . . . . . . . . . . . 111

7.1.9 Softwares . . . . . . . . . . . . . . . . . . . . . . . . . . . 112

7.2 Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 112

7.2.1 Cell Culture Methods . . . . . . . . . . . . . . . . . . . . 112

7.2.1.1 Transient Transfection of Mammalian Cells . . 113

7.2.2 Incorporation of Analogs into the Cells . . . . . . . . . . 113

7.2.2.1 Incorporation by Electroporation . . . . . . . . 113

7.2.2.2 Liposome Transfection Using DOTOP . . . . . . 114

7.2.3 Cell viability Assays . . . . . . . . . . . . . . . . . . . . . 114

7.2.3.1 AlamarBlue Assay . . . . . . . . . . . . . . . . . 114

xii CONTENTS

7.2.3.2 Luminescent Cell Viability Assay . . . . . . . . 1157.2.4 Molecular Biology Methods . . . . . . . . . . . . . . . . . 115

7.2.4.1 Cell Lysate Preparation . . . . . . . . . . . . . . 1157.2.4.2 Immunocytochemistry for γ-H2AX Staining . . 1167.2.4.3 Autophagy Sensors (LC3B-RFP) Assay . . . . . 117

7.2.5 Microscopy Methods . . . . . . . . . . . . . . . . . . . . . 1177.2.5.1 Wide-field Frequency Domain FLIM . . . . . . 1177.2.5.2 Laser Scanning Confocal Microscopy . . . . . . . 119

7.2.6 Statistics and Data Analysis . . . . . . . . . . . . . . . . 1207.2.6.1 Colocalization Analysis . . . . . . . . . . . . . . 120

Abbreviations 123

Bibliography 129

Chapter 1

Introduction

Biological organisms are the most complex systems on the earth ranging fromsimple prokaryotes to the highly evolved eukaryotes. One of the reasons for thisincredible complexity is the diversity of interactions among biomolecules thatvary from simple ions to macromolecules like proteins and nucleic acids. Allthese diverse molecules operate in concert to accomplish a plethora of cellularfunctions. Major categories of biomolecules in cells are proteins, carbohydrates,nucleic acids, and lipids. However, there are other small biomolecules that areequally essential for cellular homeostasis. These molecules predominantly con-tribute as the precursors of the cellular constituents or regulatory factors e.g.in signaling.

In this thesis, the cellular dynamics of nucleotides is investigated. Nu-cleotides are a special class of low molecular weight molecules that exhibit var-ious specialized roles in cells. Nucleotides typically constitute of nitrogenousbases (purine or pyrimidine), a five carbon sugar and phosphates. Flavin adeninedinucleotide (FAD), nicotinamide adenine dinucleotide (NAD+), and adenosinetriphosphate (ATP) are prominent examples of nucleotides. Nucleotides serveas the monomeric precursors for DNA and RNA biosynthesis. Adenine nu-cleotides like FAD, NAD+ and coenzyme A (CoA) act as coenzymes neededfor the proper functioning of the enzymes. Dinucleotide NAD+ is implicated inelectron transfer in various cellular redox reactions. Purine nucleotide triphos-

2 Introduction

phates especially ATP act as intermediate energy sources. ATP is known as theuniversal energy carrier of the cells and is central to the cellular metabolic path-ways. Nucleotides also act as metabolic regulators e.g. cAMP plays a role inhormonal signaling and ATP is used in protein phosphorylation and subsequentsignaling. Nucleotides also serve as activated intermediates in certain biosyn-thetic pathways e.g. UDP-glucose during glycogen synthesis. One aim of thisthesis is to develop methods to follow the hydrolysis of ATP in the respectivereactions.

Posttranslational modifications (PTMs) of proteins are physical or chem-ical modifications of proteins after they are synthesized. PTMs regulate thebiological activity of proteins by changing their properties. This results in themodulation of protein interactions with other partners giving rise to their diversefunctional roles. Some nucleotides act as donors for protein modifications e.g.s-adenosylmethionine acts as a methyl donor in the methylation. Both ATP andNAD+ act as substrates in the posttranslational modifications of proteins. ATPacts as a donor in case of protein phosphorylation. NAD+ acts as a donor forthe process of Poly(ADP-ribosylation) (PARylation), a posttranslational modi-fication that is essential for the initiation of DNA damage response. This thesisalso aims at the visualization of PARylation in DNA damage response.

Investigating the dynamics of the biomolecular interactions is paramountto understanding the basic biological functioning. Numerous conventional meth-ods have been devised to detect and monitor these interactions. This allowedthe the characterization of the molecules and their interaction pathways. Thesemethods are based on biochemical or biophysical techniques which are ofteninvasive and provide an ensemble picture of a molecular process. The visu-alization of these processes in real-time and in cells could provide a profoundunderstanding of the molecular events. Direct monitoring of the binding kinet-ics, subcellular localization and, mobility at molecular level will be beneficial inexploring the interactome of a biomolecule in a given biological process. Flu-orescence microscopy has been at the forefront of biological research for a fewdecades. It has a unique advantage of deciphering the mechanisms of different

3

functions at cellular and molecular levels. The technological advancements inthe field of fluorescence microscopy are driven by both, progress in the synthesisof fluorescent probes and methodological progress in microscopy. The advance-ment in the synthesis of biosensors and fluorescent probes has led to a revolutionin imaging the molecular dynamics in living cells. The availability of fusion flu-orescent proteins like green and yellow fluorescent proteins (GFP and YFP) hasincreased the possibilities to monitor and quantify specific molecular interac-tions with high precision and resolution. At the same time, modern techniqueslike FRET, FCS, FLIM and FRAP have been extensively used in biologicalresearch to study the molecular diffusion, interactions and, dynamics in cells.Fluorescence lifetime imaging microscopy (FLIM) is a powerful and sensitivetechnique that is being extensively used for imaging in biology. FLIM imagingis based on Förster resonance energy transfer (FRET) between the fluorophoresin close proximity. FRET leads to changes in the fluorescence lifetime whichis the characteristic for each fluorophore, thus providing contrast for imaging afluorophore undergoing FRET. FLIM imaging can be performed in the complexenvironment of the cell to monitor molecular interactions in real-time.

This thesis aims at exploring the potential of novel nucleotide based flu-orescent markers which were synthesised in the group of Prof. Dr. A. Marxat the University of Konstanz in fluorescence microscopy experiments of the in-tracellular hydrolysis of ATP and the involvement of the PTM PARylation incellular DNA damage response. These modified nucleotides were incorporatedinto the cells and imaged in association with EGFP fusion proteins. The uti-lization of these fluorescent analogs in the cells results in the change in theirspectral properties including their intensities and fluorescence lifetimes. Thesefluorescence characteristics were monitored with FLIM microscopy and used tostudy the mechanistic details of the pathways they are associated with.

This thesis is based on the imaging of two fluorescent analogs i.e. ATPanalog (Ap4) and NAD+ analog (TMR-NAD) in living cells. The hydrolysisof the ATP analog in living cells was monitored in real-time. The hydrolysiswas observed to occur in the lysosomes. Also, the correlation of Ap4 hydrolysis

4 Introduction

was seen with cellular autophagy. The protein PARylation in DNA damageresponse was visualized by the utilization of NAD+ analog in combination withthe fluorescence microscopy. The EGFP fusion proteins were used to ascertainthe kinetics of the two proteins, ARTD1 and mH2A, by FLIM imaging.

The first part of the thesis deals with the theoretical background of thefluorescence microscopy and its application in studying the molecular dynamicsin biology. Also, ATP hydrolysis in cells and DNA damage response is reviewed.The second part covers the experimental results and discussions related to theATP analog hydrolysis and PARylation. The third part presents the experi-mental methodology and the materials utilized in the work.

Chapter 2

Molecular Imaging in Living

Cells

2.1 Microscopy in Biological Research

Microscopy has been at the forefront of biological observations since the 17thcentury. With the inventions made by Anton Van Leeuwenhoek, the beginningof imaging in biology was marked. These simple optical devices allowed us thevisualization of the tiny organisms which he referred to as animalcules. In 1665,Robert Hooke made an important discovery after observing a thin slice of corkusing his device and the word “cell” was used for the first time. [1] However, itonly referred to a basic physical unit of living organisms at that time. Later, inthe 19th century the studies by three scientists, Matthias Schleiden, TheodorSchwan, and Rudolf Virchow described the cell as a physiological unit of lifethat lead to cell theory, a fundamental unifying theory in biology. [2]

The history of cell biology has been closely related to the developmentof the microscopy. Light microscopy is especially well suited for imaging cellsbecause the size of the subcellular structures matches the resolution limit of theconventional microscopes and also using light for imaging is largely harmlessfor biological samples. In the late 19th century, the increase in the develop-ment of optical instruments on the industrial scale led to the improvement of

6 Molecular Imaging in Living Cells

the resolution and illumination. This led to the discovery of previously invis-ible cellular components. The improvement in the quality of lenses with timeresulted in the decrease in the optical aberrations. This facilitated the visual-ization of the structural details of biological samples using conventional brightfield microscopy. However, most of the cellular structures are not colored, do notabsorb much light and have the similar refractive indices. This makes it hardto distinguish between different cellular structures. New specimen cutting andstaining techniques increased the specimen contrast. Staining of the samplesusing chemical dyes e.g. hematoxylin (binds to proteins) and eosin (binds toDNA) were used to make the different cellular constituents distinguishable. [3]

The development of additional optical methods that provide contrast assistedthe progress for imaging living cells. Two commonly used microscopy techniquesare phase contrast microscopy and differential interference contrast microscopy.These techniques allow the phase shift of light while passing through the cellsto be seen which results in a clear contrast within different cell organelles.

The subsequent development of fluorescence microscopy revolutionizedthe whole imaging field. Initially these microscopes relied on the autofluores-cence of samples. The development of fluorescent dyes like fluorescein, rho-damine and Texas Red made the fluorescence microscopy more powerful andversatile. These fluorophores absorb and emit light at a certain wavelengths thatare different. The emitted wavelength is collected to make the image whereasthe excitation wavelength is filtered out by specific filters which leads to imageswith high signal-to-noise ratios. Initially, the fluorescent labels were taggedto antibodies against specific cellular targets and the cells were stained withit. This was an ideal approach for imaging specific cellular structures. Theseimmunofluorescence techniques were applicable for fixed cells only. Moreover,some proteins were also tagged in vitro with a fluorescent dye and then in-jected the cells for microscopic studies.e.g. actin polymerization. [4] A majorbreakthrough in fluorescent labelling was the discovery of green fluorescent pro-tein, [5] it’s structural analysis, [6] and the expression of fusion proteins withGFP. [7] The discovery and development of green fluorescent protein (GFP) wasa breakthrough which had an everlasting effect on biological research. GFP

2.1 Microscopy in Biological Research 7

is a natural fluorescent protein found in a jellyfish (Aequorea Victoria) whichhas been characterized. The cloning of GFP resulted in the development of itsmany variants and other fusion proteins. The genetic expression of GFP fu-sion proteins promoted the monitoring of the desired subcellular locations anddynamic behavior of molecules in the cell, giving rise to a myriad of biologicalapplications.

With the advent of fluorescence microscopy, contrast and resolution ofcellular images improved remarkably. But fluorescence from features that areabove and below the focal plane of the specimen blurred the images due to theout-of-focus light. The princile of confocal microscope that was given by MarvinMinsky in 1957 [8] avoids this problem by detecting the fluorescence only fromone point at a time and thus allowing us to visualize molecules in one planeto create a sharp three-dimensional image. Confocal microscopy thus provedan invaluable tool for imaging cell structure and function by providing an in-crease in the resolution and optical sectioning capability. Lately biologists usemore advanced techniques like structured illumination microscopy (SIM), light-sheet microscopy and, total internal reflection fluorescence (TIRF) microscopywhich permit the live cell imaging with even better three-dimensional resolu-tion over longer time. The last three decades have also seen the development ofnew types of nonlinear optical microscopes such as multiphoton excitation mi-croscopy. These techniques are compatible with the imaging of thick tissues andeven organisms as they use near-infrared (NIR) wavelengths which have deeperpenetration depths and are comparatively less damaging for the living specimen.Recent emerging technological advancements in optics and electronics has givenus super-resolution techniques like Stimulated Emission Depletion Microscopy(STED) and Stochastic Optical Reconstruction Microscopy (STORM) whichcircumvented the diffraction limits of conventional light microscopy. With someof these techniques, it is possible to resolve cellular features of up to 20 nmin the XY plane and 50 nm in the Z plane. These techniques are technicallychallenging and need very sophisticated instrumentation. Nevertheless, they areincreasingly used to resolve the ultrastructure of cellular components. [9] The de-velopment of Förster resonance energy transfer (FRET) microscopy paved theway for imaging the dynamics of molecular interactions. [10] During the last 20


years, it has been widely applied to understand the molecular proximity, confor-mation, and interaction. Spectroscopic techniques like fluorescence correlationspectroscopy (FCS), fluorescence recovery after photobleaching (FRAP), andFörster resonance energy transfer (FRET) paved the way for monitoring theinteraction and mobility of biomolecules in cells.

2.1.1 Principles of Fluorescence Microscopy

Fluorescence is the light emitted by a substance after it absorbs light of a certainwavelength. The light is emitted because of the transition of electrons from theexcited electronic state to the ground electronic state. Fluorescence was firstobserved by a Spanish botanist, Nicolas Monardes in 1565. [11] Robert Boyle, in-spired by Monardes’ observation investigated this system further and concludedthe presence of some essential salt in the wood to be responsible for this effect.In 1845, John Herschel observed fluorescence from quinine sulphate. In 1852,George G. Stokes published his work on quinine sulphate and used the termdispersive reflection to describe the phenomenon. He described his observationthat red light was emitted by the substance when illuminated with ultraviolet(UV) light. [12] He coined the term “fluorescence”. His ideas subsequently led tothe application of fluorescence for analytical purposes. It was in 1930s whenfluorochromes were first utilized to stain biological samples for investigation.

Upon the illumination of a fluorophore with a particular wavelength, ex-citation of an electron takes place from its ground state S0 to the excited stateS1 or S2. The excited electron quickly loses its energy to return to the groundstate (S0) or first excited state (S1). The process in which a molecule emits lightfrom an electronically excited state is called luminescence. The molecules cango into excited states by physical or chemical means, and the luminescence gen-erated by a molecule that is excited by photons is called photoluminescence. [13]

The electronic transitions that occur after photo-excitation of a molecule canbe comprehensively summarized by the Jablonski diagram (Figure 2.1). Thediagram is named after the Polish physicist Alexander Jablonski who has pi-oneered many important concepts of fluorescence spectroscopy. [14] Depending


on the mechanism, the emitted light can be categorized into fluorescence andphosphorescence. Fluorescence is the transition of a molecule from an electronicexcited state (S1) to ground state (S0) with the emission of a photon. It hap-pens at the picosecond to nanosecond time scale. Alternatively, the moleculecan undergo intersystem crossing (ISC) to the triplet state (T1). The numberof electrons undergoing intersystem crossing is lower as they compete with thealternative pathways of fluorescence and internal conversion. In this case, thetransition from T1 to S0 results in phosphorescence. Since this transition isforbidden, the phosphorescence occurs over a longer time scale of the order ofmicroseconds to seconds. Excited molecules may also undergo non-radiativetransitions because of internal conversion to lower electronic states. Electronicstates are accompanied by vibrational energy levels which have relatively smallenergy differences.

Figure 2.1: Jablonski Diagram

Light emitted in phosphorescence has a longer wavelength and lower en-ergy as compared to the light absorbed because a part of the energy is released innon-radiative decay process. This shift towards the longer wavelength is calledthe Stokes shift. Excited fluorophores can also transfer their energy simply by


collisions to other molecules. This causes non-radiative decay and leads to fluo-rescence quenching. Electronic transitions occur in a variety of substances withconjugated double bond systems. The efficiency of the conversion of absorbedto emitted light by a fluorophore can be estimated in terms of its quantumyield. Fluorescence quantum yield is the ratio of photons absorbed to the pho-tons emitted through fluorescence. A fluorophore can undergo 106 cycles ofexcitations and emissions before it is photobleached.

2.1.2 Types of Microscope Setups

Based on the configuration of the main elements of a microscope, different typesof microscopes have been developed that vary in their operations. All types ofsetups offer advantages and disadvantages and are used depending on the desiredrequirements of the observation. Widefield and confocal are the two prominentlyused microscopy methods that are discussed below:

2.1.2.1 Widefield Microscopy

Widefield microscopy (WFM) is a form of bright field microscopy where thewhole specimen is illuminated by the light simultaneously. It mainly consistsof a light source, a camera, and a microscope unit. Initially, gas arc lamps likethe mercury arc-lamp and the xenon-arc lamp were used as the light sources inwidefield microscopes. Nowadays, light-emitting diodes (LED) and high inten-sity lasers are also used as fluorescence light sources. The camera is based onsemiconductor detectors and most commonly used sensors are charge-coupleddevices (CCD) and the complementary metal oxide semiconductors (CMOS).Widefield microscopes exist in two configurations: upright in which the lightilluminates the specimen from below and inverted, in which the specimen isilluminated from above. The upright configuration is generally suited for fixedsamples whereas inverted microscopes are used to image living samples. Appro-priate filters are used for the illumination with the excitation light and detectionof emission fluorescence. The spatial resolution of a microscope is the minimumdistance between two points in a specimen that can be perceived separately. Theresolution limit is governed by the numerical aperture of the objective used, thewavelength of the excitation source, and the refractive index of the medium be-


tween object and objective. In a perfectly aligned light microscope under idealconditions, the resolution limit is approximately 200 nm in the XY plane for anumerical aperture of 1.4 and 500 nm excitation. The major drawback of wide-field fluorescence microscopy configuration is a relatively high background. It isbecause the entire specimen is illuminated at once and the light from differentregions other than the focal plane contributes to the signal to obscures somefeatures (Figure 2.2 a).

Widefield imaging is suitable for imaging dynamic processes in living cellsover longer periods. It is more favourable for the visualization of thin samplese.g. monolayers of cells with a reasonable signal-to-noise ratio. The advantageof the widefield microscopy is that the signal recording requires lesser exposuretimes because the sample is entirely exposed to the light that maximizes thelight gathered by the objective. Widefield microscopy is used to image a widevariety of samples including bacteria, yeast, adherent human cells, and thintissue sections. [15]

2.1.2.2 Confocal Microscopy

Laser scanning confocal microscopy has the advantage as it utilizes a pinholeto filter out the out-of-focus light. A focused laser beam at a single point il-luminates the sample and out-of-focus light is eliminated before the emissionis collected by the objective. Each point of the sample is scanned individuallyin a sequential manner and only the light coming from the pinhole is recorded,thus building the image point by point. [16] This gives an advantage for imagingrelatively thick and dense samples and to obtain better three-dimensionally re-solved images. Lasers are used as the excitation sources in confocal microscopy.The emitted light is collected and detected using photomultiplier tubes (PMTs).The photons collected are converted into electric signal that corresponds to anintensity measurement for each pixel. This is displayed as a complete imageafter computer processing. The major disadvantage of the laser scanning con-focal microscopy is that it is relatively slow since an image is formed by rasterscanning of the specimen. The comparative high intensities per pixel makes thesample prone to photobleaching. Moreover, the high-intensity lasers used can


damage living samples especially when the samples are imaged for prolongedtime. Another variant of confocal microscopy called the spinning disk confocalmicroscopy is used widely nowadays. It uses a disk with multiple pinholes to scanmultiple points in focus simultaneously and the image is generated by emissionfrom all these points. A sensitive electron-multiplying charge-coupled device(CCD) is used to capture the image. [17] (Figure 2.2 b) This microscopy methodis faster as compared to laser scanning and can image hundreds of frames persecond. It is widely used for imaging more dynamic cellular processes at theprotein and organelle level. e.g. dynamics of microtubule in mammalian cellshave been imaged using this method. [18]

Figure 2.2: Schematics of the microscope setups; (a) Widefield setup (b)

Confocal setup.

Technological advancements have led to the Integration of multiple elec-tronic systems including detectors, lasers, and computers in combination withimage processing algorithms to produce a complete imaging confocal unit. Con-focal microscopy has revolutionized the imaging of biological samples by in-creasing the signal-to-noise ratio. This has led to a tremendous increase in thepopularity of confocal imaging in recent years mostly among biologists.

2.2 Imaging Molecular Interactions and Dynamics 13

2.2 Imaging Molecular Interactions and Dynam-

ics

Transient interactions between various macromolecules like proteins, nucleicacids, sugars, cofactors, metal ions, etc are of great importance for organiz-ing the complex and wide range of functions in the living cells. Detection andmonitoring of these interactions is crucial for unraveling the underlying mecha-nisms of many cellular processes. [18] Understanding the assembly of molecularcomplexes and their regulation is important to decipher various physiologicaldisorders and to devise potential therapeutic strategies. Many different tech-niques have thus been developed over time to probe biomolecular interactionsprimarily involving proteins. [19] Examples of prominent molecular biology andbiochemical techniques are coimmunoprecipitation assays, protein arrays, X-raycrystallography, and mass spectrometry. These techniques have some inherentdisadvantages as they are not applicable for the analysis of molecular dynamicsand interactions in a living cell. Instead, these methods demand the destructionof the native cellular structure and function and cannot distinguish between di-rect and indirect interactions. Special in vitro conditions are needed to facilitatethe potential interactions to perform these assays which sometimes creates theexperimental the artifacts. [20] Recent advancements in microscopy have broughta revolution in the methodology for studying the dynamics of interactions in thecells. Some of these methods offer the live cell imaging that efficiently comple-ments the conventional biochemical and biophysical methods for observationsin vivo.

The highly dynamic interactions that occur at the time scale of microsec-onds can be visualized and quantified in living cells using contemporary mi-croscopy techniques in combination with the fluorescent fusion proteins andfluorescent bio-analogs. The cloning of fluorescent proteins with their widespectra of emission and excitation wavelengths as fusions with proteins of inter-est has proved to be an efficient way to follow specific cellular functions. [21,22]

Widefield and confocal microscopy approaches have for example been used tostudy the subcellular localization of nuclear receptors. Their interacting part-


ners and their regulatory mechanism have been extensively studied using theseapproaches. [23,24,25] Confocal microscopy can be used for the colocalization andtracking of molecules to understand the positioning of the molecules in sub-cellular compartments and their interactions. To this end, one can colocalizedifferent molecular species in confocal microscopy images of the same samplearea recorded at different wavelengths. Thus colocalization have been used ex-tensively to study cellular trafficking mechanism, [26] membrane protein dynam-ics [27] and receptor localization. [28] A problem of colocalization is the fact thatthe resolution limit of confocal microscopy is far worse than the typical size ofproteins. Therefore, other microscopy techniques are widely used to determinethe protein-molecule interaction. These include Förster resonance energy trans-fer (FRET), [29] fluorescence recovery after photobleaching (FRAP), [30] Fluo-rescence Correlation Spectroscopy (FCS), [31] bioluminescence resonance energytransfer (BRET), [32] and bimolecular fluorescence complementation (BiFC). [32]

Apart from monitoring interactions on a protein length scale, one of the impor-tant advantages these imaging techniques offer over the traditional methods ispossibility to visualize of molecular interactions specifically in living cells. Thelive cell imaging tools provide the functional details including the conformationalchanges and the dynamic behavior of molecules in real-time and non-invasively.Especially methods based on FRET have proven to be effective methods forvisualizing the molecular interactions in a living cell. In combination with theexpression of fluorescent proteins, this has helped in the elucidation of the kinet-ics of crucial processes like transcription, translation and metabolism. [33] FRETis discussed thoroughly in the next section.

2.2.1 Förster Resonance Energy Transfer

An emitting fluorophore can be described as an oscillating dipole. It can transferits energy directly and non-radiatively to another chromophore in case of theirclose proximity. This transfer of energy from donor to acceptor fluorophore in-duces electronic excitation of the acceptor without the involvement of emissionof photons. This phenomena is called Förster resonance energy transfer andthe pair of molecules involved in such interaction is called a FRET pair. Carioand Frank first observed resonance energy transfer in 1922. [34,35,36] In 1927 first


theoretical explanation was given by J. Perrin and a few years later, his sonF. Perrin stated the theory of resonance energy transfer. [37,38] Theodor Förstercontributed to the theoretical understating of the resonance energy transfer.In 1946, he postulated that the energy transfer depends on the intermoleculardistance and the overlap of their emission and absorption spectra. The termFörster radius associated with FRET has been named after him. [39,40]

For a measurable FRET to occur, certain prerequisites need to be ful-filled. These include the close proximity between the two fluorophores i.e. thedonor and acceptor (typically 1 to 10 nm), a significant spectral overlap betweenthe donor emission and the acceptor absorption, their appropriate dipole ori-entations, and the compatible fluorescence lifetimes. The limitation of FRETto distances below 10 nm means that its observation confirms colocalization onsize scales of a typical protein. It thus qualifies to monitor protein-moleculeinteractions. The efficiency of FRET is dependent on the inverse sixth power ofthe distance between the donor and acceptor. It can be stated by the followingequation:

E =R6

0

R60 + r

(2.1)

R0 is the Förster radius and it is the distance at which the energy transfer is50 percent efficient and r is the experimental distance between the two. For thepractical applications of FRET, the Förster radius R0 needs to be calculated.R0 can be calculated by the equation:

R60 =

9000 ln 10 ΦD K2

128π5 N n4

∫ ∞0

FD(λ) .εA(λ) .λ4 .dλ (2.2)

Here, ΦD is the donor quantum yield, N is Avogadro’s number, n is therefractive index of the medium, and lambda is the wavelength. K2 is the orien-tation factor that depends on the alignment of donor and acceptor dipoles. [41]

The orientation factor ranges from 0, in case both dipoles are perpendicular to 4where both dipoles occur parallel. Assuming the random orientation of dipolesbecause of the rapid movement of molecules, the orientation factor is considered


to average to 0.67. [42]

FRET can manifest itself in different forms that can be monitored usingvarious separate approaches. As a consequence, many diverse techniques havebeen employed to quantitatively measure and analyze FRET. [43,44,45] Some ofthe widely used approaches to measure FRET are discussed below.

2.2.1.1 Fluorescence Intensity based FRET

In this method, FRET is determined from the intensity of the fluorophores thatcan be quantified in various ways. The most straightforward method is measur-ing donor quenching. It is based on measurements of fluorescence intensities ofthe donor after excitation in absence of the acceptor and in presence of accep-tor. In case of FRET in presence of an acceptor, the donor intensity decreases.Since the intensity has to be measured in two different samples, any differencein the experimental conditions between two samples makes this method proneto errors. To make the measurement more reliable, fluorescence intensity hasto be averaged over a large number of samples. Thus the method cannot beused on single-cell level. The approach is only suitable for measuring FRET onensemble level e.g. in cytofluorimetry and spectrofluorimetry. [46,47] The FRETefficiency in case of intensity based approach can be calculated by the equation:

E = 1 − IDAID

(2.3)

Here IDA is the fluorescence intensity of the donor and ID is the intensity ofthe donor in presence of acceptor.

FRET thus results in the change in intensities of the two fluorophoresinvolved based on the distance between them. The ratiometric intensity valuesof donor and acceptor can be quantified and then be used for the estimationof the distances. This principle is widely employed in biology for studying thebiomolecular interaction after their fusion with the fluorophores. [48,49]

In acceptor photobleaching, the fluorescence intensity of the donor is


measured before and after photodepletion of the acceptor fluorophore. The in-tensity of the donor will increase in case of FRET assuming the photobleachingabolishes the acceptor absorption. This increase in intensity of the donor givesthe estimation of FRET. This method is time-consuming and destroys the sam-ple such that the measurement cannot be repeated. Moreover, the possibility ofincomplete photodestruction of acceptor and generation of dark acceptor speciesmay lead to variable results. This method is mostly prefered for the analysis offixed samples. [50]

One of the most reliable methods is the sensitized emission method. Inthis method, the donor fluorophore is excited by the appropriate wavelengththat results in the energy transfer and the excitation of the acceptor. The spec-trally resolved emission fluorescence of the acceptor is determined to measureFRET. [51] A significant problem with this method is the fluorescence crosstalkbetween the FRET pair. Thus extensive control experiments are needed toestablish quantitively accurate FRET measurements. The corrective measuresinclude measuring separately the intensities of donor only, acceptor only, and thecombination of both. Despite these complications, this method is widely usedfor FRET measurements. It is best suited for the analysis of in vitro sampleswith predetermined ratios of the donor and acceptor. However, this method isstill used for dynamic FRET measurements in living samples.

2.2.1.2 Fluorescence Anisotropy based FRET

In this approach, the variation in the anisotropic nature of the emission from amolecule is measured by using excitation with polarized light. [52] When a sampleis excited with polarized light, the emitted light remains largely anisotropic i.e.polarized in the same direction as excitation. In case of FRET, the emitted pho-ton from the acceptor is polarized based on acceptor orientation. This causes adecrease in the anisotropy which is measured to estimate FRET. The data fromanisotropy measurements can be interpreted instantaneously after the acquisi-tion without complex image analysis procedures. Thus, measurements of thistype take relatively little time. Anisotropy is comparatively insensitive and thusis better suited for the quantitative analysis of large clusters of molecules. [53,54]


2.2.1.3 Fluorescence Lifetime based FRET

Fluorescence lifetime measurements for FRET are based on the decrease in thefluorescence lifetime of donor fluorophore when it is in close proximity of theacceptor. It can be monitored with fluorescence lifetime imaging (FLIM).

2.2.2 Fluorescence Lifetime Imaging Microscopy (FLIM)

The average time of a molecule in the excited electronic state after absorptionof light is known as its fluorescence lifetime (τ). The fluorescence lifetime canbe defined as the inverse of the sum of all relaxations processes after absorption:

τ =1

kr + knr(2.4)

Here, kr is radiative rate constant and knr is non-radiative rate constant. Incase of FRET, the fluorescence lifetime (τ) decreases and can be given by:

τ =1

kr + knr + kFRET(2.5)

The decay of the excited state of a population with respect to time after exci-tation with a pulse is given by:

dn(t)

dt= −(kr + knr) n(t) (2.6)

The population of fluorophores in the excited state (n) decays with time becauseof both radiative as well as non-radiative processes. Integration and substitutionof the excited state population n(t) with the emission intensity I (t) leads to theequation:

I(t) = I(0) e−(t/τ) (2.7)

Here, I(t) is the intensity at any time t, n(t) is number of excited moleculesat time t, and τ is fluorescence lifetime. The FRET efficiency E in terms offluorescence lifetimes is calculated as:

E = 1 − τDAτD

(2.8)


Here τDA is lifetime of donor in presence of acceptor and τA is the lifetime ofdonor in absence of acceptor.

Fluorescence lifetimes are characteristic for each fluorophore and it typ-ically takes nanoseconds for a fluorophore to return to the ground state. Whentwo molecules undergo FRET, the energy of the electron in its excitation stateis nonradiatively transferred to the acceptor. This additional mechanism forthe dissipation of energy in a fluorophore results in a decrease in its fluores-cence lifetime. This decrease in the lifetime is measured to quantify the FRET.FLIM offers multiple advantages over the conventional intensity-based FRETmeasurements. [55,56] Fluorescence lifetimes are intrinsically dependent on thechemical and physical properties of a fluorophore. They are relatively insen-sitive to the change in fluorophore concentration and the intensity of the ex-citation light. This leads to a significant advantage for estimating FRET invivo where the fluorophore concentration is not well defined. Also, artifacts dueto autofluorescence of the sample and spectral bleed-through can be avoidedunlike in intensity-based measurements since only the imaging of the donor isrequired. [54,57] FLIM imaging provides the unambiguous quantification of FRETefficiency because the contrast of the images is lifetime based and not the inten-sity based. Fluorescence lifetime imaging is applied to determine FRET usingtwo methods discussed below.

2.2.2.1 Time-Domain FLIM

This method is based on measuring the fluorescence decay in time domain. Afluorophore is excited by a short laser pulse and the subsequent time of arrivalfor each single photon is measured. The process is repeated several times toacquire the photon distribution over time in the form of a histogram. The distri-bution is fitted to an exponential decay function and the representative lifetimeis extracted from the curve. The decay follows first-order kinetics for simple flu-orophores, however for some fluorophores, the decay is more complex. [56,58,59]

Time-correlated single photon counting (TCSPC) FLIM can be performed ona spectrofluorometer, a flow cytometer or a microscope. In the contemporarymicroscopy setups, time domain FLIM is performed by using a laser scanning


confocal microscope, where each point is excited sequentially with a laser andthe data is acquired pixel by pixel. The lifetimes are calculated for each pixelseparately and a FLIM image of the sample is created. This approach requireslong acquisition times and it can take tens of minutes for the construction of animage. [60] However, TCSPC-FLIM offers a high dynamic range, high sensitivity,and has the best signal-to-noise ratio. [61] It is reliable in measuring the photonarrival time and gives accurate FRET measurements (Figure 2.3 a). [62]

2.2.2.2 Frequency-Domain FLIM

In the frequency domain method, a modulated light source with a certain fre-quency and a modulated detector are used. The fluorescence emission is depen-dent on the modulation pattern and thus, the phase shift and the modulationof the emitted waveform will be governed by the lifetime of the fluorophores.The shorter the lifetime, the smaller will be the phase shift. The time delay be-tween excitation and emission causes demodulation of emitted fluorescence sig-nal. Thus, the longer the lifetime, the greater will be the demodulation. Thesephase delays and the modulation ratios are measured simultaneously (Figure 2.3b). Calculation of the lifetime of the fluorophore measuring the phase delay (φ)

leads to the phase lifetime (τφ) and the calculation based on measurement of thedecrease in amplitude (m) yields the modulation lifetime (τm). The followingequations give the estimation of the fluorescence lifetimes:

τφ =1

ωtanφ (2.9)

τm =1

ω

√( 1

m

)2− 1 (2.10)

In the above equation, the change in the amplitude is given by:

m =E/e

A/a(2.11)

For a single exponential decay, the phase lifetime (τφ) and the modulationlifetime (τm) will be same at all modulation frequencies. But in case of mul-tiexponential decays, τφ < τm with the values depending on the modulation


frequencies. In order to calibrate the FLIM system, a dye with a known fluo-rescence lifetime under standard conditions is used.

Frequency domain FLIM can be accomplished by using both widefieldapproach where the modulated cameras are used or by a laser scanning ap-proach where modulated point scanners are used. In case of widefield frequencydomain FLIM, gain modulated CCD cameras which are modulated at the samefrequency as the excitation light are used after the wide-field excitation. A se-ries of images at different phases with respect to the excitation light from 0◦

to 360◦ is acquired. The modulation frequency used depends on the lifetime offluorophore and is usually 10 to 100 MHz for the measurement of nanosecondlifetime decays. With the widefield illumination, fast acquisition of lifetimessimultaneously from all the pixels is possible, thus this method is preferred forreal time biological imaging applications.

Figure 2.3: Fluorescence lifetime measurement; (a) In time domain (b) In

frequency domain.

The frequency domain FLIM data can be analyzed by phasor plots thatprovide a direct two dimensional representation of lifetime distributions of allthe pixels. In this plot each pixel is mapped as a single point called “phasor”.Each phasor is a vector whose length is determined from the modulation (M ),


and angle is defined by the phase delay (Φ). Lifetimes that fall on the semicirclerepresent the single exponential lifetimes. [63,64] The coordinates of each pointare given by:

x = M(ω) cos Φ(ω) (2.12)

y = M(ω) sin Φ(ω) (2.13)

This universal semicircle represents the lifetime trajectory for any single lifetimecomponent. The fluorophores with one lifetime component will fall directly onthe semicircle and those with multiple lifetime components will fall inside thesemicircle. The components with longer lifetimes will shift towards the left onthe semicircle (Figure 2.4 a). [65] Frequency domain FLIM imaging has variousadvantages over the time domain method. It does not require high energy pulsedlasers thus photobleaching and phototoxicity of the sample can be avoided. Sinceit is camera based, the image acquisition is fast and thus suited for visualizingdynamics in live cell imaging applications.

Figure 2.4: Mapping of fluorescence lifetimes using phasor plot.


2.2.3 Applications of FLIM FRET Imaging

FRET is an effective method to visualize the dynamic molecular interactions inliving cells with improved resolution and sensitivity. The advancement in themicroscopy and the characterization of the various types of fluorescent probeshave allowed this technique to be used in biological research and as well asin biomedical imaging. FRET imaging has been utilized to evaluate crucialcellular functions like signal transduction, vesicular transport and gene expres-sion. [33,66,67] FRET and related imaging techniques have been used extensivelyfor studying the structure and conformational changes of proteins and nucleicacids. [68,69,70] This method has been also used to understand the multiple in-teractions in proteins complex. e.g. three-chromophore fluorescence resonanceenergy transfer (3-FRET) has allowed the analysis of three different interactionswithin a molecular machinery using three different fluorescent labels. [71]

In FLIM imaging, the measurements of only the donor can be carriedout independently of the acceptor, thus it is ideal in situations where a darkquencher is used as the acceptor. Also, it is better suited for the investigationof the molecules with intrinsic fluorescence in biological systems like NADH,FAD and tryptophan. [72] For the last decades FLIM in the time domain hasbeen used for diverse applications in biological research and clinical diagno-sis. [73] A fine example of the detailed methodology of the application of FLIMin both time domain as well as in frequency domain has been demonstrated byusing dimerization of transcription factor calcium channel associated transcrip-tional regulator CCAT as a model system. [74] FLIM in frequency domain hasfound some major applications in the field of biosensors. [75,76] FLIM has beensuccessfully used to measure the change in pH, oxygen concentration and ionconcentration in vivo. [77,78,79] Recently frequency domain FLIM-FRET has beenused to visualize the status of protein methylation in cells using methylation re-porter. [80] and receptor phosphorylation on plasma membrane. [81] FRET-basedbiosensors have been successfully employed to monitor the enzyme activity ofprotein kinase A. [82]

Chapter 3

Kinetics of ATP Hydrolysis

3.1 Introduction

Adenosine triphosphate (ATP) is one of the essential and ubiquitous biomoleculesin the living world. ATP is a nucleotide that serves as primary energy currencyin the cell. All the sources of energy that organisms utilize in the form of foodare eventually converted into the ATP, which in turn provides energy for all theendergonic functions in an organisms. [83]

ATP is widely known to be discovered in 1929 by Karl Lohman, a Germanchemist working at the Kaiser Wilhelm Institute of Biology in Berlin. However,many simultaneous studies including the one by Cyrus Hartwell Fiske with hisgraduate student Yellapragada Subbarow of Harvard Medical School played im-portant role in the discovery of ATP. They primarily discovered the role ofthis molecule in muscle contraction. [84] In 1935, the structure of ATP was pro-posed by Katashi Makino and it was later confirmed by Alexender Todd andBasil Lythgoe at the University of Cambridge. [85] From 1939 to 1941, Fritz Lip-mann’s work led to the confirmation that energy is mainly stored in the form ofATP in the cells. In 1964, Paul D. Boyer proposed the enzymatic mechanism ofATP synthesis by ATP synthase that he later discovered experimentally in1973.The mechanism came to be known as the “binding change mechanism”. John

3.1 Introduction 25

E. Walker, a British chemist, isolated and studied the mitochondrial membraneproteins and gave the structure of ATP synthase. John E. Walker elucidatedthe structure of ATP synthase published in 1994. [86,87] He shared a noble prizein Chemistry with Paul Boyer in 1997 for the discovery and elucidation of themechanism of ATP synthase.

ATP is ubiquitously present in all living organisms. It is critical for mul-tiple functions in cells ranging from simple prokaryotes to complex mammaliansystems. The energy that is derived from different nutrients by a cell is not avail-able in the appropriate quantity for the utilization in various energy-consumingpathways. This energy from the chemical bonds is translated into the highenergy phosphate bonds of ATP in the cellular mitochondria. [88] Around 100to 150 moles of ATP are constantly recycled by the human body per day forvarious cellular activities. At a given time, there are approximately one billionATP molecules present in each cell and these molecules are completely recycledwithin a few minutes. Since there are hundreds of trillions of cells in an averagehuman body, about 1023 ATP molecules normally exist in a human body. Foreach ATP molecule, the terminal phosphate is removed and added 3 times inone minute. [89] A molecule of ATP contains one nucleotide, adenosine, and threephosphates. Three of the oxygen atoms are shared either by two phosphates orthe first phosphate and carbon group (Figure 3.1). [90,91] The ATP hydrolysisresults in the release of high amounts of energy from each phosphoanhydridebond and this hydrolysis is coupled to endergonic biological reactions. Withthe hydrolysis of β-γ phosphoanhydride bond, about 30 kJ/mol of free energy isreleased, and the hydrolysis of α-β bond releases about 32 kJ/mol of energy. [92]

ATP is a relatively stable molecule and thus hydrolysis takes place in presenceof a catalyst that in most cases is a specific enzyme.

Understanding the mechanism of the functioning of ATP was fundamen-tal to unraveling the biochemistry of metabolism in cells. The negative chargeof the phosphates is responsible for the sequestration of the ATP molecules intocompartments enveloped by phospholipids and also for their stability towardschemical attack. [93] The chemical stability combined with the thermodynamic

26 Kinetics of ATP Hydrolysis

Figure 3.1: Chemical structure of adenosine triphosphate (ATP) nu-

cleotide.

instability makes this molecule suitable for biochemical and physiological rolesin living organisms. [94] ATP is used to fuel the millions of metabolic chemicalreactions that require energy in the cell. It is the source of energy for the me-chanical mobility of organisms e.g. muscle contraction including the skeletalmuscles for movement and the heart muscles for blood circulation. [95] In lowerorganisms, the energy is used for the moment of cilia and flagella. [96] Moreover,active transport of molecules like ions through the cell membrane against con-centration gradients also utilizes ATP. Transport of cargo inside a cell along thecytoskeleton components with the help of motor proteins also requires ATP. [97]

However, apart from providing energy, ATP is also involved in other type ofphysiologically important mechanisms. As an example, the process of phospho-rylation wherein a protein gets a phosphate group attached to it in response ofa stimulus involves ATP. These phosphorylation and dephosphorylation eventsact as on/off switches that regulate many cellular pathways including signal-ing. [98,99] Phosphorylation and dephosphorylation reactions are catalyzed byprotein kinases and phosphatases. [100] Derivatives of ATP are involved in cellu-lar signalling. Cyclic adenosine monophosphate (cAMP) is a derivative of ATPand is widely employed in the regulation of metabolic and signaling pathwaysin eukaryotes and bacteria. [101] The nucleobase adenine, a part of adenosine is a

3.2 Cellular ATP Hydrolyzing Enzymes 27

constituent of nucleic acids, DNA, and RNA. ATP acts as a precursor for thesenitrogenous bases.

3.2 Cellular ATP Hydrolyzing Enzymes

Living organisms possess numerous enzymes that hydrolyze ATP to performdiverse cellular functions. Some of the essential processes include cellular activetransport, nucleic acid biosynthesis, protein biogenesis and degradation, mem-brane biogenesis, signal transduction, and gene expression regulation. Someprominent cellular ATPase systems are discussed below.

Transport ATPases are transmembrane proteins that are responsible formaintaining the ion gradient across biological membranes. These are dividedinto three general categories:

1. P-type ATPases are a large group of ion pumps that are evolutionarilyconserved and found in both prokaryotes and eukaryotes. Their regula-tory mechanism is based on autophosphorylation. [102] Some of the com-mon examples of this type of ATPases are the sodium-potassium pump(Na+/K+-ATPase), the proton-potassium pump (H+/K+-ATPase), thecalcium pump (Ca2+-ATPase) and the plasma membrane proton pump(H+-ATPase).

2. V-type ATPases, also called proton pumps, mainly transport protonsacross the intracellular and plasma membrane in eukaryotic cells. Theyuse ATP to produce the proton gradient. They are found also on themembranes of endosomes and lysosomes. A variety of cells with special-ized functions also have V-type ATPases on the plasma membranes thatinclude macrophages, neutrophils, sperm cells, and osteoclasts. [103] [104]

3. F-type ATPases usually work as ATP synthases in cells. They synthesizeATP from the proton gradient across the membrane but occasionally alsohydrolyze ATP. They are found in the plasma membrane of bacteria and


the inner mitochondrial membrane and chloroplast of eukaryotes. Theyconsist of a soluble F1 and a membrane-embedded F0 portion. [105] [106]

Motor proteins are a fascinating class of protein complexes that act as mechani-cal nano-machines and are associated with cellular cytoskeleton. These proteinsmove in a unidirectional manner along the tracks of cytoskeletal filaments totransport cargo while using the energy from ATP hydrolysis. Kinesins anddyneins are the motor proteins that walk along the microtubule network in thecells and transport cargo. These proteins typically consist of two domains, anATP binding domain responsible for ATP hydrolysis to generate energy and acargo binding domain. [107,108] Kinesins transport the large molecules, vesiclesand cellular organelles like mitochondria unidirectionally from the center of thecell towards the surface. They are responsible for the movement of chromosomesduring the cell division. [109,110] Dyneins, on the other hand, transport cargo fromthe cell periphery towards the center. An important example is transport of or-ganelles from the distal region to the cell body in axons. [111] Myosin, a skeletalmuscle protein responsible for the muscle contraction was the first motor pro-tein identified. The head domain of the myosin is the most conserved motif thatcontains the binding site for actin and for ATP. Each myosin binds to the actinfilament and slides across the filaments thus converting the chemical energy ofATP into mechanical movement. [112] The actin cytoskeleton is also involved inthe crawling movements of some specific cell types. Examples of this type ofmovement includes the migration of cancer cells during metastasis, migration ofembryonic cells during developmental stages and of the white blood cells duringimmune response. [113,114,115]

ATPases associated with diverse cellular activities (AAA proteins) area family of ATPases with diverse functions and play a crucial role in proteinhomeostasis. This superfamily of proteins is mainly involved in membrane fu-sion, disassembly of protein aggregates, microtubule dynamics, transcriptionalregulation, protein folding and DNA replication. [116,116]

Kinases are an important class of enzymes that carry out the phospho-

3.2 Cellular ATP Hydrolyzing Enzymes 29

rylation by transferring the γ-phosphate of ATP to hydroxyl groups of a widerange of substrates including lipids, sugars, and amino acid residues of the pro-teins. The phosphorylated substrate can donate a phosphate group by a processcalled dephosphorylation catalyzed by phosphatases. Phosphorylation plays apivotal role in crucial cellular processes including metabolism, cell signaling,protein homeostasis, and cellular transport regulation. In prokaryotes, some ofthe kinases involved in signaling are His-Asp kinases, eukaryotic protein kinase(ePK), eukaryotic-like kinase (ELK). [117,118] In eukaryotes, the kinases are clas-sified on the basis of the amino acid residues they phosphorylate, which includetyrosine specific kinases, serine/threonine kinases or tyrosine and threonine dualspecificity kinases. [119,120]

Some of the prominent families of kinases are:

1. Cyclin-dependent kinases (CDKs) well known for their role in cell cycleregulation. Each of the four stages of the cell cycle is tightly regulated bythe phosphorylation of specific cyclins which is carried out by this classof serine/threonine kinases. The level of cyclins is regulated at each stageand binding to CDKs releases the checkpoints for the cell cycle progres-sion. [121] [122]

2. Mitogen-activated protein kinases (MAPK) are constituents of the signaltransduction systems. They are serine/threonine type of kinases that per-ceive the extracellular signals and mount response by regulating the geneexpression. MAPK pathways form a cascade of kinases and each substratephosphorylation catalyses the transmission of signal usually to the cellulartranscriptional factors. Some of the prominent transcriptional factors di-rectly or indirectly regulated by the MAPK are Myc, c-Myc, c-Jun, Max,Elk-1, Max, ATF-2 and p53. These transcription factors regulate impor-tant cellular events like differentiation, proliferation and apoptosis. [123,124]

3. Sugar kinases are a group of kinases that phosphorylate various carbohy-drates. They play a pivotal role in the metabolism of carbohydrates e.g.in glycolysis. Some of the examples are glucokinase, hexokinase involved


in glucose metabolism, and phosphofructokinase (PFK) involved in theglycolysis pathway. [125] [126]

4. Lipid kinases are responsible for the phosphorylation of lipids in cells. Thephosphorylation of lipids on the plasma membrane is primarily involvedin the cellular signal transduction. Lipid kinases like phosphatidylinosi-tol 3-kinases (PI3K), diacylglycerol kinase and sphingosine kinases play acrucial role in the regulation of cellular intracellular trafficking, motilityand differentiation. [127]

3.3 Methods for Studying ATP hydrolysis

ATP hydrolysis is crucial for the normal cellular functioning and impairment inATP hydrolysis activity has enormous effects on the physiology of an organism.Monitoring the ATP hydrolysis in cells is desirable for understanding the variousphysiological processes and in elucidating the underlying regulatory mechanismof the enzymes involved. Also, the estimation of the kinetics of ATPase hy-drolytic activity is beneficial for studying enzyme functionality and inhibitoryeffects of certain molecules. Thus methods to investigate ATP hydrolysis wouldbe valuable tools for biochemical and physiological investigations. Therefore, avariety of assays has been developed to determine ATP hydrolysis activity invarious contexts. These methods have many contrasting advantages and disad-vantages and none of them singly fulfills all the requirements for ideal studies.Some of the widely used approaches based on both imaging and non-imagingmethods are discussed below.

3.3.1 Assays Based on Inorganic Phosphate Detection

In most of the energy transducing ATP hydrolysis reactions, inorganic phos-phate is released as a product. Secondary reactions are carried out with thefree phosphate that directly or indirectly create colored complexes which areobserved by using a colorimeter of a spectrophotometer. One such widely usedmethod is the classical molybdate method based on the formation of a coloredcomplex after phosphomolybdate reacts with the basic dye malachite green. The

3.3 Methods for Studying ATP hydrolysis 31

formation of this complex can be followed by measuring the absorbance at 600nm to 660 nm with a spectrophotometer. This method has been employed regu-larly for the detection of alkaline phosphatase activities. [128,129] Although theseassays are simpler, easy to perform, and cost-effective, but the major drawbackis that only static endpoint measurements can be performed with them. There-fore, they are less sensitive, not suitable for quantitative studies, and cannot beperformed in living cells.

3.3.2 Radioactivity Based Assays

Radioactivity based assays have been used for decades to measure the activityof ATPases. These are highly sensitive assays. However, the safety issues as-sociated with these assays make them least favorites. In addition, these assayscannot be used for continuous detection and need the separation of the radioac-tive substrate from the radioactive end product to measure the ATPase activity.Various approaches have been devised in different studies to achieve this goal.

As an example, the radioactive assay has been used to study rep AT-Pase, a helicase involved in the E.coli DNA replication. This assay uses theγ-32P ATP. On hydrolysis, 32P is released and the two are separated by thinlayer chromatography (TLC) and radioactivity of different bands is measuredwhich gives an estimate of ATPase activity. [130] A γ-32PATP assay has also beenrecently used to measure the activity of protein kinase C. The assay is based onthe measurement of the amount of radioactivity incorporated into the substrateover time. [131,132] In a similar assay, radioactively labeled α-32P ATP is usedto monitor the hydrolysis activity of Cas-3 protein, an ATP dependent single-strand DNA nuclease. Here the ATP and ADP in the reaction mixture wasseparated by TLC on a polyethyleneimine-cellulose plate and the radioactivityis measured. [133]

3.3.3 Enzyme Coupled Assyas

These are the spectrophotometric assays where the rate of ATP hydrolysis isindirectly estimated by measuring the enzymatic activity of another enzyme.A widely used method is based on the coupling of nicotinamide adenine din-


ucleotide hydrogen/ nicotinamide adenine dinucleotide (NADH/NAD+) redoxreaction to the ATP hydrolysis process. With each ATP hydrolyzed, pyruvatekinase (PK) converts one molecule of phosphoenolpyruvate (PEP) to pyruvateand the process leads to the formation of ATP from ADP. The released pyru-vate is converted to lactate by the enzyme lactate dehydrogenase (LDH). In thisreaction, NADH is oxidized to NAD+. NADH shows the maximum absorptionat 340 nm but its oxidation product NAD+ does not absorb at this wavelength.Hence the decrease in absorbance at 340 nm gives an estimate of ATP hy-drolysis. [134] It is evident that this assay is not appropriate for pyrophosphateproducing reactions. Another similar method relies on the coupling of ATPhydrolysis to the nucleoside phosphorylase reaction using 7-methylguanosine asfluorescent substrate. 7-methylguanosine is fluorescent and is converted to 7-methylguanine, which is not fluorescent, thus the fluorescent readout is used tomonitor the progress of the reaction. [135]

Methods of this type allow continuous measurements of ATP hydrolysisactivity. Since these assays involve many other enzymes other than the ones tobe studied, the effect of other enzymes on the detection has to be taken intoconsideration. So, they are not preferably used in living cells.

3.3.4 Protein Based ATP Sensors

Recently some fluorescence protein sensors have been developed that can begenetically encoded into the cells to monitor ATP and activities associated withATP hydrolysis. They are convenient to use with no extra cell preparation orincorporation of any reagent and can be employed to monitor the activity inliving specimens with high sensitivity and resolution.

An eminent example is the FRET-based indicator ATeam (Adenosine 5-Triphosphate indicator based on Epsilon subunit for Analytical Measurements).It is a fusion protein developed by Imamura et al. and consists of variants of cyanfluorescent protein (CFP) and yellow fluorescent protein (YFP). Upon bindingto ATP, a conformational change in the sensor leads to FRET which is used toquantify ATP in cells. ATeam sensor has been used to study the changes in ATP

3.4 Modified ATP Analogs 33

levels in living cell under different conditions. [136] The same method was used tostudy ATP metabolism in podocytes. [137] ATeam was also used to image extra-cellular ATP dynamics in real-time in Neuro2A cells (a mouse neuroblastomacell line) [138] Depoli and coworkers have used fluorescent ATP biosensors basedon ATeams to evaluate the metabolic state at single cell level. The study hasfocused on the visualization of the dynamics of ATP levels and the characteri-zation of metabolic activities in mitochondria mainly in cancer cells. [139] ATPdependent conformational change of K+ channel has been visualized in Humanembryonic kidney (HEK) cells by using a cyan and yellow fluorescent fusionprotein as a FRET pair. [140] Lobas and coworkers have developed a geneticallyencoded fluorescent sensor named as iATP neurotransmitter sensing fluorescentreporters (iATPSnFRs). The sensor has been used for imaging and quantifyingthe ATP in specific cellular organelles. [141]

3.3.5 Assays Based on Fluorescent ATP Analogs

Finally, ATP hydrolysis sensors based on the consumption of artificially synthe-sized fluorescent ATP analogs by cellular enzymes. This activity can be easilymonitored by tracking the fluorescence properties of the analog. Based on thisprinciple, a variety of ATP analogs have been developed. Some of these analogsexploit the FRET between two dyes that form a FRET pair attached at two sitesof the analog. [142] The chemical modification of the analog generally changes thechemical properties of the analog including the binding kinetics to its enzyme’sactive site. Thus, great care needs to be taken while the development of sucha molecule along with the validation of the assays. Additionally, these analogsare often cell membrane-impermeable and thus special techniques like electro-poration, lipid transfection, or microinjection are necessary to incorporate theminto the cells. The procedure is often physiologically detrimental to cells. Theapproach of ATP detection based on modified analogs is elaborated below.

3.4 Modified ATP Analogs

Numerous ATP analogs have been designed and synthesized until today to ad-dress various questions related to the ATP hydrolysis activity in the cell. One


approach is to modify the phosphate chain of ATP to change the rate at whichthe analog would be hydrolyzed. In these reactions, ATP is hydrolyzed betweenβ and γ phosphate groups. Thus slowly hydrolyzing or even non-hydrolyzingATP analogs can be produced. Non-hydrolyzing analogs have been used in thecrystallographic studies of proteins as these analogs form stable complexes afterbinding to the active sites of enzymes. [143,144] Based on the fact that ATP actsas the phosphate group donor for kinases, ATP analogs have been synthesizedand used to study different kinases. e.g. ATP γ-S is hydrolyzed at a muchslower rate as compared to ATP. So, the proteins that are thiophosphorylatedcannot undergo rapid dephosphorylation and form stable complexes which canbe analyzed subsequently. [107]

As pointed out before, fluorescence spectroscopy based methods are im-portant tools for biological studies.Thus many fluorescent ATP analogs havealso been synthesized with diverse properties and used for monitoring of theATP hydrolysis in the living cells. However, these synthesized analogs are oftennot taken up by the cells and thus need to be incorporated into the cells usingdifferent invasive methods. Also, the stability of these analogs in the cellularenvironment and their acceptance by the native enzymes is hard to predict andneeds to be evaluated properly. In general, ATP can be fluorescently modifiedby conjugating fluorophores to its ribose, the adenine base or the phosphatechain. Some of the ATP analogs that have been developed with fluorophorederivatives at ribose moiety are 2’(3’)-O-(2,4,6-trinitrophenyl)-ATP [145] 2’(3’)-O-(N-methylanthraniloyl)-ATP (MANT-ATP). [146]

N-methylanthraniloyl derivatives of ATP, e.g. MANT-ATP have beenwidely used for studying various kinases, myosin ATPase, and sodium-potassiumATPases. These analogs are intrinsically fluorescent and exhibit maximum flu-orescence emission at 440 nm and show a considerable substrate specificitywith a broad range of ATP hydrolyzing enzymes, thus allowing the direct mea-surement of ATPase activity. [147] A blue emitting pyrene fluorophore has beenlinked to ribose moiety of ATP coupled with a butyryl linker to synthesizehighly sensitive fluorescent probe for studying myosin ATPase. This analog has

3.4 Modified ATP Analogs 35

proved helpful in monitoring the binding kinetics and ATPase activity of myosinmolecules. [148] Two fluorescent analogs developed by coupling the Cy3 and Cy5derivatives with 2’(3’)-O-[N-(2-aminoethyl)carbamoyl]ATP/ADP (EDA-ATP)are Cy3-EDA-/ADP and Cy5-EDA-ATP/ADP. [149] The 2’- and 3’-O-isomershave been separated and used for single molecule studies of actin-myosin inter-actions. These analogs have been used in combination with TIRF fluorescencemicroscopy for studying the kinetics of motility in myosin filaments in vitro. [150]

An interesting approach to modify ATP for studying the hydrolysis ac-tivity is the labeling of ATP analogs with two dyes that are suitable to undergoFRET. These analogs exhibit a change in their fluorescence properties thusmaking them ideal candidates for the monitoring of ATP hydrolysis activity ofenzymes.

Modified ATP with Sulfo-Cy3 and Sulfo-Cy5 dyes attached to γ andC2 or O3’ positions were synthesized. The probe undergoes FRET in an in-tact molecule because of the close proximity of the FRET pair. FRET is thenterminated with the hydrolysis of the molecule. This change in the FRET ismeasured to estimate the progress of the reaction. The analog has been success-fully tested for the phosphodiesterase - I from Crotalus adamanteus commonlyknown as SVPD. [142,151] A doubly labeled ATP analog modified at N6 positionand at the γ position of phosphoanhydride chain with two fluorophores thatwork as a FRET pair have been synthesized as a FRET sensor. The analog, re-ferred to as time-resolved ATPase sensor (TRASE), was successfully employedto monitor the activity of UBA1, a constituent of the ubiquitin proteasomepathway in real-time. [152]

Another modified analog of diadenosine triphosphate (Ap3A) was de-veloped as FRET sensor. Ap3A is an exceptional nucleotide that plays a rolein cell signaling and is associated with tumor suppression. It is the substrateof enzyme Fhit hydrolase. [153,154] The developed analog was shown to be wellaccepted by Fhit enzyme and was used to study the enzymatic activity of tu-mor suppressor Fhit enzyme in vitro. [155] A modified analog of diadenosine


tetraphosphate (Ap4A) has also been developed as a FRET sensor. Ap4A is aspecial nucleotide that in involved in the gene expression regulation pathwaysduring stress in many organisms. Nudix hydrolases (NUdT2) enzyme regulatesthe level of Ap4A. [156,157] The analog has been stably used to study the activ-ity of NUdT2, a human diadenosine tetraphosphate hydrolase in living cells byfluorescence microscopy. [158]

Chapter 4

Protein PARylation in DNA

Damage Response

4.1 Protein Modifications and Interactions

The human proteome is more complex then the human genome. The estimatednumber of genes in the human genome is approximately 25,000 but on the otherhand, the number of proteins is estimated to be over a million. [159] This largediscrepancy between the two numbers is explained by the fact that one gene en-codes more than one protein. Some of the mechanisms that explain this anomalyare differential transcriptional regulation, alternative splicing, genetic recombi-nation, and post-translational modifications (PTMs). [160] PTMs increase thecomplexity of the proteome by introducing more functional diversity. PTMsare the chemical modification of the fully synthesized polypeptide chain by thecovalent addition of various functional groups or by its proteolytic cleavage.Proteolytic cleavage is the removal of a segment from the inactive form of thepolypeptide to form an active protein. PTMs are commonly responsible for theregulation of activity, localization, and the interaction of proteins in the cell.Some of the protein PTMs are briefly discussed below:

1. Phosphorylation is the addition of a phosphate group to the proteins ontheir serine, threonine, or tyrosine residues by kinases. It is the most

38 Protein PARylation in DNA Damage Response

crucial PTM for the existence of any organism and is widely involved incellular signal transduction and cell cycle regulation. [161]

2. Acetylation is the transfer of the acetyl group from acetyl-CoA to theamine group of lysine by the enzyme N-acetyltransferase. Acetylation canbe reversed by deacetylase enzymes. Acetylation at N-termini is very com-mon in eukaryotes and plays a role in gene expression regulation e.g. bythe condensation and decondensation of histones. [162]

3. Methylation is the transfer of one methyl group to oxygen or nitrogenknown as O-methylation and N-methylation respectively. Methylation ismediated by methyltransferases where S-adenosyl methionine (SAM) isthe donor. Methylation at multiple sites on histones affects the processof transcription by modulating the availability of DNA and thus it is theprimary mechanism of epigenetic gene regulation. [163]

4. Ubiquitination is the addition of the polypeptide ubiquitin to the aminegroup of lysine of the protein. Ubiquitin consists of 76 amino acids andattaches to the target protein by its glycine residue of the C-terminal.Ubiquitin tags are identified by the cellular proteasome system that sig-nals the degradation of the targeted protein. [164]

5. Glycosylation is the attachment of sugar moiety to the protein and basedon the nature of the linkage, it can be N- or O- glycosylation. In N-glycosylation, the sugar moiety is covalently bound to the nitrogen on theasparagine residue. O-glycosylation occurs on the hydroxyl group of serineor threonine residues of the proteins. Glycosylation takes place usually inmembrane proteins and plays a role in cell-cell interaction, signal trans-duction, immune response, endocytosis. [165]

6. Lipidation is the process of addition of a lipid moiety to proteins, particu-larly the membrane proteins. Based on the lipid moiety that is appended

4.1 Protein Modifications and Interactions 39

to the protein, lipidation can be C-terminal glycosyl-phosphatidylinositol(GPI) anchor, N-terminal myristoylation, S-myristoylation, and S-prenylation.Each of the lipid groups confers different affinities to the proteins andthus localizes proteins to different membranes i.e. the plasma membrane,vesicular membranes (lysosomes, endosomes), mitochondrial membrane,or endoplasmic reticulum. [166]

Proteins with PTMs function prevalently by modulating their interac-tions with other cellular components including other proteins. These PTMs cre-ate a vast array of specific interaction domains that act as binding sites for differ-ent binding partners. For example, transcription factors and signal-transducingmolecules with phosphotyrosine-binding domain binds to the activated receptortyrosine kinase that signals the activation of many diverse pathways. [167] His-tone PTMs such as methylation and acetylation regulate chromatin dynamicsto facilitate differential gene expression. The modified residues are recognizedby protein modules like chromodomains, which interact with methylated lysineand bromodomains, which further interact with acetylated lysine. These con-served domains act as recruiters for various transcription factors. [168] In thiswork, experiments on the influence of the PTMs on DNA damage response willbe reported. DNA damage response is a complex process that involves multipleprotein factors. PTMs play a dynamic role in DNA damage response by thetransformation of the protein properties already synthesized to respond instan-taneously in case of the damage. Modified proteins often act as a scaffold forthe interactions of various factors involved. Some of the prominent PTMs in-volved in DNA damage response are acetylation, methylation, phosphorylation,ubiquitination, and SUMOylation.

4.1.1 Protein Poly-ADP-ribosylation

Poly-ADP-ribosylation (PARylation) is another PTM with an important influ-ence on DNA damage response. It consists in the addition of adenosine diphos-phate (ADP)-ribose moiety to a protein. The transfer of ADP-ribose is catalyzedby the enzyme poly-adenosine diphosphate-ribose polymerase (PARP) fromNicotinamide adenine dinucleotide (NAD+) to a variety of amino acids of the


target protein. ADP ribosylation can be classified as mono (ADP-ribosyl)ation(MARylation), oligo(ADP-ribosyl)ation, or poly(ADP-ribosyl)ation (PARyla-tion) depending on the number of ADP-ribose units added to the target pro-tein. [169,170] There are 17 known members of the PARP family in mammalsand each enzyme has a defined role. PARP1 is the most prominent enzyme ofthis class and is responsible for most of the PARP activity. [171,172] PARP1 isknown to PARylate itself as a primary target. This phenomenon is known asautoparylation. PARylation is reversible and ADP-ribose units can be removedby the enzyme Poly(ADP-ribose) glycohydrolase (PARG). PARG cleaves thePAR chains to release free ADP ribose units. [173] PARylation is a primary stepin DNA damage response as it initiates the response by acting as a scaffold formultiple repair factors.

Some other functions attributed to protein PARylation are its role intranscription, replication, apoptosis, chromatin remodeling and metabolism.PARPs can induce apoptosis by the production of PAR that stimulates therelease of mitochondrial apoptosis-inducing factor. [174] PARPs also promotenecrosis by causing the depletion of ATP levels in cells. ATP is depleted byPARP activation in an attempt to repair the damaged DNA. [175] The antiviralactivity of PARP-13 by inhibiting the production retroviral RNA has also beendiscovered. PARP-1 has been implicated in the immune response by modula-tion of NF-κB activity [176] and in chromatin remodeling and gene expression.Histones as well as non-histones get PARylated to influence the nucleosomestructure and chromatin binding. [177] PARP-1 alters the functions of transcrip-tional regulatory machinery to modulate the transcriptional activity and generegulation. [178,179]

4.2 DNA Damage and Repair

DNA damage is a fundamental cellular problem. It consists in the change in thebasic chemical or physical structure of the DNA. A damaged DNA can resultin impairment of essential cellular processes like replication and transcriptionthat can eventually perturb the normal functionality of cells. It may arise by

4.2 DNA Damage and Repair 41

endogenous metabolic processes in the cell or due to exogenous or environmen-tal factors. If a damaged DNA is not repaired, it could result in mutations. Ineukaryotes, DNA is protected inside the nucleus and other organelles like mito-chondria and chloroplasts because of the importance of DNA for the functioningof the cell. Cells have evolved with a comprehensive response mechanism to notonly protect the DNA but also to correct the damage by various repair mech-anisms. Both exogenous as well as endogenous DNA damaging factors exerttheir genotoxic effects by the same mechanisms. There are many different typesof DNA damages, some of which are comprised below.

4.2.1 Types and Causes of DNA Damage

1. Oxidative DNA damage is mainly caused by free reactive oxygen species(ROS) like superoxide radicals ( O –

2 ), hydrogen peroxide (H2O2) and thehydroxyl radical ( OH–) that are e.g. generated as bi-products of oxidativerespiration in mitochondria. Normally, the cells have an indigenous mech-anism of protection from ROS by separating the oxidative process spatiallyin mitochondria and antioxidant enzymes like catalase and superoxide dis-mutase. [180] The hydroxyl ( OH–) radical, an electrophile generated as abyproduct of Fenton reaction, attacks double bonds of bases and sugarsto form products that result in erroneous pairing and mutations. e.g.hydroxylation of thymine forms thymine glycol and hydroxylation of C-8guanine forms 8-oxo-Gaunine. [181] Oxidation of lipids by hydroxyl-radicalsgenerates aldehydes like 4-hydroxynonenal that react with bases to formharmful mutagenic adducts. [182]

2. Alkylation of DNA refers to the addition of alkyl group to specific basesfrom an alkylating agent. It results in the formation of products like O2

and O6 alkylthymine, O6 methyl guanine and O6-ethylguanine. Methyla-tion is the most common form of alkylation of the nitrogenous DNA basesthat is carried exogenously by agents like betaine, choline, and certainchemicals presumably found in environmental pollutants and food. [183]

Oxygen methylated DNA bases like O6-methylguanine and O4-methylamine


are highly harmful as they produce detrimental nucleotide transition mu-tations. DNA alkylation is involved in cancer formation e.g. colon andcolorectal carcinoma.

3. Dimerization is the formation of dimers primarily between two pyrimidinebases. It is mostly caused by high energy electromagnetic radiation usuallyin the UV range. Depending on the wavelength and dose of radiation, twodifferent types of dimers are produced, pyrimidine 6-4 photoproducts (6-4PPs) and cyclobutane pyrimidine dimers (CPD). [184] Translesion poly-merases introduce base transitions during the replication on encounteringthese bulky dimers, thus contributing to the mutagenesis. [185]

4. Bulky adducts are formed by chemical crosslinking of various componentsof DNA. Various compounds including polycyclic aromatic hydrocarbons(PAH) and benzopyrene diol epoxide which are very potent adduct form-ing agents found in environmental pollutants like smoke. Also, chemicalslike trioxsalen, formaldehyde, mitomycin C, and cisplatin (crosslinkingagents) and, ethidium bromide, (intercalating agents) contribute to theformation of adducts in DNA. These compounds are commonly found inindustrial solvents, disinfectants, cleaning solutions, and petroleum deriva-tives. Bulky adducts of this type alter the physical structure of DNA, getinto the way of the translation and transcription machinery, and stall theDNA polymerases. [186] This may eventually lead to replication arrest andcell death.

5. DNA strand breaks (DSBs) can occur by both endogenous and by exoge-nous factors. They occur by the disruption of the phosphodiester bondbetween the adjacent deoxyribose residues. Single strand breaks (SSBs)are more common and occur as byproducts of the normal DNA manip-ulation processes like replication, transcription, and relaxation of DNAsupercoils. [187] SSBs can also occur as a result of oxidative stress and dur-ing the repair of abasic sites as these sites are readily converted into SSBsby the base excision repair (BER) pathway to repair them. A mixture of

4.2 DNA Damage and Repair 43

SSBs and DSBs among other lesions are caused by ionizing gamma andultraviolet (UV) radiations. [188] Doublestrand breaks are less common andare formed when two damage sites on both the DNA strands are in closeproximity. DSBs are also induced directly by ionizing radiation. DNAbreaks induced by radiation will be discussed in detail.

4.2.2 Mechanisms of DNA Repair

For the perpetual safeguard of the genome integrity and stability, a complexand sophisticated network of DNA damage repair mechanisms has evolved incells. A multitude of mechanisms is involved in the repair of different types ofDNA damages. Failure of any repair mechanism is implicated in a variety ofgenetic disorders, carcinogenesis and aging. Some of the DNA damage repairmechanisms are briefly described below:

1. Base excision repair (BER) occurs when the individual bases have under-gone chemical alterations like oxidation. The damaged base is recognizedby DNA glycosylase which mediates its removal with the help of endonu-cleases resulting in the formation of an abasic site (apurinic/apyrimidinicsite). This site is filled by the action of polymerase-β that adds one nu-cleotide and then ligases seal the nick to complete the repair. [189]

2. Nucleotide excision repair (NER) on the other hand repairs the bulkyadducts and lesions like pyrimidine dimers and other photoproducts thatdistort the physical structure of DNA. NER involves multiple steps andmore than 20 enzymes. NER can operate through two distinct pathway:Global genomic NER (GG-NER) that removes lesions anywhere in thegenome, even the regions that are not transcriptionally active and thetranscriptionally coupled NER (TC-NER), that repairs lesions in tran-scriptionally active genes to resume the stalled transcription. In GG-NER, the damage is recognized by the xeroderma pigmentosum com-plementation group C / UV Excision Repair Protein RAD23 HomologB (XPC/hHR23B) protein complex while in TC-NER, it is RNA poly-merase that senses the damage with the help of Cockayne syndrome fac-


tors A and B (CSA and CSB). [190,191] The subsequent steps of both thepathways converge in which the helicases unwind the DNA duplex and en-donucleases XPG (Xeroderma Pigmentosum Complementation Group G)and ERCC1-XPF (Excision Repair Cross-Complementing Rodent RepairDeficiency, Complementation Group 1- Xeroderma Pigmentosum GroupF-Complementing Protein) cleave the DNA generating around 30 oligonu-cleotide lesion. The gap is repaired by DNA polymerases and ligases tocomplete the repair. [192]

3. The mismatch repair (MMR) pathway is essential for the correction ofbase-base mismatches, insertions and deletions loops (IDL) that arise dur-ing the replication from polymerases. In MMR, mismatch, insertion, ordeletion is recognized by MutS. This initiates the recruitment of otherreplication factors to form MutH-MutS-MutL ternary complex. Then ex-onuclease 1, DNA polymerase-δ in association with additional factors likeproliferative cell nuclear antigen (PCNA) and replication protein A (RPA)resynthesize the new strand and excise the nascent strand. [193]

4. Double strand break repair (DSB) is the most serious and difficult repairmechanism in the cell because unlike other repair mechanisms, the com-plementary DNA strand is not available for the faithful replication of thedamaged strand since both the strands are damaged. Cells have evolvedtwo distinct types of DSBR pathways: homologous recombination (HR)pathway and non-homologous end-joining (NHEJ) pathway.

The HR pathway requires a homologous sister chromosome to retrieve theinformation and thus repairs DSBs in an error-free manner. It is initiatedby formation of an MRN complex consisting of RAD50 (ATP-binding cas-sette–ATPase), NBS1 (phosphopeptide-binding Nijmegen breakage syn-drome protein 1) and MRE11 (meiotic recombination 11 homolog). Thedamaged ends are processed to produce overhangs. RAD51 nucleopro-tein screens for the homologous strand on the sister chromatid. A DNAduplex is formed from the damaged and undamaged strands. Then the

4.3 PARylation in DNA Damage Response 45

DNA polymerase forms the new strand resulting in the formation of atemporary heteroduplex structure. This recombinant intermediate is re-solved to yield two separate double strands. [194] NHEJ repair on the otherhand is an error-prone mechanism since it does not utilize a homologoustemplate for repair. In this mechanism, the breaks are recognized byKu70/Ku80 heterodimeric protein to initiate the process. This is followedby the recruitment of DNA-dependent protein kinase (DNA-PK), X-rayrepair cross-complementing factor (XRCC4) and DNA Ligase IV. Sub-sequently, MRN protein complex (MRE11/Rad50/NBS1) and nucleasesincluding artemis are recruited. Finally, DNA ligase IV is involved inligation to complete the process. [195]

4.3 PARylation in DNA Damage Response

4.3.1 PARP1: Structure and Function

The poly(ADP-ribose) Polymerase (PARP) superfamily consists of 17 enzymesthat play a critical role in various cellular homeostasis processes including tran-scriptional regulation, cell cycle regulation, cell signaling, and DNA repair. [171]

PARP1, also known as ARDT1 (ADP-ribosyltransferase diphtheria toxin-like1), is the most prominent and best-studied member of the PARP family. It isresponsible for synthesis of the majority of PAR in cells. PARP1 essentiallycatalyzes the covalent attachment of ADP-ribose subunits via an ester bondfrom NAD onto acidic residues (aspartate or glutamate) of target proteins toform a poly(ADP-ribose) polymer. PARP 1 has a molecular weight of 116kilodaltons (kDa) and consists of multiple independently folded domains con-nected by free linkers. The three major functional domains are: Amino-terminalDNA binding domain (DBD), a central auto-modification domain (AMD) andthe carboxyl-terminal catalytic domain (CAT) (Figure 4.1). The DNA bindingdomain consists of two Cys-Cys-His-Cys (CCHC) zinc finger motifs (ZF1 andZF2) that bind to DNA and modulate DNA dependent activities and a bipar-tite nuclear localization signal (NLS). The AMD domain consists in a BRCA1(Breast cancer type 1 susceptibility protein) carboxy-terminal domain (BRCT)which mediates the interaction of PARP1 with its interacting partners during


DNA repair. [196] The catalytic domain (CD) at the carboxyl-terminal is themost conserved domain and it contains WGR motif consisting of three aminoacids, [tryptophan (W), glycine (G), and arginine (R)] that mediates PARP1activation, an alpha helical PARP regulatory domain (PRD) and catalytic do-main which binds to NAD. [197]

Figure 4.1: Domain architecture of Poly(ADP-ribose)polymerase-1

(ARTD1). [198]

Most of the PARylating enzymes can transfer only a single ADP-ribose tothe target protein, a process called mono(ADP-ribose)ylation (MAR). PARP1,2and 3 are involved in the PARylation in DNA damage response. [169] PARP1 isknown for PARylating itself and during this autoPARylation, the ADP-ribosepolymer can grow up to 200 units. Moreover, these PAR molecules can adopthighly branched patterns. PARP1 has a wide range of interacting partnersthat are the constituents of DNA repair complexes and many of them getPARylated. Other than DNA repair, it plays major roles in some other cel-lular functions. PARylation of the target protein contributes to the changesin its properties mainly through the charge effect. This results in the modula-tion of protein-protein interaction, protein-DNA interactions and the enzymaticactivity of target proteins. [197] PARP inhibitors have been widely used as ther-apeutic agents for human diseases including cancer, e.g. olaparib. Their modeof action depends on the obstruction of DNA repair capability to promote celldeath. PARP1 regulates the transcription by modulating the chromatin struc-ture. Auto-modification of PARP1 imparts a negative charge that helps in theloosening of chromatin which in turn increases the transcriptional activity. [177]

PARP1 has also been shown to PARylate the histones H1 and H2B, again loos-ening the chromatin and giving access to the transcriptional machinery.

4.3 PARylation in DNA Damage Response 47

4.3.2 Role of PARP1 in DNA Repair

The most prominent role of PARylation is in DNA damage response. 17 mem-bers of PARP family have been identified until now and out of them, the roleof three members (PARPs 1,2 and 3) has been recognized in DNA damage re-sponse. Among these, PARP1 is predominantly involved. PARP1 has beenshown to initiate and modulate various types of DNA repair pathways. PARP1interacts with XRCC1, a break repair factor in case of single strand break re-pair (SSBR) and base excision repair (BER). This promotes the assembly ofthe other repair factors needed. [199] PAR chains act as scaffolds to recruit andassemble the DNA repair complexes. PARP1 also helps in strand break re-pair and base excision repair by interacting with 8-oxoguanine glycosylase 1, aDNA glycosylase. Some of the prominent DNA repairing factors that interactwith PARP1 are DNA ligase III, PCNA, DNA polymerase-β (DNAP). The in-teraction of PARP1 with DNA damage-binding protein 2 (DDB2) and ALC1(Chromodomain-helicase-DNA-binding protein 1-like) regulate the NER path-way. UV lesions are recognized by DDB2 and PARP1 is recruited, leading tothe chromatin remodeling by ALC1. [200]

PARP1 acts as a sensor to recognize SSBs and DSBs and the bindingof PARP1 to the damaged DNA significantly activates its enzymatic activity.HR is activated by single strand DNA resection and PARP1 is involved in thisrepair pathway. [201] It also plays an important role in SSB repair by facilitatingthe recruitment of some HR repair factors to the damage site. Many studiesindicate the role of PARP1 in both the classical and alternate pathways ofNHEJ recombination in case of double strand break repair (DSBR). PARP1stimulates the classical non-homologous end joining (cNHEJ) by interacting withthe DNA-dependent protein kinase (DNA-PKcs). [202] A complex of PARP1,DNA-PKcs, and Ku70-Ku80 dimer promotes the spatial rearrangement of thesynapsis to undergo recombination. The role of PARP1 has been implicated inthe recruitment of multiple protein factors for the alternative NHEJ (aNHEJ)pathway as well. [203] Chromatin associated PARP1 has been shown to recruitALC1 protein, a well-known chromatin remodeler, to the DNA damage site.ACL1 is a member of SNF2 ATPase superfamily (chromatin remodeler) that


mediates DNA repair mechanism by regulating chromatin structure. [204]

4.4 Methods for Studying DNADamage Response

Many diverse strategies are being used to investigate DNA damage and repaireach with their advantages and disadvantages. In response to the damage inDNA, various repair proteins are accumulated and these proteins act as impor-tant biomarkers for the evaluation of DNA damage. Some of the assays basedon various strategies are discussed here.

4.4.1 PCR Based Methods

Polymerase chain reaction (PCR) is a well established technique used for invito analysis and quantification of DNA. It is used to detect the breaks andlesions of the DNA in combination with agarose gel electrophoresis. The TaqDNA polymerase is blocked at the site of DNA damage resulting in the decreaseof the quantity of PCR product formed. The PCR product is visualized afterrunning on the agarose gel that gives an estimation of the damaged template.This principle has been employed to detect and quantify the DNA damageusing quantitative PCR in many different studies. [205,206] Certain variants ofPCR have been developed specifically for detection of the DNA damage, e.g. astraightforward technique called ligation mediated PCR (LMPCR) can be usedfor the analysis of photoproducts developed after UV treatment of the DNA.Another technique that combines the PCR and antibody detection is calledimmune-coupled PCR (ICPCR). This technique can detect the primer-dimersforming in cells upon UV-radiation. [207] These techniques can only be used invitro or after the DNA extraction from cells. Live cell monitoring of the damageor repair process is not possible.

4.4.2 Molecular Marker Based Method

The phosphorylated form of the histone variant H2A family member X (H2AX)known as γH2AX is a widely used marker for DNA damage detection. The in-creased levels of phosphorylation indicate the appearance of DNA strand breaks.They are detected by immunofluorescence after using specific fluorescent anti-

4.4 Methods for Studying DNA Damage Response 49

body against serine-139 of gamma H2AX. [208] The DNA strand break damagecan be visualized after fixing the cells. Since the XRCC1 protein also plays a rolein many pathways for the repair of strand breaks in mammals, the expression ofXRCC1 has also been implicated for the estimation of different aspects of DNAbreaks and repair. [209] Some other repair factors that are being used as markersin immunofluorescence and western blotting are BRCA1, KU70, RAD52, PCNAand CDC 25A.

4.4.3 Fluorescence Based Methods

A simple and efficient method for evaluating DNA damage in cells is the cometassay. It is based on the gel electrophoresis of a single cell where the cleavedDNA fragments migrate out of the cell by applying an electric potential andthe undamaged DNA remains within the cell. This results in the shape of acomet after staining the gel. Other than detecting the single strand and doublestrand breaks, the method is also used to detect pyrimidine dimers, oxidativeand alkylation damage with the aid of lesion specific enzyme that removes thespecific damaged bases. [210] Terminal deoxynucleotidyl transferase (TdT) dUTPnick-end labeling (TUNEL) assay is an established fluorescence based assay usedfor the visualization of DNA fragment on DNA damage and apoptosis. In thisassay, the enzyme terminal deoxynucleotidyl transferase (TdT) is used to cat-alyze the incorporation of deoxynucleotides tagged with the fluorophores ontothe 3’-hydroxyl end of DNA fragments. The damaged DNA can be visualizedby fluorescence detection. [211]

Fluorescence in situ hybridization (FISH) is used for in situ labeling ofaberrations in DNA like SSBs, DSBs, and single nucleotide changes. In this tech-nique, fluorescent nucleotide probes are designed that target specific genomicsequences. The hybridization of these probes can be conveniently visualized incells. It offers a better resolution as compared to fluorescent antibody basedmethods and is applicable for quantitative live-cell imaging. [212] This methodis widely used in clinical diagnosis with applications in the detection of geneticalterations associated with cancer. It can be performed in fixed cells and hasan inherent disadvantage of higher background fluorescence that compromises


its sensitivity. All these methods have immensely contributed to the knowledgeof the DNA damage response. However, monitoring of damage response in liv-ing cells is impossible with these methods. Monitoring the damage response inreal-time would help in ascertaining the dynamic interplay of the repair factorsinvolved in the process.

4.5 Modified NAD Analogs

To understand the molecular mechanism of PARylation and its role in regu-lation of biological functions like DNA repair, modified NAD molecules havebeen developed. These analogs molecules have resulted in the development ofefficient methods to gain valuable insights into the PARylation and its relatedprocesses. Most of these methods rely on analysis of the products using gelelectrophoresis or visualization by fluorescence microscopy. A fluorescent NADanalog, nicotinamide 1,N6-ethenoadenine dinucleotide has been synthesized byJorge and coworkers. The analog has an emission wavelength of 410 nm andits fluorescence intensity increases 10 fold upon its hydrolysis by phosphodi-esterase I. [213] Jiang and coworkers have used an NAD analog to fluorescentlylabel and visualize PARP-1 and tankyrase-1 in gel. They have identified vari-ous potential PARP-1 interacting proteins by using a biotin affinity tag. [214] Inanother strategy, non-reacting NAD analog has been used for elucidation of theX-ray crystallography structures of the NAD-utilizing enzyme complexes. Theapproach was used to study the X-ray structure of SIRT3 and SIRT5 (a mem-ber of the mammalian sirtuin family of proteins). [215] The same carba-NAD hasbeen used by Langelier and coworkers to elucidate the binding and occupancyof an NAD analog to PARP1 during DNA damage response by crystallographystudies. [216] Various NAD analogs have been developed for the visualization ofribosyl transferase activity both in vitro and in vivo. Wyang and coworkershave synthesized an ATP analog by introducing an alkyl group that leads toearly chain termination while being incorporated by ADP-ribosyl transferase.It was used to study the activity of ADP-ribosyl transferase and its substratebinding. [217] An NAD analog called Bio-NAD (6-biotin-17-nicotinamide-ade-nine-dinucleotide) has been used for the isolation of ribosylated proteins and forthe quantification of PARP activity in cells. [218,219] An orthogonal NAD ana-

4.6 Fluorescence Microscopy for Studying Damage Response 51

log has been used in combination with genetically engineered members of theARTD family to label and identify specific target proteins. [220] Recently manymodified NAD analogs have been employed to visualize PARylation in livingcells using FLIM microscopy. [221,222]

4.6 Fluorescence Microscopy for Studying Dam-

age Response

Microscopy has emerged as a robust technique for the visualization of cellularstructure, biomolecular interactions and biomolecular localization. With theadvancement in the field of fluorescence microscopy in the last few decades,researchers have been able to dissect complex cellular processes on cellular aswell as single molecule levels. DNA repair is an intricate process that involvesthe interplay of various repair factors and their interaction with DNA to formmultifactor complexes. Although DNA repair has been studied using conven-tional biochemical, biophysical and cell biology techniques, the dynamics andthe underlying regulatory mechanisms of different processes are still elusive to alarge extent. It has been challenging to answer such questions by conventionalbiochemical and biophysical techniques. With the emerging powerful fluores-cence microscopy techniques, direct visualization of DNA repair factors andtheir interactions has become possible. This has resulted in the quantitativedetermination of the concentration, diffusion rate, localization, and kinetics ofmolecules in the real-time.

To study DNA damage response, the primary step is to induce DNA dam-age. Various kinds of physical and chemical methods have been used to induceDNA damage depending on the demand of the experiment, the kind of dam-age intended, and the method of visualization. Chemical agents that are beingused to induce DNA damage are methylating agents e.g. ethyl methanesul-fonate (EMS), N-methyl-N‘-nitro-N-nitrosoguanidine (MNNG) methylmethanesulfonate (MMS), cisplatin, bleomycin, camptothecin, nitroquinoline 1-Oxide(4-NQO) and, hydroxyurea. [223] Physical DNA damaging agents are also usedfor the induction of DNA damage. Ionizing radiations are the most common


DNA damage inducing agents that exert their effect directly or indirectly. γ-radiation, X-rays, UV radiation, infrared radiation, β-radiation, and α-particlesare some of the radiation types that have been used to induce DNA damage.

4.6.1 DNA Damage by Laser Microirradiation

The application of a fluorescent microscope for monitoring the DNA damageresponse after inducing DNA damage has emerged as a robust method forstudying the process of DNA damage response. Among various methods bywhich the damage can be induced, chemical agents and radiation damage arethe most prominent ones. The disadvantage of the damage induced by chemi-cal agents that it often is asynchronous and affects the DNA globally in a cell.As an alternative, laser microirradiation with specific wavelengths is a powerfultool for inducing calibrated DNA damage. Various types of damages includingSSBs, DSBs, and base lesions can be induced reliably by using a wide range ofwavelengths. [224] This method has emerged as a favorite for studying the DNAdamage response and has led to the possibility of studying the localization andaccumulation at sub-cellular level.

Electromagnetic radiations over the whole spectrum has been used forinducing DNA damage in various studies. Radiation in the UV region results indamage because of the linear absorption by DNA. UV radiation mostly inducethe formation of photoproducts and base modifications. In 1970s, UV irradia-tion was initially used to study its effect on DNA in cells. Cramer and coworkersused a UV laser beam (257 nm) to establish the formation of UV induced pho-tolesions. They reported the local microirradiation damage of the nucleus inChinese hamster cells. [225,226] Subsequent investigations led to the developmentof protocols for the induction of UV photoproducts, oxidative damage, singleand double strand breaks and lesions. Also, the effect of sensitizing agentslike bromodeoxyuridine (BrdU) and Hoechst dyes with the radiation on DNAdamage was revealed. [227] The effect of UV and visible light on DNA damagewas further calibrated with the advent of sophisticated microscopy and lasersetups. The relation between different wavelengths and the base modificationswas quantitatively established. UV-A in combination with FISH was first used


by Bonner and coworkers to introduce DSBs in human DNA. They identifiedvarious repair factors that accumulate at the damage site. [228]

Lately, multiphoton excitation has been introduced to manipulate a con-fined, three-dimensional volume of approximately 1 µm3 in a single cell. Thiskind of irradiation results in the absorption of multiple photons. [229] Ultrafastlaser micro irradiation is predominantly used because of its ability to reducenon-specific absorption and thus to minimize photo damage of other cellularcomponents. In order to limit the average power deposited in a particular cellvolume, ultra-short laser pulses (femtosecond or picosecond) of high intensityare delivered onto the sample. Moreover, this type of irradiation is convenientfor performing time-resolved imaging. [230] Various potential mechanisms havebeen proposed regarding the damage of DNA using ultra-short high-energy laserpulses. This radiation can create thermal effects by increasing the collisionsbetween free electrons thus catalysing the DNA damage. [231] The high energyultra-short pulses also prompt photoionization events and the formation of plas-mas that result in the generation of single and double strand breaks. [230] Mul-tiple studies have reported the application of near infrared and infrared pulsedlaser for the introduction of double strand breaks in DNA. This is essential asUV-A lasers that were being used for this purpose required a sensitizing agentthat could potentially affect the repair process in cells. Also, sensitizers cancause non-specific damage to other cellular structures like mitochondria. Cellsabsorb the least energy in the IR region thus causing minimum damage becauseof phototoxicity and thermal effects. [232]

It has been demonstrated that irradiation with pulsed NIR lasers (800nm wavelength) in absence of sensitizers can produce DSBs in DNA. The accu-mulation and kinetics of DS repair pathway NHEJ proteins (Ku70/80) at thedamage site was also evaluated after the induction of DS breaks by NIR ra-diations. [233] Near IR femtosecond microirradiations have been used to inducechromosomal alterations and DNA photproducts in order to visualize rapid ac-cumulation of the repair factors. The damage is based on the three-photonabsorption and has been used to visualize the accumulation of EGFP-PCNA in


cells. [232] Schmalz and coworkers have recently demonstrated the utilization ofa wide range of wavelengths to induce various specific types of DNA damagesby using femtosecond laser pulses. [234]

4.6.2 Imaging of DNA Damage Response

Numerous studies have been done to visualize the interplay of the diverse repairfactors involved in the DNA damage response in various organisms from bacte-ria to mammals. Most of the methods are based on the traditional approachesof molecular biology, biochemistry, and genetics. The results of these methodsare based on the response exhibited by a large population of the repair factorsinvolved. Thus often the end point ensemble picture of the whole process isrecorded. With the advent of highly sensitive imaging techniques, it has beenpossible to continously follow the molecular events at single cell level. Fluores-cence microscopy in combination with fluorescent biological markers have madeit possible to monitor multiple molecular processes in the cell. Many of thesemethods have been employed to study DNA damage response. Initially, fluores-cence microscopy was used to visualize the organization of DNA repair complexin fixed cells. The cells were fixed at different time points after the initiation ofDNA damage and fluorescence readout was recorded. [235] While this approachhas led to the understanding of the different factors involved, an insight into thedynamics and the precise sequence of events was not possible. Live cell imaginghas emerged as a powerful technique by which the DNA damage response canbe followed in a spatiotemporal manner after the controlled initiation of DNAdamage. Monitoring the DNA damage response factors in living cells wouldreveal the sequence of molecular events involved.

Various aspects of molecular dynamics that contribute to DNA damageresponse have already been evaluated by different live-cell imaging techniquesincluding studies of the movement of the repair factors and their assemblyat damage site. With the application of robust approaches like fluorescencecorrelation spectroscopy (FCS) and fluorescence recovery after photobleaching(FRAP), enormous information has been gained about the mobility and recruit-ment of repair factors during DNA damage response. FCS is a technique used to


determine the diffusive movement of molecules based on the autocorrelation ofintensity fluctuations. It has been used to follow the complex diffusion processesof DNA repair factors and their binding in cells. [236] FRAP has been used toestimate the dynamics of molecules at a specific cellular location by measuringthe rate of replenishment of fluorescent proteins in a photobleached region. Thismethod has been used to study the recruitment of ERCC1 endonuclease to therepair sites during the NER. [237] The dynamics of two important mismatch re-pair proteins, MutS and MutL, during MMR pathway have also been studiedin E.coli. The complex formation was observed in real-time after fluorescentlabeling of the two proteins. [237] Another aspect of DNA damage response un-raveled by live cell imaging is the dynamics and mobility of chromatin to aidin the accessibility of the repair machinery. The rearrangement of chromatindomains during the DSBR has been studied by using fluorescent-tagged dou-ble strand break marker 53BP1 (tumor suppressor p53-binding protein 1). [238]

The increase in the chromatin mobility facilitates the DNA repair. A localizedchromatin expansion has been visualized to occur at DSBs by using a GFPtagged histone H2B. [239] PARylation that primarily initiates the DNA damageresponse, has also been monitored in real-time at DNA damage sites after mi-croirradiation with a near-infrared laser. [222] Two proteins UvrA and UvrB thatare involved in the NER repair have been studied by using multicolor fluores-cence imaging after labeling the proteins with quantum dots in bacteria. Theinteraction of the protein factors with DNA has been visualized during NER atthe single molecule level. [240]

Advanced super-resolution techniques like photoactivated localization mi-croscopy (PALM) and stochastic optical reconstruction microscopy (STORM)provide the resolution of up to nanometers and offer an advantage to studyDNA damage with comparatively high resolving power. Structured illumina-tion microscopy has been used to elucidate repair mechanism at DSBR site. [241]

These techniques have been immensely useful to monitor the movement of singlemolecules in complex cellular environment with high precision. A critical inves-tigation based on the super-resolution microscopy was the imaging of telomeresto understand the mechanism by which they are self-protected from the DNA


repair machinery. [242] Fluorescence lifetime imaging microscopy (FLIM) basedFöster resonance energy transfer (FRET) has been widely used for visualizingchromatin architecture and its dynamics during DNA repair. [243] Since the life-time of a fluorophore is intrinsically influenced by the microenvironment, e.g.its local concentration, FLIM imaging ideal for visualizing the condensation anddecondensation of chromatin during DNA repair. [244,245]

Chapter 5

ATP analog hydrolysis in

living cells

The importance of ATP hydrolysis and ATP hydrolyzing enzymes in cellularfunctioning is evident. It is important to discern a specific cellular processassociated with ATP hydrolysis and specific hydrolyzing enzymes. Conven-tional biochemical methods do not provide this information as they are mostlyendpoint-based or require the fixation of cells before evaluation. It would bevaluable to monitor ATP hydrolysis in real-time in living cells.

The objective of this project was to visualize the activity of ATP hy-drolysis by using fluorescence microscopy in combination with fluorescent ATPanalogs. The ATP analogs were synthesized in the group of Prof. Dr. AndreasMarx (Department of Chemistry, University of Konstanz). Two nucleotides,adenosine triphosphate (Ap3) and adenosine tetraphosphate (Ap4), were mod-ified with the rhodamine derivative Atto-488 at the terminal phosphate and anon-fluorescent quencher group at the adenosine base at the N6 position. Ap3consists of three phosphates whereas Ap4 consists of four phosphate groups.In an intact molecule, the fluorescence of the dye is quenched due to FRETbecause of the spatial proximity between Atto-488 and quencher. Here, thedye acts as a donor and the quencher eclipse acts as an acceptor. The energytransfer is abolished by the enzymatic cleavage resulting in an increase in the

58 ATP analog hydrolysis in living cells

fluorescence intensity as well as an increase in the fluorescence lifetime. Moni-toring the change in the fluorescence intensity and fluorescence lifetime is usedto quantify the hydrolysis of analogs in our experiments. The change in theintensity was monitored by laser scanning confocal microscopy and the changein the fluorescence lifetime was detected using FLIM microscopy. The princi-ple for the detection of ATP hydrolysis using this analog is depicted in figure 5.1.

Figure 5.1: Schematics of the Ap4 analog and its hydrolysis. Visualization

of Ap4 analog with an increase in its fluorescence intensity and lifetime on

hydrolysis.

The modification on the phosphates as well as on the N6 position aretolerated by the various enzymes in vitro as well as in vivo. [155,246,158,247] Thecleavage of the analog as a substrate was monitored after treating it by snakevenom phosphodiesterase I (SVPD) a phosphodiesterase from C. adamenteus.SVPD is an enzyme known to cleave a broad range of modified nucleotides. [248]

The reaction mixture was analyzed by HPLC to check its acceptance by theenzyme.This experiment was done by Daniel Hammler from the Marx group.Moreover, the increase in the fluorescence lifetime of the compound on its hydrol-ysis was confirmed by time-resolved fluorescence measurement. After treating

5.1 Stability and Enzyme Specificity of Ap4 59

the analogs with SVPD, time-correlated single photon counting (TCSPC) wasused to confirm the cleavage of the analogs. Since SVPD is very tolerant withrespect to modifications of its substrates, the hydrolysis of the analogs was thenconfirmed under more restrictive conditions. To this end, the kinetics of thecleavage of the analogs was monitored in HeLa cell lysates using a frequency-domain wide-field FLIM microscope and an increase in the fluorescence lifetimewas observed over time. [249]

5.1 Stability and Enzyme Specificity of Ap4

Figure 5.2: (a) Stability of Ap4 analog at different pHs (b) Hydrolysis of

Ap4 in lysate at different temperatures. Blank represents the hydrolysis in

reaction buffer. Error bars represent the standard deviation of mean.

Before using the analogs for the observations in living cells, the stabilityof the analogs under various conditions was evaluated. The pH stability of theAp4 analog was evaluated by monitoring the fluorescence intensity at differentpH conditions, i.e. pH 4, pH 7 and, pH 10 over the period of 3 hours at 37 ◦

C. No fluorescence increase was observed with time indicating that the analogwas stable over a wide range of pH ranging from 4 to 10. The absence of anydegradation indicates that the synthesized ATP analogs were suitable for incellulo studies. Analog treated with SVPD was used as a positive control. In


order to make sure that the Ap4 hydrolysis is enzyme specific, the hydrolysisof Ap4 was monitored in the lysate at 4◦ C and at 37◦ C. Hydrolysis wasalso monitored in pure lysis buffer as a control (blank). At 4◦ C, there wasno significant increase in the fluorescent intensity as compared to that at 37◦

C. This is because of the inactivity of the enzymes at the non-physiologicaltemperature (Figure 5.2).

5.2 Ap4 hydrolysis In Vitro

The hydrolysis of Ap4 and Ap3 analogs in the HeLa cell lysates was monitoredby the intensity increase using a plate reader assay for 1 hour at 37◦ C. The as-say was set up by adding the analogs at the concentration of 20 µM to the lysatein a 96-well plate. The protein concentration in the lysate was kept constantat 1 mg/ml. The incubation was started in the pre-warmed plate reader. Theincrease in the intensity of the cleaved analog would give the readout of theirhydrolysis. The observed increase in the hydrolysis of the Ap4 over time washigher than that of Ap3. The hydrolysis activity of Ap4 and Ap3 was comparedin the lysates of several different cell lines. In all types of cellular lysates, theAp4 hydrolysis was seen to be significantly higher than that of Ap3. (Figure 5.3)

It has been shown previously that the analogs having longer chain termi-nal phosphate modifications are more readily accepted by the enzymes, e.g. incase of DNA polymerases and the ubiquitin-activating enzyme UBA1. [250,251]

This is believed to be due to the longer phosphate chain making the analogmore flexible to fit within the binding pocket of the enzymes by keeping themodification away from the binding site. On the other hand, the small chainof three phosphates makes the analog comparatively rigid when fitting in theactive sites. Moreover, the decrease in charge by the addition of the dye is alsocompensated by the additional phosphate groups. [250] We observed that the Ap4analog is utilized more efficiently in vitro as well as in cells as compared to Ap3analog. Thus, only the Ap4 analog was used for the following experiments.

To get an insight into the cellular pathways that are responsible for the

5.2 Ap4 hydrolysis In Vitro 61

Figure 5.3: (a) Continous monitoring of Ap4 and Ap3 hydrolysis in HeLa

cell lysates by measuring the increase in the fluorescence intensity. Blank

represents the hydrolysis in reaction buffer.(b) Hydrolysis of Ap4 and Ap3

in different cell lysates. Error bars represent the standard deviation from

mean.

utilization of the Ap4 nucleotide analog, an in vitro inhibitor assay was per-formed. In this experiment, specific inhibitors of various prominent cellularATP consuming pathways were used to inhibit the hydrolysis activity of Ap4 incell lysates. The inhibitors used were ouabain, an inhibitor of sodium-potassiumATPase pump; [252] oligomycin and sodium azide, the inhibitors of the mitochon-drial F0-F1 ATPase system; [253] staurosporine, a protein kinase inhibitor; [254]

sodium orthovanadate, an inhibitor of phosphatases and, kinases and sodium py-rophosphate which act as a competitive inhibitor of ATPases on the whole. [255]

The hydrolysis of the Ap4 analog was monitored in the plate reader assay at37◦ C. Cell lysates were prepared from two different cell lines HeLa and HEK293T (human embryonic kidney cell line). A wide range of concentrations ofinhibitors was used and the kinetics was monitored. The increase in the con-centration of inhibitors in each case resulted in the inhibition of the hydrolysisin different proportions. The average relative inhibition of the Ap4 hydrolysisactivity indicates that the enzymes involved in diverse cellular pathways utilizethe Ap4 analog (Figure 5.4).


Figure 5.4: Relative inhibition of Ap4 analog turn-over in HeLa and HEK

cell lysates after treating with the various inhibitors. Error bars represent

the standard deviation from mean.

5.3 Incorporation of the Ap4 analog in living cells

After monitoring the hydrolysis of the ATP analog in vitro in cellular lysates, theobjective was to visualize the hydrolysis of Ap4 analog in living cells. For thispurpose, it was necessary to incorporate the Ap4 analog into the cells withoutaffecting the physiology of the cells. Most of the synthesized chemical analogsare polar and thus are impermeable for the cell membrane. Diverse strategieshave been used to synthesize analogs that can freely diffuse into the cells. Sincethe Ap4 analog itself is not cell-permeable, two different methods were used tointroduce the analog into the cells: Triton-X membrane permeabilization andelectroporation. In the first strategy, HeLa cells were treated with a mixtureof 0.01% Triton X-100, a non-ionic detergent and 100 µM of the Ap4 analog in1X phosphate-buffered saline (PBS) for 12 minutes at 4 ◦ C. After permeabi-lization, the cells were washed with 1X PBS and then imaged. It is known thatlow concentrations of Triton-X-100 make the plasma membrane permeable byinsertion into the lipid bilayer. [256] With this approach, the Ap4 was internalizedand dispensed non-homogeneously as punctate structures in the cells. However,

5.3 Incorporation of the Ap4 analog in living cells 63

most of the cells treated with Triton-X-100 collapsed during the experiment andthe overall viability decreased as could be seen by a gradual disintegration ofthe plasma membrane.

.

Figure 5.5: Incorporation of Ap4 analog into HeLa cells by electroporation.

Increase in the incorporation of Ap4 observed with the increase in the con-

centration of the analog in the electroporation medium and with the increase

in pulse duration; scale bar: 30 µm.

Another approach tested for the internalization was electroporation. Cellswere treated with the Ap4 analog at the desired concentrations and short electricpulses of 160 V were applied with a specific pulse duration. Viability assays ofthe cells electroporated with Ap4 analog showed an insignificant decrease in via-bility a few hours after the treatment. After electroporation, the cells exhibitedlittle morphological changes and imaging was possible for an extended period.It was observed that with the increase in the pulse duration and the concentra-tion of the analog in the medium, the Ap4 incorporation increased (Figure 5.5).


Electroporation was found to be the most efficient method since the number ofcells with the analog incorporated was comparatively high and the distributionof the analog was homogeneous in the cells. In subsequent experiments, wetherefore employed electroporation as the method for incorporation of the Ap4analog.

.

Figure 5.6: Comparison the the cells after electroporation and Triton X-

100 permeabilization; scale bar:10 µm

The cells were electroporated in presence of different concentrations ofthe Ap4 analog. After two hours of incubation, the cells were treated with theAlamarBlue reagent as discussed in chapter 5. Cells without electroporationwere used as control. The percentage of viability was estimated with respect tothe control. No considerable reduction in the viability was observed at Ap4 con-centrations up to 200 µM at the time scale of our experimental procedure. Wehave used the concentration of 100 µM in our experiments with cells (Figure 5.7).

5.4 Imaging of Ap4 Hydrolysis in Living Cells 65

Figure 5.7: Viability of cells after electroporation with different concen-

trations of Ap4. The values represent the mean of three independent exper-

iments performed in triplicates. Error bars represent the standard deviation

of the mean.

5.4 Imaging of Ap4 Hydrolysis in Living Cells

The hydrolysis of the Ap4 analog results in the increase in the fluorescence life-time of the dye attached to the phosphate chain of the analog. This is becauseof the increase in the fluorescence quantum yield of Atto-488 donor fluorophoreafter the termination of FRET between the dye and the quencher, after enzy-matic cleavage of the Ap4 analog. This principle was used for performing FLIMmicroscopy of the Ap4 analog hydrolysis in the living cells. FLIM imaging isbeneficial because the detection is less prone to artifacts due to photobleaching.It is more apt for the detection of FRET in the complex environment of a livingcell as it offers various other advantages over the conventional intensity basedimaging. [56,55] Thus it was possible to follow the kinetics of enzymatic processesin which the Ap4 analog is cleaved by monitoring fluorescence lifetime changesas a function of time.

5.4.1 Wide Field FLIM Imaging of the Ap4 Hydrolysis

For the FLIM imaging using the widefield microscopy setup, the incorporation ofthe analog was peroformed by Triton-X-100 permeabilization with the concen-tration of 100 uM analog in permeabilization buffer. Before permeabilization,


HeLa cells were incubated for 30 min at 4◦ C to slow down their metabolismin order to make sure that the compound is not already cleaved before we startimaging the cells. The fluorescence lifetime was measured on a custom buildwide-field frequency domain FLIM setup equipped with a CCD camera and amodulated intensifier. Changes in the fluorescence lifetimes of the Ap4 analogwhich were recorded for each pixel, are directly related to the hydrolytic conver-sion of the Ap4 analog. The widefield approach gives a high temporal resolutionbecause of the simultaneous data acquisition for all pixels. An increase in thefluorescence lifetimes, both the phase lifetime as well as the modulation lifetime,was seen all over the cell. However, the change in the lifetime in the nuclearregion was higher than in the cytoplasm. After 30 minutes, the lifetimes inboth the regions matched. Vesicular structures with higher fluorescence life-times were also found in the cytoplasm. This indicated the hydrolysis of theanalog in different cellular compartments. The relative increase in the phaselifetime was more prominent (from 1.5 ns to 2.2 ns) as compared to the increasein modulation lifetime (2.5 ns to 2.9 ns) The increase in the lifetime saturatedafter 1 hour (Figure 5.8).

5.4.2 Confocal FLIM microscopy of the Ap4 hydrolysis

To achieve a better spatial resolution, FLIM images were also recorded usinga fast confocal microscope (Leica SP8 FALCON) in which FLIM was readoutby time correlated single photon counting (TCSPC). This approach is based onconfocal scanning with pulsed laser excitation. The measurement was done atroom temperature over a period of 1 hour. For this experiment, the Ap4 ana-log was incorporated into HeLa cells using electroporation. A pulse durationof 10 µs was used at the constant voltage of 160V and 10 pulses were applied.The concentration of the analog in the electroporation buffer was 100 µM. Wefound that Ap4 hydrolysis started as soon as it was introduced into the cells andreached a steady-state in about 60 minutes during which the average fluores-cence lifetime rose from around 2.6 ns to 3.2 ns (Figure 5.9). The experimentswere done at Leica Microsystems in Mannheim, Germany.

The enzymatic hydrolysis of the ATP analog could thus be monitored in

5.4 Imaging of Ap4 Hydrolysis in Living Cells 67

Figure 5.8: (a) Representative widefield FLIM images of a cell at different

time points; scale bar:10µm. (b) Quantitative evaluation of the phase and

modulation fluorescence lifetimes (No. of cells=10). Error bars represent

the standard deviation of mean.

the living cells with high temporal resolution. More experiments were performedin the following in order to characterize the cellular processes involved in thehydrolysis and utilization of the Ap4 analog.


Figure 5.9: Live cell imaging of Ap4 hydrolysis by TCSPC FLIM using

a confocal microscope setup. (a) Representative FLIM images of a cell at

different time points. (b) Amplitude weighted mean fluorescence lifetimes

plotted with respect to the time (N=8). Error bars represent the standard

deviation of the mean; scale bar:10 µm.

5.5 Ap4 hydrolysis in lysosomes.

In order to identify the sub-cellular location associated with the Ap4 hydrolysisin living cells, confocal fluorescence microscopy was performed and the imagesrevealed the fluorescence of the Ap4 analog appearing in punctuate structuresin live cells. For confocal imaging, HeLa cells were electroporated with Ap4 atthe concentration of 100 µM analog in the electroporation medium. After elec-troporation, the buffer was replaced with the complete medium for the live cellimaging experiments. The appearance of fluorescence indicated the hydrolysis of

5.5 Ap4 hydrolysis in lysosomes. 69

the analogs. Various fluorescent organelle trackers were used for colocalizationexperiments to find the cellular compartment associated with the Ap4 hydroly-sis. Mitotracker red is a reduced probe oxidized in the cells to form a fluorescentproduct that is sequestered into the mitochondria. Lysotracker deep red is aweakly basic probe that is specifically accumulated in the lysosomes because oftheir acidic nature. These dyes passively diffuse into the cells when incubatedwith the cellular media. The cells were treated with these fluorescent trackers.The cells were washed with 1X DPBS and Ap4 incorporation was done usingelectroporation. The cells were fixed using paraformaldehyde and imaging wasperformed.

A relatively weak correlation was found between the Ap4 analog hydroly-sis and mitochondria (Pearson’s correlation factor 0.77). However, a strong cor-relation (Pearson’s correlation factor 0.91) was found between the Ap4 analoghydrolysis puncta and lysosomes(Figure 5.10). This implies that the hydrolysisof Ap4 analogs takes place in lysosomes.

To test further the hypothesis of localization of Ap4 hydrolysis in lyso-somes, the cells were treated with β–lapachone that damages lysosomes andinduces necrosis by elevating the levels of free radicals. [257,258]The cells wereelectroporated with the Ap4 analog after incubation with increasing concentra-tions of β–lapachone for 2 hours. A significant decrease in punctuate fluorescentstructures was observed in the cells treated with β–lapachone. Also, the cellswere treated in parallel with increasing concentrations of oligomycin, an in-hibitor of mitochondrial ATPase. No effect on the Ap4 hydrolysis was observedon treatment with oligomycin (Figure 5.11). Moreover, the disintegration oflysosomes could also be observed after staining the β–lapachone treated cellswith lysotracker (Figure 5.12). This is a second indication that lysosomes areprominently involved in the hydrolysis of the Ap4 analog.

To further elucidate whether the hydrolysis of the Ap4 analog occurs inlysosomes, two lysosomal inhibitors were used. Chloroquine is a lysosomotropicagent that inhibits the enzymatic activity by increasing the pH inside the lyso-


Figure 5.10: Colocalization of Ap4 hydrolysis spots with (a) mitochondria

(b) lysosomes using fluorescent trackers: Colocalization of both compounds

is shown in yellow. A scatterplot of red and green pixel intensities is shown;

N=20; scale bar: 10 µm.


somes. The monoprotonated form of chloroquine diffuses into the lysosomeswhere it is deprotonated and is trapped, thus changing the lysosomal pH andthereby inhibiting the lysosomal activity. [259] By contrast, bafilomycin A1 is amacrolide antibiotic that is used as an inhibitor of lysosomal H+ ATPase. [260]

Bafilomycin prevents the acidification of endosomes and lysosomes and thus in-hibits lysosomal functioning. In this experiment, the cells were treated withincreasing concentrations of the respective inhibitors overnight, the Ap4 analogwas added and internalized via electroporation before the cells were imaged.The use of either inhibitor led to a drastic decrease in the hydrolysis of the Ap4analog in living cells (Figure 5.13). These experiments confirm the involvementof lysosomes in the Ap4 hydrolysis.

Lysosomes require a large amount of energy to maintain their low intra-luminal pH of 4.2 to 5.3. [261] Moreover, lysosomes are the centers for crucialcellular functions like metabolism, immune defense, and signaling. Lysosomalmembranes harbor a wide range of transporters that are involved in the ac-tive translocation of various substances like heavy metal ions and oligoscca-harides. [262], [263] Copper transporters ATP7A and ATP7B are the P-type AT-Pases that couple with the ATP hydrolysis for the Cu2+ ion transport fromthe cytoplasm into the lysosomes. [264] Multivesicular body (MVB) pathway isessential for the sorting and transport of proteins for degradation and recyclingin lysosomes. ATP hydrolyzing enzyme VPS-4 acts in concert with the ESCRT(endosomal sorting complex required for transport) complex that mediates themembrane remodeling during endosome-lysosome fusion. [265] The data largelyleads us to the speculations that Ap4 analog is utilized in one or more lysosomalfunctions.

In order to make sure that the increase in the fluorescence intensity isindeed an enzymatic process and not an artifact or a nonspecific phenomenon,the time-lapse confocal imaging of the living cells was done after incorporationof the Ap4 analog and an increase in the fluorescence intensity was seen overtime, indicating that the hydrolysis of the Ap4 analog is an enzyme dependentprocess (Figure 5.14).


There was the possibility that Ap4 was being recognized as a foreignmolecule by the cell that could result in its non-specific degradation. Ap4could be taken up and removed by the cellular endosomal pathway in a non-specific manner. To investigate this possibility, we did the colocalization anal-ysis of Ap4 with the 70 kDa Dextran-rhodamine B, a widely used endocytosismarker. [266,267] The very poor colocalization of Ap4 with endosomes as comparedto that of lysosomes shows that Ap4 is not taken up by the cell non-specificallyby the process of endocytosis and is not hydrolyzed by the endosomal pathwayas such (Figure 5.15 a and b).

In another control experiment, cells were also electroporated with nativeunmodified Atto-488 dye. The dye can be seen diffused all over the cell anddoesn’t get accumulated in specific cellular structures unlike Ap4 analog. Itindicates that the Ap4 analog is specifically localized and metabolized in cellswhich is not the case with native Atto 488 (Figure 5.15 c).


Figure 5.14: (a) Representative images of the increase in the Ap4 fluo-

rescence intensity with respect to time; scale bar:20 µm (b) Plot showing

quantitative average increase in the corrected total cell fluorescence (CTCF)

with respect to time. The values indicate the average of 20 cells and the

error bars indicate the standard deviation of the mean.


Figure 5.11: Effect of oligomycin (a) and β-lapachone (b) on Ap4 hydrolysis

at different concentrations.(c) and (d) The change in the hydrolysis activity

is plotted w.r.t. the concentration. Corrected total cell fluorescence (CTCF)

is plotted. Error bars represent the standard deviation of the mean; N = 20;

scale bar:10 µm.


Figure 5.12: Fluorescence images showing the effect of β-lapachone on (a)

Ap4 hydrolysis and (b)lysosomes. Surface plot profile showing the intensity

distribution in the cell. Increase in the fluorescence distribution in lysosomes

observed because of their disintegration; scale bar : 10 µm.


Figure 5.13: (a) Effect of bafilomycin on Ap4 hydrolysis at different con-

centrations.(b) Effect of chloroquine on Ap4 hydrolysis at different concen-

trations.(c) and (d) Plots representing the decrease in the Ap4 hydrolysis

after treating with bafilomycin and chloroquine respectively. Error bars rep-

resent the standard deviation of mean. Statistical significance compared

to control.(N=20 to 25). Ordinary one-way ANOVA, ns : not significant,

***P<0.001,****P<0.0001; scale bar:10 µm.


Figure 5.15: (a) Representative images of the colocalization of Ap4 hy-

drolysis with endosomes (b) Quantification of colocalization. The values

correspond to the mean of 25 cells. Unpaired t test ***P < 0.001. Error

bars represent the standard deviation of the mean; scale bar : 10 µm. (c)

Comparison of fluorescence distribution in cells after electroporation with

Ap4 and native Atto 488 dye; scale bar : 10 µm.


5.6 Utilization of Ap4 Analog in Autophagy

After observing that the Ap4 analog is localized and hydrolyzed in lysosomes,we investigated whether monitoring its fluorescence can be utilized to follow anyspecific cellular processes. One of the fundamental functions in which lysosomesplay an important role is the process of autophagy. Autophagy is a regulatedcatabolic pathway for the turnover of non-functional proteins, macromolecularaggregates and damaged organelles by lysosomal degradation. [268] Autophagybegins with the formation of a double-membrane vesicle (autophagosome) thatengulfs cellular constituents such as protein aggregates and damaged organelles.The outer membrane of the autophagosome then docks and fuses with the lyso-some to deliver the sequestered cargo. The inner membrane of the fused vesicle(autolysosome) along with the interior contents of the autophagosome is de-graded by lysosomal hydrolases to generate nucleotides, amino acids, and freefatty acids that are recycled to provide raw materials and energy to the cells.

Lysosomes play a fundamental role in the process of cellular autophagy.It is known that macrophage requires ATP for autophagy activation, whereaschaperon mediated autophagy requires ATP for chaperon-substrate binding andsubstrate unfolding. [269] Finally, microautophagy uses ATP to induce membraneinvaginations and cargo sequestering. [270] Therefore, we investigated the possi-bility of the Ap4 analog being utilized during the process of autophagy. To thisend, we transiently transfected LC3B-red fluorescent protein (LC3B-RFP) inHeLa cells and the protein was expressed in the cells within 24 hours. The LC3Bprotein normally resides in the cytosol. During the process of autophagy, it isassociated with phagophores after the attachment of phosphatidylethanolamine.This makes it appropriate as a marker for the process of autophagy. [271] In thisexperiment, cells were treated with 200 nM of Ink-128 (sapanisertib) which is anautophagy inducer or 100 µM of chloroquine which is an autophagy inhibitor.Cells without any treatment were used as control. Sapanisertib is a potent andselective inhibitor of mTOR signalling pathway that induces autophagy by thedysregulation of certain metabolic pathways. Cells were imaged after electro-poration with the Ap4 analog and the hydrolysis of Ap4 was quantified andits colocalization with LC3B-RFP was quantified. Colocalization of LC3B-RFP

5.6 Utilization of Ap4 Analog in Autophagy 79

puncta and Ap4 analog puncta was observed as shown in figure 5.16.

Figure 5.16: (a) Representative image of Colocalization of Ap4 hydrol-

ysis and LC3B-RFP fluorescence. HeLa Cells were transfected with the

autophagy sensor LC3B-RFP and cultured for 24 hours (red). Cells were

then treated with Ink-128 for 4 hours before electroporation with the Ap4

analog (green); scale bar: 10 µm. (b) Line profiles for pixel intensities of the

colocalized Ap4 and LC3B-RFP.

In chloroquine treated cells, LC3B-RFP puncta formation increased. Weexplain this by the blocking of autophagy, resulting in the accumulation of au-tophagosomes and the decrease in Ap4 hydrolysis. Eventually, this leads to adecline in colocalization. Upon induction of autophagy by Ink-128, the colocal-


ization between Ap4 puncta and LC3B-RFP puncta increases as compared tocontrol. This is due to the increase in autophagy activity and the Ap4 hydroly-sis. This experiment indicates that Ap4 might be utilized during later stages ofautophagy (Figure 5.17).

Figure 5.17: (a) Quantification of the colocalized spots. (b) Tukey’s multi-

ple comparison tests for the colocalization data and its statistical significance

with adjusted p values. The values represent the mean of 20 cells. Error

bars represent the standard deviation of the mean.

5.7 Summary and Outlook 81

5.7 Summary and Outlook

The importance of the ATP and its hydrolysis in the biological functioning isundisputed. ATP hydrolysis is largely enzyme-dependent in various cellularprocesses. To get an insight into the molecular mechanism involved in thesepathways, it is important to monitor the enzymatic hydrolysis activity of ATP.This project aimed to visualize the hydrolysis of ATP and the related pathways.For this purpose, a novel ATP analog named AP4 was designed and synthesized.The analog has a fluorescent dye (Atto-488) attached to one terminal and aquencher attached to the other. In an intact molecule, the fluorescence of thedye is quenched due to Förster resonance energy transfer (FRET) because of thespatial proximity between donor and quencher. Here, the dye acts as a donorand the quencher acts as an acceptor. The energy transfer is abolished by en-zymatic cleavage resulting in an increase in the fluorescence intensity as well asin an increase in the fluorescence lifetime. The change in intensity and lifetimewas monitored by confocal and FLIM imaging to quantify the hydrolysis of theanalog. Since the cell membrane is impermeable to the analog, two methods,i.e. Triton X-100 treatment and electroporation were utilized to incorporate theanalog into living mammalian cells. Electroporation was preferred for most ofthe experiments because it was less detrimental for the cells.

Confocal imaging was performed to visualize the localization of analoghydrolysis in combination with the fluorescent organelle trackers. By the colo-calization experiments, it was observed that the analog is mainly hydrolysed inthe lysosomes. However, the possibility of hydrolysis by other cellular processescan not be ruled out as background fluorescence is seen in other parts of thecell. Moreover, the utilization of AP4 in the process of cellular autophagy wasinvestigated by using the autophagy marker LC3B simultaneously with the ana-log together with stimulation and suppression of autophagy. Since lysosomesare involved in autophagy and the process requires a large amount of energy,the utilization of AP4 as a source of energy is highly likely.

The increase in the enzyme specificity of the analog would be desirable in


future to visualize activity of a specific enzymes in cells. This approach wouldbe especially valuable for the high throughput screening of the chemicals relatedto lysosomal and autophagy inhibition with clinical relevance. Our experimentsindicate the possibility of using this analog to quantify the autophagic vacuoledistribution profiles and autophagosome-lysosome fusion. This approach couldprovide detailed insights into the mechanism of autophagy induction and flux.Also, the usage of fluorescently-tagged autophagy markers in combination withthe analog will be beneficial for further monitoring the process of autophagy.

Chapter 6

Protein PARylation in DNA

Damage Response

The genetic material of all organisms is prone to various kinds of damagesfrom both intrinsic as well as external environmental factors. The implica-tions of DNA damage can be far-reaching ranging from genetic disorders tocarcinogenesis. To maintain the DNA in its indigenous form is a challenge andcells have evolved with a complex mechanism of multiple pathways for DNAdamage response. A fundamental step in the DNA damage response is theprocess of protein PolyADP-ribosylation at the site of DNA damage. ProteinPARylation is key to the initiation of the DNA damage repair process and poly(ADP-ribose) polymerase (or ARTD1) is prominently involved in the processof PARylation. [272] Innumerable studies have been carried to study the mecha-nism and the regulation of the process of PARylation. PAR binding interactionshave been demonstrated in various studies so far by using techniques like surfaceplasmon resonance (SPR) and electrophoretic mobility shift assays (EMSA). [273]

Since DNA damage repair is a dynamic process and takes place in response toa variety of stimuli, the techniques that are used in vitro and in the fixed cellsdo not give an appropriate picture of the molecular events involved. Methodsfor the direct observation of poly PAR in living cells are still nonexistent andthus it has been largely impossible to observe the dynamic events in cells.


This study aimed at visualizing the protein PARylation in real-time afterthe induction of DNA damage by microirradiation. We have used a NAD analogmodified with the fluorescent dye tetramethylrhodamine (TMR) in combinationwith the FLIM microscopy to monitor the process of PARylation. The fluores-cent NAD analog has been characterized and its utilization by ARTD1 has beenstudied previously in vitro as well as in cells. [222,274]. An EGFP fusion to theprotein of interest was used to follow its recruitment to the damaged site. DNAdamage was performed in a spatially well-defined region by laser microirradia-tion. The imaging is based on the FRET between the two fluorophores whenthe EGFP fusion protein and the NAD analog are in close proximity. Thiscauses a decrease in the fluorescence lifetime of the donor fluorophore which ismeasured to visualize the protein PARylation and the associated interactions.The schematics of the experimental approach is depicted in figures 6.1 and 6.2.

Figure 6.1: Schematics of the experimental procedure.

6.1 Incorporation of TMR-NAD Analog into the Living Cells 85

Figure 6.2: Visualization of protein-specific PARylation in cells using

TMR-NAD analog and FLIM-FRET imaging.

6.1 Incorporation of TMR-NAD Analog into the

Living Cells

Most of the chemical probes are polar in nature and thus are impermeable forthe cell membranes. So various methods have been used to introduce theseprobes into the cells. These methods having varying efficiencies are used to fa-cilitate their transport into the cells. We have tried four different methods thatare discussed below:

1. Electroporation: In this method, an electric field is applied across the cellsto create transient pores in the cell membrane that allows the diffusion ofanalogs into the cells. It has been used diversely from bacteria to mam-malian cells. [275] This method needs thorough testing and standardizationof parameters including voltage, pulse duration, and the number of cyclesto maximize the intake of probes and minimize the cellular toxicity.


Electroporation was used for the incorporation of TMR-NAD into the cellsand it was performed in ZAP buffer as a medium. Rectangular pulses witha constant voltage of 160 V were used. Variation of parameters includ-ing the number of pulses (10-15) and the pulse duration (300-1500 µs)were used to optimize the conditions. Homogeneous distribution of theanalog was achieved with electroporation and the intensity of the cells in-creased as a function of pulse duration as well as the concentration of theanalog in the medium. However, it was seen that the normal physiologyof the cells is compromised after electroporation and that damage to thecell health increase considerably with an increase in the pulse durationduring electroporation. The recruitment of DNA repair factors is gov-erned by the parameters used for the damage. The precise conditions atwhich the expected kinetics of the protein accumulation is achieved with-out compromising the cell physiology during the imaging period neededto be standardized for this method.

2. Carrier peptide: Several peptides have been designed which have provedefficient in mediating the internalization of chemical probes while main-taining cellular integrity. These peptides are termed protein transductiondomains (PTDs) and they cross the membrane without the help of anytransporters thus facilitating the transport of cargo. [276] We have used acarrier peptide known as pep-1 cysteamine for the delivery of TMR-NADanalog into the cells (sequence: KETWWETWWTEWSQPKKKRKV)This peptide is not toxic to the cells and forms a non-covalent complexwith the cargo to be transported. [277] The transport mechanism of thispeptide is largely unknown. However, it has been shown that the hy-drophobic domain of the peptide forms a helical structure when in closeproximity to the lipids. Hydrophobic interactions between the peptideand lipid membrane are mediated by tryptophan residues that promoteits insertion into the membrane. The conditions were standardized andthe best results in terms of the cell health and the kinetics were obtainedat a concentration of 100 µM of Pep1. The reaction was carried out in PBSwith the optimum concentration of TMR-NAD as 50 µM and 1mM DTT.

6.1 Incorporation of TMR-NAD Analog into the Living Cells 87

Great care has to be taken while using the peptide carrier because theyform fluorescent aggregates. This causes inhomogeneous distributions ofthe analog in the cells.

3. Liposome transfection using DOTAP: Liposomal transfection is a conven-tional method of incorporating DNA in cells. This method has been usedrecently to introduce certain probes into the cells. DOTAP ((1,2-Di-(9Z-octadecenoyl)-3-trimethylammonium propane methylsulfate)) is a cationiclipid that has been used for the transfection of mammalian cell lines withhigh efficiency. It can be used directly into the cell culture medium andhas relatively few cytotoxic effects. [278] For the incorporation of analogusing this approach, the analog and the DOTAP were suspended in themedium and the liposomes were allowed to form for some time. The cul-tured cells were treated with the transfection mixture and incubated for 1to 4 hours with 100 to 200 µM concentrations of TMR-NAD analog. Onehour of incubation and 200 µM of TMR-NAD concentration was found tobe optimum and were used for further experiments. A variety of fluores-cence distributions was obtained with different cells. The cells with themost uniform distribution of fluorescence were used for the experiment.Most of the experiments in this study were done after the incorporation ofTMR-NAD with DOTAP transfection because of the better cell viability.

4. Cell squeezing: In this method, the cells are mechanically squeezed througha fine capillary in the media with the analog. The analog gets into thecells through the temporary pores that are formed during the process.The analog is not homogeneously distributing in the cells. For this pro-cedure, the cells have to be detached from the dish and suspended in themedia. The cells need to adhere again to the surface before imaging canbe performed. Thus the method is comparatively time consuming andcumbersome.

All the above mentioned methods were used for the incorporation of the NADanalog into the cells and were tested for the advantages and the disadvantages


they offer. The different methods affect the incorporation of analog in differ-ent ways. Also, the cell viability was seen to be dependent on the method ofincorporation. Cells that were electroporated exhibited a homogeneous analogconcentration but were morphologically compromised. The quantitative resultsobtained earlier showed a considerable decrease in the viability of cells withan increase in electroporation pulse duration and concentration. On the otherhand, the treatment of cells with lipid reagent and carrier peptide resulted in adiscrete and punctuated distribution of analog but healthy morphological fea-tures of the cell. Moreover, the cells treated with different methods showeddifferent behaviors upon DNA damage induction (Figure 6.3 a.)

We also performed cell viability assay based on luminescence to com-pare the cells transfected with DOTAP and the cells that are electroporatedat different concentrations of the analog. For this purpose, the CellTiter-Gloluminescent assay was used which works on the principle of the quantificationof ATP in the metabolically active cells. [279] Cells without transfection wereused as control and the viability was calculated as a percentage with respectto the control. There was a decrease in the cell viability with the increase inthe concentration of the NAD analog in both cases. A reduction of nearly 20%was observed in the cell viability at the highest analog concentration of 400 µMin case of DOTAP transfection but it was nearly 50% in case of electropora-tion.(Figure 6.3 b) We therefore have used only upto 200 µM of analog in allour experiments.

6.2 Laser induced DNA damage 89

Figure 6.3: (a) Comparison of the cells incorporated with NAD analog

using electroporation and DOTAP. Scale bar:10 µm (b) Cell viability assay

after NAD analog incorporation using electroporation and DOTAP. Error

bars represent the standard deviation of the mean.

6.2 Laser induced DNA damage

In order to visualize protein PARylation in living cells, DNA damage was in-duced in a well-defined linear region of the nucleus for the activation of ARTD1.The microirradiation was done using short laser pulses. This method of in-


ducing DNA damage offers the advantage of having a distinguished point ofreference at which visualization of the DNA damage response can be started.The widefield FLIM setup was originally established in the laboratory by Dr.Annette S. Indlekofer. [249] Later on, the titanium–sapphire near infrared laserwas integrated into the setup by Tobias Löffler. He standardized the variousparameters for the induction of DNA damage. [280] The emission wavelength ofthe laser could be tuned from 700 nm to 980 nm. The frequency of the laserpulses was 76 MHz. The pulse picker was introduced into the setup in orderto control the total amount of energy incident on the sample by adjusting thefrequency. The average energy could be varied by changing the frequency of thelaser pulses without decreasing the energy per pulse. A humidified chamber wasused to maintain the physiological conditions of 37◦C temperature and 5% CO2

for the cells during the whole procedure of inducing damage and imaging.

The damage was written in a linear region of 10 µm length by movingthe stage at 1 µm per sec with an average laser power of 1.7 mW incident onthe sample. In the experiments discussed here, we used short pulses of 780 nmwavelength with pulse duration of 280 femtoseconds to induce damage. The fre-quency of the laser pulses was reduced to 3.8 MHz in our experiments with thehelp of a pulse picker. The average laser power using these parameters was mea-sured to be 1.3 mW. Using these conditions, the optimum DNA damage responsewas observed. A decrease in the average power would result in an insufficient re-sponse to be measured and an increase in the laser power would damage the cellsinstantly. For the DNA damage induction, the cells were carefully chosen basedon the presence of the fluorescence from the expression of EGFP fusion proteinand that of the incorporated analog. The cells having the optimum concentra-tions of both the fluorophores responded well to the damage induction. Cellswith higher expression of EGFP fusion protein showed no response to the DNAdamage. we speculate that the higher expression of a protein induces high stresslevels in the cells which might result in a decrease in the DNA damage response.

The non-linear multi-photon absorption by laser pulses prompts varioustypes of DNA damages depending on the total average energy absorbed by the

6.3 Nature of the DNA damage Induced 91

cell over a particular time for which a cell is irradiated. [231,281]

6.3 Nature of the DNA damage Induced

The variety of DNA damage that can be induced with microirradiation has beenpreviously discussed in detail in chapter 3. Two primary types of DNA damagesthat are induced by the radiations are double strand breaks and the formationof photoproducts. We have used two assays to determine the type of damagesinduced in our experiments. The γ-H2AX assay is widely used for estimatingthe DNA damage response to double strand breaks (DSBs). H2AX is a variantof core histone H2A that gets phosphorylated on serine-139 upon double strandbreak induction by PI3-kinases. The phosphorylated form is called γ-H2AX andit localizes at the site of DNA repair to recruit other repair factors. [282,283] Theantibody is used against the γ-H2AX to detect the characteristic DNA doublestrand breaks. For this experiment, the cells were grown on a plate with a gridmarked on the bottom surface. The damage was induced by microirradiationand the positions of the cells were noted. The cells were fixed after the inductionof damage and the antibody staining was performed. At the end, the predefinedcells were recognized by the positions on the grid and were imaged using fluo-rescence microscopy.

Another prominent DNA damage is the formation of cyclobutane pyrim-idine dimers (CPD) mostly induced by UV radiation. Pyrimidine dimers areformed by the covalent bonding of two consecutive nucleotides in a DNA strand.The CPD immunoassay is an antibody-based assay used for the detection andquantification of CPD. [281] A similar strategy as in the case of γ-H2AX stainingwas used to perform CPD staining as well. The CPD staining was done in theBioimaging Centre, University of Konstanz. The cells were stained with theanti-CPD antibody after the microirradiation and then fixed. The cells weretreated with a secondary antibody conjugated with FITC and immunofluores-cence was performed.

These assays exhibited a variety of damages that are induced when dif-


ferent parameters for the microirradiation are used. The laser power of 1.7 mWwith a repetition rate of 3.8 MHz and pulse duration of 280 femtoseconds wasused on the nuclei for damage. It leads to the combination of pyrimidine dimerformation and DNA DSBs. The intense DSBs observed are not only restrictedto the damaged area but are also over the surrounding regions in the nucleus.Pyrimidine dimers were seen restricted to the area that was micro irradiatedas evident from the CPD assay (Figure 6.4). The average laser power was de-creased with all other parameters constant by using the pulse picker. Withless average power on the cells, the damage was decreased for both, DSBs anddimers, and confined to the region where the damage was written. However,the damage response in terms of the given protein accumulation was reducedmaking it difficult to measure the notable FRET.

Figure 6.4: Brightfield and fluorescence images indicating the formation of

DNA primer-dimers and double strand breaks after the CPD and γ-H2AX

assays. The damage was induced in the HeLa cells with the incident laser

power 1.7 mW, repetition rate of 3.8 MHz, and pulse duration of 280 fem-

toseconds on the sample. Scale bar: 10 µm.

6.4 Kinetics of ARTD1 PARylation 93

6.4 Kinetics of ARTD1 PARylation

In order to visualize the process of PARylation using live cell imaging, EGFP-ARTD1 protein was overexpressed in cells. Later, NAD analog was introducedinto the cells using one of the discussed methods. The modification of ARTD1with fluorescently labeled PAR was observed directly using the fluorescence life-time based FRET measurements. ARTD1-EGFP fluorescent protein acted asdonor and TMR-NAD acted as an acceptor. The covalent binding of fluorescentADP-ribose units to ARTD1 during the formation of polymer chain results inFRET between the two fluorophores attached to the molecules. The measure-ment of FRET using the fluorescence lifetime of EGFP allows the visualizationof PARylation. Since FRET is exclusively based on the spatial proximity ofdonor and acceptor, both the covalent interactions of PAR chains as well as thenon-covalent interactions can be directly monitored.

The kinetics of the recruitment of ARTD1 protein to the DNA damagesite was studied after the ARTD1 fused to EGFP was overexpressed in HeLacells one day before the experiment. The EGFP-ARTD1 protein overexpressionwas observed after 24 hours. The TMR-NAD analog was incorporated into thecells using liposome transfection with DOTAP 24 hours after plasmid transfec-tion. The DNA damage was made by microirradiation of NIR laser pulses overa linear region using NIR pulses with 1.5 mW of power on the sample. Thedamage was induced in the nucleus in a linear region rather than at one pointto differentiate it from possible artifacts. The damage by microirradiation offersan advantage as the local concentration of the PAR binding proteins increasesmaking it easy to detect and quantify the fluorescence intensity. Moreover, theincrease in fluorophore concentration makes the fluorescence lifetime signal easyto be detected and thus the specific protein-PAR interaction can be observedwithout the background from non-interacting proteins. The cells were imagedimmediately after the microirradiation. Fluorescence lifetime images were takenevery 10 seconds using modulated 488 laser as the source of excitation for theEGFP. The intensity of the imaging laser (488 nm λ) was 50 µW. The increasein the intensity of EGFP-ARTD1 indicates the recruitment of ARTD1 to thesite of damage. The intensity was measured over time in the damaged andthe surrounding regions after the incorporation of TMR-NAD into the cells.


The representative time-lapse images of the intensity and lifetimes are shownin figure 6.5. The signal intensities in the damaged areas were corrected forthe background related to the surrounding areas in the nucleus. The ratio ofthe corrected signal was normalized to the intensity before irradiation and thenthese intensities were plotted against the time. The relative intensity is givenby the following equation:

RelativeIntensity =

(IDmg − IBkg

ISur − IBkg

)(IDmg0 − IBkg0

ISur0 − IBkg0

) (6.1)

In the above equation, IDmg is the intensity at the damaged region, ISuris the intensity of the area surrounding the damaged region and IBkg is thebackground intensity. IDmg0, ISur0 and IBkg0 are the same at time 0, i.e. beforethe induction of the damage.

Cells transfected with EGFP-ARTD1 without the TMR-NAD incorpo-ration were used as control. There is an increase in the intensity that reachesa maximum in around 10 seconds indicating the instantaneous accumulationof ARTD1 at the damaged site. The intensity gradually declines soon after itpeaks. The decrease is likely due to the diffusion of ARTD1 after it is recruited.The observation is in accordance with studies that present the time scale of therecruitment of ARTD1 on the DNA damage. [284,285] The kinetics of accumula-tion in control cells is almost similar to that of experiment. This indicates thatthe introduction of TMR-NAD analog using these conditions does not affect thephysiological response of the cells to the DNA damage (Figure 6.6).

In control experiment, the accumulation of TMR-NAD analog was visu-alized without the overexpression of any fluorescent donor protein. The HeLacells were transfected with the TMR-NAD analog using DOTAP with 200 µM


Figure 6.5: Representative images for the changes in intensity and flu-

orescence lifetime (phase lifetime) of HeLa cells with EGFP-ARTD1 and

TMR-NAD analog after microirradiation. Decrease in the lifetime at the

site of DNA damage indicated with the false colour can be observed upon

DNA damage: Scale bar:10 µm.

Figure 6.6: Relative intensity changes in cells with EGFP-ARTD1 and

TMR-NAD analog (experiment) and the cells without TMR-NAD analog

(control) with respect to time. The error bars represent the standard error

of the mean (SEM).

of analog in the transfection mixture. The relative increase in the intensity ofthe analog was imaged after the DNA damage by laser microirradiation. A


gradual increase in the intensity of the analog can be seen because of the accu-mulation of the analog in response to damage. The accumulation is because ofthe PARylation of various wild type proteins at the damage site. Moreover, theunfused EGFP construct without any repair protein was overexpressed in thecells and the change in intensity was quantified after DNA damage induction.There was no intensity increase in comparison to the cells with EGFP-ARTD1fusion protein. On the other hand, a clear increase in the relative intensity wasseen in the cells with EGFP-ARTD1 on the damage induction (Figure 6.7).

Figure 6.7: (a) Plot representing the increase in the accumulation of TMR-

NAD analog at the damage site after the DNA damage in absence of any

EGFP fusion protein (N=20). (b) Accumulation of the EGFP-ARTD1 fusion

protein seen after DNA damage. No accumulation seen in case of the cells

overexpressed with unfused EGFP (N= 15). The error bars represent the

standard error of the mean.

In order to visualize the protein interactions in the context of PARylation,FLIM imaging was performed immediately after the microirradiation. The pixelcolor in the representative FLIM images corresponds to the fluorescence lifetimeof the donor EGFP and the intensity of color corresponds to the intensity of thefluorescence (figure 6.5). Average fluorescence lifetimes of the donor in the dam-aged area and the surrounding area were evaluated in the experiment as well as


in the control. There is a considerable decrease in the fluorescence lifetime ofthe surrounding areas as well. This is due to the significant photobleaching ofthe whole area during imaging that changes the fluorescence lifetime. Since themodulation lifetime (τm) is more sensitive to changes, we have used the phaselifetime (τφ) for all the practical purposes in this study. Note that the FRETefficiencies also depend on the position of PAR chains on a protein relative tothe position of EGFP tag on the protein. The closer the two dyes, the moreefficient the FRET would be.

In absence of TMR-NAD, there is no significant difference in the flu-orescence lifetimes between the site of damage and the surrounding areas ofthe nucleus. However, in presence of TMR-NAD, the fluorescence lifetime ofEGFP-ARTD1 shows a significant reduction exhibiting the spatial proximityof the donor and acceptor. This occurs because of the activation of ARTD1following the DNA damage by microirradiation. The activation leads to theformation of PAR polymer chains on the ARTD1 itself that results in FRET. Inthis case, it is the auto-PARylation of ARTD1 that is exhibited by the decreasein the fluorescence lifetime. The lifetimes exhibit a gradual decrease over the 5minutes because of the decrease in the PARylation. The decrease could be theeffect of the diffusion of PARylated ARTD1 after the first wave of DNA dam-age response and the deparylation of ARTD1. [286] PARylation is subsequentlyfollowed by rapid dePARylation which is necessary for the next steps of DNAdamage response. [287,288]

The fluorescence phase lifetimes were used to calculate the FRET efficien-cies in the regions of damage as well as in the surroundings. FRET efficienciescan be calculated from fluorescence lifetimes of donor in presence of acceptorfluorophore (τDA) and absence of acceptor fluorophores (τA) by the equation2.8. The FRET efficiencies were calculated as percentages. In the damagedregion, there is significantly higher FRET efficiency over time as compared tothe surrounding areas and the control (Figure 6.8 a). In case of ARTD1, FRETefficiency increases from 1% to 6% in the damaged region. A small increase inthe FRET efficiency can also be seen in the surrounding areas. This is due to


the fact that DNA gets damaged to some extent in the surrounding areas aswell with microirradiation as shown by the previous damage assays (Figure 6.8b).

Figure 6.8: (a) Decrease in the fluorescence lifetimes of EGFP-ARTD1 in

the damaged region and the surrounding region of experiment and control

with respect to time. (b) Increase in the FRET efficiencies plotted. The error

bars represent the standard error of the mean (SEM). In case of smaller error

bars than symbols, they are not shown (N=21).

In case of control, where no TMR-NAD is incorporated, there is a de-crease of about only 50 ps in lifetime as compared to that of about 150 ps in theexperiment. Also, more increase in the FRET efficiency can be seen in the ex-periment as compared to control (Figure 6.9). In order to rule out any artifactsin the lifetime measurements, an unfused EGFP construct was overexpressedin the cells and the change in lifetimes was observed after inducing the DNAdamage. There is a significant difference between the lifetime changes betweenthe EGFP and EGFP-ARTD1 in cells. However, a small decrease in the lifetimecan be seen in the case of unfused EGFP. Similarly, there is a small increasein calculated FRET efficiencies in case of unfused EGFP as compared to theEGFP-ARTD1 (Figure 6.10).


Figure 6.9: (a) Comparison of fluorescence lifetimes of EGFP-ARTD1 of

experiment (with TMR-NAD) and control (without TMR-NAD) after dam-

age. (b) Comparison of the FRET efficiencies plotted. The error bars rep-

resent the standard error of the mean (N=21).

Figure 6.10: (a) Comparison of the decrease in the fluorescence lifetimes

of EGFP-ARTD1 and unfused EGFP after the DNA damage induction. (b)

Comparison in the FRET efficiencies of EGFP-ARTD1 and only EGFP plot-

ted. Error bars represent the standard error of the mean (N=20).

This decrease in the lifetime in the controls can be attributed to thephotobleaching of the EGFP-ARTD1 while imaging overtime. The effect of


photobleaching on the fluorescence lifetimes has been previously demonstratedin these experiments and the increase in the FRET efficiency because of thiseffect has been discussed. [280]. A fraction of photobleached EGFP creates ashift of the average lifetime towards the shorter time. Thus with the increase inphotobleaching over time, an unusual change in lifetimes is detected. Moreover,iris effect is also responsible for the small change in unfused EGFP lifetimes. Inthe wide field microscopy setup, there is a variation in the optical path lengthsbetween different pixels. This leads to the offset in the phases of different pixelsestablishing different fluorescence phase lifetimes in different regions of a givenimage. This is known as the iris effect and is one of the factors responsible fordifferent average lifetimes in different areas of control. [249]

6.5 Kinetics of Macro H2A

The dynamics of histones plays an important role in the process of gene expres-sion regulation and DNA repair. Posttranslational modifications of histones in-fluence these processes by changing the chromatin structure. One of the majorchromatin modifications include the substitution of canonical histones by histonevariants. These histone variants differ from the canonical histones not only instructure but also in spatial and temporal expression given the various biologicalcontexts. [289] Histone families H2A and H3 have the most prominent variants.H2A is the largest family with structurally and functionally diverse variantslike H2A.X, H2A.Z, macroH2A (mH2A), H2A.B, etc. MacroH2A (mH2A) con-sists of an N-terminal domain similar to histone H2A and an extra C-terminalnon-histone domain (NHR) or macro domain. [290] Two subtypes of mH2A havebeen identified, named mH2A1 and mH2A2. mH2A1 has two splice variants,mH2A1.1 and mH2A1.2. [291]

The role of mH2A in the maintenance of heterochromatin architectureand DNA damage response has been demonstrated in many studies. DuringDNA repair, the reorganization of chromatin is accomplished by histone mod-ifications. These modifications together with the exchange of histone variantscontrol chromatin structure and its access to repair factors, thus allowing the

6.5 Kinetics of Macro H2A 101

repair. [292] mH2A1.1 has a PAR binding domain that promotes its interactionswith the DNA repair and chromatin remodeling machinery after it senses thePAR activation. However, mH2A1.2 is not involved in PAR binding and chro-matin remodeling. mH2A has also been shown to bind to PAR and its associ-ated metabolites. Various proteins containing macrodomain are recruited to theDNA damage sites through their interactions with PAR. [293] Moreover, the roleof mH2A1.1 has been shown in the inhibition of PARP1 to limit the PARylationof the chromatin to stop the depletion of the cellular energy pool.

Due to the relevance of mH2A in DNA damage response, we used FLIMimaging for studying the mH2A dynamics and its interactions with PAR chainsat the DNA damage site. For this purpose, the HeLa cells were overexpressedwith macro H2A1.1-EGFP by transient transfection. The TMR-NAD analogwas incorporated into the cells as described previously. The cells were exam-ined directly by wide field FLIM microscope after the induction of DNA damageby microirradiation. The recruitment of mH2A was visualized instantaneouslyafter the damage and the kinetics was determined. The representative time-lapseimages of the intensity and lifetimes are shown in figure 6.11. Cells expressingEGFP-mH2A without the incorporation of TMR-NAD were used as control.The intensity of the EGFP-mH2A peaked instantaneously after the DNA dam-age induction. The intensity then decreased gradually within the measurementtime. The accumulation of mH2A in the control was less as compared to theexperiment although the kinetics of the accumulation did not vary. This relativedecrease in intensity indicates that the incorporation of TMR-NAD affects theresponse of mH2A in DNA damage response (Figure 6.12).

An instantaneous decrease in the average fluorescence lifetime of mH2A-EGFP can be observed in the damaged area of the experiment from about 2.6ns to 2.2 ns. The decrease is significant as compared to that of the surroundingregions which decreases from 2.7 ns to 2.5 ns. This indicates the interactionof mH2A with the fluorescently labeled poly(ADP-Ribose). The change in thelifetime of the EGFP-mH2A happens at a shorter time scale as compared tothat of EGFP-ARTD1. This is possible due to the interactions between multi-


Figure 6.11: Representative images for the change in intensity and fluores-

cence lifetime (phase lifetime) of HeLa cells with EGFP-mH2A and TMR-

NAD analog with time. The decrease in lifetime at the site of DNA damage

observed upon DNA damage. Scale bar:10 µm.

Figure 6.12: Relative normalized intensity change in cells with EGFP-

mH2A and TMR-NAD analog (experiment) and the cells without TMR-

NAD analog (control) with respect to time. The error bars represent the

standard error of the mean.

ple mH2As with many different PAR chains at the damaged site. The mH2A1.1contains a macro domain that has a high binding affinity for the PAR chainsand thus interacts with repair factors through this PAR binding domain. [294]

Macro H2A binds to the PAR chains and blocks their further polymerization. It


induces the global inhibition of PARylation after the initial PARylation eventsto restrict the energy consumption of the cell.

After the sudden decrease in the fluorescence lifetime of EGFP-mH2A,the lifetime stays almost constant over a long time indicating the prolonged as-sociation of mH2A with the PAR chains. In case of control cells, i.e. the cells inwhich the NAD analog was not incorporated, the initial fluorescence lifetimesare lower although the lifetime shows a similar decreasing trend with time (Fig-ure 6.13 a). In the case of mH2A, there is an instantaneous rise in the FRETefficiency to 17% at the beginning of the measurement. It decreases slightly withtime because of the slight recovery in the lifetime. A clear difference is visiblebetween the FRET efficiencies of the damaged areas and the surrounding areas(Figure 6.13 b) Thus the non-covalent interactions between the mH2A and thePAR chain could be visualized using this approach.

Figure 6.13: (a) Decrease in the fluorescence lifetimes of EGFP-mH2A in

the damaged region and the surrounding region of experiment and control

with respect to time. (b) Increase in the FRET efficiency plotted. The error

bars represent the standard error of the mean (SEM). In case of smaller error

bars than symbols, they are not indicated (N=25).


It is noteworthy that DNA damage response is often accompanied bythe decondensation of chromatin and thus the change in the proximity of donorand acceptor dyes may take place. Since histones are actively associated withchromatin, this principle has been exploited to study the chromatin architectureand dynamics by FLIM imaging. [295,245] In our experiments, it is highly likelythat the chromatin remodeling also contributes significantly to the fluorescencelifetime change of the EGFP-mH2A. Chromatin relaxation is a prerequisite forDNA repair and this relaxation can cause an increase in fluorescence lifetime ofEGFP-mH2A. It could negatively influence our lifetime measurements.

The imaging of the EGFP-ARTD1 and EGFP-mH2A was also performedfor 40 minutes after the DNA damage induction to monitor the kinetics of theseproteins for a longer time. The kinetics of the accumulation of these proteindoes not show any significant dynamics after the initial 5 minutes. The decreasein the intensity is relatively slow in case of mH2A as compared to ARTD1. Therapid decrease in the ARTD1 is in agreement with previous studies. [272,296] Thismeans that mH2A stays at the DNA damage site for longer time (Figure 6.14a). The decrease in the lifetime of ARTD1 is slower. It takes about 10 minutesin the case of ARTD1 to saturate the decrease while the decrease is rapid in just4 minutes in mH2A. Despite the contribution of photobleaching to the decreasein the lifetime, mH2A shows a slight increase. This indicates the termination ofthe transient interactions of mH2A with the other PARylated proteins involvedover a longer time (Figure 6.14 b).

The comparison of the change in relative intensities and the change inlifetimes gives us an idea about the relative kinetics of the two processes i.e.the accumulation of a protein and its PARylation. In case of ARTD1, the twocurves peak at the same time but the decrease in the intensity is relatively fasteras compared to the decrease in lifetime. This indicates that the recruitment ofARTD1 to the site of DNA damage and its auto-PARylation take place almostsimultaneously. We are able to monitor the gradual diffusion from the damagesite overtime (Figure 6.15 a).


Figure 6.14: Change in the (a) relative intensities and (b) the fluorescence

lifetimes of EGFP-ARTD1 and EGFP-mH2A after the DNA damage ob-

served for 40 minutes. The error bars represent the standard error of the

mean (N=5).

Figure 6.15: Relative change in the intensities and lifetimes of (a) EGFP-

ARTD1 and (b) EGFP-mH2A after DNA damage. The error bars represent

the standard error of the mean.


In case of mH2A, the two processes are simultaneous. There is a rapidaccumulation of mH2A accompanied by the rapid decrease in lifetime. However,both the parameters stay constant overtime indicating the mH2A localizationand its interactions with PAR chains remain for longer time than the time scaleof our imaging (Figure 6.15 b). This may have to do with the fact that mH2Ais known to play a role in DNA damage response by mediating the formation ofDNA repair complex after the first wave of PARylation. [297,298]

6.6 Summary and Outlook

DNA damage response is a fundamental cellular event that is imperative formaintaining genome integrity. Cells have evolved with a complex process thatinvolves a plethora of protein factors to repair the damaged DNA. Various con-ventional molecular biology techniques have been used to study the mechanismof this process. But the investigation of the real-time details of the molecularevents was possible only after the advent of fluorescence microscopy. ProteinPARylation is a primary step involved in DNA damage response. The goal ofthe project was to visualize the protein PARylation in real-time by live cellimaging. To this end, a novel NAD analog modified with a fluorescent dye (tetramethylrhodamine) was synthesized. NAD is utilized as a substrate in thePARylation of proteins and thus the process can be monitored by using fluo-rescent NAD analog in combination with microscopy. The FLIM was used tomonitor the process of PARylation. FLIM imaging is based on the Förster res-onance energy transfer (FRET) between the two fluorophores when the proteinand the NAD analog are in close proximity. The decrease in the fluorescencelifetime of the donor fluorophore is measured to visualize the protein PARyla-tion and the associated interactions. Various methods were used to incorporatethe analog into living cells but the lipid transfection using DOTAP was foundto be the most efficient.

We have used the EGFP fusion proteins of ARTD1 and mH2A to fol-low their recruitment to the DNA damage site. An efficient method of lasermicroirradiation was used to induce the damage within a pre-defined region in

6.6 Summary and Outlook 107

a precisely controlled manner. The change in the intensities of the EGFP fu-sion proteins was used to monitor their recruitment to the site of DNA damage.The PARylation of an EGFP fusion protein results in FRET because of theproximity of the donor EGFP and acceptor NAD analog. The decrease in thefluorescence lifetime of donor EGFP fusion protein is monitored.

We have specifically visualized the recruitment of ARTD1 and mH2A inreal-time. Also, the kinetics of their PARylation was determined based on theFLIM imaging. Our study shows that FLIM-FRET imaging can be an appropri-ate tool for visualizing the covalent and non-covalent interactions in cells. Theinteractions of specific proteins with PAR chains can be visualized in a cellularcontext with high spatial and temporal resolution. It would be worthwhile todetermine the dePARylation activity of enzymes in the context of this analogto have a comprehensive understanding of DNA damage response using thisapproach. This method can find potential applications for the characterizationand screening of the poly ADP-ribose inhibitors in vitro. The approach can beuseful for the development of pharmaceuticals to target carcinogenesis.

Chapter 7

Materials and Methods

7.1 Materials

7.1.1 Buffers and Solutions

Table 7.1: Composition of buffers and other solutions used in this work.

Buffers and solutions Composition

10x PBS 26.7 mM KCl, 14.7 mM KH2PO4, 1.38 M NaCl, 80.6 mM Na2HPO4.7 H2O

4% paraformaldehyde (PFA) 4 w/v (% ) PFA in MQ water, pH 7.4

Cell lysis buffer (Hypertonic)1 % Triton X-100, 300mM NaCl, 5 mM EDTA,

1× Protease Inhibitor in 1X PBS

Hypotonic lysis buffer 50 mM HEPES pH 6.8, 5mM MgCl2 and 1x protease inhibitor in PBS

SDS-PAGE running buffer 25 mM Tris pH 8.3, 192 mM glycerol, 3.45 mM SDS

SVPD Buffer 100 mM Tris-HCl (pH 8.7),100 mM NaCl, 15 mMMgCl2

SDS-PAGE running buffer 25 mM Tris pH 8.3, 192 mM glycerol, 3.45 mM SDS

SVPD buffer 100 mM NaCl, 100 mM Tris-HCl (pH 8.7), 15 mM MgCl2

Western Blot buffer 25 mM Tris pH 8.3, 192 mM glycerol, 20% -V methanol

ZAP buffer 10 mM K2HPO4/KH2PO4, 1 mM MgCl2, 250 mM sucrose, pH 7.4

7.1 Materials 109

7.1.2 Mammalian Cell Lines and Bacteria

Table 7.2: Cell lines and bacteria used in this work.

Cell line Origin Description

HeLa Derived from human cervical cancer cells Group of Prof.Dr. Bürkle, University of Konstanz

HeLa S3 Human cervical cancer cell line Group of Prof. Dr. Dietrich, University of Konstanz

HEK 293T Human embryonic kidney cell line Group of Prof. Dr. Brunner, University of Konstanz

H1299 Human cell lung carcinoma cell line Group of Prof.Dr. Bürkle,Universität Konstanz

E.Coli NovaBlue Competent Cells for plasmid transformation Novagen

7.1.3 Media

Table 7.3: Media and other related constituents used in this work.

Media component Manufacturer Catalog Number

Agar Carl Roth 9002-18-0

Ampicillin Carl Roth 69-52-3

Chloramphenicol Carl Roth 56-75-7

Ampicillin Carl Roth 69-52-3

Dublecco’s modified eagle medium (DMEM) Thermo Fisher Scientific 31053028

Fetal bovine serum (FBS) Biochrom S0615

L-glutamine Thermo Fisher Scientific 25030081

Opti-MEM Thermo Fisher Scientific 31985062

Penicillin-Streptomycin Thermo Fisher Scientific 15140-122

Synth-a-Freeze Cryopreservation Medium Thermo Fisher Scientific A1254201

TrypLE Express Invitrogen 12604013

110 Materials and Methods

7.1.4 Plasmids

Table 7.4: Recombinant plasmids used in this work.

Plasmids Description

EGFP Group of Prof. Dr. Hauck, University of Konstanz

EGFP-PARP Sascha Beneke, Group of Prof. Dr. Dietrich, University of Konstanz

EGFP-ARTD1 Group of Prof. Dr. Dietrich, University of Konstanz

XRCC1-EGFP Group of Prof. Elisa May(Bioimaging Center) Universität Konstanz

PCNA-EGFP Group of Prof. Elisa May(Bioimaging Center) Universität Konstanz

XPC-EGFFP Group of Prof. Elisa May(Bioimaging Center) Universität Konstanz

mH2A1.1-CT-EGFP Addgene plasmid [291] RRID No. 45167

7.1.5 Reagents and Chemicals

Antiphospho-histone H2AX (Pser 139) antibody produced in rabbit (Sigma, #H5912), Anti-Rabbit IgG secondary antibody, Alexa Fluor 555 (Thermo Fischer,#A-31572) AlamarBlue (Thermo Fisher Scientific, #A50100), Bafilomycin A1(Abcam, #88899-55-2), BSA (Sigma-Aldrich, #A7030), Cholera Toxin SubunitB Alexa Fluor 647 Conjugate (Thermo Fisher Scientific, #C34778) CellTiter-Glo Luminescent Cell Viability Assay (promega, #G7571 chloroquine (Sigma-Aldrich, #C6628), Coumarin 6 (Sigma-Aldrich, #546283),DOTAP transfectionreagent (Biontex, #T010–1.0 ) DMSO (Sigma-Aldrich, #D8418), D(+)glucose so-lution (Sigma-Aldrich #G8769) glycerol (VWR, #24386.298 and Sigma, #G2025),Dextran, Rhodamine B; 70,000 MW (ThermoFisher Scientific,#D1841), Dul-becco’s Phosphate-Buffered Saline (DPBS) (Gibco,# 14190144) HEPES solution1 M (Thermo Fisher Scientific, #15630-056), Immersion oil for microscope (Le-ica, #11513859) Lipofectamine 2000 ransfection Reagent (Thermo Fisher Scien-tific, #11668019), B- Lapachone (Cayman Chemicals,#4707-32-8), Lyso-Tracker(Thermo Fisher Scientific, #L7526), Mito-Tracker Red (Thermo Fisher Scien-

7.1 Materials 111

tific, #M7514), PFA (Merck, #1.04005), poly-L-lysine (Sigma-Aldrich, #P4707),protease inhibitors EDTA-free (Sigma-Aldrich, #4693159001), Protein ladder(Thermo Fisher Scientific, #26617), Oligomycin (Merck, #1404-19-9), OuabainOctahydrate (Sigma,#11018-89-6) , LysoTracker Deep Red (Thermo Fisher Sci-entific, c L12492), Rhodamine 6G (Radiant dyes) Rapamycin (AlfaAesar, #53123-88-9), Sapanisertib (INK 128) (Cayman Chemicals, #1224844-38-5), TransferrinAlexa Fluor-555 Conjugate(Thermo Fisher Scientific, #T35352).

7.1.6 Kits

Plasmid DNA Purification Mini Prep Kit (Genaxxon, # S5369)CellTiter-Glo 2.0 Cell Viability Assay (# Promega)Premo Autophagy Sensor LC3B-RFP (Thermo Fisher Scientific, # P36236)

7.1.7 Equipments

Biophotometer D30 (Eppendorf), Centrifuges 5424 R, and 541R (Eppendorf),ChemiDoc Imaging System (Bio-Rad), CO2 incubator for cell culture (Binder),confocal microscope TCS SP5 (Leica Microsystems CMS), counting chamber(Neubauer), electroporator ElectroCell S20 (βtech), gel imaging system GelDoc-XR (Bio-Rad), microscope DMIL LED (Leica Microsystems CMS) with LEDlamp (CoolLED), mini Trans-Blot Cell for Western Blotting (Bio-Rad), mini-PROTEAN tetra vertical electrophoresis cell for SDS-PAGE (Bio-Rad), platereader (Synergy HT), stage incubator chamber for FLIM imaging setup (TokaiHit,INUBG2E-GSI) sonifier-250 (Branson), spectrophotometer ND-1000 (Pe-qLab), UV/VIS spectrophotometer (Agilent), wide-field frequency domain FLIMsetup (diode laser LDM488.20.A350 (Omicron), DMI 6000B inverted microscope(Leica Microsystems CMS), PIMAX4:1024i RF CCD camera coupled to a GenIII intensifier (Princeton Instruments), water bath (Thermo Fisher Scientific)

7.1.8 Consumables

1.5 ml reaction tubes (Stein Labortechnik), microscopy chambers 8-well ibiTreatµ-Slides (Ibidi), 35 mm imaging dish with imprinted cell location grid (Ibidi,#81166), plastic labware (10 cm dishes, 96-well plates, serological pipettes, cry-


ovials; Corning). 35 mm imaging dishes with polymer bottom and 500 µm celllocation grid (Ibidi #81166)

7.1.9 Softwares

EndNote (version X8.2, Clarivate Analytics), GraphPad-Prism 7 (GraphPadSoftware Inc., CA, USA.), ImageJ (version 1.5, National Institutes of Health,Bethesda,USA), Inkscape (version 0.92), Leica LAS X(version 3.5.5, Leica Mi-crosystems CMS GmbH), Lightfield (V 5.2, Princeton Instruments), MATLAB(version R2013 b and R2015 a; MathWorks, Inc.), SnapGene (version 5.1, GSLBiotech LLC), SimFCS (version 3.0, Laboratory for Fluorescence Dynamics,University of California, Irvine, USA)

7.2 Methods

7.2.1 Cell Culture Methods

Mammalian cells were grown in sterile conditions in an incubator at 37 ◦C and5 % CO2. Cells were grown in 10 cm Petri dishes in Dublecco’s modified ea-gle medium (DMEM) supplemented with 10 % FCS (fetal calf serum) and 1 %penstrip (Penicillin-Streptomycin). HeLa cells were supplemented additionallywith 2 mM L-glutamine. Cells were passaged regularly after every two or threedays depending on the confluency. For passaging, cells were washed with 10ml of DPBS and detached from the surface of the dish by treating with 1 mlof trypsin for 1 to 10 minutes depending on the type of cell line. The freshmedium was added to deactivate the trypsin and cells were resuspended in thefresh media in a new Petri dish.

For imaging experiments, cells were seeded in the appropriate dishes ac-cording to the requirements of the experiment.We have used 8-well dishes (8-wellµ-Slide, Ibidi) for most of the imaging experiments with confocal and widefieldmicroscope setups. The Ibidi dishes were treated with poly-L-lysine soultion (20 % volume in water) for 1 hour at 37 ◦C. For the imaging experiments, 20,000to 30,000 cells were seeded in each well of the 8-well Ibidi dish. The cells were

7.2 Methods 113

counted using Neubauer counting chamber in suspension after detaching fromthe surface. The cells were treated with 10 mM HEPES buffer for imaging onthe microscope in order to maintain the physiological pH of the media for longertime.

7.2.1.1 Transient Transfection of Mammalian Cells

HeLa cells were transfected with transfection reagent Lipofectamine 2000. 1µlof Lipofectamine and 1ug of plasmid DNA each were separately mixed with 50µL Opti-MEM media. The two solutions were incubated at room temperaturefrom 3 to 5 minutes and then mixed together by pipetting. The mixture wasincubated at room temperature for 10 to 15 minutes so that the liposomes areformed. The transfection mixture was added to the cells already seeded in 8-well µ-slide (20,000 cells in 200 µl media). The transfection mixture was removedfrom the cells and replaced with the fresh media after 3 to 4 hours. The cellswere incubated for at least 24 hours for the optimal expression of the protein ofinterest.

7.2.2 Incorporation of Analogs into the Cells

7.2.2.1 Incorporation by Electroporation

For the incorporation of fluorescent analogs into the mammalian cells, electro-poration was used. HeLa cells were seeded in 8 well Ibidi dishes before a day ofexperiment. The desired concentration of the analog was prepared in the ZAPbuffer. The cells were washed with DPBS and 150 µl of analog mixture wasadded to each well. Electroporation was performed using the electrodes thatfitted the half of the well. A constant voltage of 160 V was applied for 10 pulsesand a cycle duration of 1000 ms. The duration of each pulse was varied from1000 to 1500 µsec depending on properties like the molecular weight and thepolarity of the analog to be incorporated. The electric pulses were monitoredby an oscilloscope. After electroporation, the ZAP buffer mixture was removedand cells were washed three times with DPBS. The medium was again addedand imaging was performed.


7.2.2.2 Liposome Transfection Using DOTOP

For the incorporation of the TMR-NAD analog into the cells, the liposome trans-fection was performed using DOTAP (1,2-Di-(9 Z-octadecenoyl)-3-trimethyl-ammonium-propane methyl sulfate) (Biontex). 2.3 µls of the DOTAP wereadded to 100 µls of Opti-MEM medium (15 µM). 100 µM to 200 µM of TMR-NAD solution was also prepared in Opti-MEM medium. The solutions weremixed thoroughly and incubated for 15 minutes at room temperature. Themixture was added to the cells and incubated for one hour at 37 ◦C and 5 %CO2. The mixture was exchanged with regular pre-warmed medium and cellswere imaged. For imaging, the cells were kept in DMEM without phenol red.

7.2.3 Cell viability Assays

7.2.3.1 AlamarBlue Assay

The viability of the cells was determined after electroporation for the incorpo-ration of Ap4 analog using AlamarBlue. AlamarBlue contains resazurin, whichis converted to the fluorescent product resorufin by a reduction reaction in themetabolically active cells. The fluorescence intensity is proportional to the cel-lular metabolic activity which is the measure of cell health and viability. [299]

Cells were seeded in 8-well µ-slides in triplicates (30,000 cells per well)and allowed to grow overnight at 37 ◦C and 5 % CO2. Next day the cells wereelectroporated in presence of increasing concentrations of Ap4 as explained pre-viously. As controls, cells were not electroporated at all or electroporated withZAP buffer in absence of analog. After two hours of incubation, the resazurinsolution was added to the cells and incubated for 60 minutes at 37◦C and 5 %CO2. The AlamarBlue solution was prepared as 30 % V in complete mediumand added to the wells. The reagent mixture in the wells without cells was usedas blank. After incubation period, the reagent solution was transferred to a96 well plate and fluorescence was read using a plate reader (Synergy HT) at25◦C. For excitation, a 530/25 bandpass filter was selected and for reading theemission, 590/35 bandpass filter was selected. The wells with only the reagentwere used as blanks. The fluorescence intensity values of treated cells were nor-

7.2 Methods 115

malized with that of the control wells to calculate the percentage of viability.The experiment was performed in triplicates in at least three independent as-says. The viability was calculated as percentage with respect to the control aftersubtracting background fluorescence from each value.

7.2.3.2 Luminescent Cell Viability Assay

The effect of the NAD-TMR analog on cell health and viability was assessedusing CellTiter-Glo luminescent cell viability assay. The assay is based on thequantification of luminescent signal from the lysed cells based on the amountof ATP present. The luminescence is exhibited by the luciferase activity thatdirectly depends on ATP and thus the metabolic activity of the cells. [300] Cellswere seeded in 8-well Ibidi plates (30,000 cells per well) and electroporationor the DOTAP transfection was performed the next day for the incorporationof the analog with the increasing concentrations. After one-hour incubationat 37 ◦C and 5 % of CO2, the CellTiter-Glo reagent mixture was added tothe wells for cell lysis. The plate was placed on a shaker for 10 minutes forthe homogeneous lysis of cells. Control wells were prepared without cells formeasuring the background luminescence. A plate reader was used to recordthe luminescence. The experiment was repeated three times each in triplicates.Background luminescence from blank was subtracted from each reading and theviability was calculated as percentage with respect to the control.

7.2.4 Molecular Biology Methods

7.2.4.1 Cell Lysate Preparation

For the preparation of the cell lysates, cells were grown in 10 cm Petri dishestill they were 80 % to 90 % confluent. The cells were treated with 1 ml oftrypsin for 5 minutes to detach them from the surface of the dish after wash-ing them with DPBS. DPBS was added and cells were transferred to a conicaltube and centrifuged for 600 rpm for 5 minutes. The supernatant was removedand cells were washed two more times in the same manner. After removing thesupernatant completely, the cells were treated with appropriate volumes of thecell lysis buffer depending on the amount of cell pellet. The mixture was ho-mogenized by pipetting up and down a few times and incubated at 4◦ C for 30


minutes. The mixture was centrifuged at 14000 rpm for 30 minutes at 4◦ C toseparate the cell debris from the soluble fraction in supernatant that containedthe proteins. The protein concentration of the supernatant was measured usinga photometer (BioPhotometer) and then used for the experiment or stored at-80 ◦ C for further use.

For the in vitro cell lysate assays, a hypotonic buffer lysis buffer was used(50 mM HEPES pH 6.8, 5mM MgCl2 and 1X protease inhibitor in PBS). Theprotein concentration of the lysates was adjusted to 1 mg/ml with 1X PBS andthe concentration of 20 uM of ATP analogs (Ap3 or Ap4) was added. The totalvolume of the reaction mixture of 100 µl was maintained in each well of a 96well plate. The fluorescence was recorded over time with excitation filters of485/20 and emission filters of 528/20. Each experiment was done in triplicatesand repeated at least three times. For the hydrolysis of the ATP analog bySVPD, the ATP analog was incubated with and without SVPD (3.3 mU/µL)for 45 minutes at 30 ◦C in the SVPD buffer. For the inhibitor assays, a widerange of different concentrations were used in the in vitro assays to determinethe optimum inhibiting concentration of each inhibitor. The reaction kineticswas recorded after the addition of inhibitors in the same manner and the averagedecrease in the hydrolysis activity was determined with respect to the controlwith no inhibitor.

7.2.4.2 Immunocytochemistry for γ-H2AX Staining

For staining the cells after microirradiation, cells were grown in round bottomedµ-dish marked with the location grid at the bottom (Ibidi). The position of thecells was noted and the cells were fixed for 30 minutes after washing once withDPBS. After fixation, the cells were washed twice with DPBS and permeabi-lized by treating with 0.2 % Triton X-100 in PBS for exactly 5 minutes. Thecells were washed again three times and incubated with the primary antibody,Anti-phospho-Histone H2AX (Pser 139) (Sigma, # H5912) for overnight at 4◦ C(buffer: 0.1 % BSA in PBS with 0.1 % TritonX-100) at the dilution of 1:5000.Next day, the cells were washed again and incubated with secondary antibody(anti-Rabbit IgG Secondary Antibody, Alexa Fluor 555) for 1 hour at room

7.2 Methods 117

temperature at the dilution of 1:500. The cells were washed again three timesfor 5 minutes each with PBS. The cells were covered with DPBS, and stored at4◦ C until microscopy was performed.

7.2.4.3 Autophagy Sensors (LC3B-RFP) Assay

The Premo Autophagy Sensor Kit consists of an LC3B-red fluorescent pro-tein (RFP) chimera with an efficient viral transduction system, BacMam (Bac-ulovirus with a Mammalian promoter). For the expression of LC3B-RFP inHeLa cells, cells were seeded in the 8-well Ibidi plate with 20,000 cells in eachwell for doing the transfection next day. The volumes of the LC3B-RFP and theviral particles to be used were calculated according to the following equation:

Volume of LC3B - RFP (ml) =(number of cells) (MOI)

1 × 108(7.1)

Here MOI (multiplicity of infection) is the number of viral particles perwell, and (1 × 108) is the number of viral particles per ml of the reagent. Wehave used a MOI of 50 for 20,000 cells and thus, 10 µl of LC3B-RFP were addedto the cells directly and mixed by pipetting. The cells were incubated overnightfor the expression of LC3B. The cells were used for the desired experiments nextday. Subsequently, the cells were imaged using 552 nm wavelength for excitationand emission was collected from 570 to 600 nm using the appropriate filters.

7.2.5 Microscopy Methods

7.2.5.1 Wide-field Frequency Domain FLIM

A custom-built microscopy setup was used for FLIM imaging. The setup wasinitially built by Dr. Annette S. Indlekofer [320] and later modified by TobiasLöffler. The setup primarily consists of an inverted microscope (DMI 6000B, Le-ica Microsystems CMS), a diode laser with 488 nm (LDM488.20.A350, Omicron)modulated at 70 MHz and a highly sensitive RF CCD camera (PIMAX4:1024iPrinceton Instruments) coupled to a GenIII intensifier (Princeton Instruments).


The modulated 488 wavelength laser is used for the excitation of the EGFP fu-sion protein and the fluorescent ATP analog. The laser beam is cleaned by usinga filter (ZET488/10, Chroma). A cleanup filter (488/10 ZET) was used for the488 excitation laser. A telescope and a collimator is used to increase the diame-ter of the beam and collimate it respectively and the beam is carried through amulti-mode fiber (UM22-600, diameter = 600 µm, NA = 0.22, Thorlabs) to themicroscope. The beam is coupled into the back aperture of the objective anddirected onto the sample. An oil immersion objective (HC PL APO 100x 1.40NA Oil CS2, Leica Microsystems CMS) is used to illuminate the sample andcollect the emitted fluorescence. The emission wavelength is separated from theexciting light by using a suitable dichroic mirror. The diameter of the emittedbeam is increased by using a tube lens to make it comparable with the size ofthe chip of the camera. A bandpass filter (Semrock and 515/30 ET, Chroma)is used to clean the emission wavelength. A filter wheel is placed between theemission port of the microscope and the camera to switch between the filtersin order to image various emission wavelengths simultaneously. It consists of abandpass filter (ET Bandpass 515/30, Chroma) and a longpass filter (EdgeBa-sic long pass 561, Semrock). The CCD camera with an intensifier is used as adetector to form the image. The chip of the camera is maintained at -25◦C toreduce the background noise.

Another solid-state laser of wavelength 561 (Cobolt Jive), was coupledinto the same beam path and a dichroic mirror (561/10 ZET, Chroma) was usedfor its clean up. This laser was used to visualize the FRET donor. The twolasers were selected using mechanical shutters. A custom designed Lab Viewprogram was used to control the laser shutter. This allowed the simultaneousimaging of two fluorophores. The laser shutters (Thorlabs) were controlled by acustom-built Lab View program and the laser intensity was controlled by neu-tral density filters for each laser beam. Omicron laser controller software wasused to operate the 488 laser. The Lightfield software was used to operate thecamera and to modulate the laser excitation. To maintain the proper physio-logical conditions for the cells while imaging, the microscope is equipped witha stage (Tokai Hit) where the temperature, CO2 concentration and humidity

7.2 Methods 119

can be controlled. To control the movement of the microscope stage during theDNA damage induction, SwitchBoard software (Märzhäuser Wetzlar) was used.

In order to induce DNA damage to the cells, an NIR pulsed laser (Mira900,Coherent) was also coupled into the setup. A pulse picker was introduced intothe beam path to control the frequency of the NIR laser excitation. FLIM imag-ing was done with 70 MHz frequency of the camera with the intensity gain of90 and pixel number of the image was set 512 x 512. Each image consists of 12frames, each frame acquired at the phase angles of 0 °, 131 °, 262 °, 196 °, 327 °,65 °, 295 °, 33 °, 164 °, 98 °, 229 °, 0 ° with the same order. The images are ac-quired at random order of phases to eliminate the effect of photobleaching. [301]

The shutter and the camera was controlled using custom-built application withLabView program for the fast and synchronized imaging. Dr. Peyman Zerakdeveloped the script for controlling and automation of the shutters.

Before acquiring the FLIM images, the setup was calibrated with thestandard fluorophores of known fluorescence lifetimes. We used a solution ofCoumarin 6 in ethanol (100 uM) and a solution of Rhodamine 6G in water (10uM) and the dyes were excited using modulated 488 nm laser with the powers of10 µW and 100 µW, respectively. The two dyes have lifetimes of 2.5 ns and 3.9ns respectively. [302,303] The images were corrected for the background and sortedfor the different phases with the Lightfield software. The simFCS software wasused to determine the lifetimes of the reference fluorophores before imaging thecells. The data were analyzed and the fluorescence intensity, phase lifetimes andmodulation lifetimes were derived using the MATLAB scripts. The MATLABscripts were initially developed by Dr. Waldemar Schrimpf (group of Prof.Dr. Lamb, Ludwig-Maximilians-Universität, Munich, Germany). The derivedlifetimes were used to calculate the FRET efficiencies in different experimentalconditions.

7.2.5.2 Laser Scanning Confocal Microscopy

Laser scanning confocal microscopy was performed using a TCS SP8 uprightmicroscope from Leica Microsystems with a 63 X oil immersion objective lens


(1.40 NA HCX PL APO CS, Leica Microsystems). The LAS X software was usedfor acquiring the images. For live cell imaging, the Ap4 analog was excited by 488nm laser and the emitted fluorescence signal was detected with a photomultipliertube (PMT) from 500 nm to 530 nm. LysoTracker Red was excited with 552nm wavelength and emission was collected at 560-600 nm using appropriatefilter sets. The autophagy sensor (L3B-RFP) was excited using 552 wavelengthlaser and the fluorescence was collected between 570 nm and 600 nm. Dextran-rhodamine B was also excited by 552 wavelength and the emission was collectedfrom 580 nm to 620 nm. Transmission images were also acquired using thetransmitted light detector after illumination with 638 nm laser. Images wereacquired with a resolution of 1024 × 1024 pixels, at the scanning speed of 100lines per second and the line average of 4. Images were processed using ImageJ software.

7.2.6 Statistics and Data Analysis

The statistical evaluation of the data was done using GraphPad Prism 7.05software. Statistical significance was determined by using one-way analysis ofvariance (ANOVA), Tukey’s multiple comparison test or the unpaired t-test fortwo groups, as appropriate, separately in each figure. The confidence intervalwas set as 95 % for all tests. The degree of significance was given as ns (notsignificant) p > 0.05, * p < 0.05, ** p < 0.01, and *** p < 0.001. All data areexpressed as mean ± standard deviation (SD) or the ± standard error of themean (SEM).

7.2.6.1 Colocalization Analysis

All the confocal images were processed and evaluated quantitatively using im-age analyzing software Image J (National Institutes of Health, Bethesda, USA).The colocalizaton analysis was done by image J plugin, Coloc 2. It performsthe intensity correlation over all the pixels and calculates Pearson’s correlationcoefficient. The two images from different channels or the two ROIs were se-lected and the algorithm was run. The numerical results and the scatter plotswere obtained as output. Quantification of colocalized spots was done by Fiji’splugin, ComDet v.0.50. The plugin finds the bright intensity spots that are colo-

7.2 Methods 121

calized in a color composite image. The appropriate particle size and intensitythreshold to be detected were entered manually and the localized spots weredisplayed in overlayed rectangles. The result table with the various parametersand coordinates of all the particles was provided.

Corrected total cell fluorescence (CTCF) was calculated using Image J.After selecting a region of interest, integrated intensity and the area were mea-sured. From a different area, background fluorescence intensity was calculated.CTCF was calculates by the following equation:

Integrated density - (Selected area x Mean background fluorescence) (7.2)

Abbreviations

6-4 PPs Pyrimidine 6-4 photoproducts

AAA ATPases Associated with diverse cellular Activities

ADP Adenosine diphosphate

ARTD1 ADP-ribosyltransferase diphtheria toxin-like 1

ATeam Adenosine 5-Triphosphate indicator based on Epsilon subunitfor Analytical Measurements

ATP Adenosine triphosphate

BER Base excision repair

BiFC Bimolecular fluorescence complementation

BRCA1 Breast cancer type 1 susceptibility protein

BRET Bioluminescence resonance energy transfer

cAMP Cyclic adenosine monophosphate

CAT Carboxyl-terminal catalytic domain

CATS Calcium channel associated transcriptional regulation

CCD Charge Coupled Device

CCHC Cys-Cys-His-Cys

CDKs Cyclin dependent kinases

124 ABBREVIATIONS

CMOS Complementary Metal Oxide Semiconductors

CPD Cyclobutane pyrimidine dimers

CSA and CSB Cockayne syndrome factors A and B

DBD DNA binding domain

DNAP DNA polymerase

DOTAP 1,2-Di-(9Z-octadecenoyl)-3-trimethylammonium propane methyl-sulfate

DSBR Double strand break repair

DSBs DNA strand breaks

EDA-ATP 2’(3’)- O-[N-(2-aminoethyl)carbamoyl]ATP

ELK Eukaryotic-like kinase

ePK Eukaryotic protein kinase

ERCC1 Excision Repair Cross-Complementing Rodent Repair Defi-ciency, Complementation Group 1

ESCRT Endosomal sorting complex required for transport

FCS Fluorescence correlation spectroscopy

FISH Fluorescence in situ hybridization

FLIM Fluorescence lifetime imaging microscopy

FRAP Fluorescence recovery after photobleaching

FRET Förster resonance energy transfer

GFP Green fluorescent protein

GPI Glycosyl-phosphatidylinositol

GPI glycosyl phosphatidylinositol

H1299 Human lung carcinoma cell line

HEK Human embryonic kidney

HeLa Cervical cancer cells from Henrietta Lacks

HEPES 2-[4-(2-Hydroxyethyl)piperazin-1-yl]ethanesulfonic acid

ABBREVIATIONS 125

HR Homologous recombination

IDL Insertions and deletions loops

IR Infrared radiations

IRF Total internal reflection fluorescence

ISC Intersystem crossing

LC3 Light chain 3

LDH Lactate dehydrogenase

LED Light emitting diodes

LSCM Light scanning confocal microscopy

MANT-ATP 2’(3’)- O-(N-methylanthraniloyl)-ATP

MAPK Mitogen-activated protein kinase

MAPK Mitogen-activated protein kinases

MMR Mismatch repair

MRE11 Meiotic recombination 11 homolog 1

MVB Multivesicular body

NAD Nicotinamide adenine dinucleotide

NADH Nicotinamide adenine dinucleotide hydrogen

NBS1 Phosphopeptide-binding Nijmegen breakage syndrome protein1

NER Nucleotide excision repair

NF-κB Nuclear factor kappa-light-chain-enhancer of activated B cells

NHEJ Non-homologous end-joining

NIR Near Infrared

NLS Nuclear localization signal

NudT2 Nudix hydrolases

PAHs Polycyclic aromatic hydrocarbons

PALM Photoactivated localization microscopy

126 ABBREVIATIONS

PARG Poly (ADP-ribose) glycohydrolase

PARG Poly(ADP-ribose) glycohydrolase

PARP Poly (ADP-ribose) Polymerase

PCNA Proliferative cell nuclear antigen

PCR Polymerase chain reaction

PEP Phosphoenolpyruvate

PFA Paraformaldehyde

PFK Phosphofructokinase

PI3K Phosphatidylinositol 3-kinases

PKA Protein kinase A

PMTs Photomultiplier tubes

PTM post-translational modifications

RFP Red fluoescent protein

ROS Reactive oxygen species

RPA Replication protein A

SAM S-adenosyl methionine

SEM Standard error of the mean

SIM Structured illumination microscopy

SnFRs Neurotransmitter sensing fluorescent reporters

SSBs Single strand breaks

STED Stimulated Emission Depletion Microscopy

STORM Stochastic Optical Reconstruction Microscopy

SVPD Snake venom phosphodiesterase

TCSPC Time correlated single photon counting

TdT Terminal deoxynucleotidyl transferase

TL Thin layer chromatography

TMR Tetramethylrhodamine

ABBREVIATIONS 127

TRASE Time-resolved ATPase sensor

TUNEL Terminal deoxynucleotidyl transferase dUTP nick end labeling

UV Ultraviolet radiations

WFM Widefield microscopy

XPC Xeroderma pigmentosum, complementation group C

XPF Xeroderma Pigmentosum Group F-Complementing Protein

XPG Xeroderma Pigmentosum Complementation Group G

XRCC X-ray repair cross-complementing protein 4

YFP Yellow fluorescent protein

128 ABBREVIATIONS

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