Ryan -Thèse Martin

De

Keywords: DNA damage, (6-4) photoproduct, DNA repair, photolyase, photorepair, photoactivation, UV-Vis femtosecond spectroscopy, photoinduced electron transfer

We studied the light-induced reactions of the (6-4) photolyase, a flavoenzyme of the cryptochrome/photolyase family that repairs the UV-induced (6-4) photodamage in DNA with the aid of blue light. We studied this photorepair reaction as well as the light-induced cofactor reduction called photoactivation that the enzyme uses to bring itself to a repair-active state in the (6-4) photolyase from Xenopus laevis. We have studied the photoactivation of the FAD ox cofactor of the enzyme using femtosecond polarised transient absorption spectroscopy. We observed a sub-picosecond electron transfer (~400 fs) after excitation of the FAD ox cofactor. We were able to characterise a tryptophan residue as the electron donor. We sought to differentiate the spectroscopically identical but differently oriented tryptophan residues within the protein's photoactivation site by transient anisotropy measurements. Our results suggest that the photoactivation mechanism is not fully compatible with the mechanism thought to be conserved among photolyases: an electron transfer mechanism via electron hopping along a chain of three highly conserved tryptophan residues. Using series of single turnover flashes, we have found that the repair reaction proceeds by a successive two-photon mechanism. The first photon converts the (6-4) lesion into a metastable intermediate X, the lifetime of which is ~2 min. Absorption of a second photon within the lifetime of X results to the restoration of intact nucleobases. In light of our findings, the reaction was also studied by femtosecond transient absorption spectroscopy. We have observed the electron transfer from the excited FADH -to the (6-4) lesion in 247 ps.

Etudes sur les mécanismes primaires de la photolyase (6-4) : photoactivation et photoreparation de l'ADN Résumé :

Ce travail concerne les mécanismes photoinduits d'une flavoprotéine appartenant à la famille des cryptochromes et photolyases (CPF) : la photolyase . En utilisant de la lumière bleue cette protéine répare un dommage de l'ADN induit par les UV, le photoproduit . Nous avons étudié cette photoreparation ainsi qu'une autre réaction photoactivée que la photolyase utilise pour réduire son cofacteur flavine, la photoactivation. Nous avons faits nos études en utilisant la photolyase de Xenopus laevis. Nous avons étudié la photoréduction du chromophore FAD ox de l'enzyme par spectroscopie d'absorption transitoire femtoseconde polarisée. Nous avons observé un transfert d'électron ultrarapide (~400 fs) après excitation du chromophore FAD ox . Nous avons caractérisés un résidu tryptophane comme réducteur. Nous avons cherché de distinguer entre les différents tryptophanes présents dans le site de photoactivation par des mesures d'anisotropie. Les résultats obtenus suggèrent que le mécanisme de transfert d'électron dans la photolyase n'est pas compatible avec le mécanisme supposé chez les photolyases qui consiste des transferts d'électron de long d'une chaine de trois résidus tryptophane. Grâce à des trains d'impulsions courtes, nous avons démontré que la photolyase (6-4) répare l'ADN par un mécanisme à deux photons successif. Le premier photon sert à convertir le défaut en un intermédiaire métastable, X, qui a une durée de vie de ~2 min. Un second photon absorbé pendant cette durée de vie permet d'achever le cycle de réparation. Nous avons aussi étudié la réaction par la spectroscopie UV-Vis femtoseconde. Nous avons observés le transfert d'un électron provenant de FADH -au défaut en 247 ps.

Mots clés : Défauts ADN, photoproduit , réparation de l'ADN, photolyase, photoréparation, photoactivation, spectroscopie UV-Vis femtoseconde, transfert d'électron photoinduit Acknowledgement This thesis will not see its completion without the help of people whose support and encouragement have always been present along the way. To them, a debt of gratitude is due.

First and foremost I thank my adviser Pascal Plaza. Thank you for your confidence in me. Most especially, thank you for your patience with me all these years. It was not a smooth sail throughout but I know it is for my own betterment. I sincerely thank you for your guidance and instruction. I thank Lahouari Krim, Thomas Gustavsson, and Marten Vos for graciously accepting the responsibility of judging this work. I also thank you for your comments and pieces of advice during my defence.

I would also like to thank my colleagues for always being present when I needed them. Their support is not limited to the technical aspect of my thesis but as well as daily motivational words and precious company during breaks. I thank Fabien Lacombat for the technical help in my experiments especially during the extended hours of acquisition, for patiently explaining to me the workings of our experiments, and most especially the daily pep talk I get especially during hard times. To Agathe Espagne who despite her busy schedule is always present to help with any questions I pose and for performing some experiments in my stead. And to Nadia Dozova also for always being generous in giving me a helping hand whenever I get stuck treating my data and most especially tea.

To our collaborators: Klaus Brettel for his confidence in me and for his precious pieces of advice and support. To Junpei Yamamoto for entrusting me with his precious samples and for his encouragement. And to Pavel Müller to whom all credit with W D is due, I thank you for your friendship and even technical help when needed.

I big thank you to the people who taught me precious scientific techniques that were of great use in my thesis. To Raphaël Guerois of CEA for his aid and instruction on homology modelling. To the group of Chris

Bowler at ENS Biology especially Gerald Zabulon for teaching me the indispensable skill of protein expression and purification.

On a more personal note I would like to take this chance to express my gratitude to people who have been behind me through these past few years: my long suffering friends and family. This work is dedicated to you.

As a consequence of ultraviolet (UV) irradiation, DNA molecules obtain lesions in the form of highly mutagenic and cytotoxic cross-links between two adjacent pyrimidine bases. Two major forms of such lesions exist: the cyclobutane pyrimidine dimer (CPD) and the pyrimidine(6-4)pyrimidone photoproduct (or simply photoproduct). In a vast number of organisms, Nature has devised an elegant way to reverse these light-induced damages by using blue light and photoactive repair enzymes called photolyases. There are also two types of photolyase; one for each type of lesion [1].

Since the discovery of the photolyase several decades ago, knowledge concerning the repair mechanism of the CPD photolyase has significantly advanced to the point of now having a general picture of how the protein repairs the CPD lesion with the aid of light. On the other hand, the (6-4) photolyase repair mechanism is less understood owing in part to the reaction's complexity. No agreement on the repair mechanism has been reached in literature [1][2][3][4][5][6].

Photolyases bind a flavin chromophore that allows them to use light as an energy source in performing repair; more specifically UV and blue light. Flavins have three redox-active forms: the oxidised form (FAD ox ), the semiquinone form (FADH • ), and the fully reduced form (FADH -). The form of the flavin that is active in the photorepair of photolyase is the fully reduced form (FADH -) [1].

Whenever the flavin is not in its repair-active fully reduced state, photolyase may reduce the cofactor via a mechanism also induced by light. This mechanism is called photoactivation. The photoactivation of the CPD photolyase has been extensively studied by the groups of K. Brettel and M. Vos using time-resolved spectroscopy [7]. The reduction of the flavin in the CPD photolyase involves the transfer of electrons through several specific redox-active residues in the vicinity of the chromophore.

My work aims to elucidate the mechanism by which the phtotolyase from Xenopus laevis repairs the photoproduct. It also aims to study the photoactivation mechanism that photolyase employs to render it capable of (6-4) photoproduct repair. When I started my thesis, the details regarding these two reactions are not known. The repair mechanism of photolyase was at the time debated in literature. Several models to explain the mechanism have been put forward by both theoreticians and experimentalists yet it appears that the only general agreement is that the repair proceeds by the absorption of a single photon. Regarding the mechanism of photoactivation, the exact pathway of electron transfer that the protein uses to reduce its oxidised cofactor has not been explicitly demonstrated. The studies performed on these subjects aim to specifically address these two questions. Since the nature of the species involves the study of short-lived species created after the absorption of light, the main tool implemented was the technique of time-resolved spectroscopy.

The studies were performed using the combined techniques of time-resolved pump-probe spectroscopy, organic synthesis, and biochemistry to characterise the two photoreactions. To implement the said multi-disciplinary approach to the problem, collaboration has been made with the groups of Klaus Brettel (iBiTecS, CEA Saclay) and Junpei Yamamoto (Osaka University). The studies resulting from this collaboration is presented in this manuscript. This manuscript is divided into four chapters.

The first chapter is a bibliographic review of the status of the literature concerning the photolyase. It begins with the products of UV irradiation on DNA molecules: pyrimidine dimers.

Following is a review on how Nature deals with such problems in certain organisms: a look the photolyases. A review of a group of closely related proteins called cryptochromes is also presented. The next section focuses on the various theories found in literature on the mechanism of photorepair and photoactivation by photolyase. This section gives emphasis on the mechanisms of photolyase. The first chapter is concluded with a brief review of the enzyme cofactor with which the photolyase is able to effectuate DNA repairthe FAD molecule.

The second chapter is dedicated to the mechanism by which photolyase reduces its flavin cofactor to its active form for photorepair with the aid of light. We begin the first chapter by a study of our protein whose structure we obtained via structure modelling due to the unavailability of its crystal structure. The analysis of the protein's structure is essential to the subject of photoactivation since the reaction involves specific residues in the protein. In this chapter, we present the results of our spectroscopic characterisation of the photoreduction of the oxidised flavin cofactor of photolyase.

Therein, we identify the primary electron donor and we attempt to determine the path of electron transfer within the protein during reduction.

The studies performed on the 64PP repair by photolyase are presented in the third chapter.

We commence by answering a fundamental question of the repair mechanism: how many photons does it need for photolyase to repair the (6-4) photoproduct? We believe that answering this question will help in the elucidation of the real mechanism of photorepair. The reaction was further studied using femtosecond broadband transient absorption spectroscopy. To date, this is the second ultrafast study of the photorepair mechanism. Also in this chapter is our photorepair study of a lesser studied form of the photoproduct: the T(6-4)C photoproduct.

Introduction

It is well known that UV radiation induces particularly harmful lesions in DNA. These lesions are in the form of dimers between two adjacent pyrimidine bases on a DNA strand. There are two major types: the cyclobutane pyrimidine dimer (CPD) and the pyrimidine(6-4')pyrimidone photoproduct (or simply photoproduct). A review on the UV-induced lesions and their biological effects can be found in Cadet 2005 [1]. In many organisms, the UV-induced lesions can be repaired by photoactive enzymes called photolyases.

A number of reviews on the subject of photolyases are available. Among these, the 2003 review by A. Sancar is the most widely cited and relatively the most comprehensive [2]. Other general reviews on the subject of photolyases can be found in Weber [3], Falciatore et al. [4], and Müller et al. [5]. A more specialised review on the spectroscopy of photolyases can be found in Kao et al. [6] Schleicher et al. [7]. On the subject of photorepair, the following reviews will be helpful: Essen [8], Brettel et al. [9], Kneuttinger et al. [10].

Photolyases are flavin-containing proteins (flavoproteins) that use blue light to repair UV induced DNA damages [2]. Noteworthy, photolyases directly repair the lesions and restore the original intact nucleobases in a single light-powered operation. This is called photorepair. Such process contrasts with the more complex, step-wise and ATP-consuming mechanisms of nucleotide excision repair or base excision repair [11]. Let us note that placental mammals, among which humans, are devoid of photolyases [12]. Two types of photolyases exist that are specific to the repair of either the CPD dimer or the photoproduct. These are the CPD photolyase and the photolyase. The latter type is the subject of this thesis. Photolyases are related to another family of flavoproteins, called cryptochromes, that mostly do not repair DNA but have different signalling functions. More information on cryptochromes be found in the reviews by Partch et al. [13], Cashmore et al. [14,15], and Lin et al. [16,17]. Photolyases and crytochromes together form the "cryptochrome photolyase family" or simply CPF.

Proteins belonging to this family generally have a high degree of homology, in both sequence and structure, and all are photoactive. This chapter aims to provide the state of the art of research in the field of photoinduced reactions of (6-4) photolyases, namely DNA photorepair and the so-called photoactivation reaction. Indeed, for photorepair to take place, the enzyme must be in a particular active state. The state of the photolyase is defined by the redox state of its flavin cofactor. The catalytically-active form of the flavin is the fully reduced form: FADH -. Photoactivation refers to the reduction of the flavin cofactor into its fully reduced form whenever it is oxidised. This process is also mediated by light and is hence called photoactivation.

Photoactivation and photorepair are the main subjects of my thesis.

In the first section, the various lesions of DNA induced by UV light on adjacent pyrimidines will be introduced. We will be presenting how these various mutagenic lesions are formed in DNA.

The second part of the chapter is devoted to the CPF family of proteins. This section reviews the general structure of the CPF proteins, using the photolyase as a model, as well as the different classifications of the CPF proteins.

The third section reviews the photoactivation and photorepair mechanisms of the photolyases in detail. We will also present a brief look at the photoreduction of cryptochromes. A bibliographic review of the different models proposed for the mechanism of these two reactions will be presented.

The photochemistry of the CPF family of proteins is based on its flavin cofactor that acts as the photon receptor in the protein. This chapter closes by outlining the basic photophysical properties of flavin cofactors found in CPF proteins.

UV-Induced DNA Lesions

The exposure of DNA molecules to UV light gives rise to reactions between two adjacent pyrimidine nucleobases present in the sequence. These photoinduced reactions lead to the dimerisation of these nucleobases. The resulting species is called a DNA photoproduct or photolesion. These products are highly mutagenic and exist in two major types: the cyclobutane pyrimidine dimer (CDP) and the pyrimidine(6-4')pyrimidone photoproduct (64PP).

Cyclobutane pyrimidine dimer, CPD

A [2π + 2π] cycloadition between two C5-C6 double bonds of the pyrimidine ring gives rise to a species known as the cyclobutane pyrimidine dimer or CPD. Here, the two pyrimidine rings are bridged together by the four-membered cyclobutane ring. The CPD accounts for the majority of the formed pyrimidine dimers after UV irradiation in cells [1]. A [2π + 2π] cycloaddition between the C5-C6 and the C4 carbonyl (imine in the case of cytosine in its keto form) will form a dimer bridged by a four-membered heterocyle (oxetane for thymine and azetidine for cytosine). This photoreaction between an alkene and a carbonyl group is also called the Paternò-Büchi reaction [18]. The oxetane/azetidine species, being thermodynamically unstable, will split into a structure consisting of two rings connected by a single covalent bond. The 5'part of the lesion will be a pyrimidine derivative whose C6 position is covalently attached to the C4 position of the 3' pyrimidone. In their study of the formation of the (6-4) photoproduct, Marguet and Markovitsi showed that the oxetane has a lifetime of 2 ms in water [19]. This final product is called the pyrimidine pyrimidone photoproduct or simply the (6-4) photoproduct or (6-4)PP. Although the (6-4)PP is the less abundant photoproduct formed, it is believed to be more cytotoxic than the CPD [20,21].

T-T [2 + 2] cycloaddition CPD dimer

T-T T(ox)T T(6-4)T [2 + 2] cycloaddition

Figure 3. Formation of the photoproduct between two thymine bases Further illumination of UV light promotes another photoreaction in the (6-4) photoproduct. The 3' pyrimidone ring undergoes a photo-induced isomerisation (via 4π electrocyclisation) to yield the highly strained Dewar isomer [22].

UV-A/B UV-C

Photolyases and Cryptochromes

Presented here is a review of the general properties of the two major classes of photolyases and the related group of cryptochromes.

Structure of the cryptochrome and photolyase family 1.2.1.1 Phylogenetics

We begin our discussion of the cryptochrome and photolyase family (CPF) by a description of its members from the point of view of evolution. To aid us in this task phylogenetic analysis, i.e. the construction of a phylogenetic tree, allows to visualise the evolutionary links between many different types of CPF proteins. Recent phylogenetic trees, reproduced from Geisselbrecht et al. [23] and Juhas et al. [24] are shown in Figure 5.

A Figure 5. Phylogenetic tree of several representative proteins of the cryptochrome photolyase superfamily. A: figure reproduced from Geisselbrecht et al. [23]; B: figure taken from Juhas et al. [24] (the scale bar represents amino acid substitutions per site).

Figure 5 shows that the CPF proteins are further divided into several subfamilies. For the photolyases there are the CPD photolyase (categorised into class I, II and III) and the (6-4) photolyase.

For cryptochromes there are the plant cryptochromes, animal cryptochromes, CryPro cryptochromes, and the DASH cryptochromes. Each of these will be discussed in the succeeding chapters.

As we will see in later sections, there is often a high degree of homology among members of the CPF family [25,26]. In the event of the discovery of a new CPF protein, its classification has essentially been based up to now on its ability of photorepairing DNA. If it is capable of it, it will be classified as a photolyase. Otherwise, it will be categorised under the more generic term of cryptochrome [2,27]. It should however be noted that some cryptochromes also posses photorepair capabilities and are referred to as dual-function cryptochromes [28,29].

We have performed a similar phylogenetic analysis for the proteins that are mentioned in this thesis, particularly in Chapter 3. The resulting phylogenetic tree is given in Annex A.

Primary and secondary structure

Photolyases are proteins whose primary structure consists of a sequence of around 400-600 amino acids. The 150 amino acids of the protein's C-terminal exhibit the highest degree of homology among photolyases [30]. For this reason, this region has initially been thought to be the flavin cofactor binding region of the protein [31]. This fact has been established by the succeeding studies on the protein's structure [32][33][34][35]. There is a limited degree of homology between plant/animal photolyases and microbial B photolyases [36]. From this difference stems the division of the photolyases into three classes: class I for microbial photolyase, class II for the photolyases of higher multicellular eukaryotes but also is archaebacteria, eubacteria, and unicellular algae [37] and the class III photolyases which are more phylogenetically related to plant cryptochromes [27].

The crystal structures of some photolyases have been determined: Class I CPD photolyases from E. coli (here abbreviated EcCPD; PDB code: 1DNP [31]) and Anacystis nidulans (1TEZ [38]), Class II CPD photolyase from Methanosarcina mazei (2XRY [39]) and from Oryza sativa (3UMV [40]), photolyases from Drosophila melanogaster (3CVU [41]), Arabidopsis thaliana (3FY4 [42]) and Agrobacterium tumefaciens (4DJA [43]). As far as cryptochromes are concerned, let's mention: Cry1

from Drosophila melanogaster (4GU5 [44]) and Mus musculus (4KOR [START_REF] Czarna | Structures of drosophila cryptochrome and mouse cryptochrome1 provide insight into circadian function[END_REF]), Cry2 from Mus musculus (416E [START_REF] Xing | Scffbxl3 ubiquitin ligase targets cryptochromes at their cofactor pocket[END_REF]), Cry3 from Arabidopsis thaliana (2J4D [START_REF] Klar | Cryptochrome 3 from arabidopsis thaliana: Structural and functional analysis of its complex with a folate light antenna[END_REF]), CryDASH from Synechocystis sp. (1NP7 [START_REF] Brudler | Identification of a new cryptochrome class: Structure, function, and evolution[END_REF])

and Arabidopsis thaliana (2IJG [START_REF] Huang | Crystal structure of cryptochrome 3 from arabidopsis thaliana and its implications for photolyase activity[END_REF]). We will start describing the well-known structure of EcCPD in some details. It is represented in Figure 6. EcCPD is a 471-residue long globular protein. EcCPD contains two distinct regions: the Nterminal and C-terminal domain [34]. Amino acids 1 to131 comprise the N-terminal α/β domain. This domain contains 6 α-helices and 5 β-sheets. The C-terminal α domain is composed of residues 201-471.

These residues only form helices (14 α-helices and 5 π-helices). The two domains are connected by a loop formed by the residues 132 to 200 that winds around the α/β domain. The C-terminal domain contains the amino acids that interact with the FAD chromophore. The residues that interact with a second chromophore, MTHF (see §1.2.1.3), are shared by both the C-terminal and N-terminal domains. The chromophores are non-covalently linked to the protein. The chromophores will be discussed separately in the next section.

The crystal structure of the 543 residue long (6-4) photolyase from D. melanogaster is shown in Figure 6 (PDB: 3CVU [41]). The secondary structures found in the sequence highly resemble that of CPD photolyase. The domain of residues 1 to170 contains 6 α-helices, 5 β-sheets, and 1 π-helix. The region containing the residues 190 to 504 contains 16 α-helices and 5 π-helices. The latter region also contains the residues that interact with FAD. No second chromophore was found in the crystal structure.

Chromophores

Photolyases carry two non-covalently-bound chromophore cofactors. The first one is always FAD and the second is either methenyltetrahydrofolate (MTHF), 8-hydroxy-7,8-didemethyl-5-deazariboflavin (8-HDF) or 6,7-dimethyl-8-ribityllumazine. The structures of these chromophores are shown in Figure 8. FAD is the essential cofactor for the DNA binding and photocatalytic functions of photolyases [START_REF] Sancar | Action mechanism of escherichia-coli DNA photolyase .1. Formation of the enzyme-substrate complex[END_REF][START_REF] Jorns | Identification of a neutral flavin radical and characterization of a 2nd chromophore in escherichia-coli DNA photolyase[END_REF]. The repair-active form of FAD is its fully reduced anionic state, FADH -. The flavin cofactor is discussed separately in Section 1.4.

The second chromophore is not necessary for photocatalysis and does not affect the enzymesubstrate binding [36]. This chromophore is referred to in literature as the antenna chromophore. The presence of MTHF can be identified by the presence of an intense band at 385 nm (ε = 25 000 M -1 cm -1 ) [36]. Under limiting light (i.e. low photon density), the antenna chromophore enhances the repair rate of up to 100-fold depending on the wavelength used because of its higher absorptivity than FADH -. The antenna chromophore absorbs a photon and it transfers the excitation energy to the flavin via a Förster mechanism. MTHF is contained mainly in microbial class I photolyases and DASH cryptochromes. 8-HDF (also called F 0 ), on the other hand, is mostly found in archaeal photolyase although it has been recently reported to be present in the photolyases of higher eukaryotes and plays [START_REF] Kiontke | Structural and evolutionary aspects of antenna chromophore usage by class ii photolyases[END_REF]. The MTHF chromophore also dissociates readily from photolyase and is present in substoichiometric amounts in purified samples [START_REF] Hammalvarez | Role of enzyme-bound 5,10methenyltetrahydropteroylpolyglutamate in catalysis by escherichia-coli DNA photolyase[END_REF][START_REF] Hammalvarez | The folate cofactor of escherichia-coli DNA photolyase acts catalytically[END_REF][START_REF] Hammalvarez | The presence and distribution of reduced folates in escherichia-coli dihydrofolate-reductase mutants[END_REF]. In purified EcCPD samples, MTHF is present only in 20-30 % of the total photolyase present [START_REF] Hammalvarez | The presence and distribution of reduced folates in escherichia-coli dihydrofolate-reductase mutants[END_REF]. However, MTHF can be reconstituted by incubating the apoenzyme in solutions with high MTHF concentrations [START_REF] Johnson | Identification of the 2nd chromophore of escherichia-coli and yeast DNA photolyases as 5,10methenyltetrahydrofolate[END_REF]. The antenna chromophore in (6-4) photolyase has not yet been found in purified samples although MTHF was observed in a semi-purified sample [START_REF] Todo | Flavin adenine dinucleotide as a chromophore of the xenopus (6-4)photolyase[END_REF]. A binding pocket that strongly binds F 0 has been identified in D. melanogaster photolyase [START_REF] Glas | The archaeal cofactor f-0 is a light-harvesting antenna chromophore in eukaryotes[END_REF].

In 2013, the crystal structure of the photolyase from the prokaryote Agrobacterium tumefaciens has been resolved by Zhang et al. (PDB: 4DJA [43]). This marks the first time that a prokaryotic (6-4) photolyase was identified. It has been assumed before that the (6-4) photolyase was restricted to eukaryotic cells. Interestingly, the enzyme contained 6,7-dimethyl-8-ribityllumazine as the antenna chromophore as well as an Fe-S cluster. To date, no other study on prokaryotic (6-4) photolyase has appeared.

Photolyases

We briefly examine in the following the two general classes of photolyases: the CPD photolyase and the (6-4) photolyase. A more detailed review of their photoinduced reactions is given in Section 1.3.

CPD photolyase

The photolyase repairing the CPD lesion is called the CPD photolyase. This is due to the fact that this is the first photolyase identified in 1949 [START_REF] Kelner | Effect of visible light on the recovery of streptomyces griseus conidia from ultraviolet irradiation injury[END_REF]. Due to this, it had been referred to simply in literature as photolyase [2]. Despite the high degree of homology among CPD photolyases (generally confined to the FAD binding site), animal and plant photolyases show a limited homology to microbial photolyases [12]. From this limit stemmed the further classification of CPD photolyases into class I (microbial) and class II (amimal) subfamilies [START_REF] Yasui | A new class of DNA photolyases present in various organisms including aplacental mammals[END_REF]. A third class of photolyases (class III) has been identified in the bacteria C. crescentus [27]. Although also closely related to class I photolyase, class III photolyase is more closely related to plant cryptochromes.

(6-4) photolyase

The discovery of the photolyase that repairs the (6-4) photoproduct was only reported in 1993 by Todo et al. [START_REF] Todo | A new photoreactivating enzyme that specifically repairs ultraviolet light-induced (6-4)photoproducts[END_REF]. Though the existence of the (6-4) photoproduct was known before the discovery of the photolyase, the only substrate known to photorepaired was the CPD [START_REF] Todo | A new photoreactivating enzyme that specifically repairs ultraviolet light-induced (6-4)photoproducts[END_REF]. As far as DNA repair is concerned the photolyase has two photocatalytic functions: (i) it is the enzyme responsible for the photorepair of the (6-4) photoproduct and (ii) it photoconverts the Dewar valence isomer back to the photoproduct form thus promoting repair [START_REF] Glas | Crystal structure of the t(6-4)c lesion in complex with a (6-4) DNA photolyase and repair of uv-induced (6-4) and dewar photolesions[END_REF][START_REF] Glas | DNA (6-4) photolyases reduce dewar isomers for isomerization into (6-4) lesions[END_REF].

The photolyase has been believed to only exist in eukaryotes until the identification of the A.

tumefaciens photolyase was reported by Zhang et al. in 2013 [START_REF] Zhang | Crystal structure of a prokaryotic (6-4) photolyase with an fe-s cluster and a 6,7-dimethyl-8-ribityllumazine antenna chromophore[END_REF]. The most recent phylogenetic analyses of the CPF proteins have not yet revealed subfamilies among the (6-4) photolyases [24,[START_REF] Kiontke | Structural and evolutionary aspects of antenna chromophore usage by class ii photolyases[END_REF][START_REF] Oliveri | The cryptochrome/photolyase family in aquatic organisms[END_REF].

Cryptochromes

A class of proteins related to the photolyases is called the cryptochromes. They bear high sequence homology and structural similarity to photolyases. Cryptochromes are also blue light photoreceptors but lack, in general (see below), the photorepair capabilities of photolyases.

Cryptochromes play a role in signal relay in both plants and animals. In 1993, the first gene coding for a cryptochrome in Arabidopsis thaliana was discovered by Ahmad and Cashmore [START_REF] Ahmad | Hy4 gene of a-thaliana encodes a protein with characteristics of a blue-light photoreceptor[END_REF]. We have seen in Figure 5 that cryptochromes are classified into several subfamilies: plant cryptochromes, animal cryptochromes, DASH cryptochromes, and CryPro [23,[START_REF] Oliveri | The cryptochrome/photolyase family in aquatic organisms[END_REF]. We shall see that the classification of the CPF proteins evolve continuously; new classifications are identified. A recent and more detailed review of this subject can be found in Kiontke et al. [START_REF] Kiontke | Structural and evolutionary aspects of antenna chromophore usage by class ii photolyases[END_REF], Oliveri et al. [START_REF] Oliveri | The cryptochrome/photolyase family in aquatic organisms[END_REF] and Juhas et al. [24]. We shall look at the different subfamilies briefly in the following.

Plant cryptochromes

As reported by M. Ahmad, et al. the cryptochrome 1 of A. thaliana (AtCry1) is one of the photoreceptors taking part over light mediated responses [START_REF] Ahmad | Cryptochrome blue-light photoreceptors of arabidopsis implicated in phototropism[END_REF]. More specifically, it inhibits hipocotyl elongation [15]. AtCry1also acts as a regulator of gene expression during plant photomorphogenesis [START_REF] Wang | Direct interaction of arabidopsis cryptochromes with cop1 in light control development[END_REF][START_REF] Zeugner | Lightinduced electron transfer in arabidopsis cryptochrome-1 correlates with in vivo function[END_REF].

Animal cryptochromes

In 1996, Todo, et al. identified, on separate occasions, two blue light photoreceptor proteins in humans: hCry1 and hCry2. These proteins have a very high degree of homology (48% and 73 % identity respectively) to the (6-4) photolyase of D. melanogaster [START_REF] Hsu | Putative human blue-light photoreceptors hcry1 and hcry2 are flavoproteins[END_REF][START_REF] Todo | Similarity among the drosophila (6-4)photolyase, a human photolyase homolog, and the DNA photolyase-blue-light photoreceptor family[END_REF]. This marks the discovery of the first animal cryptochrome.

Though similar in structure to the plant cryptochrome described earlier, animal cryptochromes play a part in the regulation of the circadian rhythm or the biological clock. They act as the photoreceptors that synchronise the molecular circadian clock to solar light thereby allowing the perception of day and night [START_REF] Sancar | Cryptochrome: The second photoactive pigment in the eye and its role in circadian photoreception[END_REF]. In mammals however, the circadian rhythm is not governed by cryptochromes but by melanopsin, a protein using retinal (vitamin A) instead of flavin for photoreception [START_REF] Sancar | Regulation of the mammalian circadian clock by cryptochrome[END_REF].

The ability of migratory animals (e.g. birds, monarch butterfly, etc) to perceive the earth's magnetic field is also due to cryptochomes present in their eyes [START_REF] Rodgers | Chemical magnetoreception in birds: The radical pair mechanism[END_REF][START_REF] Gegear | Animal cryptochromes mediate magnetoreception by an unconventional photochemical mechanism[END_REF].

DASH cryptochromes

In 2000, the first description of a bacterial cryptochrome was reported by K. Hitomi, et al. and this led to the birth of a new cryptochrome family: the DASH cryptochromes (CryDASH) [START_REF] Brudler | Identification of a new cryptochrome class: Structure, function, and evolution[END_REF][START_REF] Hitomi | Bacterial cryptochrome and photolyase: Characterization of two photolyase-like genes of synechocystis sp pcc6803[END_REF]. The abbreviation DASH represents the first organisms where this cryptochrome was found: Drosophila, Arabidopsis, Synechocystis, and Homo.

The DASH cryptochrome from Synechocystis is a regulator of genetic transcription by acting as a transcription repressor [START_REF] Brudler | Identification of a new cryptochrome class: Structure, function, and evolution[END_REF]. Like photolyases, the Synechocystis CryDASH has also been reported to bind to DNA and possesses a weak DNA photorepair activity [29,[START_REF] Hitomi | Bacterial cryptochrome and photolyase: Characterization of two photolyase-like genes of synechocystis sp pcc6803[END_REF]. It has also been shown that DASH cryptochromes from A. thaliana, X. laevis, and V. cholerae can specifically repair single stranded DNA containing a CPD lesion [START_REF] Selby | A cryptochrome/photolyase class of enzymes with single-stranded DNA-specific photolyase activity[END_REF][START_REF] Pokorny | Recognition and repair of uv lesions in loop structures of duplex DNA by dash-type cryptochrome[END_REF]. More recently in 2013, Castrillo et al. reported that the DASH cryptochrome from the fungi Fusarium fujikuroi participates in the regulation of the fungi's secondary metabolism [START_REF] Castrillo | Light-dependent functions of the fusarium fujikuroi cryd dash cryptochrome in development and secondary metabolism[END_REF].

Dual-function cryptochromes

In 2009, a CPF protein, PtCPF1, from the diatom P. tricornutum has been reported to posses both receptor and DNA repair capabilities [28]. A similar dual-function CPF proteins, OtCPF1 has also been identified in the alga O. tauri [29] in 2010. Phylogenetic analysis shows that PtCPF1 and OtCPF1 are closely related to photolyases..

CryPro cryptochromes

In 2012, a new subfamily of cryptochromes was identified by Geisselbrecht et al. [23] in proteobacteria and cyanobacteria. This new subfamily is called the CryPro cryptochromes. The authors analysed CryB from Rhodobacter sphaeroides belonging to this class. This protein is a photosynthesis gene expression regulator. The crystal structure of the protein (PDB: 3ZXS) revealed that the protein utilizes 6,7-dimethyl-8-ribityllumazine as an antenna chromophore and it contains an Fe-S cluster. In terms of the cofactor content, it bears similarity to the prokaryotic (6-4) photolyase reported by Zhang et al. [START_REF] Zhang | Crystal structure of a prokaryotic (6-4) photolyase with an fe-s cluster and a 6,7-dimethyl-8-ribityllumazine antenna chromophore[END_REF].

The reaction mechanisms of photolyase

DNA photolyases recognise the pyrimidine lesions in DNA and bind to them. Unlike DNA binding proteins that recognise a specific sequence when binding to DNA (e.g. polymerases), photolyase binding is not sequence specific. That is to say, it is only governed by the presence of the pyrimidine dimer [2]. For instance the specific binding constant (K S ) of the CPD photolyase to T<>T lesion in DNA (<> symbolises a CDP lesion) is 10 -9 M and the non-specific binding constant for undamaged DNA (K NS ) is ~10 -4 M. This translates to a selectivity factor (K NS /K S ) of ~10 5 [START_REF] Husain | Binding of escherichia-coli DNA photolyase to a defined substrate containing a single t reversible t dimer[END_REF]. The rate of substrate binding (k on )

has been reported to be in the range of 10 7 M -1 s -1 for CPD photolyase [36] while the dissociation constant

(k off ) is 0.05 s -1 [2].
Catalysis by photolyase follows the classic Michaelis-Menten kinetics:

E + S → ES → EP → E + P
The only deviation from, albeit, an important one is the ES to EP conversion is absolutely lightdependent. The reaction begins by the light-independent binding of the enzyme to the substrate. During this step, the lesion is flipped-out of the DNA double helix and into the repair cavity of photolyase to obtain the enzyme-substrate complex, ES. The repair takes place with the aid of light then the photolyase dissociates from the product [2].

Apart from the repair of pyrimidine dimers, photolyase is capable of another photoinduced reaction called photoactivation. Photoactivation is a mechanism that the enzyme can use to transform an oxidised flavin cofactor (FAD ox or FADH • ) into the repair-active FADH -form. This is such since Sancar reports that even photolyases produced by cells cultured in the dark contain FADH - [2]. Hence it is possible that photolyase employs another mechanism, independent of light, that can reduce the flavin whenever it is in its oxidised form.

Photorepair

Photorepair by CPD photolyase

The classical scheme of CPD photorepair by CPD photolyase has been reviewed by Sancar in 2003 [2]. It is described in the following. The photorepair begins with the ES complex of the CPD lesion and the photolyase. The first step involves the absorption of a blue photon by MTHF (if present) or by FADH -. Photon absorption by the former is followed by the transfer of excitation energy to the latter.

This transfer is achieved over a distance of 16.8 Å at a rate of FRET). The now excited FADH -transfers an electron to the CPD lesion at a rate of 7 10 9 to 2 10 10 s -1 over 5-10 Å [START_REF] Antony | Theoretical study of electron transfer between the photolyase catalytic cofactor fadh(-) and DNA thymine dimer[END_REF][START_REF] Hahn | Conformation of the flavin adenine dinucleotide cofactor fad in DNA-photolyase: A molecular dynamics study[END_REF][START_REF] Medvedev | DNA repair mechanism by photolyase: Electron transfer path from the photolyase catalytic cofactor fadh(-) to DNA thymine dimer[END_REF]. The CPD radical then splits between the C5 -C5' and the C6 -C6' bonds via an asynchronous concerted [2+2] cycloreversion to generate a pyrimidine and a pyrimidine radical. A back transfer of the excess electron from the pyrimidine to the flavin to gives rise to two intact pyrimidine bases and the regeneration FADH -. This final step completes the repair cycle. The repair yield was also believed to be close to 100% [START_REF] Kim | Time-resolved epr studies with DNA photolyase -excited-state fadh(0) abstracts an electron from trp-306 to generate fadh-, the catalytically active form of the cofactor[END_REF]. The mechanism is illustrated in Figure 9. In 2011, Thiagarajan et al. reported a study of the kinetics of CPD dimer splitting [START_REF] Thiagarajan | Kinetics of cyclobutane thymine dimer splitting by DNA photolyase directly monitored in the uv[END_REF]. The authors directly monitored the restoration of the intact bases during repair. The main difference from the previous report by MacFarlane and Stanley is an improvement in experimental conditions that allowed them to better monitor the repair. The authors report claim that the They observed a biexponential rise in the signal that they measured for the restoration of the intact thymine with time constants of 200 ps and 1.5 ns in the kinetics. They assigned these constants to the repair of the dimer (200 ps) and to the return of an electron to the flavin to regenerate FADH -. The latter has not been observed by MacFarlane and Stanley. In a separate report, Thiagarajan et al. also quantified the quantum yield of repair to be 50% [START_REF] Thiagarajan | DNA repair by photolyase: A novel substrate with low background absorption around 265 nm for transient absorption studies in the uv[END_REF].

In their experiments, the authors were not able to detect the asynchronous splitting of the C5-C5' and the C6-C6' bonds. This mechanism is shown in Figure 11. on the CPD repair [START_REF] Liu | Dynamics and mechanism of cyclobutane pyrimidine dimer repair by DNA photolyase[END_REF]. In their version of the mechanism, shown in Figure 12, the authors claim that an electron from the excited flavin is injected to the lesion in 250 ps. This step is followed by the sequential split of the C5-C5' and C6-C6' bonds in less than 10 ps and in 90 ps respectively. This is followed by the return of an electron to the flavin to regenerate FADH -in 700 ps. Of note in their mechanism is the presence of a slow back electron transfer (to the flavin) occurring in 2.4 ns in the absence of the C6-C6' bond split. It has been pointed out by Brettel and Byrdin however that the back electron transfer should only be moderately slow and not as slow as 2.4 ns as Liu et al. had claimed [START_REF] Brettel | DNA photolyase: Is the nonproductive back electron transfer really much slower than forward transfer[END_REF][START_REF] Zhong | Reply to brettel and byrdin: On the efficiency of DNA repair by photolyase[END_REF]. They report that all electron transfer in A. nidulans CPD photolyase involves the mediation of the adenine. This electron transfer does not take place via a two-step transfer but via a superexchange mechanism. The authors also believe that the adenine is responsible for the fast splitting of the CPD dimer that they observed in their 2011 report [START_REF] Liu | Dynamics and mechanism of cyclobutane pyrimidine dimer repair by DNA photolyase[END_REF].

Photorepair by (6-4) photolyase

The (6-4) photoproduct, being more complex than the CPD lesion, is expected to be repaired in a more complex mechanism distinct from the CPD repair. Direct splitting of the 6-4' bond would indeed result instead to two damaged bases. To repair the (6-4) photoproduct, both the splitting of the C6 -C4

bond and the transfer of a hydroxyl group is required. Owing to this complexity, the repair mechanism of the (6-4) photolyase is debated upon in the literature; as we will see in the following. The repair yield of the (6-4) photoproduct has been reported to be 11% [START_REF] Hitomi | Role of two histidines in the (6-4) photolyase reaction[END_REF]. This low yield may reflect the complexity of the repair mechanism. It is of note that the (6-4) photolyase is also responsible for the repair of the Dewar isomer of the (6-4) photoproduct [START_REF] Glas | DNA (6-4) photolyases reduce dewar isomers for isomerization into (6-4) lesions[END_REF]. The repair of the Dewar lesion takes place by converting the Dewar photoproduct back to the (6-4) form and is followed by the repair of the (6-4) photoproduct.

In 1993, Todo, et al. reported photoproduct repair by a photolyase from Drosophila melanogaster [START_REF] Todo | A new photoreactivating enzyme that specifically repairs ultraviolet light-induced (6-4)photoproducts[END_REF] This has been the first reported case of the (6-4) photoproduct-specific repair in the literature. Soon thereafter, the existence of this enzyme in other organisms was discovered [START_REF] Kim | Characterization of (6-4)photoproduct DNA photolyase[END_REF][START_REF] Nakajima | Cloning and characterization of a gene (uvr3) required for photorepair of 6-4 photoproducts in arabidopsis thaliana[END_REF].

Analysis of the primary structure of this then new photolyase showed high identity with the CPD photolyase [25,26,[START_REF] Todo | Similarity among the drosophila (6-4)photolyase, a human photolyase homolog, and the DNA photolyase-blue-light photoreceptor family[END_REF]. Further structural analysis revealed even more similarities with the CPD photolyase such as their common flavin cofactor [START_REF] Todo | Flavin adenine dinucleotide as a chromophore of the xenopus (6-4)photolyase[END_REF]. From these observations, it has been speculated that the two photolyases partly share the same repair mechanism as far as the bond splitting from a fourmembered ring moiety is concerned. We have seen that the (6-4) photoproduct is more complex than the CPD so an additional step is required before one can see the parallel with CPD. This step is the conversion of the (6-4) photoproduct to the oxetane bridged dimer. This step takes place in the dark (i.e.

without the aid of light). This early repair scheme is shown in Figure 13. Upon binding of the enzyme to the lesion site, the (6-4) photoproduct is flipped-out into the active site of the enzyme and repair takes place [START_REF] Zhao | Reaction mechanism of (6-4) photolyase[END_REF]. In a step distinct from the CPD repair, the (6-4) photolyase thermally converts the For convenience, we will rename His358 and His354 as HisA and HisB, respectively. This will allow us to easily draw parallels with other works that will be described. The initially unprotonated HisA abstracts a proton from the hydroxyl group of the pyrimidine ring on the 5' side of the lesion.

Concomitantly, an initially protonated HisB donates a proton to 3'-N on the 3' pyrimidone ring of the lesion. The net effect is the creation of a charged pair that drives the formation of the oxetane. This (cf. Figure 7) [41]. In their report, they showed that the T(6-4)T photoproduct was present in the active site as an unmodified species; no oxetane structure was detected. The protein crystal was then illuminated with visible light and crystallographic analysis showed the presence of the restored T-T bases. In 2009, Domratcheva et al., using quantum mechanical (QM) calculations, deemed that the mechanistic pathway involving the formation of oxetane is energetically unfavourable due to its high activation barrier of 33.2 kcal/mol [START_REF] Domratcheva | Electronic structure of (6-4) DNA photoproduct repair involving a non-oxetane pathway[END_REF]. They proposed a non-oxetane pathway involving the direct transfer of the hydroxyl group to the pyrimidone (5') ring. This mechanism is shown in Figure 17.

A B

According to Li et al., the photoexcited FADH -in photolyase injects an electron to the lesion in 225 ps (characteristic time constant of a stretched exponential) creating a radical anion species of the photoproduct. This radical state can undergo a back electron transfer in 50 ps to give back the initial ground state; effectively rendering the reaction futile. The determining step in this mechanism is a successful proton transfer from a protonated HisB to the radical anion state that occurs in 425 ps. This proton transfer gives rise to a transition state (Structure III in Figure 18) whose structure is evocative of the oxetane moiety. On this basis, this mechanism has been named the "transient oxetane" mechanism.

This transient state serves as the bridge to enable the transfer of the hydroxyl group from the 5' to the 3' base. After this hydroxyl transfer, bond rearrangement gives rise to the repaired bases. This is followed by the return of the proton to HisB and the return of the electron to the flavin to regenerate FADH -. This occurs in more than 10 ns. Of note, in this reaction HisB is in its protonated form.

In contrast with all preceding proposals, Sadeghian ) there is a higher probability for HisA to be protonated. A proton transfer from HisA is possible. This proton transfer will result to the protonation of HisB to yield the histidine protonation configuration which is the configuration that is considered by the majority of reports found in literature. Subsequently, a proton transfer to HisB is possible to yield the consensus state where HisB is protonated.

Another theoretical study on the protonation states of the histidine was reported by Shahi and Domratcheva in 2013 [107]. Their computational studies maintain that for the active site to remain active, HisB should be in its neutral state. They also claim that the protonated histidine HisBH + is capable of serving as an electron acceptor. This implies that if HisB were protonated, it would compete with the (6-4) photoproduct over the electron coming from the excited FADH -. This explains why, according to the authors, HisB should be unprotonated for repair to proceed.

Photorepair of the T(6-4)C photoproduct

Another form of the (6-4) photoproduct is the dimer formed between a thymine as the 5' base and cytosine as the 3' base. This form of the lesion is called the T(6-4)C photoproduct and is shown in Figure 21. All studies presented previously were centred on the T(6-4)T photoproduct and studies on the T(6-4)C form are scarce.

T(6-4)C

Figure 21. T(6-4)C photoproduct

There is a lesser amount of published work on the repair of the T(6-4)C photoproduct. It is likely attributable to the difficulty in the synthesis of the substrate for this lesion. The procedure applied to the laboratory synthesis of the T(6-4)T lesion cannot be directly applied to the synthesis of the T(6-4)C lesion as reported by Mizukoshi et al. [108]. In contrast to its in vitro synthesis, the T(6-4)C lesion is more easily formed in cells due to its higher formation quantum yield over the T(6-4)T lesion: 0.74 for T(6-4)C over 0.5 for T(6-4)T as reported by Lemaire and Ruszicka in 1993 [109]. More recently in 1997, Douki et al. reported that the product yield (i.e., final product over starting material) for the T(6-4)C photoproduct is almost three times higher than the T(6-4)T lesion [20].

Zhao et al. in 1997 report that photolyase does not discriminate between the T(6-4)C and T(6-4)T forms of the photoproduct. Both are bound and repaired in the same manner by the classical repair mechanism shown in Figure 13.

In 2009, Glas et al. resolved the crystal structure of the T(6-4)C lesion bound by D. melanogaster photolyase [START_REF] Glas | Crystal structure of the t(6-4)c lesion in complex with a (6-4) DNA photolyase and repair of uv-induced (6-4) and dewar photolesions[END_REF]. Apart from the structural differences arising from the substitution of thymine by cytosine, no difference was observed with the crystal structure of T(6-4)T photoproduct bound to D. melanogaster photolyase that the same group published earlier (cf. Figure 7). Once more, this refutes the classical repair mechanism where the oxetane (or in the case of T(6-4)C, azetidine) is formed upon enzyme binding.

In the literature the mechanism of T(6-4)C repair is assumed to be the same as that for T(6-4)T photoproduct [START_REF] Glas | Crystal structure of the t(6-4)c lesion in complex with a (6-4) DNA photolyase and repair of uv-induced (6-4) and dewar photolesions[END_REF]110].

Photoactivation

Photoactivation of CPD photolyase

As mentioned above, the active form of the flavin for DNA photorepair is its fully reduced state FADH -. FADH -is prone to oxidation by molecular oxygen found in cells. It is oxidised into either of its two inactive forms: FAD ox or FADH • . In vitro, the purified enzyme contains the inactive form caused by cell lysis and protein purification in aerobic conditions [111,112]. To regenerate FADH -photolyases are capable of a photoinduced reduction mechanism * . This reaction is also called photoactivation.

In 1986, P. F. This was performed in order to identify which residue(s) was/were involved in the photoactivation. Using transient absorption spectroscopy, they were able to identify W306 as this said residue [115]. This study however was not able to unambiguously identify which form of the tryptophan is involved. [START_REF] Kim | Time-resolved epr studies with DNA photolyase -excited-state fadh(0) abstracts an electron from trp-306 to generate fadh-, the catalytically active form of the cofactor[END_REF], addressed this ambiguity by performing time-resolved EPR spectroscopy on EcCPD and its isotopically labelled W306F mutant [START_REF] Kim | Time-resolved epr studies with DNA photolyase -excited-state fadh(0) abstracts an electron from trp-306 to generate fadh-, the catalytically active form of the cofactor[END_REF]. They found that the protonated radical cation tryptophan (WH •+ ) form is the primary reducing oxidised transient species.

The crystal structure of EcCPD was resolved in 1995 by Park et al. [31]. From this structure the distance between W306 and the flavin was found to be 1.34 nm. Furthermore, two more tryptophan residues (W382 and W359) were discovered between the flavin and W306. W306 is the residue closest to the protein surface and W382 is closest to the flavin with W359 in between. The three tryptophans are shown in Figure 22. For brevity, W382, W359, and W306 are renamed W A , W B , and W C , respectively. (Sancar 2003) that even cells that were cultured in the dark already contained FADH -Hence there must also be another mechanism, which is light dependent, that is capable of converting the flavin to its active form. † Of historical note is an alternative pathway proposed by Park et al. in their 1995 report. They proposed a pathway where electron transfer involves a nearby phenylalanine residue as the primary electron donor followed by W C . This had been "supported" by studies of Saxena et al. in 2004 [116] and Wang et al. in 2005 [117] both from the group of D. Zhong.

However, it appears this view has been abandoned in 2008 [118].

An even older view is the direct long-range electron transfer between an excited FADH -and W C . This involved the conversion of an excited FADH -(in doublet state) to a quartet by intersystem crossing. The excited quartet has a lifetime long enough to abstract an electron from W C . This view was maintained until 2003 by Sancar in his review.

excitation wavelengths were chosen to selectively excite the FADH • . They showed that the photoreduction of FADH • proceeds as follows (and diagrammed in Figure 22):

1. Upon excitation of the FADH • cofactor, the excited state decays in 30 ps by abstracting an electron from WH A producing FADH -and WH A •+ .

In less than 10 ns, WH

WH C

•+ gives off a proton to the solution in 300 ns. takes place in less than 30 ps.

We will refer to the mechanism that was elucidated up to this point as the "Brettel-Vos mechanism". It is summarised in Figure 24, along with energetic data reported by Byrdin et al. [127]. ) to be at 0.8 ps for EcCPD and 0.5 ps for At64. These sub-ps timescales for the primary reduction are much faster than that for FADH In their model, the initial electron transfer from W A to the flavin occurs in 0.8 ps. This is in competition with electron transfer from the adenine that occurs in 19 ps, and to a lesser extent with the nearby W A' that occurs in 80 ps. WH A The reaction scheme is presented in Figure 26. In 2014, Liu et al. followed up their study by reporting the photoactivation mechanism of semireduced EcCPD photolyase [137]. They reported the initial electron transfer from W A to take place in 50 ps. This is followed by the reduction from W B that takes place in 2.5 ps and the subsequent electron transfer from W C in 100 ps. This scheme is illustrated in Figure 27. The first time constant (initial electron transfer from W A ) is comparable to the value of 45 ps that we see in the Brettel-Vos mechanism illustrated in Figure 24. This is the same for the 2.5 ps transfer from W B which was estimated to be less than 9 ps by the Brettel-Vos model. The difference is the rate for the last step: estimated to be less than 30 ps in the model and was reported to be 100 ps by Liu et

Photoactivation of (6-4) photolyase

In contrast to the CPD photolyase, the (6-4) photolyase contains the fully oxidised flavin. To be converted to FADH -, FAD ox needs a proton and two electrons. The reduction is in fact a sequential process that involves the prior reduction of FAD ox to FADH • and then to FADH -. The photoactivation of (6-4) photolyase (i.e. photoactivation from the oxidised flavin) has not been as well characterised as that of the CPD photolyase. This may be due to the fact that CPD and (6-4) photolyases sharing the same structural features it was assumed that they share the same reaction mechanisms.

In 2002, a paper of the photoactivation of (6-4) photolyase was published by Weber et al. [138].

The authors studied the photoreduction of the FAD ox cofactor using time-resolved EPR spectroscopy. The authors concluded that the species observed 1. Then, the more distant from the flavin is the oxidised tryptophan the slowest was assumed to be the charge recombination process, and the slowest its reduction by the next tryptophan on the chain. Since (6-4) photolyase contains FAD ox in its in vitro resting state (compared to FADH • in CPD photolyase), photoactivation to FADH -requires protonation in addition to electron transfer. The proton donor for (6-4) photolyase is currently undetermined. It is believed that the residue closest to the N5 position of the flavin plays a role in the protonation. Most CPF proteins have an aspartic acid or asparagine residue in this position and insect cryptochrome have a cysteine [140]. The residue at this position is known to determine the stability of FAD •-. In proteins where this position is occupied by aspartic acid or asparagine, FAD In the same timescale, partial proton transfer from the tryptophanyl radical to the nearest environment is hypothesized. Plant cryptochromes possess a high concentration of histidine residues in their flexible loops as compared to photolyases. Fast proton transfer from the tryptophanyl radical to a deprotonated histidine would promote the stabilisation of the FAD

The flavin cofactor of the (6-4) photolyase

Flavins are one of the most common cofactors found in biological systems. It exists in various redox forms as illustrated in Figure 30: FAD ox , FADH • , and FADH -.

. We have already seen that the active form of the flavin present in photolyases is FADH -[111].

This section is dedicated to presenting the various properties of FAD that will aid us in the study of the photorepair and photoactivation mechanisms of (6-4) photolyase.

Flavins in solution

At neutral pH 7, FAD has three redox forms: the fully oxidised FAD ox , the semi-reduced FADH • , and the fully reduced FADH -. Although unstable in solution form, we have seen in the previous sections that FADH • is the form of FAD in the in vitro resting state of EcCPD. FADH -on the other hand is easily oxidised to FAD ox in the presence of molecular oxygen.

The absorption spectrum of FAD ox exhibits two peaks in the visible region centred at 445 and 375 nm. The molar absorption coefficient (ε) at the absorption maximum is higher than 10 4 M -1 cm -1 , which is indicative of π→π* transitions [156]. The aqueous solutions of FAD ox exhibit an intense yellow-green fluorescence whose spectrum contains a single peak at around 520 nm. The absorption spectrum of FADH -on the other hand does not show any maximum. The spectrum is characterised by a strong absorbance in the UV region that decreases in the visible until it reaches zero at 500 nm. The emission spectrum has a single broad peak at around 440 nm. These spectra are shown in Figure 31.

Flavins in photolyase

FAD is found in CPF proteins in a closed, often called U-shaped, conformation [36]. It can be found in three redox states: FAD ox , FADH • , and FADH -. As has been presented in the earlier sections, the repair-active form is the fully reduced FADH -. Under physiological conditions, FAD is synthesised as FAD ox and is introduced in the apoenzyme as such [36]. It has been presented previously that photolyase converts the non-active forms to the active FADH -via a light-induced reduction called photoactivation.

However, even cells grown in the dark contain FAD in its fully reduced FADH -form. This indicates that photoactivation is not the only reaction by which photolyase converts the flavin to FADH -in vivo.

The chromophore is tightly bound by non-covalent interactions with 14 amino acid residues present in the α domain of the protein and are well conserved in the CPF family of proteins [31]. In E. coli and A. nidulans photolyase, FAD is released only after mild denaturation of the protein [32,161].

Conclusion

In this chapter, the current state of knowledge on the photoinduced reactions of photolyases has been presented. In particular, emphasis has been given to the mechanisms of DNA repair and cofactor activation. This provides a ground upon which the studies in the succeeding chapters will be based upon.

Our work has been focused on the reactions of (6-4) photolyase.

Our first objective is to establish the repair mechanism of the (6-4) photoproduct by photolyase. We have seen that the mechanism of repair is debated upon in the literature. In our studies on the repair mechanism, we aimed to answer the fundamental question on the number of excitations required for repair to take place. Studies of the mechanism can be divided into two groups: single photoexcitation and double photoexcitation mechanisms. The majority of the published works generally assume that repair proceeds with a single photoexcitation. Though divided in the exact mechanism, the common point of most is that the active flavin chromophore (FADH -) injects an electron only once to the (6-4) lesion and that this is enough to promote repair. All experimental work on photorepair assumes that this is the case. On the other hand, one theoretical work suggests that the mechanism involves two electron injections from FADH -. We believe that addressing this question is essential in order to elucidate the correct repair mechanism.

Our second goal is to study the photoactivation mechanism of the fully reduced cofactor in photolyase. The photoactivation of the fully oxidised flavin is much less studied than the semiquinone form found in EcCPD photolyase. We have seen in the models presented that the photoactivation mechanism of CPF proteins vary from one type of protein to the other. Hence, the current model for the CPD photoyase may not hold true for the (6-4) photolyase.

We addressed the issues presented using our combined techniques of transient absorption

Introduction

It has been presented in Chapter 1 that the flavin cofactor of the (6-4) photolyase may exist in various redox states but only one form, the fully reduced FADH -, is the active form for photorepair.

During the purification of the enzyme, the flavin chromophore is oxidised to FAD ox due to exposure to oxygen. This oxidised form does allow photorepair. Aside from photorepair, photolyase is capable of performing another photoinduced reaction, a photoreduction that converts FAD ox to FADH • , and then FADH • to FADH -in two separate photoreactions. This mechanism is called photoactivation [1].

The photoactivation mechanism of semi-reduced E. coli CPD photolyase (noted EcCPD, found to contain FADH • after purification) has been proposed by the groups of K. Brettel and M. Vos to involve a chain of three conserved tryptophans. Photoreduction of the flavin is believed to be achieved by the hopping of electrons along the tryptophan triad, from the first Trp to the flavin, then from the second Trp to the first Trp and finally from the third Trp to the second Trp [2]. The first step is the rate limiting step and the succeeding transfers were not experimentally resolved. Recent studies on EcCPD by Liu et al. in 2013 [3] showed that the active site for photoactivation contains two additional Trp residues that also play a role in the electron transfer mechanism. The studies are reviewed in greater detail in Chapter 1.

In contrast to the semi-reduced photolyase, the first electron transfer is not the rate limiting step for the fully oxidised photolyase. In 2008, Kao et al. observed a sub-picosecond time constant in the photoactivation of oxidised photolyases: EcCPD and A. thaliana (6-4) photolyase [4]. They assigned this to the first electron transfer to an excited FAD ox from the tryptophan triad. In 2010 Brazard, et al.

performed broadband femtosecond transient absorption spectroscopy on a (6-4) photolyase from

Ostreococcus tauri (Ot64) [5]. The authors measured three time constants describing the photoreduction of FAD ox to FAD •-. They assigned each of these to one elementary step of electron hopping along the conserved tryptophan triad, accompanied by a competing charge recombination process. The sub-ps component was assigned to reduction of the flavin by the proximal Trp and the following components to electron transfers between Trp's. It was claimed that the more distant from the flavin is a tryptophanyl radical, the slower is its reduction by the following Trp and the reduced flavin. This electron hopping mechanism, akin to the Brettel-Vos mechanism, has not been explicitly shown/proven. The authors were not able to justify that the kinetic phases they saw in their isotropic spectra were due to an electron donation by a distinct tryptophan in the triad. At this point we ask ourselves whether it is valid to assign each kinetic phase to an electron transfer step; as had been done by the others before. If so, we must be properly demonstrated. If not, then they must be explained. By doing so, a more precise image of the photoactivation mechanism would be obtained. This is the main goal of our study of the photoactivation mechanism. The work presented in this chapter is an improvement in the work of Brazard et al. Here we use better tools that would allow us to follow the electron flow from one residue to the other. This would allow us to answer the question of assigning an observed kinetic phase to each electron transfer step.

As we have seen in Chapter 1, the photoactivation of the (6-4) photolyase is less frequently studied in literature. In our work, we studied the photoactivation of the (6-4) photolyase from Xenopus laevis (noted Xl64). This photolyase contains FAD ox after purification. Xl64 photolyase is not very studied in literature. There are two existing reports on Xl64 photoactivation: EPR study reported by Weber et al. [6] and the FTIR study by Yamada et al. [7]. None of these reports investigate the Xl64 photoactivation in the sub-ps timescale.

We must also note that Liu et al., in their 2013 study of the photoactivation of oxidised EcCPD [3], were able to identify new redox-active residues that play a role in the electron relay during the reaction. We wish to identify if we can identify the same residues in the photoactivation site of Xl64.

Furthermore, we want to know if it is possible to identify new redox-active residues that might play a role in the photoactivation of Xl64. For instance, at least one of the photoactivation studies on Xl64 explicitly mentions that a tyrosine residue is involved in the reaction. This chapter opens with just that.

The first part of this chapter deals with the structural analysis of the active site of the photoactivation in Xl64. This analysis was done in order to identify the potential redox active residues in the vicinity of the reaction site. We begin by a sequence alignment of Xl64 with other CPF proteins, which allowed the identification of the "canonical" tryptophan triad. Sequence analysis is followed by the identification and localisation of all potentially redox-active residues in the vicinity of the photoactivation active site. An X-ray crystallographic structure of the enzyme not being available at the time of this work, a 3D model was obtained using homology modelling.

The second part of the chapter is devoted to the ultrafast polarised pump-probe spectroscopic study of the photoactivation reaction of the X. laevis (6-4) photolyase. We begin by identifying the kinetic phases in both the isotropic and polarised spectra. We then move on to our goal of what to assign theses kinetic phases to. We have also mentioned that one of our objectives is to establish the flow of electrons through the different tryptophans and possibly other residues that participate in photoactivation. As the various tryptophan residues have similar spectral signatures in their oxidized state, telling them apart based on the transient absorption spectra is not possible. These residues are however oriented differently with respect to the flavin. Looking at transient absorption anisotropy is expected to help differentiating them. Polarised measurements (i.e., parallel and perpendicular transient absorption spectra) allowed us to construct anisotropy spectra and kinetics. We first used an experimental scheme where the parallel and perpendicular spectra were independently measured. This scheme has the drawback of adding up the errors caused by independent fluctuations of the excitation laser energy during each measurement. We tried to circumvent this problem by recording the parallel and perpendicular spectra simultaneously. The findings are discussed in light of the homology model mentioned above.

The last part of this chapter is dedicated to the investigation of the photoactivation reaction on a longer timescale, using nanosecond real-time transient absorption spectroscopy. We in particular focused on measuring transient absorption anisotropy data in order to determine the nature of the oxidized tryptophan (tryptophanyl) found beyond the observation window accessible with the ultrafast set-up, in the ns regime. The findings were also discussed in light of the homology model. The deprotonation kinetics of the tryptophanyl was also investigated. These experiments were performed in the laboratory of Klaus Brettel in CEA Saclay.

Structural analysis of the X. laevis (6-4) photolyase

To identify the potential electron donors around the flavin cofactor of the photolyase, a crystal structure is ideally needed. To date, no such structure for Xl64 exists. To approximate this, we resorted to the construction of a 3D model using the known primary structure of Xl64 (NCBI ref: NP_001081421.1 [8]). This 3D structure is based from the existing structures of related proteins and is called a homology model. The homology model of the X. laevis (6-4) photolyase was obtained using the SWISS-MODEL platform [9][10][11][12]. The model was created by using D. melanogaster (6-4) photolyase (PDB: 3CVU [13]) as the templatethe structure from which the model is based. The two proteins share 57% identity * and 78% similarity † in sequence. This procedure is elaborated in Chapter 4, Section 6 Homology modelling.

This has been done with the assistance of Raphaël Guerois of the Laboratoire de Biologie Structurale et

Radiobiologie, iBiTec-S, CEA Saclay.

Sequence alignment

To start with, Table 1 reproduces an alignment of the primary structures of several CPD photolyases (PHR), (6-4) photolyases (64PHR), and cryptochromes (Cry), made by Biskup et al. [14] (supplementary information). These proteins come from the following organisms: Escherichia coli (Ec, bacterium), Anacystis nidulans (An, cyanobacterium), Thermus thermophilus (Tt, eubacterium), We performeda phylogenetic analysis of these CPF proteins and the result is shown in Annex A. The analysis was performed mainly to serve as supplementary information on the classification these proteins. The results of this analysis are not discussed here.

Sulfolobus
Table 1. Sequence alignment of several proteins from the cryptochrome/photolyase family. Letters in white correspond to identical residues and letters in red with a blue frame correspond to high similarities. The tryptophan residues discussed in the text have been highlighted and labelled.

Looking at Table 1, we can observe the conservation of the "canonical" tryptophan triad and W319 (in agreement with a previous identification [15]. Among the two additional tryptophans reported by Liu et al [16] to be active in the photoreduction of EcCPD (W384 and W316; generically noted W A' and W B' , respectively), one is also conserved across all the series, namely W B' ; it is W329 for Xl64. W A' is only present in the photolyases and animal cryptochromes; it is W398 for Xl64. The alternative electronic donor reported by Biskup et al. [14] and Immeln et al. [17], noted W C' , is present only in StPHR (archaea), SsCRYD (bacteria), AtCry1 and AtCry2 (plant), and CcCry (algae); Xl64 does not have it. Upon closer inspection of the structure, the third tryptophan of the conserved triad (W319) is found to lie at close distance (4.4 Å) to a fourth tryptophan (W370), also located on the surface of the protein (see below). Since this distance is comparable to the intermolecular distances between the tryptophans of the conserved triad, one can hypothesize that W370 might, potentially, extend the electron transfer chain and be the final electron donor. To date, this residue has not been mentioned in any of the existing literature on photoactivation to the best of our knowledge. Neither have we found its mention in the involvement on any other reactions of the CPF proteins. Thus, if proven, the concept would be of novelty. It is however not excluded that the residue does not have any catalytic function at all. This could also be an explanation for its absence in present literature on CPF proteins. It is referred to as W D in generic terms and is drawn in orange. Based on the sequence alignment of Table 1, let us note that W D is found only in the (6-4) photolyases and cryptochromes of animals. In the existing model of photoactivation [19] the final electron donor is a residue, tryptophan or tyrosine, exposed to the outside environment of the protein and capable of being reduced by an external reductant. The fulfilment of this criterion by W D can easily be verified with the homology model by looking at the protein surface. Shown in Figure 2 is a rendering of the protein surface of Xl64 photolyase. The blue region is the part of the surface where W C is found and the orange region is where W D is.

Homology model

Shown in green in

Both residues are accessible from the surface of the protein and are hence capable of being reduced by an external reductant. This implies that, for the photoactivation reaction, W D is also capable of having the terminal role in the electron relay along the tryptophan chain. However, based on Figure 1, it cannot be excluded that the relay simply ends with W C . As to whether or not X. laevis (6-4) photolyase uses W D during photoactivation cannot be deduced on looking at the structure alone. This question was also addressed in our time-resolved spectroscopic experiments. The last part of this chapter is dedicated specifically to answer this question.

Spectroscopic study of the wild type X. laevis (6-4) photolyase

Our study aims at understanding the primary events of the photoactivation reaction of Xl64. We seek to identify the events directly following the excitation of the oxidised flavin chromophore. This entails the identification of the transient species produced in the excitation. We then wish to follow the transfer of electrons within the protein to determine the pathway by which FAD ox is reduced. We have seen in the models of photoactivation that the mechanism is claimed to involve the hopping of electrons along a chain of tryptophans (cf. Chapter 1). In the structural analysis of the homology model, we have seen that there are six tryptophan residues in the proximity of the flavin, as shown in Figure 1, that are capable of playing a role in the relay of electrons to FAD ox . In order to establish the primary events of the photoactivation reaction, we excited the oxidised flavin cofactor and followed the events following this excitation for 1.5 ns. We followed the evolution of the system by measuring its transient absorption spectrum at multiple time delays after the excitation. It is expected that the spectroscopic signature of the oxidised radicals corresponding to the six tryptophans of interest would be similar due to their identical chemical nature. We tried to differentiate between them by looking at a property that is dependent on their orientations: anisotropy. Let us note that two types of transient absorption setups have been used, both measuring the polarised transient absorption spectra. One method employs recording the parallel and perpendicular spectra one after the other. As the parallel and perpendicular spectra are measured separately, the fluctuations of the spectra due to the (small) energy fluctuations of the laser are independent from each other. The anisotropy spectra that is calculated from these measurements (se Eq. 3.2 in §3.2.4 below) therefore suffers from adding up (quadratically) these decorrelated fluctuations, which gives rise to maximal uncertainties. We tried to eliminate this source of error by employing a method that implements the simultaneous measurement of the parallel and perpendicular signals to obtain spectra that are well correlated with regards to the pump-induced signal fluctuations. An added benefit of this method is the shortened acquisition time since the two spectra are measured simultaneously. The acquisition of both the parallel and perpendicular spectra is effectively reduced by half thereby allowing more spectra to be measured within the sample's lifetime. This translates into a better signal-to-noise ratio. The drawback on the simultaneous method is that the experimental setup is significantly more complex and more difficult to control. It in fact appeared that a systematic error slightly tainted the data, which we tried to correct.

The photoactivation of

All this is explained in more detail in Chapter IV. To distinguish between the two methods the labels 2z and 3z are used for the alternating and simultaneous setups, respectively. The experimental set-ups are also described in Chapter IV. Among various measurements of different quality, two sets of 2z recordings and one set of 3z recording were finally retained for the present work. They are referred to as 2z kin1 , 2z kin2 and 3z kin . The index 'kin' recalls that these recordings were made with full scanning of the temporal window accessible to the setup. Two complementary recordings limited to three pump-probe delays only were also made, in 2z and 3z configurations; they are named 2z 3T and 3z 3T .

Steady-state spectra

We begin by analysing the steady-state absorption and emission spectra of Xl64 photolyase. The absorption spectrum of Xl64 photolyase consists of two major absorption bands. The overall structure resembles the spectrum of FAD ox . This is not surprising as FAD ox is known to be the chromophore bound by purified Xl64 [21][22][23]. The first of these major absorption bands is found around 360 nm. Two local maxima are found in this region. The first one is at 356 nm and the second is at 373 nm. This band is assigned to the S 0 →S 2 transition of FAD ox [22]. The second band is located around 450 nm. It is assigned to the S 0 →S 1 transition of the FAD ox chromophore. This band has a maximum absorption at 447 nm. It is also structured. Two shoulders are observed: at 425 nm and 473 nm. As compared with the spectrum of free FAD ox in aqueous solution (cf. Annex B), the two bands are relatively more structured in Xl64 due to the less polar and more rigid nature of the protein's binding pocket. Weak non-zero absorption is observed for all wavelengths above 500 nm due to the small absorbance of FAD ox (ε ~ 0.1 mM -1 cm -1 ) and the scattering of the solution. The emission spectrum of the protein exhibits at maximum at 530 nm with a Stokes' shift of 80 nm (to ~19 000 cm -1 ) from the 450 nm absorption band (S 0 →S 1 ). The structure of the emission spectrum is different from the S 0 →S 1 band. The emission band is relatively unstructured (shoulderless) compared to the 450 nm band of Xl64 photolyase. This possibly indicates that the fluorescence we observe does not come from the main absorber, that is, bound FAD ox .

As we will see that the excited-state lifetime of bound FAD ox is extremely short (sub-ps) its fluorescence quantum yield is very low. It could be that the present fluorescence spectrum comes from a very small fraction of free FAD ox , released from the protein. Free FAD ox in solution is indeed know to have a longer excited-state lifetime and, hence, a higher fluorescence quantum yield. Since the flavin chromophore can exist in various redox states (cf. Chapter 1), we fitted the absorption spectrum of the Xl64 sample for transient absorption measurements to determine the distribution of the various states present. We did this by using previously published spectra of FAD ox , FADH • , and FADH -from X. laevis of Schleicher et al. [22]. These spectra have previously been extracted from this publication by Johanna Brazard and are shown in Figure 4A. The blue line in Figure 4B represents the best fit. Our analysis indicated that the sample is virtually composed of FAD ox only.

Isotropic femtosecond transient absorption spectroscopy

The measurements of transient absorbance have been carried out on concentrated solutions of typically 100 µM in a quartz cell of small volume (50 µL) and path length (0.1 cm). The protein was kept in a 100 mM TRIS buffer containing 100 mM NaCl and 25% (v/v) glycerol at pH 8.0 and the temperature was maintained at 4°C. It has been observed that a high concentration of glycerol in the sample enhances its stability (D.P. Zhong, private communication). In preliminary experiments performed with 5% (v/v) glycerol, the protein solution exhibited signs of aggregation in as early as the first few hours of data acquisition wherein the sample significantly diffuses light. Upon increasing the glycerol concentration, the sample remained free of aggregation (to the naked eye) for the duration of the experiment which is typically in the order of 20 hours.

The excitation wavelength was tuned at 475 nm.

In the present section we analyse the isotropic transient absorption spectra of oxidised Xl64, reconstructed from the polarised spectra. We used the standard definition of the isotropic signal given in Equation (3.1).

Isotropic spectra are free from rotational diffusion and intrinsic polarisation effects that are otherwise visible in the parallel and perpendicular spectrum. They therefore allow for a direct comparison with steady-state spectra of known photoproducts. They are thus best suited for the eventual identification of the transient species that are formed. The identification of these species will be discussed in §3.2.2.5. All spectra shown in this chapter have been properly corrected for chirp (see § 4.3, Chapter 4).

Initial transient absorption spectrum

To start with, let us analyse the isotropic transient absorption spectrum of Xl64, measured at an early pump-probe delay of 100 fs, shown in Figure 5. This is the spectrum of the bound flavin chromophore immediately after excitation, before any significant photoreaction occurs. The time of 100 fs is the earliest delay that can be presented as earlier pump-probe delays contain an artefact resulting from the cross-phase modulation (XPM) of the pump and the probe beam (this artefact is further described in § 3 of Chapter 4). It is nevertheless short enough to ensure that no excited-state reaction or decay has significantly begun, as will become apparent in the following. The region around the excitation wavelength (475 nm) was cut to avoid errors introduced by the scattering of the pump beam in the sample cell. This is seen as the gap in the spectra. We can classify the observed bands according to their sign. A positive transient absorption signal is found in the regions λ < 410 nm, 503 nm ≤ λ ≤ 518 nm, and λ ≤ 585 nm. In these regions, the transient absorption of the excited FAD ox is the predominating signal. Two negative bands are found in the spectral region 413 nm ≤ λ ≤ 448 nm and 518 nm ≤ λ ≤ 585 nm. In the 413 nm ≤ λ ≤ 448 nm region, the negative band is centred at 450 nm. This signal is attributed to the bleaching of the ground state absorption of FAD ox . The bleaching of the 450 nm band of FAD ox is the dominant signal in this region. The bleaching of the 475 nm band (negative contribution) is masked by the predominant transient absorption signal in the region below 410 nm. The 518 nm ≤ λ ≤ 585 nm region is dominated by the stimulated emission of the excited state. The minimum of the negative band is found at 535 nm.

Spectral dynamics of the isotropic transient absorption

We now present the isotropic transient absorption spectra of Xl64 photolyase, in the 3z kin recording between 0.1 and 1500 ps, in Figure 6. This recording is shown because of its superior signal-tonoise ratio over the 2z kin recording that is shown in Annex C.

- The evolution of the transient spectra can be divided into three phases. These phases are actually based on the time constants found in the global fitting of the data, which will be presented later:

-From 0.1 to 2.2 ps, there is large decrease in the amplitude of the simulated emission, between 500 and 660 nm. This negative signal decays into a flat and relatively featureless positive band at 2.2 ps. This indicates the rapid loss of stimulated emission of the excited state. A temporary isosbestic point is observed at 660 nm. Meanwhile there is a change in the transient absorption band between 360 and 410 nm. The formation of a shoulder at 400 nm is observed in this transient absorption band. A small decrease in amplitude is observed in the negative bleaching band between 410 and 450 nm.

-Between 2.2 and 220 ps, a sharp decrease in amplitude is observed for all the bands. This occurs for the positive transient absorption band between 360 and 410 nm and the negative bleaching band between 410 and 450 nm with no significant change in structure. The positive band at λ < 500 nm decays to a slightly more structured band with a maximum at 625 nm.

-In the third phase, between 220 and 1500 ps, there is a small decrease in amplitude of all the bands. No change in spectral form is observed. The signal does not decay to 0 at the end of the time window (1500 ps).

Global kinetic analysis of the isotropic transient absorption spectra

The above isotropic transient absorption spectra could be globally well fitted using a sum of three exponentials function plus a plateau, convoluted by a Gaussian function representing the instrument response function (IRF). After a preliminary step were the common "time zero" and width of the IRF were determined, the spectra were in fact truncated below 200 fs to get rid of spurious contaminations due to our imperfect fitting of the XPM artefact. The following time constants, τ i , have been found: τ 1 = 0.44 ± 0.01 ps, τ 2 = 36 ± 3 ps, τ 3 = 114 ± 9 ps, and τ 4 = ∞ (imposed), with a mean fit residue of 2.1 × 10 -5 (in OD units). The corresponding decay-associated difference spectra a given in Figure 7A. Quite similar results were obtained with both our 2z kin measurements (See Annex C).

In order to compactly summarize the evolution of the transient absorption spectra, we calculated the so-called evolution-associated difference spectra (EADS), as explained in detail in Chapter 4. Briefly put, the EADS are the spectra associated to the states of a formal cascading system, with a 100% reaction yield for each step that decays with the time constants found in the global analysis. These spectra have benefited both from deconvolution of the IRF and noise filtering from the global analysis. The underlying sequential model per se is not intended to represent any real population kinetics; it is just a convenient tool. The EADS for the 3v data are presented in Figure 7B. The first spectrum (EADS1; blue) is that of the initial excited state, extrapolated to t = 0. It decays in 0.44 ps (time constant) to give EADS2 (green). EADS2 evolves in 36 ps to give EADS3 (orange), with a clear population loss as seen in particular by the strong decrease of the bleaching band around 450 nm.

400
EADS3 further evolves in 114 ps and gives the final EADS4 (red), with also a population loss. EADS2, EADS3 and EADS4 bear close resemblance to each other. The positive transient absorption band in the region below 410 nm and the bleaching band visible from 410-450 nm are similar in shape for all three of them. They also all exhibit a similar broad positive band in the spectral region above 500 nm. These similarities suggest that the chemical species corresponding to these spectra are most likely analogous. It may be noted that the red absorption band of EADS2 is significantly broader and flatter than that of the following EADS, in line with the observation made above that the transient absorption spectra narrow and get more structured in this spectral region between, 2 and 200 ps. These issues will be further discussed when we go on to the identification of the transient species in §3.2.2.4.

It is now worth mentioning that a slightly better fit could be obtained with a sum of 4 exponentials plus a plateau. We got the following time constants: With this improved fit the main qualitative difference is that the spectral narrowing phase seen in the red absorption band of the product states (36 ps with the preceding fit) is now split in two steps: 8 ps to go from EADS2 to EADS3 and 51 ps to go from EADS3 to EADS4. The last two EADS of the new fit look similar to the last two EADS of the old fit.

These results should however to be taken with some care as the benefit of adding one exponential amounts to reducing the average residue of fitting by only 6 × 10 -6 (OD units). One could hypothesize that small and slow spectro-temporal fluctuations of the spectra would have here unduly been included in the fit. Comparable results were nevertheless obtained with our 2z kin2 recording (τ i = 0.46 ± 0.01 ps, 16 ± 4 ps, 62 ± 11 ps, 376 ± 172 ps and ∞), with similar DADS/EADS. The benefit from the 4 th exponential was however in this case even smaller, ca. 2 × 10 -6 , almost negligible. No satisfactory fit with four exponentials was obtained for the 2z kin1 measurement, possibly because the noise level of this set of data was significantly larger. We thus conclude that although our best recordings possibly reveal the existence of four exponential decays, corresponding to actual molecular events, a set of three exponentials followed by plateau seems to provide a somewhat more stable and reliable basis for the kinetic description of our data -even if it entails a slight loss of information.

Nature of the photoproducts

In this section, we aim to qualitatively analyse the EADS obtained in the above section in order to identify the transient species produced after the excitation of FAD ox . For the sake of simplicity, we will here essentially deal with the EADS associated to the global fit with 3 exponentials and a plateau (Figure 7). The arguments developed in this section can be transposed to the case of the fit with 4 exponentials and a plateau (Figure 8) without much change. Only a brief specific comment will be made at this respect.

It is clear that the characteristic shape of EADS1 is similar to that of excited FAD ox in solution [24]. It thus simply corresponds to excited FAD ox (FAD ox * ) before any reaction occurs -as expected for the transient absorption spectrum extrapolated at t=0 (meaning of EADS1). The real issue is of course to identify EADS 2 to 4.

According to the literature (cf. Chapter 1), upon excitation the bound FAD ox is reduced by a nearby residue through the transfer of an electron and a proton. From the structural analysis of the active site of Xl64 photolyase (cf. § 3.1) we hypothesize that this reduction may arise from the proximal tryptophan, W 396 . . An electron transfer to the excited flavin from tryptophan would yield FAD •-and WH •+ . If protonation of FAD •-were fast enough to fit our observation window we could observe the spectral signature of FADH • . If WH •+ were to deprotonate fast W • would be observed. Reduction by a tyrosine would yield YOH •+ and further deprotonation of the tyrosyl radical (likely because its pKa lies below 0 [25,26]) would yield YO • .

In order to help our assignment we used simulated difference spectra, previously calculated by Brazard et al. [5] and recalled in Figure 9. They correspond to different putative photoproducts obtained after excitation of FAD ox . These photoproducts are radical pairs formed by choosing the reduced species among FAD •-(Figure 9A) or FADH • (Figure 9B) and the oxidised species among WH 9C. One here implicitly assumes the FAD spectra correspond to pure redox species, as opposed to mixtures of states, which might be questioned. It is clear from Figure 9 that the global shape of EADS 2, 3, and 4 corresponds best to the FAD •- /WH •+ radical pair (Figure 9A, black curve,). Figure 10 shows a superposition of the last three EADS with the simulated difference spectrum of the FAD •-/WH •+ product. All spectra were normalised at the maximum of the bleaching band, at 447 nm. The overlap is obviously not perfect but common spectral features can be observed. The intense absorption maximum on the blue side of a unique bleaching minimum characterises the presence of FAD •-. Differences in shape and relative intensity of this band are likely due to our insufficient knowledge of the actual spectrum of FAD •-in Xl64. The simulated spectrum has indeed been constructed from the spectrum of FAD with previous reports in other photolyases [28,29]. This might explain the presence of residual absorbance in the transient absorption spectra beyond 580 nm. As far as FADH • is concerned, whether it is excited or reduced to FADH -by a tryptophan, one would expect pronounced negative features about 500 and 600 nm [30,31], which do not seem to be present in our EADS. One could finally evoke the possibility of a contribution of free, or somehow redox inactive, FAD ox . FAD ox * is indeed known to have a positive transient absorption band above 630 nm [24]. The expected corresponding negative stimulated emission contribution (between 500 and 600 nm) is however not clear in our EADS, which gives little credibility to this hypothesis.

If we now focus on EADS2, we observe that the mismatch with our best simulated spectrum is larger, because of the very broad, structureless absorption band EADS2 has above 500 nm. Different interpretations may be proposed. As apparent in Figure 9A, the flat absorption profile of EADS2 between 500 and 550 nm might arise from an overlap of Y • and/or W • contributions on top of the dominant WH •+ contribution. The simultaneous presence of WH •+ and W • was indeed reported by Immeln et al. [17] after excited-state decay of FAD ox in A. thaliana cryptochrome 1 (AtCry1). Although discussed in plant cryptochrome [17,32,33] ultrafast proton transfer from WH •+ has never been claimed for photolyases.

What can be said in disfavour of this hypothesis is that the aspartic acid thought to mediate this reaction in AtCry1 [32,33] is not present in Xl64. The question would also remain to explain why these putative Y • or W • radical seem to decay or turn into WH •+ in EADS3 and EADS4. This explanation also hardly explains the far red tail of EADS2 as the contributions of Y • and/or W • are very weak above 600 nm.

Alternatively, one could consider that the WH •+ radical produced immediately after electron transfer in a hot vibrational state. This hypothesis has been proposed by Immeln et al. [17] in their study on AtCry1.

This could possibly explain the broad absorption structure of EADS2. Immeln et al. claim that vibrational cooling, with spectral blue shift and narrowing, would take place in the sub-100-ps regime. If such a view were applied to our results the transition from EADS2 to EADS3 with a time constant of 26 ps might, hypothetically, be assigned to cooling. One can however not rule out that vibrational cooling simply accompanies another reaction, namely the Brettel-Vos electron hopping mechanism [31], possibly including additional pathways as in Liu et al [16].

We may incidentally recall that the spectral narrowing and structuring observed in the transition from EADS2 to EADS3 in 36 ps becomes biphasic with the 4-exponential plus plateau fit. A first phase of 8 ps sees the beginning of the process, with mostly a slight narrowing of the red absorption band, while the 51-ps phase completes the process and makes the full structures appear (Figure 8B). The cooling interpretation given above can be maintained within this framework but is hard to say at this level if any of the two phases corresponds to a pure cooling process or also involves a chemical reaction (electron transfer).

Returning to the 3-exponential plus plateau fit we finally favour the following interpretation of EADS 2 to 4. We propose they mainly arise from the pair of FAD Let us at this point make a slight digression and recall the FTIR study of Xl64 photoactivation by Yamada et al. [34]. These authors were able to observe the spectral signature of FAD •-, trapped at 200 K after excitation of FAD ox . Their steady-state difference spectrum is shown in Figure 11. It is consistent with the formation of FAD •-based on the positive peak centred at 382 nm. Of note is that the FAD •-peak at 382 nm is similar our own findings (EADS2-4) although with smoother structures. The authors assigned the weak positive band with a maximum at 503 nm to either W • or YO • .

This last option would incidentally be fairly compatible with our simulated difference spectrum of Figure 9A (green curve). It would also apparently support the early result of Weber et al. [6] on the photoactivation of oxidized Xl64, claiming that a tyrosine is the ultimate electron donor. A similar situation was reported for the photoactivation of A. nidulans CDP photolyase reported by Aubert et al. [2],

a tyrosine residue being oxidized by a tryptophanyl radical with a half-life of 50 µs. However the Yamada spectrum has been recorded under steady-state irradiation at very low temperature. These particular conditions might, somehow artificially (as compared to room temperature), extend the lifetime of the tryptophanyl radical and leave it enough time to be reduced by a tyrosine. This could also explain the marked difference of this spectrum with our EADS.

Elementary kinetic model

We will now try to build an elementary kinetic model potentially able to describe the actual population dynamics of the system. We will base our construction under the simplifying assumption that each EADS of a given fit (let us recall they just summarize the evolution of the spectra) in fact corresponds to a unique well-defined chemical species. Recalling how EADS were constructed, this hypothesis means that the system is supposed to follow a cascading scheme involving n inter-converting species, with no back reaction involved in the process; n is the number of exponentials decays (plateau included) considered in the fit.

For the sake of simplicity we will start with the case of 3 exponentials followed by a plateau.

Figure 12A illustrates the corresponding cascade, involving 4 species (sequentially called S1 to S4). In such a scheme the conversion rates k i between species S i and S i+1 are simply the reverse of the experimental time constants  i . Since we have seen that losses of transient population occur as EADS2 evolves to EADS3, and EADS3 evolves to EADS4, we have included ground-state recovery processes at each step of the cascade, in competition with inter-species conversion as also shown in Figure 12A. Note that this is by no means the only possible kinetic model applicable to our data; it is however one of the simplest. The intrinsic difference spectra associated with each species will be noted SADS (speciesassociated difference spectra). Our model contains 3 unknown parameters, the quantum yields: ϕ i of the conversion from S i to S i+1 . They will be estimated by using additional constraints as follows. The SADS are the intrinsic difference spectra of the species, to within a common multiplying constant linked to the (unknown) initial excited-state concentration. They are therefore expected to contain the same amount of pure bleaching contribution (for one molecule in any transient state one molecule is missing for the initial ground state).

In the model of Figure 12A the SADS are simply proportional to the corresponding EADS and thus all exhibit a marked negative band around 447 nm, dominated by bleaching. We have assumed in first approximation that the value of transient absorption at the maximum of this band may be used as a measure of the amount of pure bleaching, at all times.

This strong constraint implies that, at this wavelength, the pure transient absorption contributions of all molecular species involved in the reaction are identicalproportional to their respective abundances. Within the interpretation framework of the preceding section this hypothesis is reasonable as far the evolution from EADS2 to EADS3 and then to EADS4 is concerned. We have indeed assumed the nature of the species in presence do not change in this process and remain the FAD •-and WH •+ radicals.

Although the (positive) absorption contribution of FAD •-at 447 nm is far from being negligible (see Figure 9C) it may be considered constant, relatively to the pure bleaching contribution of FAD ox , because it always arises from the same reduced flavin. Figure 9C shows the contribution of WH •+ can be neglected at 447 nm, which completes the proof. It is less obvious to justify our constraint for the evolution from EADS1 to EADS2. Within our interpretation framework this corresponds to going from FAD ox * to FAD •-. These two species in fact have much in common because, in simplified molecular orbital language, they both involve the presence of a single electron in the LUMO of FAD ox . One can thus expect that transitions promoting this electron to higher lying orbitals would be essentially identical.

Indeed when one compares EADS1 and EADS2 in Figure 7B, one sees that the transient absorption band below 400 nm does not change drastically. We thus assume that, correspondingly, only small changes occur at 447 nm when FAD ox * is reduced to FAD •-.

With the above justifications in mind, we finally established the following rule: all SADS must have the same value around 447 nm. This constraint was applied by normalizing EADS2, EADS3 and EADS4 to EADS1 at 447 nm. The yields were obtained, namely: ϕ 1 =96%, ϕ 2 =55%, and ϕ 3 =46%. These numbers should of course be taken with much care as they highly rely on our set of constraints. The final SADS are presented in Figure 13A. As discussed in §3.2.2.4 about the normalized EADS (which were already the SADS), the similar shapes of SADS3 and SADS4 are compatible with chemically identical species, namely the pair of FAD •- and WH •+ radicals. Differences between them could then be assigned to changes of the particular identity of the contributing tryptophans (W A , W B , W C …). As far as SADS2 is concerned, we already discussed that its notably broader and less structured shape in the red could be due to the contribution of vibrationally hot WH •+ radical. Let us, by the way, note that very similar results exhibiting the same spectral trends and similar conversion yields were observed upon analyzing our other measurements.

We next present in Figure 13B the normalized population dynamics of the system with the above model, deconvoluted from the IRF. The initial excited-state population (value of the blue curve at t=0) has been arbitrarily set to 1 (thereby transferring the real initial excited-state population as a common multiplier of all the SADS).

The same work can of course be made on the basis of the global fit with 4 exponentials followed by a plateau. Figure 12B shows the cascade of 5 states hypothesized to represent the dynamics of the system in this context. Figure 14A displays the corresponding SADS and Figure 14B the normalized population dynamics. The difference between SADS2 and SADS3 is quite small: above 500 nm SADS2 is in fact slightly broader and flatter than SADS3, the 8-ps phase corresponding thus to a narrowing process which, as already noted above (in §3.2.2.4 about EADS), further proceeds during the 51-ps phase leading to SADS4 and could be assigned to a cooling process.

Polarised femtosecond transient absorption spectroscopy

As stated above the polarised, parallel and perpendicular, transient absorption spectra were recorded in all of our experiment. For the sake on compactness of this chapter these raw data are only given in Annex D. It should be stressed that the set of parallel and perpendicular data is completely equivalent to the set of magic and anisotropy data. Since the anisotropy data will be discussed in section 3.2.4 it is not strictly necessary to dwell on the parallel and perpendicular spectra. We however provide here a few elements that complement the isotropic results given above and introduce the tools used in the anisotropy section.

We here present a global fit of both the parallel and perpendicular spectra (3z kin recording), performed simultaneously, with the same sum of three exponential functions plus an imposed plateau as above (we will comment on that in the coming section). The following time constants have been found: τ 1 = 0.44 ± 0.01 ps, τ 2 = 39 ± 4 ps, τ 3 = 120 ± 14 ps, and τ 4 = ∞ (imposed), with a mean standard error (σ) of 2.5 × 10 -5 . These time constants agree very well with those found from the analysis of the isotropic data alone ( §3. At this point let us mention that some differences between our different measurements could be detected in the simultaneous global analysis. The difference essentially appears as small variations of the polarisation-dependent EADS and has essentially consequences when anisotropies are calculated from these (see below). The time constants remain largely reproducible.

As for the isotropic data we found that slightly better fits could be obtained with a sum of four exponentials plus a plateau. Here are the obtained lifetimes for the 3z kin recording: It should be stressed that the 4-exponential (plus plateau) fit appears to be particularly helpful when the anisotropy is considered. As will be seen below it is straightforward to calculate "raw" anisotropy out of the polarised transient absorption spectra. On the other hand it is also easy to deduce the anisotropy corresponding to the global simultaneous fit of the same data. We observed that, for the 3z kin recording, the 3-exponential function produced a slightly poorer fit of the anisotropy whereas the 4exponential function provided an excellent match. The difference was not so clear for the 2z kin2 recording although the 4-exponential simultaneous fit was possible and yielded results comparable to those of 3z kin (as previously observed for the isotropic data). Table 2 below summarizes our findings for our best two measurements, 2z kin2 and 3z kin . Let us finally mention that polarised SADS corresponding to the kinetic model of Figure 12 can be obtained in a straightforward manner. From the polarised EADS one first calculate isotropic EADS with Equation (3.1), then the normalisation procedure described in §3.2.2.4 is applied with no change. This gives rise to conversion yields (ϕ i ), very similar to those of the isotropic case, which are in turn used to calculate the polarised SADS. It is important to note that, within the models of Figure 12, each SADS is simply proportional to the corresponding EADS and that the proportionality factor (linked to the ϕ i ) is strictly identical for parallel and perpendicular polarisations. The polarised SADS are not shown here but anisotropy spectra corresponding to them will be provided below.

Femtosecond transient absorption anisotropy

In section 3.2.2, we have identified FAD •-and WH •+ as the predominant species produced, within 1.5 ns after photoreduction of FAD ox . A kinetic model (Figure 12) satisfactorily describing the data has been devised. It is composed of four or five species depending on the number of exponentials considered in the global fitting procedure, the initial excited state and three or four products, simply connected in a sequential fashion with ground-state recovery channels at each step.

The question that we now ask is if it is possible to attribute to these kinetic phases a definite chemical meaning, precise elementary steps of the global photoreduction reaction, typically electron transfers along the Trp chain. In order to simplify this task we will here reduce our scope to the results attached to the 3-exponential-plus-plateau fit. As we have seen, this fit provides a satisfactory and reproducible description of all our data even if it involves a slight loss of information for the 3z kin recording.

Within this framework a simpler question naturally arises, as to whether one can associate to each of the kinetic phases one elementary electron transfer along the Trp triad. The shortest time constant (0.44 ps) would be associated with the transfer from W A (see notations in §3. This is exactly the point of view adopted by Brazard et al. in their study on Ot64 [5]. It is also in first approximation the picture built by Liu et al. [16] for the photoreduction of oxidized EcCPD. In this latter work the kinetic model is certainly much more complex, as it involves back-reactions and two additional tryptophans (W A' and W B' ) but, if one only keeps the main channels, one gets a reduced scheme somewhat similar to the one proposed by Brazard et al. and the one considered in Figure 17. The difficulty with the interpretation of our data along Figure 17 is that there is no hard proof that each product species (S2, S3 and S4) of the kinetic model has the above-described meaning (FAD •-associated to W A H •+ , W B H •+ and W C H •+ , respectively). All WH •+ radicals having similar spectra, or exhibiting nonpredictable specific features, an assignment of each SADS to a particular tryptophan position is not possible.

In this section, we try to distinguish the different tryptophans potentially involved in the reaction by taking advantage of their different spatial orientation with respect to the flavin. We therefore looked at ? ? a spectroscopic property that depends on the orientation of the molecule, the anisotropy. The transient absorption (r) anisotropy is defined in Equation (3.2) as the difference between the parallel and the perpendicular signal divided by three times the isotropic signal.

For any freely rotating molecule, approximated to a sphere for simplicity, the anisotropy decays exponentially to 0 with a time constant called the rotational correlation time, τ rot , as shown in Equation (3.3) [35]; r 0 is the "intrinsic" anisotropy at t = 0.

) / exp( ) ( 0 rot t r t r    (3.3)
The rotational correlation time may be estimated by Equation (3.4), where V is the hydrodynamic volume of the molecule, η is the viscosity of the solvent at temperature T, and R is the gas constant.

For a photolyase in water at room temperature τ rot has been estimated to about of 30 ns [36]. For an aqueous solution containing 25% glycerol at 5°C we estimate τ rot should be about 120 ns. This value was estimated from the viscosity differences between a solution of 25% glycerol at 5°C and pure water at room temperature. The viscosity values differ by a factor of about 4 [37]. This clearly shows that rotational diffusion of the protein as a whole is not active within the time window of 1.5 ns of our fs transient absorption measurements. This in turn explains why the polarised data could successfully be fitted in §3.2.3 by the same function as the isotropic data, with very similar time constants. This observation further suggests that no substantial internal movement of the molecules of interest (FAD ox and relevant residues) occurs within the protein framework during our observation window. The anisotropy dynamics that is about to be presented should thus be considered as characteristic of intrinsic changes within the molecule, hence of the sole photoactivation reaction.

Sensing the orientation of the WH •+ radical

As a preamble to the presentation of our experimental results, we discuss here the question of finding the best conditions for sensing the orientation of the WH •+ radical with the anisotropy tool.

It is well known that under the conditions that a single transition is excited and a single transition is probed, the anisotropy simply depends on the angle (β) between the transition dipole moment of the excitation transition and the transition dipole moment of the probed transition detected, as given by Equation (3.5) [38].

(3.5)

It is thus most interesting to try to comply with the above conditions. In our transient absorption experiments, we solely excite the S 0 →S 1 transition of the flavin at 475 nm, which satisfies the first premise. To satisfy the second, we must determine the wavelength at which, ideally, only WH •+ absorbs, or at least absorbs predominantly. We know that WH •+ has a broad absorption maximum between 550 and 600 nm, in a region where FAD ox (bleaching in the transient spectra) does not contribute. It however appears that FAD •-has a small residual absorption in this region, as recalled in the insert of Figure 18.

We show in Figure 18 the ratio of the absorption coefficient of WH •+ and that of FAD •-as a function of wavelength. The resulting plot has two maxima at 562 and 580 nm. We chose the redmost maximum at 580 nm to even further neglect the FAD ox absorption. Choosing this wavelength allows us to directly compare our result to similar measurements found in literature such as the measurements by Liu et al. [3]. The contribution of FAD •-should in principle not be neglected as, for a pair of WH •+ and FAD •-radicals, it contributes at this wavelength to 17% of the total absorbance of the pair. The pure anisotropy of the WH •+ (r WH•+ ) could theoretically be extracted by using the standard sum rule [36] of Equation (3.6),

giving the total anisotropy (r) as a function of the contributions (A FAD•-and A WH•+ ) of each species to the total isotropic transient absorbance.

                   WH FAD WH WH FAD FAD A A A r A r r (3.6)
The drawback is that the anisotropy, r FAD•-, of FAD •-would be required. As we do not know it we were constrained to neglect the contribution of FAD •-to the total anisotropy.

Raw transient anisotropy

The "raw" anisotropy spectra, i.e. the anisotropy directly calculated from our 3z kin measurement with Equation (3.2), are shown in Figure 19 at selected pump-probe delays.

- We observe in Figure 19A that the anisotropy varies hugely between 500 and 650 nm during the first few ps, that is, during the initial excited-state decay and reduction of the flavin. This massive effect is understood by observing the isotropic transient spectra of Figure 6. At early times the initially negative band dominated by stimulated emission is being replaced by a positive band. At many wavelengths within this region the signal thus becomes null at some point in time. As is clear from Equation (3.2), when the isotropic signal (denominator of the fraction) goes to zero, the anisotropy diverges towards infinite values, which explains the large changes. The same effect is in fact visible at any given time in the spectral axis, around 416 nm. At this wavelength the isotropic signal being null, the anisotropy diverges.

At 2.2 ps, after the fast initial evolution, the anisotropy around 580 nm takes a value of about -0.07. As became clear from §3.2.4.1, we will be particularly interested by this region in the following, as it is dominated by the main photoproduct we identified, WH •+ .

The region below 400 nm is sensitive to both the presence of excited FAD ox and FAD •-. The spectrum of FAD •-exhibits a local absorption maximum at 400 nm (see Figure 18-insert). We observe that the anisotropy at 400 nm decreases from 0.17 to 0.06 between 0.1 ps and 2.2 ps. This change is assigned to the reduction of FAD ox * into FAD It should finally be noted that at 447 nm, i.e. at the minimum of the bleaching band, the anisotropy is found to be approximately constant all along the kinetics, with a value of about 0.45. This value is close to the theoretical value of 0.4 expected for a pure bleaching contribution. This indicates that the anisotropy of the transient absorption contribution at this wavelength, overlapped to the bleaching contribution, is likely rather close to 0.4. This would correspond to transition dipole moments making relatively small angles with that of the S 0 S 1 transition of FAD ox . We can in particular verify in Figure 20 that the anisotropy at 447 nm remains essentially constant during the whole kinetics, except for a slight rise in the hundreds of ps regime. In the blue spectral region dominated by the transient absorption contribution of the flavin, the reduction of FAD ox * into FAD •-at early times is only modestly appreciated at 400 nm but not at 375 nm. This phenomenon is much more obvious at 580 nm, for the reason exposed above (loss of stimulated emission, growth of transient absorption).

It may be noted in anticipation of a coming discussion that although these raw kinetic traces are fairly smooth (as expected for a 3z kin experiment where parallel and perpendicular signals were measured simultaneously), they are unfortunately not as reliable as they look. We indeed found with test samples that ill-reproducible systematic errors (probably of instrumental origin) slightly alter the anisotropy (see Chapter 4). Caution must therefore be taken upon analysing those traces. This question will be more directly tackled when we specifically deal with the kinetics at 580 nm in §3.2.4.5.

Global kinetic analysis of the raw anisotropy spectra

We present here an attempt at directly and globally fitting the above raw anisotropy spectra. It should be noted that raw anisotropy is in general relatively noisy because it suffers from adding up independent fluctuations coming from the parallel and perpendicular measurements. We in fact found in most cases that the number of exponentials required to fit the raw anisotropy data is significantly less than for the simultaneous analysis described above ( §3.2.3), some subtle kinetic details possibly being obscured by this approach. It also happened to be necessary to truncate the spectral range to a narrower window (typically between 425 and 650 nm) in order to achieve proper convergence of the fits. However, in the particular case of the 3z kin recording, it was expected that the fluctuations of the transient absorption signal in parallel and perpendicular polarisation would be highly correlated, thereby reducing the noise of the raw anisotropy to a much lower level than for 2z-type measurements. This indeed proved to be true in good part. Here are our results with a sum of three exponential functions and a plateau: τ 1 = 0.34 ± 0.02 ps, τ 2 = 12 ± 2 ps, τ 3 = 117 ± 12 ps, and τ 4 = ∞, with a mean fitting residue of 7.5 × 10 -3 (in anisotropy units). These time constants are rather close to those obtained with the same fitting function applied to the isotropic or polarised transient absorption spectra. The first time constant (0.34 ps) is somewhat shorter than for transient absorption spectra but it is not certain that the difference is significant. We note that the second time constant (12 ps) is in fact closer to the second time constant found for transient absorption spectra with four exponentials (see Table 2).

No significantly better fit was however obtained with a sum of four exponential functions and a plateau.

The extra exponential turned out to be un-physically short and likely only present to improve the fitting imperfections related the XPM artefact at t=0. The so-called "evolution-associated anisotropy spectra"

(noted EAAS) of our 3-exponential fit, which are formal equivalents of the EADS for transient absorption data, are given in Figure 21. We see more clearly see in Figure 21 that the anisotropy at 580 nm takes a value of -0.075 immediately after the initial excited-state decay and reduction of the flavin, then rises towards -0.023 in a few tens of ps (time constant of 12 ps within this fit) and return to -0.073 in 117 ps.

Species-associated anisotropy spectra

The EAAS we have just seen provide a compact description of the evolution of the raw anisotropy spectra in terms of a sum of exponentials but cannot in fact be considered as the intrinsic anisotropies associated to the species of any kinetic model, such as those of Figure 12 for instance. The reason is that in such models only the population kinetics, hence the transient absorption data, can adequately be described by a sum of exponentials. The corresponding anisotropy is thus expected to be a complex function, the ratio of two sums of exponentials (see Equation (3.2)), not a simple sum of exponentials. We here sought a way to both of describe precisely the data and provide anisotropy spectra one can licitly associate to species within the framework of a given kinetic model. As an example, Figure 22A provides the SAAS (or FAAS) of our 3z kin measurement, corresponding to the kinetic model of Figure 12A and the fit with three exponential functions plus a plateau reported in §3.2.3. Figure 22B shows the SAAS corresponding to the model of Figure 12B and fit with four exponential functions plus a plateau. The corresponding time constants can be found above in Table 2. They are only slightly (and non-significantly) different from those coming from the analysis of the isotropic data, shown in Figure 13. Two remarks should be made at this point. On one hand, moderate but non-negligible differences of FAAS were observed depending on the measurement being analysed (3z kin , 2z kin1 or 2z kin2 ). They were particularly visible for small values of the anisotropy, unfortunately like around 580 nm. It is indeed expected that small anisotropies, corresponding to comparable values of the parallel and perpendicular signals, are very sensitive to small variations of the latter. Table 3 summarises our findings for recordings 2z kin2 and 3z kin (the two best ones), analysed with 3 or 4 exponentials (the time constant are given in Table 2). The r i are the values of the SAAS at 580 nm, averaged over ~2 nm (5 pixels) in order to smooth out minor spectral fluctuations. It may be noted that r 1 is a value extrapolated at t=0, after deconvolution of the IRF; it should therefore be taken with special care. On the other hand, it appeared that for our 3z kin measurement a significantly better fit was obtained with four exponentials (instead of three) plus a plateau, as far as anisotropy is concerned (the difference is not that critical on the transient absorption data). Figure 23 exemplifies this fact at 580 nm (averaged over about 2 nm 5 pixels). The open circles represent the raw anisotropy data (as in §3.2.4.2).

The continuous green line is the anisotropy reconstituted a posteriori from the global simultaneous fit of the polarised transient absorption data with 3 exponentials and a plateau; the blue line corresponds to the 4-exponential fit. The improvement brought by the 4-exponential fit is obvious. Figure 20 above shows a few other anisotropy traces, reconstituted from the 4-exponential fit at other wavelengths. We note that the benefit brought by the 4-exponential fit for 3z kin was far from being so clear for recording 2z kin2 . For reference the direct fit of the raw anisotropy with 3 exponentials and a plateau is also indicated in Figure 23, in red; it is very close to the simultaneous 4-exponential fit (blue line).

Errors on the anisotropy

Since the anisotropy kinetics at 580 nm appeared particularly interesting for answering the question we raised at the beginning of §3.2.4, we would like here to discuss the question of how much we can trust the experimental measurements we recorded, in other words the question of the errors attached to these data.

In a first approach we present in Figure 24A The next issue is, as we have previously observed, that the 3z kin anisotropy kinetics are affected by ill-determined systematic errors and are not as reliable as they seem. We tried to correct for these errors by measuring the anisotropy kinetics of small molecules (typically the laser dye DCM) in fluid solution (acetonitrile at room temperature). We observed that the anisotropy expectedly decays according to the rotational diffusion kinetics in the sub-ns regime and tends towards a stable non-zero value, ca. -0.02 at 580 nm the same day as our 3z kin measurement. This value in fact changed from day to day in an uncontrolled fashion. As this limiting value should be equal to zero (full orientational randomisation) we took the option of subtracting -0.02 from the raw 3z kin anisotropy kinetics at 580 nm. The result is shown in Figure 24B. This ad hoc correction is unfortunately not grounded in a precise knowledge of the experimental origin of the defect. We thus acknowledge that it is not rigorous and possibly involves unknown new sources of error. It is simply expected to yield a more accurate result than without correction.

The anisotropy kinetics of Figure 24B should, in principle, be considered as our best determination but detailed comparisons with other measurements showed that it still needs to be taken with caution. On the one hand, the error bars of the 3z kin corrected kinetics are not fully compatible with those of recording 2z kin2 (larger than for 3z kin as expected for independent measurement of the parallel and perpendicular transient absorptions). For quite some time delays they do not overlap, as shown in Figure 25 (compare the green and orange data). With this limited number of times, a much larger number of accumulations was performed in order to improve the signal-to-noise ratio. Two recordings were made in this mode: one with 2z configuration (independent measurement of parallel and perpendicular data), noted 2z 3T , and one with 3z configuration (simultaneous measurement of parallel and perpendicular data), noted 3z 3T ('3T' stands for 'three times').

These measurements are plotted in Figure 25, together with estimated error bars (indeed smaller than those of the 'kin' measurements); recording 3z 3T has been corrected for the specific systematic error of the 3z configuration. We observe again that, at some times, the 2z 3T (red) and 3z 3T (blue) errors bars do not overlap, as well as the 3z kin (green) and 3z 3T (blue) errors bars.

Taking into account all the difficulties detailed above we conclude that our estimates of the error bars are probably too crude to be used as they stand, and allow us to narrow our best recording to 3z kin or 3z 3T as we hoped. However in order to give a more realistic idea of the dispersion of our anisotropy measurements we finally took the option of keeping all measurements in one graph and, to simplify things, of substituting the raw 'kin' measurement by an adequate fit of the data. Figure 26 shows this view.

Measurement 2z kin1 being of poorest quality (very large statistical error bars; not shown), it is represented in dotted line to recall that significantly less confidence should be placed in it. We assume that the true kinetic trace should be found somewhere within the limits defined by these points and lines. Figure 26 then roughly suggests that practical error bars on the anisotropy kinetics at 580 nm would be of the order of ±0.02, between 2 and 1500 ps.

Expected anisotropies based on the homology model

We will now intend to calculate the anisotropy kinetics expected at 580 nm within the sequential electron transfer interpretation of our measurements given in Figure 17, associated to the cascading The orientation of the S 0 →S 1 transition dipole moment of the protonated tryptophanyl radical, WH •+ , was taken from the DFT calculations of Crespo et al. [43]. They reported it to be 110° (error not given) at 570 nm, counted counterclockwise with respect to the short axis of the molecule. Figure 27 summarizes the orientations of transition dipole moments we retained. Using the atomic coordinates of the flavin and the tryptophan residues extracted from our homology model, the coordinates of the different transition dipole moments in the protein frame of reference have been calculated. The angles of the transition dipole moment of each tryptophan of with respect to the flavin were then calculated as well as the anisotropies associated with each. These are the theoretical anisotropies one would expect to observe at 580 nm on the assumption that only one oxidised tryptophan is present at a time (after exciting the S 0 →S 1 transition of the flavin). These values are listed in Table 4, together with estimated errors. To calculate these errors, we allowed the orientations of both transitions moments to vary within an interval of ±5° about the retained values, keeping them within the plane of the molecule. The deviations of the angle between transitions moments, and of the corresponding theoretical anisotropy, induced by those variations were then noted. Since they do not depend linearly on the variations of orientations of the transition moments, they are in fact not symmetrical about the central value. For the sake of simplicity Table 4 however provides one single error taken to be the greatest deviation (rounded to an upper value). We can readily compare the intrinsic anisotropies assigned to the various WH •+ radicals of Table 4 (let us refer to them in short as r Wa , r Wb , etc.) to the values at 580 nm of the "species-associated anisotropy spectra" (SAAS), reported in Table 3. As we have earlier explained the anisotropies relative to recording 3z kin in fact need to be up-shifted by +0.02 in order to correct for the systematic error of the 3z configuration (the correction was not made at the level of global simultaneous fit and SAAS). The corrected values (r i c ) are collected in Table 5. As suggested by Figure 26 an experimental error of the order of ±0.02 can be assigned to r 2 c , r 3 c r 4 c and r 5 c (see §3.2.4.5).

Table 5. Corrected anisotropies at 580 nm of the SAAS (FAAS) of experiment 3z kin . The systematic error of the 3z configuration has been corrected by adding an offset of +0.02 to the values of Table 3. We find that although r 2 c (-0.042) is more negative than r Wa (-0.006) it fits well within the confidence interval defined by the error. Species S2 (see Figure 9A) could then correspond to W A carrying the WH •+ radical. The comparison is much less favourable for r 3 c (+0.007) because it lies very far from r Wb (+0.25). Even counting an experimental error of the order of 0.02, r 3 c does not even get close to the confidence interval of r Wb . It is thus doubtful that W B H •+ alone could be assigned to species S3.

With r 4 c (-0.057) a good agreement with r Wc (-0.05), and its confidence interval, is achieved. Species S4 could then be associated to W C H •+ . The confidence intervals of r 3 c and r Wd (-0.17) apparently do not overlap, which should in principle allow us to rule out any role of W D in the present time scale. We will however see below that such a conclusion must be taken with care as our errors might have been underestimated.

Comparison of experiment with simulation

Using the theoretical values of Table 4

 

      i iso i iso tot i iso tot iso i i A A A A r r , , , , with , (3.7)
A i,iso are the contribution of each species to the total isotropic transient absorption, A tot,iso . They were taken from the simultaneous 3-exponential (plus plateau) fit of recording 3z kin . The anisotropy of FAD ox * (r 1 ) comes from Table 3 but it is not critical since its influence is limited to the first couple of ps.

The anisotropies r 2 / r 3 / r 4 of the tryptophanyl radicals are taken from Table 4. The result is thus a hybrid between the (hypothesized) population dynamics coming from the experimental data and the theoretical anisotropies calculated from the homology model.

As a test of this approach, we performed the same exercise with the published results of Liu et al.

relative to the photoreduction of oxidised EcCPD [16]. The authors indeed published a detailed kinetic scheme involving five tryptophans, which can easily be used to compute population dynamics, and an experimental anisotropy kinetics at 580 nm (supporting information). On the other hand the crystal structure of EcCPD is known [44], which allowed us to estimate the intrinsic anisotropies attached to each tryptophanyl radical. Table 6 lists the theoretical anisotropies we computed.

Table 6. Angles between WH •+ and FAD ox transition moments and corresponding anisotropies, associated to the different tryptophan residues involved in the photoactivation of EcCPD photolyase. The calculations were based on the known crystal structure of EcCPD [44].

Angle between WH In order to simplify the calculation we additionally assumed that the isotropic contribution of each tryptophanyl was identical. A couple additional parameters concerning the contribution of FAD ox * were grossly adjusted so as to reach a better agreement with the experimental data at short times. [16] relative to the photoreduction of oxidised EcCPD, and our simulation (blue line) of the same based on the crystal structure of EcCPD [44].

The overlap of the experimental and simulated kinetics is obviously not ideal but several qualitative elements do agree. We see that, after full decay of the initial excited state, the anisotropy takes a rather small negative values (less than -0.025), corresponding to the weak negative contribution of W A .

Then the anisotropy becomes positive for a few tens of ps, due to the positive contribution of W B and W B' . Finally the anisotropy drops and stays at a negative values imposed by W C . Globally, the shape of the experimental and simulated kinetics should be considered to agree well.

Since we do not know the experimental error in the experimental kinetics of Liu et al. we can only make conjectures about the discrepancies with the simulationand we will not discuss the kinetic scheme on which our simulation is based. Looking in particular at the plateau in the ns regime, assumed to be due to W C H •+ only, a mismatch of 0.07 is observed. It is somewhat larger than the error on r Wc of Table 6. A first explanation could be that the orientations of the transition dipole moments in the molecular frames, as set in §3.2.4.6, would somehow be incorrect. In other terms the error on r Wc , and by extension on other theoretical anisotropies, would have been underestimated. A second argument could be that the crystal structure of EcCPD we used for our simulation does not reflect the likely flexibility of the surface-exposed residues, in particular W C . Retrospectively, this discussion suggests that the distinction between W C and W D of Xl64 could becomes less clear than discussed in §3.2.4.6. If flexibility of W C and W D were invoked, as well as an additional imprecision due to the use of a homology model instead of a crystal structure, it cannot be ruled out that the angles (between tryptophanyl and flavin transition moments) attached to W C and W D may actually differ by less than 16.8° (as shown in Table 4) and cannot be experimentally distinguished with clarity.

Figure 29 now compares our different measurements of the anisotropy kinetics of Xl64 at 580 nm (summarized as explained above; blue, red, green lines and dots) to our simulation (black line). In this case the isotropic contribution of each tryptophanyl was carefully taken from the simultaneous 3exponential fit of the data within the interpretation of it along the model of Figure 12A and Figure 17. We observe that, after the decay of the initial excited state, the experimental anisotropy takes a more negative value (between -0.10 and -0.05) than the expected nearly null contribution of W A . We have however seen in §3.2.4.6 that the error on r Wa (Table 4) is large enough to explain this apparent mismatch.

Alternatively one might think of a contribution of W A' (r Wa' =-0.2) at that time. It is however not very likely as we have seen in our homology model that W A' lies rather far away from the flavin (9.3 Å); in fact farther away than in the case of EcCPD (6.3 Å) where the role of W A' was almost negligible [16]. The second qualitative observation we can make from Figure 29 is that the experimental anisotropy never takes positive values at intermediate times, as expected from a dominant contribution of W B (or W B' ). The reason could either be that the electron transfer pathway does not go through W B or W B' or, more likely, that it does go through W B or W B' but the oxidation hole does not stay long enough on these residues to be detected in the kinetics as opposed to scheme of Figure 17 were it stays on W B for ~110 ps. Finally we observe a surprisingly good agreement between the experimental anisotropy values at long times (by 1 ns) and the simulation. Since the simulation at long times is dominated by the contribution of oxidised W C , it can thus be proposed that W C is the final electron donor in the present time window, as opposed to W D .

Oxidised W D would indeed be expected to have a more negative anisotropy (-0.17). As already discussed at length this conclusion must be moderated as errors in the calculation of the theoretical anisotropies, due to imprecise determination of the orientations of the transition moments, could blur the distinction between W C and W D .

In any case the analysis of the anisotropy kinetics at 580 nm suggests that the mechanism of the photoreduction is likely more complex than the elementary cascade involving W A , W B , and W C with no back-reaction, as depicted in Figure 12A and 17. The experimental data are compatible with W A being the primary electron donor reducing the flavin in the sub-ps regime. The involvement of W A' at short times seems unlikely for a matter of distance to the flavin, according to our homology model. More importantly, the rather flat aspect of the kinetics at 580 nm between 2-3 ps and 1.5 ns, without positive overshoot, is evocative of a process where the oxidation hole, after reaching W B , would not reside long on it but very rapidly hop to W C , or possibly back W A . Within such a view the very clear kinetic phases that were observed in all our measurements should perhaps be interpreted not as well-defined elementary electron transfer steps but as accompanying processes, vibrational cooling or protein relaxation, that somehow modulate the rate of charge recombination. A precise picture unfortunately eludes us for the moment. The involvement of other residues capable of reducing the excited flavin or the tryptophanyl radicals is not excluded but it is not clear to us what would be their exact influence on the kinetics at 580 nm. We finally tentatively propose that W C is the final electron donor in the 1.5-ns time window of our fs experiments, even if W D remains a possible second choice.

"Millisecond constant"

As will be explained in Chapter 4 we observed transient signals in the negative time delays (i.e., negative time delays refer to delays where the sample is probed before excitation by the pump) in our photoactivation experiment. The signal is due to the insufficient speed of the 2D translation of a sample that was supposed to avoid re-excitation of a previously excited sample volume. As a result, were able to measure the transient absorption spectrum in the ms timescale (3 ms). The inverted spectrum of the "ms constant" is shown in Figure 30. The ms spectrum is actually an artefact that was subtracted from all Xl64 spectra that we presented in this chapter. In the "3-ms" spectrum of Figure 30, we see a peak at 382 nm with is relatively smooth and featureless as compared to the transient spectra of the Xl64 in Figure 6. There is also a positive peak centred at 510 nm which goes to zero at 560 nm. A negative band centred at 450 nm is also observed.

Looking back at Figure 9A, such a spectral form may be explained by the presence of W • or YO • with FAD •-. A precise assignment is not simple. The peak at 400 nm (Figure 9A), which would be more intense in the presence of YO • than in the presence of W • , is lacking. The band at 510 nm is also problematic as the same forms are expected for both YO • and W • species. A conclusion is hence difficult to reach.

Nanosecond transient absorption spectroscopy

In the previous section, we have proposed that the final electron donor within a time window limited to 1.5 ns is W C . There is however no certainty that the electron transfer chain ends with W C . We have in particular pointed out the possibility that W D could play the role of final electron donor. This section is dedicated to testing this hypothesis, more precisely to checking whether the anisotropy measured in the spectral region dominated by the absorption of the WH •+ radical changes between 1.5 ns and a much longer observation window of tens to hundreds of ns. To determine if W D plays a part in the photoactivation, we need to look at what happens to the system after 1500 ps.

For this we measured the real-time evolution of the polarised transient absorption of oxidised Xl64 at selected wavelengths, after 5 ns excitation at 475 nm (provided by a Nd:YAG laser), in the ns-µs time range. The time resolution of this setup was 10 ns. We monitored the protonated tryptophanyl at 562 nm and the bleaching of FAD ox at 457 nm. This experiment was performed in the laboratory of Klaus Brettel at iBiTecS in CEA Saclay. The choice of monitoring wavelengths is in fact due to the availability LED's used in the lab. However, no compromise has been made by this choice; 562 nm also lies at a maximum of WH •+ absorption (cf. Figure 18). The anisotropy observed here is directly comparable to that of 580 nm that we measured in our ultrafast experiments.

Polarised transient absorption decays

Anisotropy in the ns timescale

The anisotropy at 457 nm and 562 nm were calculated using Equation (3.5). Figure 33 shows the anisotropy decay at 457 nm which we assign to be the bleaching of the flavin's ground state. It was successfully fitted with a single exponential decay (plus a small constant taking care of experimental imperfections). The initial anisotropy has a value of 0.63 and decays to 0 (constant being substracted) with a time constant of 980 ns. Figure 34 shows the anisotropy decay measured at 562 nm. A single exponential decay (plus constant) fit was also performed. Subtracting the adjustment constant, the initial anisotropy is found to be -0.055 and decays to 0 in 773 ns. From the above results it can be deduced that rotational diffusion of the protein, responsible for the global decay of the anisotropy, takes place with a time constant of about 0.9 µs. This value is significantly longer (by a factor of ~7. 5) than what we had predicted above on the basis literature values (photolyase in glycerol-free solution at room temperature) and dependence of the viscosity on temperature and glycerol content. We tentatively explain this discrepancy by hypothesizing that the glycerol content could have accidentally been larger in this experiment and possibly reached a level around 50%.

On the other, focussing on the 562-nm decay, we observe that the initial anisotropy corresponds well to the final anisotropy measured at 1.5 ns. Recalling that the time resolution of the ns setup is 10 ns the "initial" anisotropy measure here in fact corresponds to the actual anisotropy about 10 nanoseconds after excitation. We thus deduce that the anisotropy does not change much between 1.5 ns and, say, 10 ns.

The following monoexponential decay suggests that the intrinsic anisotropy does not evolve during rotational diffusion, that is, during a couple µs. This conclusion is however to be take with care as the experimental data are rather noisy at 562 nm. Subtle changes of intrinsic anisotropy (yielding multiexponential decay) could have been missed. In line with our preceding conclusion we tentatively propose that W C keeps being the last electron donor until at least 10 ns, possibly a couple µs.

The results obtained rather point to the conclusion that W D is not involved in the photoactivation reaction. However, as we mentioned above, uncertainties coming from the orientations of the transition moments of the flavin and the tryptophanyl radical could still challenge this view. To completely rule out the role of W D , a control experiment should be performed. We are planning to repeat our measurements on a point mutant of Xl64 lacking W D . This would be done by replacing the tryptophan with the sterically similar but redox-inert phenylalanine. This mutant, W370F, is currently under preparation by Junpei Yamamoto of the Osaka University. Should the anisotropy measured using W370F be the same as that measured using the WT Xl64 photolyase, then it would be concluded that W D is not involved in photoactivation.

Isotropic transient absorption in the ns-µs timescale

To conclude this part let us briefly the isotropic transient absorption decays corresponding to the polarised measurements of Figure 31 and 32. The isotropic signal was calculated using Equation (3.1).

Figure 35 shows the kinetic trace at 457 nm, corresponding to the bleaching of the flavin absorption. It remains strictly flat during more than 20 µs, which indicates than no charge recombination occurs during this time. Figure 36 shows the isotropic transient absorption decay at 562 nm. A single exponential plus constant was used to fit it, yielding a time constant of 24 µs. At this preliminary stage of the study in the ns-µs regime this decay is tentatively assigned to the deprotonation of the tryptophanyl radical, yielding the neutral W • radical, without charge recombination (Figure 35). It is indeed seen in Figure 9C that deprotonation of WH •+ is expected to be revealed by a partial decay of the absorbance at 562 nm. The time constant of this process is very large as compared to the 300-ns deprotonation reported for semireduced EcCPD [2]. The difference could be explained by the fact that a lower temperature (5°C instead of 10°C) and large glycerol content was used in our study. These facts could justify why proton exchange with the solvent would be dramatically slowed down (as is rotational diffusion). It cannot also be excluded that specific structural reasons (accessibility of water to the last tryptophanyl) also explain the difference between the two proteins.

Conclusion

We began our study of Xl64 photoactivation by identifying the residues most likely to be participating in the photoactivation reaction (i.e. potential electron donors in the vicinity of the flavin and of the "canonical" tryptophan triad). Due to the lack of an available X-ray structure, we constructed a model of the protein by homology modelling. In the model, we have identified the various tryptophan residues that are believed to be implicated in the electron relay in the photoactivation reaction. These tryptophan residues include the conserved tryptophan triad of photolyases (W A , W B , and W C ) and the tryptophan residues identified by Li et al. (W A' and W B' ). We further found that W C in Xl64 lies within electron transfer distance to another tryptophan that is exposed to the protein surfacehence potentially capable of being the terminal electron donor of the photoactivation reaction. This tryptophan, which we called W D , has never been mentioned in any literature involving the CPF proteins. We have also identified three tyrosine residues that might play a role in the electron relay. One of these tyrosine residues is lying in the vicinity of the flavin and two are in the vicinity of W C and W D .

We followed the events following the excitation of oxidised FAD ox in Xl64 by femtosecond transient absorption spectroscopy. Using this technique, we were able to obtain the difference absorption spectra of the transient species produced after the excitation of FAD ox . We have seen that the primary species produced in the timescale accessible to our set-up (1.5 ns) are the protonated tryptophanyl radical (WH •+ ) and the flavin anion radical (FAD •-). The presence of other species has not been explicitly identified in our spectra. All our transient absorption data could be fitted reproducibly with a sum three exponentials and a plateau. We obtained a slightly better fit of our 3z kin data using a sum of four exponentials with a plateau. We have constructed a cascading kinetic model for the both the three-and four-exponential models. Charge recombination between FAD •-and WH •+ , hence recovery of the initial FAD ox ground state, is present at each step and the corresponding conversion yield at each was calculated.

A physical interpretation of our kinetic model has been sought by performing transient anisotropy measurements. We addressed the question of whether or not each of the kinetic phases we observed in the fitting of our data can be assigned to an elementary electron transfer step along the conserved Trp triad, as had been the case for the studies on the photoactivation of oxidised photolyase that were mentioned in the introduction. We have shown from the analysis of the anisotropy kinetics at 580 nm that such is not the case and that the real picture of the photoactivation mechanism may be more complex that was previously thought (i.e. electron hopping between three conserved tryptophans) for (6-4) photolyase. The simplest explanation in our case would be the presence of back reactions and other processes that would complicate the electron relay among the tryptophans. Another possibility would be the involvement of other residuesapart from the triad. Our femtosecond experiments point to the conclusion that our newly identified residue that can function as an electron donor (in the electron relay), W D , would not play a role in the reaction. The same conclusion is reached from our experiments in the nanosecond timescale.

Introduction

This chapter is dedicated to the study of the photorepair process of the (6-4) photolyase, more specifically (6-4) photolyase from Xenopus laevis. As shown in Chapter 1, the number of excitations necessary for photolyase to bring about repair is not clearly known and a on the repair mechanism has not yet been reached (cf. Chapter 1).

Determining the number of photons necessary for the repair reaction is the main objective of the first part of this chapter. More specifically, we aim to distinguish between a one-and a two-photon reaction. This question has been addressed using spectroscopic methods by gauging the response of the (6-4) photoproduct-(6-4) photolyase system to blue light illumination from a steady-state light source and single-turnover flashes from a picosecond laser. These experiments hereinafter referred to as "steady-state experiments" and "flash experiments" respectively.

Most studies done on the photorepair of the (6-4) photoproduct is focused on the photoproduct formed between two thymine nucleobases (i.e. T(6-4)T)) . We have seen in Chapter 1 that the (6-4) photoproduct also exists in the form of a dimer between a thymine and a cytosine base. This form, the T(6-4)C photoproduct, occurs more abundantly than the T(6-4)T photoproduct. Presently, the mechanism of repair of the T(6-4)C photoproduct is extrapolated from that of the T(6-4)T photoproduct.

Our main objective is to determine whether or not the T(6-4)C photoproduct is repaired in the same manner as the T(6-4)T photoproduct. We present here our preliminary study of the repair of the T(6-4)C photoproduct by photolyase.

We end this chapter by presenting results of our preliminary studies of the photorepair reaction by ultrafast transient absorption spectroscopy. The only existing ultrafast study of this reaction so far was performed by Li et al. [1], on the (6-4) photolyase of Arabidopsis thaliana. Our work concerns the photolyase of a different species, Xenopus laevis, and has been performed by using an experimental technique that brings substantial improvements for the precise identification of reaction intermediates, broadband femtosecond transient absorption spectroscopy.

Photorepair of the T(6-4)T photoproduct by (6-4) photolyase

We present in this section the results on our investigation on the repair of the T(6-4)T photoproduct by (6-4) photolyase. The experiments described below were performed in the Laboratory of Klaus Brettel in iBiTecS, CEA Saclay.

Absorption spectra

T(6-4)T photoproduct

The absorption spectra of the T(6-4)T photoproduct and its repaired form, TT, are shown on Figure 1. The spectra presented correspond to measured spectra of the isolated (6-4) photoproduct and isolated TT bases in aqueous media. The spectrum of the intact TT nucleobases is shown in black on Figure 1. The spectrum is characterized by a broad absorption band between 240 and 300 nm. The absorption maximum of is situated at 265 nm with a molar absorption coefficient, ε, of 18 000 M -1 cm -1 . Upon the formation of the photoproduct between the two nucleobases, the absorption spectrum becomes the green spectrum in Figure 1. The spectrum contains a broad band from about 260 nm to 360 nm. The absorption maximum of the T(6-4)T photoproduct is found at 325 nm with an ε of 6 000 M -1 cm -1 . The two absorption maxima are well separated from each other. Hence studying them spectroscopically is ideal since we can distinguish between the two species unambiguously.

All photoproduct substrates used in our repair experiments have been synthesised by Junpei Yamamoto of the Osaka University.

Fully reduced X. laevis (6-4) photolyase

The fully reduced enzyme (containing FADH -) was obtained by reducing the oxidised enzyme (containing FAD ox ) with continuous light of wavelength 430-800 nm in the presence of a reducing agent (cysteine). The absorption of the fully reduced (6-4) photolyase is shown in Figure 2. The spectrum of the fully reduced (6-4) photolyase consists of a very intense band centred at 280 nm (not shown) corresponding to the absorption of the aromatic residues of the protein. The tail of this absorption band is seen in Figure 1 as the intense band that goes beyond λ < 300 nm. From 300 nm, the absorption decreases to about 0 in the region > 550 nm. A local maximum is found at 375 nm.

In our experiments, we used the (6-4) photolyase from Xenopus laevis. The enzyme has been prepared by Junpei Yamamoto of the Osaka University.

Photorepair under steady-state irradiation

Observation of T(6-4)T and TT restoration

To determine the number of photons necessary for repair, we sought to observe the repair of the (6-4) photoproduct as a consequence of blue light illumination. The observation of the repair was achieved by monitoring the absorbance changes in the spectrum of a solution of the T(6-4)T photoproduct and the repair active photolyase. This is done observing the absorption bands of the T(6-4)T (325 nm) and the TT (265 nm).

Nucleobases have very high absorptions in the UV. Nucleobases flanking the T(6-4)T photoproduct in a DNA oligomer would be contributing to a very high background absorption and that would saturate the absorption at this region (i.e. Abs >>> 1). As a result, absorbance changes would be undetectable. Since the restoration of the repaired bases at 265 nm is the sole conclusive evidence for repair, interfering background absorption is undesirable. In the CPD photolyase repair studies, Thiagarajan et al. from the laboratory of Klaus Brettel (iBiTecS, CEA Saclay) reported that an effective alternative to normal nucleobases is dihydrothymine (DHT) [2]. DHT is non-absorbing at 265 nm and would hence permit the observation of the restoration of the repaired bases. The DHT substrate is discussed in Chapter 4.

During blue light illumination, the (6-4) photoproduct bound to a (6-4) photolyase (PL -T(6-4)C complex) is repaired by the enzyme. The repair results to a decrease in the absorption of T(6-4)T at 325 nm. The decrease in T(6-4)T absorption is accompanied by an increase in TT absorption at 265 nm corresponding to the restoration of the previously damaged TT nucleobase. For a repair reaction involving a single photon, the loss of T(6-4)T with each photon dose is always accompanied by an increase in TT absorption since only one photon dose is required for repair. On the other hand in a two-photon mechanism, a single photon does not lead to restoration of TT. For a biphotonic mechanism, TT restoration will not be observed until after the second photon dose. The first photon will only serve as to create an intermediate from which the intact TT will originate. With the assumption that the intermediate produced from the first photon does not absorb at 265 nm, a two-photon repair can be observed if no TT restoration is observed accompanying T(6-4)T depletion with a first photon.

Shown in Figure 3 Arrows show the evolution of the spectra with increasing illumination time. The sample contained 60 µM of the DHT substrate and 1.7 µM of X. laevis photolyase. The arrows show the evolution of the spectra with every illumination period.

We observe in Figure 3 that with every increase in illumination time, the absorption at 325 nm decreases. This is concomitant with an increase in the absorbance at 265 nm. These absorption changes observed show the behaviour that is expected when photorepair of the T(6-4)T photoproduct is taking place.

To establish the number of photons involved in the repair, we must plot the observed changes as a function of the illumination time. In a one-photon reaction, we expect that every loss in absorbance at 325 nm (i.e. ~every T(6-4)T photoproduct consumed) is commensurate to the absorbance gain at 265 nm (i.e.

TT bases restored). The plot of the two absorption changes should be linear. This shows the direct relationship between the number of photons absorbed (translates to the amount of substrate consumed) to the repair that such photon absorption creates (the amount of bases restored). However, in a two-photon model of the photorepair [3], the first photon does not directly result into repair. Rather, the first photon transforms the lesion into an intermediate. Only upon the absorption of a second photon that TT restoration takes place from the intermediate. This would be seen as a non-linear behaviour in the plot of the absorbance changes in 265 nm. More specifically, it would be the onset of the 265 nm that is non- The data in Figure 4 shows a linear absorption increase at 265 nm accompanied by the bleaching of the 325 nm. Figure 4 shows the repair curves fitted with a line. The linear fit suggests that the reaction proceeds via a one-photon mechanism. The values of the slopes at 265 nm and 325 nm give rise to repair yields of 5.9 % and 6.1 % respectively. Our calculated repair yield is significantly lower by about 50% from the 11% repair yield reported in literature by Hitomi et al. [4]. The yield estimated by the authors was obtained by analysing bands of the repaired and unrepaired substrate in an electrophoresis gel. The amount of material was estimated by the relative intensities of the band on the gel. This method is less accurate as it tends to overestimate the amount of material present in a band. Our procedure for the calculation of repair yields is given in Chapter 4.

The first point in the data (green) corresponds to an illumination tile of 100 s. With an excitation rate of about 0.13 s -1 , this corresponds to 13 photons already absorbed per each absorbing centre (i.e. the FADH -cofactor). Although the curve is apparently linear, it does not exclude the possibility that a nonlinear regime can be observed at lower excitation levels. Lower excitation would lead to small changes in absorption that the steady-state spectrophotometer (resolution ~1 mOD) would not be able to detect. To be able to observe this region, we had to use a more sensitive set-up that we describe in Section 3.1.3.

Photorepair of the normal substrate

This section is aimed to demonstrate that the DHT substrate used in the photorepair studies the T(6-4)T photoproduct that we present works as well as a normal DNA oligonucleotide strand. We use the term "normal" when referring to substrates that do not contain any modified nucleobases. This is used with regard to the DHT substratea substrate that contains hydrogenated thymines (See Chapter 4). The normal 8-mer substrate has the sequence d(CGAT(6-4)TGAT). As mentioned earlier, the normal substrate only allows the visualisation of the 325 nm due to the saturation of the absorption in the UV region from the absorption of the normal nucleobases flanking the lesion.

The photorepair of the normal substrate is shown in Figure 5. The experimental conditions are exactly the same as the experiment presented in Figure 3. The spectra shown in Figure 5 show the evolution of the 325 nm band as a consequence of blue light (384 nm) illumination. The insert of Figure 5 shows the repair curve of the spectra. The linear fit of the curve gives a slope value of -5 x 10 -5 . This corresponds to a repair yield of 5.7%. The obtained value of the repair yield from the DHT substrate is comparable to this yield.

For the estimation of the steady-state repair quantum yield ( SS ),  325 of -6.0 mM -1 cm -1 was commonly used for repair of both 10-mer DHT and 8-mer normal substrates. This value was obtained from the absorption spectrum of the 8-mer normal substrate assuming ε 260 = 64.3 mM -1 cm -1 as calculated according to Tataurov et al. [5] In the case of the 10-mer DHT substrate, direct detection of the restoration of intact thymines was also available at 265 nm;  265 for repair of the DHT substrate was obtained as follows. The absorption coefficient at 260 nm of a repaired substrate that contains a TTT sequence was calculated (according to Ref. [5]) to be 15.0 mM -1 cm -1 larger than that of the unrepaired DHT substrate, hence  260 = 15.0 mM -1 cm -1 . This value was used to scale the absorption difference spectrum for photorepair of the DHT substrate extracted from experiments as those shown in Figure 1 of the main text.

The scaled difference spectrum yielded  265 = 16.2 mM -1 cm -1 .

DMAD actinometry

Determination of the excitation rate per absorbing center (k exc ) was carried out as previously described by Thiagarajan et al. [6] by using a chemical actinometer, p-(dimethylamino)benzenediazonium tetrafluoroborate (DMAD). See Chapter 4 for further details on the estmation of k exc.

The absorption coefficient of FADH -in Xl (6-4) photolyase at 384 nm,   = 5.8 mM -1 cm -1 , was derived from photoreduction experiments (see Figure 18). Figure 6 illustrates the absorption spectra and kinetics at 378 nm of the actinometer for successive irradiations in exactly the same configuration as for photorepair. In this study, we usually obtained k exc equal to 0.

Photorepair under single-flash excitation

Single-flash excitation

To measure the behaviour of the system at low excitations, we used single turnover flashes of low energy having 100 ps durations at 355 nm. The choice of conditions is based on a report of Li et al. [1].

Therein it is indicated that the lifetime of the photoexcited flavin in the PL-T(6-4)T complex, determined by the electron transfer from the flavin to the substrate, is 225 ps. They also report that restoration of the photoactive flavin, which is the return of the electron from the substrate to the flavin, does not take place before several nanoseconds. (cf. Chapter 1) It can then be inferred that, under the conditions chosen, there is virtually no FADH -, excited by a first photon and restored to its initial state, is available for reexcitation in the span of one 100-ps pulse.

To check for the existence of a non-linear regime in the repair curve (and effectively verify the single-photon model deduced from the previous section), we must determine if there is any difference in the amplitude of the absorbance change (again at 265 nm and 325 nm) induced by a first flash † and the flashes that come thereafter on a sample that has previously received no excitation at all. For the latter, we have exposed the sample to eight successive laser flashes (355 nm, 100 ps duration, 4.78 mJ cm -1 , 4 s between each individual flash, ~0.15 photons absorbed per FADH - ‡ ). We then immediately recorded and averaged the absorbance changes induced by the next eight flashes thereafter. For the first flash signal, we measured the absorbance changes induced by a single flash on a dark adapted sample.

To verify the signals that we will be measuring at 265 nm and 325 nm for the enzyme and substrate complex, we measured signals in the absence of either the substrate (S only) or the enzyme (E only). Measuring these signals allows us to rule out any contributions that would be coming from the excitation of either just the enzyme or just the substrate. These signals were obtained in the same was at that for the enzyme -substrate complex. These "control signals" are shown in Figure 8. We observe a small signal in the form of a decrease in absorption of the S only signal at 325 nm.

This signal loss is seen in both the first flash and the successive flashes case. This indicates the loss of substrate as a result of excitation at 325 nm. This loss of substrate is attributed to the formation of the Dewar isomer of the (6-4) photoproduct (cf. Chapter 1). The E only signals also show a non-negligible absorption changes as well. This absorbance change is most probably due to a spurious long lived excited state of FADH -in the absence of the (6-4) photoproduct. In the presence of the (6-4) photoproduct, the excited state of FADH -is quenched. This was reported by Li et al. [1]. † First flash given to a dark-adapted sample. A dark-adapted sample is the photoproductphotolyase solution that has been kept in the dark (and on ice) for at least 20 min. ‡ See Section 3.1.5.3.

120

For the enzymesubstrate complex, the resulting signals are shown in Figure 7a. We then averaged a total of seven acquisitions to obtain the signals shown in Figure 7b. As expected, we observe an absorption increase at 265 nm and a bleaching at 325 nm. At 325 nm, the amplitudes of the bleaching are essentially at the same level for both the first flash and the successive flashes case (amplitude of ~ -0.4 mOD). At 265 nm, there is a striking difference between the amplitude of the two signals. The signal from the first flash (amplitude of ~ < 0.2 mOD) is significantly smaller than the signal measured in the successive flashes case (amplitude ~0.7 mOD). The results indicate that during the first flash of light on a dark adapted sample, the (6-4) photoproduct is consumed. However, there is almost no restoration of the TT bases observed at 265 nm that is compatible with the loss at 325 nm. A bleaching of about -0.4 at 325 nm should correspond to an amplitude gain of about 0.6 at 265 nm. This amplitude gain was only observed several flashes after the first flash (i.e. successive flashes signal).

These results suggest that the (6-4) photoproduct is consumed in the first flash of light but is not immediately reverted to the TT. The restoration expected was not observed until several flashes after. The results highly suggest a two-photon mechanism.

The small signal measured at 265 nm after the first flash can be explained by several reasons. The first of which is the actinic effect by the monitoring light at 265 nm. It is possible that the monitoring light excites a small portion of the sample prior to the arrival of the first flash. The excited population is then re-excited by the first laser flash and hence repair takes place. The absorption increase can be due to the absorption band of the intermediate X created by the first excitation. Both of these reasons can be explained within a two-photon repair model.

Alternatively, to account for the small signal observed at 265 nm after the first flash, one can imagine a one photon repair mechanism also taking place. This would suggest that the (6-4) photolyase is capable of both a one-and two-photon repair mechanism. If this were true, our results show that a onephoton would be a minority in the repair pathway and the major repair mechanism occurs via a twophoton excitation. Otherwise, a bigger signal would have been observed for the first flash in Figure 7A.

By more sensitive measurements of the repair curve, we have seen that a non-linear regime in the repair curve exists at low excitation rates. These results are in line with what is to be observed in the occasion of a two-photon mechanism. We have shown that the first excitation of light does not immediately bring about repair but rather, it transforms the lesion into an intermediate. For the moment, we will call this intermediate X.

"Stairs" experiments

We will now try to establish the evolution of the first flash signal to the successive flashes signal.

This will allows us to construct the curve of the restoration of TT similar. To obtain this curve we sent a train of 20 flashes (with a two-second gap between each flash) to a dark adapted sample. All the while, the restoration of TT at 265 nm was continuously monitored. The resulting signal is shown in Figure 9. The obtained signal resembles a flight of stairs. Looking closely at the evolution of each step, one can observe that the height of each step is not constant. The height of each step increases with each flash (indicated by the arrows is Figure 9) until a point at about the 10 th flash where the step height appears constant with every flash.

The behaviour of the signal cannot be explained by a one-photon model. As mentioned in Section 3.1.2, a one-photon mechanism would give rise to a signal that rises linearly. We can see with the arbitrarily drawn trend line in Figure 9 that the start of the signal is more likely quadraticone that is more likely expected in the case of a two-photon mechanism. The stairs signal is not at all contradictory to the results of the steady state experiments. In addition to the lower excitation in the flash experiments, we can confirm from Figure 9 that the absorption changes accompanying a non-linear rise in the signal are too small to be resolved by the steady-state spectrophotometer.

Characterisation of X

At this point, we now have demonstrated that the repair of the T(6-4)T photoproduct proceeds via a two-photon mechanism. The first photon gives rise to an intermediate that is non-or negligibly absorbing at 325 nm and 265 nm. The second photon then gives rise to the repaired TT bases. This section is focused on the characterisation of the intermediate produced after the first photon. We will call this intermediate X.

The observation of X

During the measurement of the stairs signals, we have noticed that all stairs measured from a dark adapted sample all yield the same form. Meanwhile when a stairs signal was measured right after another stairs signal § , a different form was obtained for the latter. This different signal has a seemingly constant height for the steps beginning the first flash; linear in behaviour contrary to what has been presented in Figure 9. This is a stairs signal that is rather expected for a single photon reaction. Furthermore when we measured two successive stairs and we increase the delay after which the second signal is measured, we observed that the non-linear regime of the stairs signal reappears. In between the two measures, the sample is kept in the dark. We hence measured stairs at different dark-adaptation times. The delay with which the non-linear regime appears was shorter than that observed in Figure 9 for stairs signals measured at dark incubation periods shorter than the dark adaptation time of at least 20 min. This observation has led to the assumption that the intermediate X formed between the first and second photons has a finite lifetime. If the lifetime of X were infinite, we would be observing a constant height of § 35 s is the fastest delay at which we can measure two stairs signals.

123 the second set of stairs' steps regardless of the delay between the two measures. Presented in Figure 10 are several stairs signals measured at four different times preceding another stairs signals. In Figure 10, we observe that measuring a stairs signal 35 s after a preceding one results to a signal wherein the height of each of the steps is comparable with each other. The linear regime stars at about the 5 th flash in this case. For the stairs signal measured 2 min after a preceding one, the linear regime begins at about the 7 th flash. For a stairs measured 5 mins after a preceding one, the linear regime starts at around the 8 th flash. In order to quantify the effects described, we will define a quantity that represents the delay with which the appearance of the linear regime on each stairs signal is observed. We will call this quantity the repair lag (see Figure 10). The repair lag is the intercept of the linear regime of each stairs signal to the x-axis (i.e. baseline). This is obtained by the extrapolation of the linear fit of a stairs' linear regime. The start of the linear regime is the first flash that produces a constant step height in the steps. The different repair lags measured for each stairs signal is shown in Table 1. To observe the variation of the repair lag with the dark adaptation time, we plot these quantities against each other in Figure 11. The curve of the repair lag shows that increasing the dark adaptation time also increases the repair lag observed. The behaviour is best fitted with a single exponential plus a plateau. The time constant obtained was 1.7 min. This time constant is attributed to the spontaneous decay of the intermediate X.

The decay of X

We have seen that the intermediate X has a finite lifetime and it decays into something with a time constant of 1.7 min. The goal of the experiments described in this section is to try to determine what the intermediate X decays into. To do this, we observed the decay at various wavelengths around 325 nm.

The wavelengths were chosen around the absorption band of the (6-4) photoproduct. This is with an initial hypothesis that X will decay back to the initial (6-4) photoproduct.

We did this by building up a population of X whose long-term absorption changes we measured at single wavelengths in a spectrophotometer. The sample consisted of 8.8 µM of photoreduced X. laevis photolyase and 33 µM of the DHT 10-mer substrate. We subjected the dark adapted sample to 12 laser flashes (5.0 mJ cm -1 within ~16 s before time zero) to create the population of X. We then transferred the sample to a spectrophotometer (~10 s were required for the transfer). The absorption changes over 14 min were monitored over 310 nm, 325 nm, 347 nm and 370 nm. Two, four, and three measures were averaged for the 310 nm, 325 nm, and 325 nm signals respectively. Only one signal was recorded at 370 nm. The signals are shown in Figure 12. In order to test the hypothesis that X decays back into the (6-4) photoproduct, we compare the decay amplitudes of the traces in Figure 12. If X decays back to the (6-4) photoproduct, the decay amplitudes at the studied wavelengths should give back the absorption of the (6-4) photoproduct. When compared with the absorption spectrum of the (6-4) photoproduct, the four amplitudes must hence coincide well with the absorption of the (6-4) photoproduct. Otherwise, it would suggest that X decays into a different species.

In Figure 13, we have overlain the amplitudes of the kinetics in Figure 12 with the spectrum of the (6-4) phtotproduct. We see in Figure 13 that the decay amplitudes of X at several wavelengths correspond with their respective absorptions in the (6-4) photoproduct spectrum. We can therefore conclude that X reverts back into the (6-4) photoproduct with a rate constant of 0.01 s -1 .

Taken altogether our data strongly suggest that X does not absorb at 325 nm. It is also non-or at least weakly absorbing at 265 nm. We recall here the experiments performed by Marguet and Markovitsi on the formation of the (6-4) photoproduct [7]. In their experiments, the (6-4) photoproduct is formed via an intermediate that does not absorb at 325 nm and 290 nm. This intermediate is the oxetane bridged dimer (cf. Chapter 1). Due to its structure which is less conjugated than the (6-4) photoproduct, the oxetane species is expected to absorb at wavelengths < 300 nm. Our data is not in contradiction with their observations. Furthermore they also report that the lifetime of the oxetane species in a photolyase-free environment is 2 ms. This lifetime is much faster than our observed lifetime of 100 s. The difference can be indicative of the stabilisation of the oxetane intermediate by the photolyase. In the proposed twophoton repair scheme by Sadeghian et al. [3], the authors proposed that the intermediate species between the T(6-4)T photoproduct and the repaired TT pair is the oxetane bridged dimer. Our results are therefore not in contradiction with the predictions of Sadeghian et al. on the identity of the intermediate in a twophoton repair reaction.

Moreover, our results also do not contradict the photorepair experiments reported by Maul et al. [8]. The authors reported that they didn't find an oxetane species bound to the photolyase in the crystal structure of (6-4) photolyase bound to a T(6-4)T substrate. Our results show that the oxetane species does not form thermically from the (6-4) photoproduct upon binding to the (6-4) photolyase as previously believed [9]. After white light illumination of the crystal, the authors were able to observe the repair of the photoproduct. This was from the structure of the illuminated (6-4) photoproduct - photolyase crystal where they found the repaired TT bases. Furthermore, they observed no oxetane moiety. This observation can be explained from the fact that they completely repaired all photoproduct present in the crystal via white light illumination. Hence at the time of the structure determination, all lesions have been repaired and there is no (6-4) photolyase caught in mid-repair (i.e., with the oxetane intermediate bound to it) during the X-ray diffraction of the crystal.

Our data also suggest that the formation of X is accompanied by the return of the electron (injected to the lesion upon excitation) to the flavin chromophore to regenerate FADH -. Should this not be the case, a contribution from the FADH • -FADH -differential spectrum should have been observed.

The difference spectrum is shown in Figure 14 and was reproduced from Thiagarajan et al. [10]. If the flavin is in the radical form upon formation of X, the decay signals in Figure 12 would not be decaying to zero as we have observed. They would have otherwise decayed to the corresponding absorptions at 310 nm, 325 nm, 347 nm, and 370 nm as found in Figure 14.

Quantitative analysis of the proposed reaction scheme **

In the preceding, we have determined that the photorepair reaction by X. laevis (6-4) photolyase is a two-photon process. In the following sections we will build a model for the reaction. Our objective is to be able to build a model that describes the signals we obtained in the previous sections. The goal of building the model is to ultimately extract the quantum yields, η 1 and η 2 , of the two photoreactions. ** This part has been taken from the Supplementary Information section of our publication on the subject [11].

The order of things in the repair of the (6-4) photoproduct begins with the binding of the photolyase to the (6-4) photolesion on a DNA strand. The binding produces the enzymesubstrate complex (noted as PL-T(6-4)T). A photon is absorbed by PL-T(6-4)T to produce the intermediate X from the T(6-4)T photoproduct. The intermediate is not released by the enzyme. The complex is transformed to the enzymeintermediate complex which we note as PL-X. The yield of the production of PL-X is noted as η 1 . As observed in Section 3.1.4, PL-X decays back to PL-T(6-4)T with rate constant of k x . A second photon is absorbed this time by PL-X that transforms X into the repaired TT bases still bound to the enzyme. This is the enzyme-product complex that we note as PL-TT. The yield of the production of PL-TT is noted as η 2 . The repair ends with the release of TT by the photolyase. This scheme is summarised below: This assumption is mainly based on a study by Kondoh et al. [12] who reported the complex formation rate constant (k on ) to be 10 5 -10 6 M -1 s -1 (to be multiplied by the substrate concentration, ~10-60 µM, to give the effective binding rate of ~1-60 s -1 in this case) and the time constant for the product release to be ~50 µs (corresponding release rate: 2×10 4 s -1 ). Both numbers are compatible with the assumption.

The substrate concentration was in excess of the enzyme concentration throughout all our experiments. For repair experiments with flash excitation, the sample typically contained 4 µM of enzyme and initially 15 µM of substrate. One sample received at most 150 flashes, each of them exciting ~15% of the enzymes and repairing ~0.01 substrate per enzyme, resulting in at most ~15% decrease in substrate excess. For repair experiments with continuous wave light (i.e., steady-state experiments), samples contained 1-2 µM of enzyme and 35-60 µM of substrate, and less than 50% of the substrate was repaired.

2.

The substrate dissociation constant (about 16 nM [13]) is much lower than the enzyme (and substrate) concentration. As the substrate is in excess, virtually all of the enzyme present is in complex with DNA (PL-T(6-4)T or PL-X).

Thus, the given reaction scheme simplifies to:

PL-T(6-4)T PL-X light (η 1 ) light (η 2 )
TT + PL-T(6-4)T k x

For convenience, the following abbreviations are introduced: d, concentration of PL-T(6-4)T;

x, concentration of PL-X; r, concentration of fully repaired DNA (TT); s, total concentration of substrate (DNA containing T(6-4)T), bound to enzyme or free); s 0 , total initial concentration of substrate; p 0 , total concentration of enzyme.

From mass conservation, one obtains:

        0 0 p x d s r x s (1)
Hence, for the changes due to light or dark reactions (indicated by ∆):

             0 x d 0 r x s
(2)

Continuous light excitation

The kinetic equations describing the evolution of these concentrations under steady-state irradiation read as follows:

                  x k r x k x k d k x x k x k d k d 2 exc dt d 2 exc x 1 exc dt d 2 exc x 1 exc dt d (3)
Here, k exc stands for the excitation rate per enzyme. For monochromatic light, k exc = σ F i, where σ F is the flavin absorption cross-section at the excitation wavelength and i is the photon irradiance, expressed in photons per unit area and unit time. No distinction is made between excitation of PL-T(6-4)T, and of PL-X, which implies that the same chromophore (FADH -) absorbs light in the two cases as theorized by Sadeghian et al [3]. Note that the last term in the first equation of System (3) comes from the fact that, once the second photoreaction has converted X to intact TT, the repaired DNA strand is quickly released from the enzyme and replaced by a strand carrying a T(6-4)T lesion. The amount of X converted to TT is thus equal to the amount of newly formed PL-T(6-4)T, at the slow timescale of interest.

A steady-state (SS) regime is reached when the concentration of PL-X does not evolve any more in time, that is, when dx/dt = 0. Replacing in Equations (3) d by p 0 -x then yields:

0 x 2 1 exc 1 exc SS p k ) ( k k x       (4)
The steady state repair quantum yield in finally obtained by dividing dr/dt by the total rate of excitations, k exc p 0 .

exc x 2 1 2 1 SS k / k         (5)
The system of differential Equations (3) combined with d = p 0 -x and the initial conditions x(0) = r(0) = 0 was exactly solved yielding

                             ) ( k k t ) ( k k ) ( k k t - exp 1 - k p / r 2 1 exc x 2 2 1 exc x 2 1 exc x 2 1 2 exc 0 (6)
Equation ( 6) was used to simulate the repair kinetics expected for a two-photon mechanism with detection at 265 nm (Figure 15). For the parameters obtained in the present study and the excitation rate, enzyme concentration and optical path as in Figure 3, the expected quadratic shape of the repair kinetics detected is well visible only during the first 10 s and the corresponding absorption changes are  1 mOD (Figure 15 insert) which is well below than what can be resolved in the steady state experiments. After these few seconds a photostationnary regime establishes and the standard linear regime develops. This simulation explains a posteriori why a quadratic behaviour was not observed and the need for the more sensitive flash experiments. . Parameters: (ε r -ε s ) 265 = 16.2 mM -1 cm -1 , p 0 = 1.7 µM,  = 1 cm, k exc = 0.13 s -1 , η 1 = 6.7%, η 2 = 83%, k X = 0.01 s -1 . The insert is an expansion of the initial part of the curve.

Single flash excitation

In the following, it is assumed that for a single laser flash of 100 ps duration and rather low energy (excitation probability of ~15%) as used in the experiments, the probability to induce two successive photoreactions (excitation of PL-T(6-4)T, production of PL-X and excitation of PL-X) can be neglected.

This assumption is built upon the report by Li et al. [14] wherein the lifetime of the photoexcited flavin within the PL-T(6-4)T complex, determined by electron transfer from the flavin to the substrate, is 225 ps. The authors further found that back electron transfer from the repaired substrate, hence restoring the initial photoactive flavin, does not take place before several ns. It can thus be inferred that virtually no FADH -, excited by a first photon, is available for a second excitation within 100 ps.

The concentrations of the species just before a flash are noted with primed symbols. Non-primed symbols refer to concentrations taken just after the flash. Index n identifies the flash number. Defining α as the fraction of enzymes excited by each flash, the concentration changes induced by a single flash n can be expressed as:

                              n 2 n n n n 2 n 1 n n n n 2 n 1 n n n ' x ' r r r ' x ' d ' x x x ' x ' d ' d d d (7)
Using Equations ( 1) and ( 2), one additionally gets:

  n 0 1 n 1 n n n ' x p ' d ' s s s             ( 7')
In first approximation  can be estimated with the standard saturation formula for short pulses,

I F e 1      [15]
, where  F is again the absorption cross-section of FADH -at the excitation wavelength and I is the irradiation fluence, expressed in photons per unit area.

Flash sequence ("stairs")

The aim of this section is to obtain an analytical expression for the concentration of repaired substrate after each flash in a flash sequence as required to perform least squares fits of experimental data of this type (cf. Figure 10).

Iteration rules governing the concentration changes between subsequent flashes need to be developed, numbered by index n. For convenience, abbreviations A = α  1 , B = α  2 are introduced and Equations ( 7) are rewritten as (d n can be obtained by d n = p 0x n and is not shown):

        n n n n n n n ' x B ' r r ' x B ' d A ' x x (8)
To account for the decay of intermediate X back to T(6-4)T between two consecutive flashes, a term is introduced to represent the fraction of the X state that survives from just after flash n-1 to just before flash n. This term is called here D and is defined as

t k x e D  



. Hence:

       1 n n 1 n n r ' r D x ' x (9)
From Equations (1) one can obtain d' n = p 0x' n . Taking this together with x' n and r' n from Equations ( 9) to rewrite equations (8), the iteration rules are obtained:

            1 n 1 n n 1 n 0 n r x B D r x ) B A 1 ( D A p x (10)
The system of Equations ( 10) are then rewritten with the following matrix notations:

         n n n r x V ,          0 A p K 0 and            1 B D 0 ) B A 1 ( D M
One then simply gets:

1 - n n V M + K = V  (11)
Leading to:

0 n 1 - n 0 k k n V M K M = V           ( 12
)
V 0 is the concentration vector before flash 1. By diagonalizing matrix M, it is straightforward to obtain its k th power and to calculate the geometrical series M k . Careful calculations ultimately lead to:

                                           n 0 x n 0 n n 0 x n 0 n U 1 U 1 D B f U 1 U 1 n U 1 D B A p r U f U 1 U 1 A p x (13)
with U = D(1-A-B). It was here assumed that some initial concentration of PL-X was present before flash 1, due to the actinic effect of the continuous monitoring beam as mentioned earlier. This was expressed by starting conditions, x 0 = p 0 f x0 and r 0 = 0, f x0 being the fraction of complexes initially in the PL-X form.

For simplicity of the model, actinic effect of the monitoring light during the flash sequence was neglected, as it has been estimated that the corresponding excitation to be less than 10% of the mean excitation made by the flashes.

The predictions of Equations ( 13) were checked to be identical to those of the elementary iteration rules (Equations 10).

Estimation of the repair quantum yields η 1 and η 2

Equations for the absorption changes

Concentration changes and quantum yields from observed absorption changes were deduced according to relation ΔA λ = Σε j,λ Δc j , where ΔA λ is the change of absorbance at wavelength λ, ε j,λ is the molar decadic absorption coefficient of species j at wavelength λ, Δc j the change of concentration of species j, and  the optical path length. Considering only DNA based absorption changes (flavin-based changes are too fast [14] to be observed at the low time resolution used here) and Equations (2), one obtains:

ΔA λ / = (ε x,λ -ε s,λ ) Δx + (ε r,λ -ε s,λ ) Δr (14)
At 265 nm, (ε x -ε s ) can be neglected (neither X nor T(6-4)T absorb at 265 nm), and (ε rε s ) = 16.2 mM -1 cm -1 (see Section 2).

At 325 nm, (ε x -ε s ) = (ε r -ε s ) = -6.0 mM -1 cm -1 (neither X nor intact thymine absorb at 325 nm).

Hence:

ΔA 265 / = 16.2 mM -1 cm -1 × Δr (

ΔA 325 / = -6.0 mM -1 cm -1 × (Δx + Δr) = 6.0 mM -1 cm -1 × Δs (16)

Determination of η 1 and η 2 from flash (first vs. successive) excitation

The quantum yield η 1 of the first photoreaction can be directly determined from the absorption change at 325 nm upon irradiation of a dark adapted sample by the first flash (ΔA 325,1st ). For the first flash, one can use s 1 = - η 1 p 0 (Equation (7') with x' 1 = 0) together with Equation ( 16). To do so, a correction is done for a small bleaching at 325 nm due to conversion of T(6-4)T to its Dewar isomer [16,17] upon direct excitation of T(6-4)T in its long-wavelength absorption tail by the 355 nm flash. This effect was observed in control samples containing only substrate, but no enzyme (Figure 7, right, traces marked "S only") † † .

The result of this correction is shown in Figure 16 (blue trace). Furthermore, a small, but non-negligible absorption increase at 325 nm was observed in control samples containing only enzyme, but no substrate (Figure 7 right, traces marked "E only"), presumably due to some spurious long-lived photooxidation of FADH -. Additional correction for this signal yielded † † The quantum yield of T(6-4)T to Dewar conversion can be estimated from the amplitude of the "S only" signal (ΔA 325 ≈ -0.06 mOD for 13 µM substrate at I = 8.54×10 15 photons cm -2 ), using σ 355,T(6-4)T = 3.2×10 -18 cm 2 (calculated from the absorption spectrum in Scheme 1 of the main text) and complete bleaching of the 325 nm band of T(6-4)T upon Dewar formation. We obtained ~3%, as compared to literature data of 2%and (8.2±2)% the red trace in Figure 16. Photooxidation of FADH -may be less prominent in the presence than in the absence of substrate, because the latter quenches the excited state of FADH - [14].

The two corrected signals shown in Figure 16 were hence considered as limiting cases for the estimation of η 1 . From the amplitudes (averaged between 0.3 s and 1 s; namely, -0.283 and -0.425 mOD)

together with α = 0.145 (for σ 355,FADH-= 1.84×10 -17 cm 2 , corresponding to ε 355,FADH-= 4.8 mM -1 cm -1 (cf.

Figure 18), and I = 8.54×10 15 photons cm -2 , corresponding to 4.78 mJ cm -2 at 355 nm) and p 0 = 4.9 µM, 6.6% < η 1 <10.0% is obtained.

The quantum yield η 2 of the second photoreaction could be estimated from the absorption change at 265 nm in the successive flash regime (SFR), using Equation ( 15) and r SFR = α η 2 x' SFR and provided that x' SFR is known. According to Equations (7') and ( 16):

ΔA 325,SFR /ΔA 325,1st = s SFR /s 1 = (p 0x' SFR )/p 0 .

Hence,

x' SFR = (1-ΔA 325,SFR /ΔA 325,1st ) p 0 (18) Inspecting the 325 nm signals in Figure 8 and taking into account the control signals and the noise, it can be excluded that the successive flash regime amplitude ΔA 325,SFR was less than 80% of the first flash amplitude ΔA 325,1st . Hence, x' SFR < 0.2 p 0 . After correction for a minor absorption increase observed in samples with only enzyme, but no substrate (Figure 7; the signal for substrate only was negligible), the averaged (0.3-1 s) 265 nm successive flash regime amplitude ΔA 265,SFR yields η 2 > 27%. Without this correction, η 2 > 28% is obtained.

Determination of η 1 and η 2 from flash sequence ("stairs")

An independent estimation of η 1 and η 2 was obtained by fitting the 265 nm absorption change trace for 20 subsequent flashes given to a dark adapted sample (Figure 9).

A fitting model was constructed with the formalism outlined in Section 3.1.5.4, essentially combining Equations ( 13) and ( 15), and additionally giving the A signal an exponential rise time (τ rise ) after each flash. The resulting fit function can be written in the following compact form:

                 tot rise n n 1 n n 1 n n / ) t t ( 265 s r 265 ) t t ( ) r r )( e 1 ( A  (19)
In this expression t n is the arrival time of the n th flash. H(x) is the unit step function, taking 0 value for x < 0 and 1 elsewhere. Further, r n is taken from Equation ( 13) with A = α  1 , B = α  2 , and For the experimental data in Figure 9, the first flash was at t 1 = 0, the flash interval t n+1 -t n was 2 s and the total number of flashes n tot was 20. As seen in section 2.4.4.1, (ε r -ε s ) 265 = 16.2 mM -1 cm -1 . The optical path  was 1 cm. τ rise was fixed at 0.1 s -1 corresponding to the response function of the detection system. k x was fixed at 0.01 s -1 , according to Figures 10 and11. The total concentration of enzyme, p 0 , was imposed to 3.9 µM. The only adjustable parameters of the fit were A, B and f x0 .

The best fit was obtained for A = 0.0128, B = 0.1587 and f x0 = 0.0188. With our estimation of α (0.191 ‡ ‡ ), values of  1 = 6.7% and  2 = 83% were deduced. The values of η 1 and η 2 obtained are in line with the prediction that for the observed signals. The yield of the first photoreaction is smaller than the second.

Quantum yield uncertainty estimation

From continuous light excitation

In Figure 17, we compare the acceptable values of  1 and  2 obtained by the analysis of three independent experiments, steady state photorepair under continuous light (Figure 3), absorption changes upon the first flash from dark and in the successive flash regime (Figure 8), and absorption changes in a series of 20 flashes from dark ("stairs"; Figure 9). ). The zones limited by red curves are deduced from the fit of the "stairs" experiment, and our simulations of it (cf. Section 3.1.5.3.3). The red dotted curve corresponds to  = 0.191 and p 0 = 3.9 µM. The (internal) thin red solid curve corresponds to  = 0.191 and p 0 varying between 3.7 and 4.1 µM. The (external) thick red solid curve corresponds to  varying between 0.156 and 0.225, and p 0 varying between 3.7 and 4.1 µM. The quantum yields obtained from the best fit of the "stairs" experiment ( 1 = 6.7% and  2 = 83%) are recalled by the red cross.

Under continuous light,  1 ,  2 and  SS are linked by Equation ( 5) which can be rewritten as: ‡ ‡ The excitation energy in the "stairs" experiment was larger than for the "first vs. successive" measurement.

1 / k / k SS 1 exc x 1 2        (20)
Taking into account the major error sources in the determination of η SS (estimated ±5% for the photon irradiance, ±5% for the absorption coefficient of FADH -at 384 nm and ±5% for the enzyme concentration), the results of steady-state irradiation (Section 3.1.2) provide two limiting values of  SS , namely 5.1% and 6.9%. According to Equation (20), each of these values, together with k x /k exc = 0.0769, in turn defines a hyperbolic line in { 1 ,  2 } space (blue curves in Figure 17).

From flash (first vs. successive) excitation

From the amplitude of the 325 nm absorption change upon the first flash from dark, we derived 6.6% < η 1 <10.0%. These first limits did not take into account potential errors in the flash energy (estimated to ±15%) and in the absorption coefficient of FADH -at 355 nm (±5%), which combine to produce a relative error on  of about 18% (taking into account the saturation formula), and the errors on the enzyme concentration (±5%). Including them, we finally obtain 5.4% < η 1 <11.8% (green dashed horizontal lines in Figure 17), to which we add η 2 > 27% from Section 3.1.5.5.3 (green dashed vertical line in Figure S17).

From flash sequence ("stairs")

One then needs to discuss the confidence corresponding to the fit of the "stairs" experiment. It should first be noted that the standard errors on parameters A and B (4×10 -5 and 1.64×10 -3 , respectively), provided by the fitting algorithm, are not relevant to our problem. The reason is that the model described in the beginning of the section assumes that the energy of each shot is constant. In the actual experiment the shot-to-shot laser fluctuations were of the order of 6% (relative value). Given that the A signal after flash n depends on the cumulated effect of the n-1 preceding flashes, proper inclusion of the laser fluctuations would require fitting 20 different values of A and B, which is not tractable.

To tackle this issue we performed simulations of the "stairs" experiment, based on the iteration rules previously described in Section 3.1.5.1, and additionally allowing parameter α (hence A and B) to randomly fluctuate from flash to flash. Since the actual "stairs" experiment was the average of two separate measurements, we only allowed fluctuations of 6% / 2 = 4.24% about an average α (noted α * ) of 0.191. 10000 simulations were performed using as generators the optimal parameters deduced from the fit of the experimental "stairs" (i.e. η 1 = 6.7%, η 2 = 83%, f x0 = 1.88%). Each simulation was then fitted by the procedure of Section 3.1.5.1. We therefore obtained a pool of 10000 optimal A, B and f x0 parameters corresponding to the chosen fixed generators, that we will now note ) is the so-called prior probability that real parameters are η 1 * and η 2 *

, not knowing the result of any fit. [18] It can be assumed to be flat in (η 1 , η 2 ) space. p(A, B) can be thought of as a normalization factor. [18] With the above general premises one can write the following proportionality relation:

p(η 1 * , η 2 * | A, B)  p(A, B| η 1 * , η 2 * ) (22)
It should be noted that our simulations directly provide p(A, B| η

) as a function of A and B for fixed η 1 * and η 2 * , but not as a function of η 1 * and η 2 * for fixed A and B.

A simplification appeared when we substituted A and B by two new variables, x and y, defined as follows:

                           B A B A 0.999962 0.00870995 - 0.00870995 0.999962 y x (23)
We empirically observed with different sets of (

where n x and n y are 2 independent normal distributions of variables x and y, centered on values (x * and y * ) deduced from A * = α * η 1 * and B * = α * η 2 * by Equation (23). Importantly, their standard deviations ( x = 0.021,  y = 0.000029) do not substantially depend on x * and y * .

We finally build a contour in (η

) space such that there is 95% probability that (η

) lies within this contour. Given the above equalities this reduces to finding a contour in (x * , y * ) space over which the integral of p(x, y| η 1 * , η 2 * ) is equal to 0.95.

In order to let x * and y * play equivalent roles, we introduce the following polar coordinates:

                 sin r / y y cos r / x x y * x * (25) With these notations, ) r 2 1 exp( 2 1 ) , | y p(x, 2 y x * 2 * 1       (26)
The desired contour will be looked for as an ellipse, centered in point (x, y) deduced from the experimental fit parameters, limited by maximal r equals to R. Integrating over this ellipse, we get:

) R 2 1 exp( 1 rdrd ) r 2 1 exp( 2 1 dy )dx , | y p(x, 2 y x 2 y x * * * 2 * 1               (27)
The value of R corresponding to 95% probability is 2.45.

The corresponding contour in (η 1 , η 2 ) space is shown in Figure 17 (red dashed line). The most probable value ( 1 = 6.7% and  2 = 83%) is recalled with a red cross.

For one fixed value of  it is straightforward to predict the effect of varying p 0 on  1 and  2 . Equation ( 13) indeed reveals that r n is invariant when: (28) This means that the same fit of the "stairs" data can provide different values A and B, bound by Equation (28), when p 0 is changed. One may note that, in our domain of interest, B takes much larger values than A, which implies that the relative variations of B are much smaller than those of A. If one completely neglects them, it appears that the relative variations of A are simply equal to those of p 0 (with opposite sign). The red thin solid line in Figure 17 represents the union of all confidence contours associated to  = 0.191 and values of p 0 found in ±5% interval around the most probable concentration (3.9 µM).

            constant c B f p constant c B A p constant c B A 3 0 x 0 2 0 1
Next, we calculated as above the confidence contours associated to limiting values of  ( = 0.156 and  = 0.225), corresponding to potential errors in the flash energy (estimated to ±15%), and of the enzyme concentration (±5%). The global confidence area, including all sources of errors, is delimited by the thick red solid lines of Figure 17.

Photorepair under solar irradiation

From equation 5, it can be deduces that the steady-state overall repair yield η ss decreases hyperbolically with the increasing value of the ratio exc x k k . A low excitation rate such as that due to the low photon density on the earth's surface can mean an inefficient repair via a two-photon mechanism. To evaluate the plausibility of the two-photon model under natural conditions, we calculated the k exc of natural light. The excitation rate k exc of FADH -in Xl (6-4) photolyase exposed to sunlight was estimated using the following formula:

         d ) ( I ) ( k 2 1
FADH exc (29) where  FADH-(λ) is the absorption cross-section of the fully reduced flavin at wavelength λ and I(λ) is the solar spectral irradiance at wavelength λ. The formula is an extension of the k exc formula we used for monochromatic light in Section 3.1.5.1.2. To calculate for the k exc of natural light, the formula was integrated over the range of the solar irradiation spectrum that corresponds to the absorption of the FADH -chromophore (i.e. 310-550 nm). For I(λ), the terrestrial solar irradiation spectrum ASTM G173 provided by the American Society for Testing and Materials (ASTM) was used [19]. This spectrum represents the annual averaged daylight terrestrial hemispherical irradiance incident on a sun-facing 37° tilted surface at clear sky conditions for the 48 contiguous United States.

The  FADH-(λ) spectrum was obtained from a photoreduction experiment. Of note, the absorption by tryptophan residues contributes significantly to the absorption spectra in Figure 18b up to ~310 nm, but I(λ) is negligible below 310 nm. Integrating over the FADH -absorption region from 310 to 550 nm, one obtains k exc = 0.37 s -1 . Within a living cell, I(λ) seen by the enzyme could be decreased due to light absorption and scattering by other constituents of the cell. On the other hand, the effective absorption cross-section of photolyase may be increased by the -yet uncertain-presence of an antenna pigment, similarly to CPD photolyases [20][21][22]. [19] were converted from W m -2 nm -1 to photons s -1 cm -1 nm -1 . (b) Absorption spectrum of Xl (6-4) photolyase with fully reduced flavin cofactor (FADH -; red line) obtained by photoreduction (see Section 1) of the enzyme containing fully oxidized flavin (FAD ox ; black line). The absorption cross sections σ (right hand scale) were obtained by scaling the spectrum before photoreduction to ε = 11.2 mM -1 cm -1 in the 450 nm peak of FAD ox which corresponds to σ = 4.28 × 10 -17 cm².

Despite these uncertainties, it appears plausible that the excitation rate k exc under natural conditions can well compete with the dark decay of intermediate X at k x ≈ 0.01 s -1 . A non-stabilized oxetane would convert back to the (6-4)PP at a rate of ~500 s -1 [23], so that the two-photon repair mechanism would be very inefficient under natural light. Stabilization of the oxetane by the enzyme appears hence to be crucial for the repair efficiency of (6-4) photolyase under natural conditions.

Preliminary study of the steady-state photorepair of the T(6-4)C photoproduct

So far, whenever we speak of the photorepair of the (6-4) photoproduct, we always speak of the repair of the dimer formed between two thymine residues: T(6-4)T. This section is dedicated to our studies on another form of the photoproduct. This form is the protoproduct between a thymine and a cytosine nucleobase. In Chapter 1, it was pointed out that the literature on the T(6-4)C repair is greatly outnumbered by that of T(6-4)T form of the lesion. Furthermore, it is assumed that the repair mechanism of the T(6-4)C photoproduct is the same as that of the T(6-4)T [24]. The repair mechanism for the T(6-4)C photoproduct is hence simply extrapolated from that of the T(6-4)T [25][26][27]. Hence it is assumed that T(6-4)C repair follows the same mechanism of T(6-4)T repair. In light of our results that the T(6-4)T photoproduct is repaired via a two-step mechanism involving two photoexcitations, we wish to verify if this mechanism holds true for the T(6-4)C photoproduct.

The interest in studying the T(6-4)C photoproduct is from the fact that it is more easily formed in vivo than the T(6-4)T photoproduct. The T(6-4)C photoproduct is hence more abundant [28]. The greater abundance of T(6-4)C formation is mainly due to its higher formation quantum yield of 0.75 compared to 0.5 for T(6-4)T [29]. The T(6-4)C photoproduct in greater depth in Chapter 1.

We begin our study of the T(6-4)C repair by observing the repair of the T(6-4)C photoproduct by (6-4) photolyase from X. laevis.

The T(6-4)C photoproduct

The T(6-4)C photoproduct as shown in Figure 19 is the (6-4) photoproduct from the [2+2] cycloaddition between an adjacent pair of a thymine (5'-side) and a cytosine (3'-side) on a strand of DNA.

T(6-4)C As mentioned in Chapter 1 (section 1.3.1.1) despite its facile in vivo formation, the T(6-4)C is not easily synthesised in the laboratory as compared to the T(6-4)T photoproduct [30]. This is the reason why the studies on the T(6-4)C photoproduct are less common. The absorption spectrum of the T(6-4)C photoproduct is shown in Figure 20. The spectrum of the T(6-4)C photoproduct contains a maximum at 315 nm corresponding to the absorption of the (6-4) photoproduct. The 315 nm maximum is blue shifted with respect to the 325 nm of the T(6-4)T photoproduct. The difference is due to the less polar amino group (less polar than the hydroxyl group of T(6-4)T) that shifts the absorption peak to the blue. The absorption band observed around 220 nm belongs to the phenyl moieties of the dimethoxytrityl (DMT) protecting group of the phosphoramidite building block (See Footnote § §).

Steady state photorepair of the T(6-4)C photoproduct by X. laevis (6-4) photolyase

The steady-state photorepair was performed under the exact same conditions as the photorepair of the T(6-4)T photoproduct described in Chapter 2. The sample consisted of about 2 µM of the photolyase 35 µM of the substrate in a 10 mM phosphate buffer containing 100 mM NaCl, 6 mM cysteine, and 5%

glycerol. The same excitation rate of 0.13 s -1 was used for the 384 nm illuminating light for photorepair.

For the substrate, we used an oligomer of 10 nucleobases containing one T(6-4)C photolesion of the sequence: d(ACAGCCGT(6-4)CGCAGGT). The result of the photorepair experiment is shown in Figure 21. § § Phosphoramidite = phosphite diester (RO) 2 PNH 2 . Precursor of the oligonucleotide containing the T(6-4)C photoproduct. In the phosphoramidite form, the species is just the (6-4) photolesion that is attached to DMT (Dimethoxytrityl, protecting group) and the phosporamidite at the 5'-and 3'ends of the lesion respectively. In Figure 4 we can observe that the DHT substrate is repaired, as seen from the absorption changes in the repair curve at 315 nm and 265 nm. The shape of the DHT repair curve is however quite different from that of the normal substrate. We see that it is no longer linear. The first few points on the curve appear approximately aligned but, after a few hundreds of seconds of illumination, the initial linearity appears to be lost. We observe that a decreasing amount of substrate is repaired with every new illumination period. The DHT repair curve is in fact best fitted with an exponential function of the form ) 1 (

) ( 0 t t k e A y    
. We obtained A 266 nm = 0.19344 for the 266 nm curve and A 315 nm = -0.07676 for the 315 nm curve. They share a value of 5.56  10 -4 s -1 for k and t 0 was fixed at 60 s. Solving for the initial slope (i.e. A×k) 1.08  10 -4 s -1 is the value obtained for the 266 nm curve and 1.08  10 -4 s -1 for the 315 nm curve. Comparing the 315 nm slope from the DHT experiment and that from the normal substrate we find that the slope from the DHT experiment is in fact 2.16 times larger than that of the normal substrate.

The above non-linear behaviour may be explained by several reasons. One of them could be the loss of the enzyme during the course of the experiment. The deactivation of the enzyme (via oxidation) over time will indeed result to less substrate being repaired and hence explain what we observe in Figure 4. The oxidation of the enzyme leads to the formation of FADH • or FAD ox , which can be easily observed in the absorbance spectra. Figure 23 shows the differential spectra for the spectra in Figure 22. Since the appearance of the absorption bands for FAD ox (λ max = 450 nm) or FADH • (broad absorption band between 500 nm and 675 nm) are not observed in Figure 23, the deactivation of the enzyme can be excluded.

250
Another reason for the observed non-linear behaviour could be the overestimation of the amount of added substrate. The ratio of the enzyme to substrate was chosen to be 1:5 to ensure that there will be substrate available for repair. Should this ratio become overestimated, the actual concentration of the substrate in the sample would be lower than what is believed. This will lead to a point in the curve where the concentration of the (6-4) photoproduct is less than the enzyme. What would be observed then is a decrease in the amount of substrate repaired over time. An overestimation of the substrate may come from impurities in the substrate solution. During data analysis of the photorepair, it has come to our attention that the T(6-4)C DHT substrate was in fact not pure. Reverse phase HPLC analysis indeed a posteriori revealed that an unidentified contaminant was present in the solution of the DHT nucleotide used for building the DHT oligomer. The amount of this unknown contaminant is undetermined. It was however "undetected" by NMR when DHT was verified prior to building the oligomer. This means that the contaminant has similar shifts as DHT in the NMR spectrum *** . The contaminant was possibly integrated into the oligomer and may be responsible for the peculiarities observed in the repair of the DHT substrate.

The said contaminant has an absorption maximum at 250 nm. The contaminant then interferes with the detection of the TC restoration at 265 nm in the photorepair experiments. The 250 nm band appears to be intermediate between the thymine absorption at 260 nm and the DHT absorption at 220 nm.

Furthermore, the contaminant is less polar than DHT. It has a retention time (t r ) of 18 min. compared to 15.1 min. of DHT in reversed phase chromatography. Since DHT is produced from the catalytic *** Possible NMR differentiation between the two may be sought using 2D NMR (i.e. 1 H COSY) but was not performed.

hydrogenation of thymine, the contaminant may then be a side product of hydrogenation. This may be the incomplete hydrogenation of the C=C bond in the pyrimidine ring or the hydrogenation of the C=O group of the ring. The latter would most likely lead to a shift in the absorption to the UV region as it leads to a decrease in the degree of unsaturation of the thymine ring. The loss of a C=O group would also decrease the polarity of the species which could explain the lower polarity observed.

Due to insufficient amount of material, the photorepair measurements for the T(6-4)C photoproduct was not repeated. Only a single experiment was afforded for each type of substrate. For a valid conclusion to be reached, the experiment needs to be repaired with a purer substrate. Furthermore, we did not calculate the repair yields for the experiments we have presented. This is because of the fact that we do not yet have a reliable value for the ∆ε at 265 nm and 315 nm.

Preliminary studies of T(6-4)T photorepair by femtosecond pump-probe spectroscopy

We present here the results of our preliminary femtosecond studies of the T(6-4)T photorepair by X. leavis means that for samples of small volume (typically 50 µL) such as ours, even though the irradiation is made on a very small surface (diameter of the order of 100 µm) and the sample cuvette is moved in two dimensions (Lissajous curve), each point of the sample is bound to be excited several times. Starting with an initially dark-adapted sample (which does not contain X), it can be foreseen that the average population of X will grow with time during the course of the experiment. A rough answer to the question at hand is provided by the following considerations. According to the results presented in §3.1 the quantum yield of the first photoreaction ( 1 ) is 6.7% and that of the second one ( 2 ) is 86%. In the worst case we will suppose that a photodynamical equilibrium is reached at each point of the sample, meaning that the concentration of X remains constant (equal amounts being formed and repaired by next excitation pulse). In that case the fraction of X (respective to the total concentration of active protein) cannot be larger than  1 /( 1 + 2 ) (see Equation 4), hence ~7%. This number completely neglects the fact that X is metastable and decays to the initial dark state with a lifetime of 100 s. Because of the complex 2D movement of the sample cuvette and the phenomenon of diffusion it is not straightforward to quantify for our set-up the time elapsed between consecutive irradiations of the same volume. It can however be inferred that the average fraction of X is likely less than 7%. This limit is also probably overestimated because full photodynamical equilibrium at each point of the cuvette may not have been actually reached.

For the above reasons we will consider that our experiments essentially probed the first photoreaction.

We will give below further details on the choice of substrate to protein ratio.

In the sections that follow, we present the transient absorption dynamics of fully reduced photolyase from X. leavis (Xl64), first in the absence of substrate, then in the presence of a T(6-4)T photoproduct. The first experiment will explore the intrinsic dynamics of excited FADH -within Xl64.

The second one, we will reveal the specific reactions due to the presence of substrate. All spectra reported in this section were measured at the magic angle, and thus correspond to rotationally isotropic transient populations.

Reference spectra

We shall begin with a presentation of a few useful results on the spectroscopy of FADH -. What we will be presenting are data reported by Kao et al. [32] and Brazard et al. [33].

Steady-state fluorescence spectra

The absorption spectrum was of the fully reduced enzyme was already described in Section 3.1.1.2 and will not be repeated here. Shown in Figure 24 are the fluorescene spectra of FADH -: free FADH -in a 12.5 mM phosphate buffer solution at pH 8.5 containing 100 mM NaCl and of FADH -in EcCPD in 50 mM Tris buffer at pH7.5 containing 100 mM NaCl, 1 mM EDTA, 5 mM dithiothereitol, and 50 % (v/v) glycerol. These spectra are reproduced from Kao et al. [32]. Fluorescence spectra of FADH -when in solution (black) and when bound to EcCPD photolyase (red) excited at 360 nm extracted from Kao et al. [32].

Shown in black is the emission spectrum of free FADH -is solution. The emission peak of the free flavin has a maximum at 450 nm. The emission band is from about 475 nm to 650 nm. The maximum of the flavin bound to the photolyase is found red shifted with a maximum at 545 nm. Another peak is also found at 515 nm. Kao et al. also reported that the fluorescence of FADH -in solution is weaker compared to when bound to photolyase, suggesting very different dynamic behaviours.

Transient absorption spectra of FADH -in solution

The transient absorption spectrum of free reduced FAD in solution (100 mM Tris pH 8, 100 mM NaCl, 10% glycerol) has been reported by Brazard et al. and is reproduced in Figure 25.

mM cysteine and 25% glycerol, with ~430-800 nm continuous light until the spectrum of the oxidised Xl64 is fully converted to that of the fully reduced Xl64. The sample was excited at 387.5 nm. A total of 21 scans were produced in the course of the experiment (see scan definition in Chapter IV, Section 4.2.2.4). To ensure the sample always contains the fully reduced enzyme, the absorption spectrum was monitored regularly during the experiment and re-reduced when the oxidised species is detected.

Initial transient absorption spectrum

Following femtosecond excitation at 387.5 nm, the early transient absorption spectrum of the fully reduced X. laevis photolyase, in the absence of substrate, is shown in Figure 26, for a pump-probe delay of 300 fs. The transient absorption spectrum of the fully reduced photolyase contains two positive bands centred at 420 nm and 575 nm. These bands are dominated by the transient absorption of the excited state of FADH -* . A minimum between the two bands is observed at 450 nm. The stimulated emission of photolyase-bound FADH -is expected to contribute negatively around 550 nm. The ground state bleaching is also expected to contribute negatively in the region below 500 nm. The absence of a negative band however, shows that the transient absorption is very dominant across all wavelengths. Negative contributions may be responsible for the spectral shapes that we observe. One may note there is an apparent red shift of the two transient absorption bands relative to the 390-nm and 520-nm peaks observed in free reduced FAD in solution. The structure of the peaks in Xl64 is also narrower than those of free FADH -. These differences may be due to the specific environment of the protein (more rigid, less polar, specific interactions of the flavin with neighbouring molecules) and/or to the particular U-shaped structure of FAD in the protein.

Spectral dynamics of the transient absorption spectrum

The chirp-corrected transient absorption spectra of the fully reduced (6-4) photolyase, for increasing pump-probe delays between 300 fs and 3.2 ns, are shown in Figure 27. The temporal evolution of the spectra can be divided into four phases. This evolution is described as follows:

350
 Between 0.3 and 1 ps (Figure 27A): increase in the amplitudes of the 420 nm and 575 nm peaks.

 Between 1 and 4 ps (Figure 27B): very small increase in the amplitudes of the 420 nm and 575 m bands. The  Between 1 and 65 ps (Figure 27C): moderate decay around 365 nm accompanied by an increase in amplitude of the 420 nm. A narrowing of the 420 nm band is hence observed. Slow decay of the 575-nm band. A slight increase in amplitude of the minimum at 450 nm is also observed.

 Between 65 and 3200 ps (Figure 27D): Uniform decay of all bands. A residual signal remains at 3200 ps.

No major change in the global spectral shape of FADH -* is observed during the temporal evolution of the transient spectra. This most likely indicates that what we observe in all Figure 27 is the complex decay of the excited FADH -. We will reserve the discussion of these observations until after we present the results of the global analysis of our data.

Global kinetic analysis

The data were best fitted with a sum of four exponential components with the following lifetimes: τ 1 = 0.35 ± 0.15 ps, τ 2 = 1.1 ± 0.6 ps, τ 3 = 22 ± 3 ps, and τ 4 = 3750 ± 55 ps. The average residue of the fit was 71 µOD. The decay-associated difference spectra resulting from the fit are presented in Figure 28A and the evolution-associated difference spectra (EADS) are shown in Figure 28B. They reveal that essentially the same species is involved at all phases of the decay. The "overall" shape of these transient spectra corresponds qualitatively well to that of the transient spectrum of free FADH -in solution (Figure 25). We hence conclude all spectra correspond to the decay of FADH -* . What we observe is hence a multi-exponential decay of FADH -. The first time constant that we measured (0.4 ps) is attributed to the general increase of amplitude in our transient absorption spectra between 0.3 and 1 ps as seen in Figure 27A. This is also observed in the evolution of EADS1 to EADS2. We also see clearly in the EADS that the other two time constants (1 ps and 22 ps) are observed as an increase in amplitude of the spectra. We can see that the 420 nm band is in effect narrowed in EADS3 as compared with EADS2 and EADS1. This is most evident from the decreasing amplitude of the blue wing of the 420 nm band from EADS1 to EADS3. Narrowing of the 575 nm band however is not evident from the spectra and a plausible explanation is difficult to reach at this point. We hence partially assign the 0.4 ps, 1.1 ps, and 22 ps to the vibrational relaxation of the excited FADH -.

We now move on to the 3750 ps time constant. It be noted that this is the main decay as we can see from the amplitude of its DADS. Compared to the time constants of free FADH -in solution reported by Kao et al. and Brazard et al., this time constant is significantly longer. This reflects the different relaxation dynamics of the bound FADH -compared to free FADH -that we have evoked while comparing the fluorescence spectra of the two species. Indeed it can be imagined that there is a difference in the butterfly-like mechanism of FADH -* relaxation ( §3. 3.1.2) between the free and bound species. The more confined and rigid environment of the protein's interior as compared to an aqueous solution can explain the longer time constant that we observed.

Transient absorption spectroscopy of fully reduced Xl64 in the presence of T(6-

4)T substrate

Before reporting the transient absorption behaviour of Xl64 in the presence of substrate, i.e.

during repair the (6-4) photoproduct, let us quickly examine one key experimental issue: the choice of substrate to enzyme ratio. It is indeed important that during the course of the experiment the concentration of active enzyme (reduced protein binding a substrate) is maintained constant in spite of the fact that repair of the substrate readily increases.

We will start with a rough calculation just to give an idea. For simplicity we will assume that photodynamical equilibrium is reached at each point of the sample cuvette. If that is not the case the effective repair yield per excitation shot will be smaller than what is estimated next. At photodynamic equilibrium the repair yield per excitation is at most equal to  1  2 /( 1 + 2 ), that is, about 6% (in fact less because of the decay of X, neglected here).

Let us suppose that the fraction of flavins excited per shot is typically 5% and the surface irradiated is about 10 -2 mm² (diameter of ~110 µm; hence excited volume of 10 -2 mm 3 ). For a total sample volume of 50 mm 3 , the effective repair fraction per excitation pulse should not be larger than 5%×6%×10 -2 /50 = 6×10 -7 of the total concentration of active enzyme. Given that 1 scan requires about 1500 shots×58delays, i.e. 87000 shots, the effective repair fraction per scan will at most be 6×10 -7 × 87000 = 5%. In other terms 20 scans would repair an amount of substrate equal to be concentration of enzyme. At least two or three equivalents of substrate must be used to be on the safe side.

In practice we used a 7:1 ratio of the substrate to the enzyme. The enzyme was photoreduced as described earlier in Section 3.3.2. The sample was in a 10 mM phosphate buffer at pH 7 containing 100 mM NaCl, 6 mM cysteine, and 25% (v/v) glycerol. A total of 27 scans were obtained for this experiment. We continuously checked during the course of the experiment the evolution of the absorption band of the substrate at 325 nm. We observed at the end of the 27 scans, we have repaired about half of the amount of substrate originally present -which is somewhat more than what was roughly predicted above. We thus conclude that the substrate to enzyme ratio was effectively large enough to keep the concentration of active enzyme constant.

Let us finally note that the substrate was used for this experiment was the normal 8-mer substrate having the sequence d(CGAT(6-4)TGAT). Since the region below 350 nm is inaccessible in our experiment, the use of the DHT substrate is not necessary.

Spectral dynamics of the transient absorption spectrum

We now report the transient absorption spectra of reduced Xl64 in the presence of T(6-4)T substrate. The evolution of the transient absorption spectrum is shown in Figure 30. The temporal evolution of the spectra can be divided into four phases and is described in the following:

350
 From 0.3 ps to 3.2 ps (Figure 30A): small amplitude decrease on the blue side of the 420 nm band, around 365 nm. Weak increase of the 420 nm peak concomitant with a small decrease of the minimum at 450 nm. There is a marked increase in the amplitude of the peak at 575 nm and a decrease of amplitude in the region between 650 and 750 nm.

 From 3 to 66 ps (Figure 30B): a sharp amplitude decrease of the plateau on the blue side of the 420-nm band is observed between 14 ps and 28 ps. A decrease of the 420-nm peak is observed only after 14 ps. There is a decrease in the amplitude of the 575-nm band, accompanied by a shift in the position of its maximum to 580 nm at 66 ps.

 From 66 to 875 ps (Figure 30C): large decay of all bands. The decay appears to give rise to a new spectral shape with a maximum at 587 nm at 875 ps.

 875 to 3200 ps (Figure 30D): the new spectral form further appears as the 420-nm band disappears. The 587-nm band continues to decay until 3200 nm where its maximum is at 590 nm.

Two other local maxima are found at 506 and 639 nm.

Global kinetic analysis

The data were best fitted with a sum of four exponential components plus a plateau with the following lifetimes: τ 1 = 1 ± 0.2 ps, τ 2 = 15 ± 2 ps, τ 3 = 247 ± 8 ps, τ 4 = 2892 ± 1851 ps, τ 5 = ∞ (plateau).

The average residue of the fit was 51µOD The DADS and EADS spectra resulting from the fit are shown in Figure 31. The evolution of the spectra in the first phase can be more clearly observed comparing EADS1 to EADS2. We observe what is in effect a narrowing of the spectral band at both the 425-nm and 575-nm bands. This results into an increase in amplitude of both bands. The decay of EADS2 leads to EADS3 without any change in the spectral form. Let us note that for this experiment, this is the main decay observed. The decay of EADS3 gives rise to a completely new spectral form, as seen in EADS4, with a time constant of 247 psindicative of the formation of a new species. The evolution of this spectrum continues until the end of the 3.2-ns time window of our experiment as is seen as the residual signal left at this time delay.

We will now compare the dynamics that we have observed here with that of the previous section in the absence of the substrate. We will compare the two experiments by looking at their DADS spectra.

The 1 ps DADS from the experiment without the substrate appears be a mixture of the 0.4 ps and 1.1 ps DADS spectra from the experiment in the presence of the substrate. The same process must be involved for the two cases which we have previously attributed to vibrational relaxation. We can verify this by looking at the spectral changes associated with these time constants. In both cases, we observe a spectral narrowing that is most evident to see in the EADS spectra. The 22 ps DADS spectrum of the experiment without the substrate is similar to the 15 ps DADS spectrum of the experiment with the substrate. This suggests that in the two cases we observe similar excited-state dynamics. The similarity of the time constants appears to indicate further that these dynamics that we describe so far are independent of the presence of the substrate. It is not surprising as they are processes that take place before the main decay.

Let us point out that for the case with the substrate, the main decay of the system is the 246 ps constant. There is a marked difference between the main decays of in the two cases. The similarity in shape of the 246 ps DADS from the experiment with the substrate and the 3750 ps DADS in the absence of the substrate (and their corresponding EADS) indicates that the same species are decaying with these time constants. We have previously seen that this species is FADH -* . The difference is that in the presence of the substrate, there is a product that is rapidly formed from the decay of the excited FADH -.

The spectrum of the product from this fast decay is given in the 2892 ps DADS of the experiment with the substrate. The spectrum bears a similarity with the 3750 ps DADS in the case without the substrate. It evokes the decay of an excited FADH -. The similar time constant also evokes the decay of excited FADH -. The absorption band is significantly broader and the peak is shifted to the red. This suggests that the spectrum cannot be solely attributed to the decay of an excited FADH -alone. This issue is addressed in the next section.

Interpretation

According to all schemes of (6-4) photolyase repair, the excited FADH -cofactor transfers an electron to the (6-4) photoproduct to commence the reaction. We thus expect the formation of FADH • as a consequence of this process. The spectrum expected from the formation of FADH • from FADH - (FADH • minus FADH -difference spectrum) was constructed from published spectra of FADH -and FADH • in Xl64 by Schleicher et al. [34] (Figure 32B). It contains four prominent bands: at 385 nm, 480 nm, 590 nm and 635 nm. This simulated difference spectrum is compared to EADS5 in Figure 32A. To aid in comparison, the spectra were normalised at 590 nm, an absorption maximum of the FADH • difference spectrum. EADS5 and the (FADH • -FADH -) difference spectrum are found to match relatively well. It should be recalled on one hand that the literature spectra may not be 100% pure. They are in any case relaxed spectra of a protein in the absence of substrate, in particular of substrate radical anion. On the other hand EADS5 is in fact an extrapolated spectrum not actually recorded in the experiment. It is supposed to correspond to what would be directly observed at a delay of about 12 ns, after full decay of the 2.9-ns component. The major peaks indicative of FADH • are present in EADS5, at 500 nm, 590 nm and about 637 nm. The presence of the 385-nm band cannot be ascertained due to scattering of the 387.5 nm pump beam in this region. A band at 420 nm indicates the presence of FADH -* . Since the 420 nm is not observed in our spectrum, we can conclude that the species present at the end of our observation window is FADH • .

We also compared EADS4 with the (FADH • -FADH -) difference spectrum. This is shown in Figure 33. The two spectra do not appear to match well although the precursors of the FADH • peaks can be identified in EADS4. In light of the discussion of §3. The question then arises to find a molecular meaning to this 2.9-ns component. It may be recalled that in the absence of the substrate, the excited state lifetime of FADH -is 3.75 ns, which can be considered as rather close to 2.9 ns. It could thus be imagined that our sample is in fact heterogeneous and contains two classes of enzyme. There may be a fraction of unreactive photolyase complex that do not undergo electron transfer from the flavin to the substrate. The origin of this population could arise from badly recognised substrate, that is, substrate bound in some improper configuration. Let's stress that the T(6-4)T lesion is in fact asymmetrical; 5'-3' and 3'-5' orientations might lead to different complexes.

Alternatively a class of structurally altered (partially denatured) protein could be present in the sample. It is unlikely that proteins not containing any substrate would be present due to the 7:1 substrate to photolyase ratio in our experiment. A second explanation we can think of is an equilibrium between the FADH -* /T(6-4)T state and the state obtained after electron transfer, FADH • /T(6-4)T •-. This would require the two states to be energetically close enough to allow facile backward electron transfer. It could further be imagined that this equilibrium (shown in Figure 34) is somehow perturbed by chemical changes occurring at the level of the reduced substrate (and escaping from the spectral window of our experiment), with a time constant of 2.9 ns. These changes would potentially be the elementary steps of actual substrate repair. Lowering the energy of the FADH • /(reduced substrate) pair would shift the equilibrium toward the FADH • product thereby explaining the experimental observation. Going back to the main decay of FADH -* in the presence of the substrate, we have observed that it occurs with a time constant of 246 ps. This is considerably faster than the 3.8-ns decay in the absence of the substrate. The 246 ps that we measured time constant is comparable with the 225 ps time constant that was observed by Li et al. in At64 [14] and is assigned to electron transfer from the flavin to the substrate.

It is finally worth stressing that, at the end of our observation window, we have not observed the decay of FADH • by back electron transfer after repair of the substrate of formation of intermediate X. We can thus conclude that this process takes significantly longer than a few ns.

The work that we just presented compliments to the landmark ultrafast study by Li et al. by providing the transient absorption spectra of the reaction. Looking at transient spectra associated with the repair process can give us a better picture of the repair mechanism. However, our results suffer from the fact that we are currently unable to access the spectral region below 350 nm. We have seen earlier that this region is a region of great importance as this is where the (6-4) photoproduct and the repaired nucleobases absorb. Looking at this region will give us information on the spectrum of the intermediate that we now know to exist due to the two-photon mechanism of the reaction. In light of the two-photon mechanism, future work should concentrate on how to better separately study the photoreactions induced by each photon. To minimise the contribution of the second reaction, a very repetition rate of the excitation pulse may be considered to allow for the decay of X back into the original T(6-4)T state.

Pursuit of the reaction in the longer timescales should also be done in order to observe what happens in the times beyond 3 ns (i.e. µs and ms). Such experiments can be done with the real-time transient absorption set-up that we used in the study of photoactivation. The second photoreaction may also be studied similarly.

Introduction

This chapter is dedicated to the description of the techniques and methods used to obtain the data reported in the preceding chapters. Presented here are the methods we used for time-resolved spectroscopic measurements, data analysis, sample preparation, and homology modelling. This chapter is mainly concerned with the femtosecond transient absorption experiments performed in the study of the photoactivation and photorepair reactions of (6-4) photolyase.

This chapter will begin by providing details on the experiments performed for the determination of the number of photons required for photorepair, although a general description of them has already been given in Chapter 3. We will also present the different substrates and the calculation of the steadystate repair yield (η ss ) from the steady-state photorepair experiments.

We will then present the set-ups used for the different spectroscopic measurements performed. We begin with the femtosecond transient absorption spectroscopy set-up. It is located in our laboratory at the Ecole Normale Supérieure in Paris. We will then present the set-up used for the nanosecond real-time transient absorption spectroscopy measurements. This set-up is located at the laboratory of Klaus Brettel at iBiTecS in CEA Saclay.

The next part will focus on how we handled our spectroscopic data. It begins with a discussion of the artefacts encountered in our spectroscopic experiments and how these were taken into account. We then proceed with describing how our data were analysed.

The last part of this chapter is dedicated to the protein itself, studied during my thesis: photolyase from Xenopus laevis. Details on how the protein was produced by Junpei Yamamoto at the Osaka University will be given. We will also describe how we prepared the samples for the spectroscopy experiments. Since the crystal structure of this protein has not yet been resolved, homology modelling was used with the guidance of Raphaël Guerois of iBiTecS, CEA Saclay in order to obtain a model structure of it. This technique is described at the end of this chapter. All substrates used in our experiments were synthesised by Junpei Yamamoto at the Osaka University. The normal substrate is an 8-mer with the sequence d(CGAT(6-4)TGAT). The DHT substrate is a 10-mer containing one T(6-4)T/T(6-4)C: d(HHHHT(6-4)TTHHH). We have shown in Chapter 2 that the two substrates are repaired with similar (steady-state) quantum yields by photolyase. Both substrates were synthesized on an Applied Biosystems 3400 DNA synthesizer, by using the phosphoramidite building blocks of the 6-4PP [2], DHT (obtained from the hydrogenation of thymidine), thymidine, and nucleosides for ultramild DNA synthesis (Glen Research) with Universal Support II PS (Glen Research). These oligonucleotides were analyzed and purified by a Gilson gradient-type analytical HPLC system equipped with a Waters 2996 photodiode-array detector. On the system, a Waters µBondasphere C18 5 µm 300 Å column (3.9150 mm) was used at a flow rate of 1 mL min -1 with a linear gradient of acetonitrile in 0.1 M triethylammonium acetate (pH 7.0) generated over 20 min.

Quantum yield determination

The steady-state photorepair quantum yield (assuming the concentration of intermediate has reached a stationary value (see Chapter 3, §3.1.5.1.2) was evaluated using the method described by Thiagarajan et al. for a unimolecular photochemical reaction [1]. The first order rate constant, k (rate of formation of repaired substrate), is determined by:

(1)

where: I is the flux of incoming photons (photons s -1 cm -2 ) σ, the absorption cross-section of the reacting molecule, σ (cm 2 ) ϕ, the reaction quantum yield. We have seen in Chapter 3 that this yield is in fact the steady-state repair yield ( SS ), simply related to the detailed parameters of the underlying 2-photon mechanism ( §3.1.5.1.2, Equation 5).

It is assumed that the solution is dilute enough so that variations of I over the optical path can be neglected. The photon flux is determined by studying the effects of the flux in question using a wellstudied actinometer (AM) of known σ AM and ϕ AM.

In this case, we used a p-(dimethylamino)benzenediazonium tetrafluoroborate (DMAD) actinometer. Writing equation one for both the actinometer and the photolyase (PL) and equating the flux, I, for both, one obtains:

(2)

And solving for φ PL ,

From the relationship between σ and ε, † we can replace (σ AM /σ PL ) with (ε AM /ε PL ). In the photorepair experiments, we measure absorbance changes as a function of illumination time. We therefore need to express the rate constants, k, in terms of absorbance changes.

For the DMAD actinometer consumed by the reaction ‡ we have:

) exp( ] [ ] [ 0 t k AM AM AM   (4)
To obtain a linear relationship we take the logarithm of both sides and obtain:

) ] ln([ ] ln[ 0 AM t k AM AM    (5)
Since the measured absorbance, A AM , is proportional to the concentration, k AM is directly obtained from the plot of ln(A AM ) vs. time.

The concentration change of the product, TT is given by:

PR k PL TT dt d ] [ ] [  (6)
Integration over the illumination period ∆t and using

  l TT A PR PL        , we get: t l k PL A PR PR PL          ] [ (7)
Where [PL] is the concentration of active photolyase, ∆ε PL is the differential molar absorption resulting from the repair of the (6-4) photoproduct, and l is the optical path length. Equation 7gives an expression for k PL . Equation 3 rearranges to: For the estimation of the steady-state repair quantum yield ( SS ),  325 of -6.0 mM -1 cm -1 was used for repair of both 10-mer DHT and 8-mer normal substrates. This value was obtained from the absorption spectrum of the 8-mer normal substrate assuming ε 260 = 64.3 mM -1 cm -1 as calculated according to Tataurov et al. [3]. This calculation allows us to predict the absorption of a given sequence of nucleobases. In the case of the 10-mer DHT substrate, direct detection of the restoration of intact † (cm²) ‡ DMAD undergoes light induced dediazotization (i.e. removal of the diazo group by N 2 evolution).

X PL PL AM AM AM PR PL A k t A              (8) Where A PL X is defined as   PL l PL    . It

Spectroscopic methods

Steady-state spectroscopy

Stationary absorption spectra were measured using two double beam spectrophotometers: Uvikon XS (Secomam) at CEA and CARY 300 (Varian) at ENS. Fluorescence spectra were measured using a Jobin-Yvon FluoroMax-3 spectrofluorometer. To avoid sample degradation, the sample holders of the spectrometers were kept at 5°C. This was done with the aid of a circulating temperature bath. The coolant mixture (water/glycerol) was circulated in the sample holders of the spectrometers. Condensation of water vapour was prevented with a constant flux of dry air on the sample holders.

Femtosecond transient absorption spectroscopy

Our investigations involve the study of transient states. We create such transient state by exciting a population of interest from the ground state using an actinic pulse of light. We then want to study the dynamics of the excited state which we know occur fast. To be able to observe these dynamics, the actinic pulse should be short compared to the (expected) characteristic dynamics of the system that we wish to observe. The observation of this excited system is done by taking its absorption spectrum. To study the excited state, or rather see what becomes of it, we take its absorption spectrum as a function of time. The absorption spectrum of the system is measured using a polychromatic beam of light. To study the evolution of the excited state, we measure the absorption spectrum of the excited state at various times (called delays) after the actinic pulse. What we obtain after is a time-lapse image of our system of interest described using its absorption spectra.

In our studies, we used a laser as our source of light. More specifically, it is a laser that delivers light pulses with duration of several tens of femtoseconds for each pulse. The femtosecond pulses allowed us to observe the dynamics of the (6-4) photolyase that we have presented in the previous chapters. The light coming from our femtosecond laser was used to generate the excitation pulse (called a pump) used to excite our sample and to generate the polychromatic light beam (obtained by non-linear optics) with which we measured our absorption spectra or more specifically our transient absorption spectra. A schematic diagram of the set-up for this type of measure will be given in Figure 3. However, we will begin by detailing its component parts in the subsections that follow.

Femtosecond laser source

The laser source that we used in our experiments has two components: (i) A mode-locked Ti:sapphire laser (Tsunami, Spectra-Physics) and (ii) a regenerative amplifier (Spitfire, Spectra-Physics).

Each component is described in the following:

Ti:Sapphire laser

Ti:Sapphire lasers are the most widely used lasers in spectroscopy due to their tunability and ability to generate ultrashort pulses. Our femtosecond Ti:Sapphire laser has an optimal operation wavelength at 810 nm. In order to satisfactorily optimise the non-linear optics required for our photolyase experiments, our laser had to be tuned at 775 nm. The name refers to the lasing medium that is a sapphire crystal (α-Al 2 O 3 ) doped with Ti 3+ ions. The crystal is pumped by another laser (Nd:YVO 4 ; Millenia, Spectra-Physics) with an energy of 4 W at a wavelength of 532 nm. To produce ultrashort pulses, the laser is mode-locked. Self mode-locking is attained through an optical Kerr effect in the amplifying crystal. The Ti:sapphire laser delivers a beam composed of 35 fs pulses with an energy of 350 mW centred at 775 nm at a repetition rate of 82 MHz.

Regenerative amplifier

The energy per pulse of the beam coming from the Ti:Sapphire laser is too low (~4 nJ) to allow the non-linear optical effects necessary for our transient absorption spectroscopy set-up. The energy of each must then be amplified. Amplification produces more powerful pulses but at a lower rate. The amplification of our pulses was done by passing the beam through a regenerative amplifier. An amplified pulse, due to its high energy, can cause damages to the optical components in the amplifier. To avoid this, the pulse is temporally stretched (to pulse duration of a few hundreds of picoseconds) before amplification. The stretching is achieved through multiple passages of the pulse through a diffraction grating. The pulse, now stretched, is said to be "chirped". The pulse is then injected into a laser cavity containing an amplifying medium. In this case, the amplifying medium is a Ti:Sapphire crystal pumped by a Nd:YLF laser (Empower 15, Spectra-Physics: 527 nm, 6 W, 1 kHz). Once the pulse has attained its maximum energy, it is ejected from the cavity and recompressed. After passing through the regenerative amplifier, the beam is now composed of 50 fs pulses having energy of 680 mW at 1 kHz. Having sufficient energy after amplification, the laser beam can now be used to generate the pump and the probe beams. The generation of these beams is described in the next subsection.

Pump beam

The pump beam is used to excite a small fraction of the sample population at one of its absorption bands. The pump beam hence needs to be tuned to this specific absorption wavelength. The pump wavelengths used in our work were generated using two methods: by optical parametric amplification (OPA) and by second harmonic generation. For the pump beam, about 230 mW of the amplified beam is used.

The pump and probe beams are focused and overlapped in the sample cell throughout the optical path of the cuvette (0.1 cm). The spatial overlap of the beams in the sample is done with parabolic mirrors that also focus the pump and probe beams to the sample. The cuvette containing the sample is constantly displaced in both the horizontal (1 mm/s) and vertical (100 mm/s) directions to avoid the re-excitation of a previously excited area. The sample is also maintained at 5°C with the aid of a water bath circulated through the sample holder. Condensation of water vapour is avoided with a constant flux of cool, dry air on both sides of the cuvette. Sample preparation is described in Section 4.5.

Data acquisition

To measure polarised transient absorption, the beam transmitted by the sample is made to pass through a polariser (analyser) oriented either parallel or perpendicular to the polarisation direction of the pump beam. The parallel and perpendicular intensities are here measured one after the other, in practice changing the analyser (two of them are mounted on a motorised translation stage). This configuration is called 2z, because two continuum beams are simultaneously detected, the polarised probe and the reference, on a CCD detector divided in two zones.

Alternatively, we implemented a set-up where the two polarisations are measured simultaneously. This was achieved by replacing the parallel and perpendicular polarisers by a Glan-Thompson polariser. This polariser splits the beam into its parallel and perpendicular polarisations thereby allowing these two components beams to be detected at the same time. This is the 3z configuration, because three continuum beams are detected in three zones of the CCD detector.

Detection of the signals is attained by sending the reference and the parallel and/or perpendicular beams to a spectrograph (SP306i, Acton) in order to disperse light and create a spectrum, which is then globally detected on a CCD detector (Pixis-100B, Roper Scientific: 100×1340 pixels, back-illuminated).

The 100 pixel height of the CCD detector is divided into two zones (2z) for the reference and parallel/perpendicular beam for the 2z configuration. It is divided into three zones (3z) for the reference, parallel, and perpendicular beams in the 3z configuration (broken lines in Figure 2).

The set-up is drawn schematically in Figure 3.

Spectral acquisition

The absorbance change ΔA(λ, t) for a pump-probe delay t, is defined as:

) ( ) , ( ) , (    np p A t A t A    (9)
Where A p and A np are the absorbances at time delay t with (p) and without (np) pump excitation. The differential absorbance is calculated as follows: The "pumped" (p pr) and "non-pumped" (np pr) intensities were measured alternatively by masking the pump pulse every two probe pulses with a mechanical chopper. Between two pumped intensity measurements there is a non-pumped intensity measure and vice versa. The readout time of the CCD taking 1.88 ms, we reduced the repetition rate of the probe beam to 333 Hz in order to record each shot. There is hence a gap of 3 ms between probe pulses and a gap of 6 ms between two pump pulses.

More precisely, there is a 6 ms gap between measuring two I p pr and two I np pr intensities.

As we are measuring the absorption of a small excited state population, the absorption that we measure is small. Furthermore, the changes in this absorption are also expected to be small. Background noise hence becomes important. To reduce the effect of the noise (i.e. maximise the signal-to-noise ratio), the process of averaging several spectra becomes essential. In our experiments, we measured 1500 spectra for each time delay which we then averaged. For the 2z 3T and 3z 3T this number was increased to 10 000 since we are measuring fewer delays. The collection of all time delays required for an experiment is called a scan. A scan normally would normally contain around 50 time delays. Each delay in the scan is an average of 1500 or 10 000 spectra. To further increase the signal-to-noise ratio, a scan is repeated until a satisfactory ratio is obtained.

Nanosecond real-time transient absorption spectroscopy

In our study of the anisotropy in the timescales longer than 1500 ps, we directly measured the real-time decay of the transient absorbance at two probe wavelengths. These experiments were performed in the laboratory of Klaus Brettel at iBiTecS in CEA Saclay.

Laser sources

Excitation pulse

The excitation pulses were generated by an Nd:YAG laser (Brilliant b, Quantel) that delivers 5 ns pulses at 532 nm, at 2 Hz. The laser is coupled with an optical parametric oscillator or OPO (Rainbow, Quantel). The OPO is used to generate the 475 nm excitation wavelength by means of non-linear optics.

Our OPO consists of an optical resonator and two non-linear crystals to produce signal and idler waves.

The polarisation of the excitation pulse is set to vertical.

Monitoring light

The monitoring light used to detect absorption changes in the set-up comes from continuous-wave lasers at specific wavelengths. This is the drawback of the system as only one wavelength can be observed at a time. Furthermore, the observation wavelengths are limited to the available commercial lasers. For our experiments on the photoactivation (6-4) photolyase, we used 50 mW cw lasers at 457 nm and 562 nm from CobaltTwist. A chopper placed before the sample is used to create a monitoring pulse of 1.5 µs duration every 30 ms out of the continuous emission of the laser. This pulse is long enough to allow recording of a full real-time kinetics in the µs regime and at the same time limits the irradiation time of the sample. The pump laser is synchronised to this chopper in such a way that it is triggered only when monitoring light is present at the sample level.

The sample

The cuvette containing the sample was placed in a sample holder thermostated to about 10°C. The sample holder is placed in a closed chamber that is constantly purged with a flux of argon to inhibit the condensation of water vapour. The sample chamber has four transparent windows allowing the excitation light and the monitoring light to pass through.

Detection system and data acquisition

The monitoring light is detected by an ultrafast detector (photodiode with rise time <500 ps, UPD 500 UP, Alphalas) connected to a digital oscilloscope (DSA 602A, Tektronix). The signal (a tension proportional to the intensity of light) is displayed (in mV) as a function of time. It is converted to ∆A using Equation 11. Where U(t<0) is the voltage before excitation (t = 0). This is analogous to the nonpumped intensities in Equations 9 and 10 that refer to the absorbance of the ground state.

White light chirp and t 0

Red light travels faster than blue light in dispersive materials (group velocity dispersion). As a consequence the white light continuum probe beam is distorted as the beam propagates through the different optical elements of the set-up (i.e. filters, CaF 2 , sample cuvette). The continuum is said to be chirped. The red end of the probe beam arrives at the sample before the blue end. As an effect, the temporal overlap between the pump beam and the probe beam varies as function of the wavelength. This temporal overlap between the pump and the probe is colloquially called "time zero" (and noted t 0 ). The t 0 artefact is corrected by first measuring how t 0 varies with wavelength. We obtain this data by measuring the transient absorption spectra of the pure solvent. By doing this, the signal that we measure is solely due to the XPM. This experiment is performed under the same conditions as the real sample (in our case, photolyase). The kinetics measured at each wavelength allows us to determine the variation of t 0 with the wavelength as a cause of the chirp. Plotting this variation we obtain a curve such as the one shown in The curve was fitted with the following function which is a sum of the group velocities (v g ) in various dispersing media in the set-up (i.e. filters, CaF 2 , cuvette,); e i is the thickness of material i and C is a constant.

   i g i v e C t ) ( ) ( 0   (12)
The fit curve is further fitted without loss of information with a polynomial of degree six to make it simpler to work with. The chirp correction is performed by time-shifting the kinetics at any given wavelength (λ) by the difference of t 0 at λ and at a reference wavelength (λ ref ), that is, by t 0 (λ)t 0 (λ ref ).

The reference wavelength is the wavelength corresponding to the median t 0 value of the t 0 values obtained.

Cross-phase modulation and stimulated Raman scattering

When beams ultrashort pulses interact in a Kerr medium, one beam may cause a change in the optical phase of the other. This phenomenon is called cross-phase modulation (XPM). In our case, the pump pulse induces a change in the refractive index of the medium of the sample. The change in refractive index causes the modulation of the phase of the chirped probe beam. This effect creates oscillations in the spectrum around t 0 . This is seen in Figure 5. The figure shows chirp-corrected spectra around 0 ps for a sample of oxidised Xl64.

In fact, at least one more artefact contributes to these oscillations. This is the stimulated Raman scattering of the solvent, and possibly the solute. This is readily observed in Figure 5 and is most evident in the -0.1 ps spectrum. The intense absorption (positive) peak at 406 nm corresponds to a Stokes line where the continuum probe plays the role of excitation beam for the Raman effect (hence is absorbed) and the pump plays the role of stimulation beam. The negative peak at 560 nm corresponds to the reverse value. Unfortunately we also observed that this value changes from day-to-day when slight adjustments of the set-up are made. We conclude that this artefact is due to the configuration of the 3z set-up. Despite our best efforts, we were neither able to correct this defect experimentally nor identify its exact source. To take this systematic error into account, we measured the anisotropies of the dye molecules for each experiment performed. We then empirically subtracted this measured anisotropy from the anisotropy value obtained for the (6-4) photolyase.

"Millisecond constant"

During the treatment of our femtosecond measurements on oxidised (6-4) photolyase we noticed a weak but quite clear transient absorption spectrum at negative time delays, as shown in Figure 8. A negative time delay means that the probe beam arrives before the pump beam. A transient absorption spectrum measured at a negative time delay should not contain any absorption bands (it should be flat at zero) since no transient species have been created yet. Yet we observed the same non-zero spectrum at all negative delays. Moreover, the spectrum resembles an inverted transient absorption spectrum typical to that of oxidised (6-4) photolyase that we have seen in Chapter II. The spectrum is shown in Figure 8. It should be recalled that the sample holder containing the cuvette is displaced in the vertical and horizontal directions to ensure that we always excite a new spot on the sample. A signal in the spectra of the negative time delays is possible if: (i) the 2D movement of the sample is not fast enough and (ii) there is long-lived decay in the sample. The largest effect will be given by the first case. If the sample is displaced fast enough, even if the second case were true, there should not be a signal detected.

The difference between two pump pulses in our experiments is 6 ms (cf. Section 4.2.2.6). Between the two pulses, the non-pumped intensity is measured (i.e. 3 ms after the first pump). This intensity was then used to divide the pumped intensity when calculating for ∆A as in Equation 10. For this reason, the spectrum we see is inverted. More importantly, what we see in Figure 8 is in effect the transient absorption spectrum of the photolyase at 3 ms albeit inverted.

In preliminary experiments for a different study of the photoactivation of X. laevis photolyase, our collaborators Klaus Brettel and Junpei Yamamoto were able to identify a lifetime of ~30 ms. This lifetime is attributed a to a long-lived charge recombination between the flavin and a tryptophanyl radical. Their experiments are not discussed here. This explains the spectrum in Figure 8.

Let us call this artefact the "ms constant" -after the fact that it is present as an offset in all transient absorption spectra. To correct for it, we subtracted the spectrum of the constant from all the transient absorption spectra. The spectrum of the constant was obtained by averaging the spectrum of the negative time delays. Only negative spectra unaffected by the XPM artefact (4.3.3) were chosen for averaging. To further minimise the spectral noise created by subtracting a noisy spectrum from another relatively noisy spectrum, the constant spectrum was smoothed using the Savitzky-Golay algorithm [7].

We used a window size of 17 points (6.8 nm) and a polynomial of degree 2. The effect of smoothing is shown in Figure 9.

Free flavin

After a pump-probe experiment, each individual scan (cf. Section 4.2.2.6) is inspected to ensure "spectral integrity". That is to say, to ensure that the form of a certain spectrum (say at 2.2 ps) over the course of the experiment does not change. We did this by choosing at a representative time delay and comparing these corresponding spectra between all scans. For our experiments with oxidised photolyase (i.e. FAD ox ), we looked at 2.2 ps. Looking at the changes in the 2.2 ps spectra over time gives an insight on the evolution of the sample over time. In particular, we are interested in changes in spectral form.

We are about to describe a particular problem that we observed in the recording of the 2z 3T data.

It should be made clear that the peculiarity that we are describing here has only been observed in the 2z 3T measurements and to a significantly smaller extent in the 3z 3T measurements. The evolution of the parallel transient-absorption spectrum at 2.2 ps over the course of data acquisition for the 2z 3T measurement is shown in Figure 10. For the 2z 3T experiment, we performed a total 53 scans. These scans were arranged in chronological order and were then grouped into 7 blocks of about 7 scans each. The spectra comprising a block were then averaged to give the spectra shown in Figure 10. Figure 10A shows that the amplitude of the 2.2 ps spectra decreases with time. This decrease in amplitude may be explained by the instability of the pump beam. Normalising the spectra at 450 nm shows no change is spectral form for the region below 450 nm. There is however a clear change in shape in the spectral region between 500 and 650 nm. This observation cannot be explained by changing pump energy. Figure 11 shows the region between 500 and 650 nm region more closely. On Figure 11, the change in spectral form is seen more clearly. There is an increase in the depth and width of the minimum at around 540 nm over time. Furthermore, the position of this minimum appears to become more shifted to the red over time to about 550 nm in spectrum 7.

To understand this effect, let's recall that a red-shifted stimulated emission band (relative to that of oxidised Xl64) is characteristic of free FAD ox in solution (cf. Annex B for FAD ox transient spectrum).

We hypothesised that the gradual red shift of the minimum over time as seen in Figure 11 arises from the contribution from unbound FAD ox progressively being released in solution from structurally altered proteins. Alternatively we also considered that the fraction of a kind of non-reactive photolyase, bearing a longer-lived excited flavin, steadily grows with time. We have indeed seen in Chapter 2 that the decay of the excited FAD ox bound to photolyase takes place in about 0.45 ps. This decay takes much longer in free FAD ox in solution: 5 to 9 ps for the closed form (80% of the population at room temperature and neutral pH) [8,9] and ~2.8 ns for the open form [9]. In case there is a mixture of bound and unbound flavin, the excited-state decay of bound FAD ox is expected to be finished at 2.2 ps while that of free FAD ox is not.

We propose it is for this reason that we observe in Figure 11 a growing contribution of what looks like stimulated emission of FAD ox . To test this idea, we took the difference (∆∆A) between spectrum 7 and 1 (non-normalised), multiplying in fact spectrum 1 by an adjustable constant (k) order chosen to better subtract the common contributions, due to the unaltered proteins. Constant k is expected to account for a reduced contribution of bound flavin in spectrum 7. The difference spectrum should then reveal the species responsible for the observed spectral changes. The ∆∆A spectrum with k=0.9 is shown in Figure 12, superimposed with the transient absorption of free FAD ox at 2.2 ps (measured under the same conditions as the photolyase, see Annex B); k was empirically chosen so as to (approximately) optimise the resemblance between the two spectra. The red spectrum in Figure 12 is the difference between the spectra 7 and 1 in Figure 10A. The difference spectrum has a transient absorption signal around 370 nm. A negative bleaching band is also seen around 450 nm as well as a stimulated emission band around 560 nm. Comparing this defference spectrum with that of free FAD ox (Annex B, blue spectrum in Figure 12) Although not identical, the two spectra bear obvious common spectral features. They in particular both share a stimulated emission band peaking around 560 nm. Differences are however also clear, like the relatively weak amplitude of the stimulated emission band in the difference spectrum (∆∆A) as compared to the bleaching or blue transient absorption bands, or the non-positive amplitude of the local maximum at 510 nm. Given the arbitrariness of choosing k it is hard to reach a safe conclusion from Figure 12. It can however be said that a species resembling free FAD ox , with an excited-state lifetime far longer than that of bound flavin (0.45 ps)

progressively appear in the solution. It could be free flavin released from partially denatured proteins. It could also correspond to altered proteins still binding the flavin, but in a non-reactive, longer-lived configuration, possibly more exposed to the solvent. We will call this species free-like FAD ox . Note that prior to performing any experiment with (6-4) photolyase, any free chromophore in solution was removed by passing the protein solution through a gel filtration column (cf. Section 4.5). It may be hypothesized that the alterations of the protein likely responsible for the rise of free-like population are due to laser irradiation.

The free-like FAD ox artefact is especially undesirable for anisotropy measurements. The anisotropy of the unreactive flavin would contribute to the anisotropy of the species we are interested in (WH •+ ). If the artefact is left uncorrected, what would be measured at 580 nm is a complex mixture of anisotropies of WH •+ and free-like excited FAD ox . This mixture would in addition change as time delay increases because the relative contribution of free-like FAD ox would decrease as its excited state would decay (80% of the decay of free FAD ox in solution occurs in about 5 ps). We indeed observed that the growth of free-like FAD ox contribution with irradiation time is much larger and clearly seen at a pumpprobe delay of 2.2 ps than at 1500 ps.

Having identified the source of the artefact, we now will go on to correcting it. To properly visualise the increase in the effect of the free-like FAD ox , it is helpful to look at all 53 spectra that make up the groups in Figure 11. Looking at the trend of how the free-like FAD ox appears may help us in its correction. Figure 14 From Figure 14, we can see that the decrease in amplitude appears to go in a linear fashion in all the wavelengths sampled. Taking account the assumptions stated above, extrapolating the linear curve to zero should give the spectrum where no FAD ox has been released in solution. The 2z 3T and 3z 3T anisotropy spectra were obtained in this manner. The anisotropy for each spectrum was first calculated using the parallel and perpendicular spectra. The anisotropy at each wavelength was then extrapolated to time zero using a linear fit to obtain the initial anisotropy. The anisotropy resulting from this extrapolation is the anisotropy of the sample when no free-like FAD ox is present. The extrapolation was performed with a Mathematica-based program.

A standard error associated to the extrapolated time-zero anisotropy (i.e. error bars in Chapter 2) was calculated. It corresponds to a confidence interval of 65%. To obtain the 95% confidence intervals that we presented in Chapter 2, this error was multiplied by 2. Prior to pump excitation, the population of the species of interest is in the ground state. The absorbance is given by the Beer-Lambert law as follows:

Data analysis

c l A a np    ) ( ) (    (14)
where ε a (λ) is the molar extinction coefficient, l is optical path length, and c is the total solute concentration.

At a time t after excitation, let us assume that some transient state i (excited state or photoproduct) is populated with a concentration c i . Its contribution to the total absorbance of the sample is given by:

  l t c t A i i e i u i     ) ( ) ( ) ( ) , ( , ,      (15a)
where ε u,i is the excited state absorption of species i and ε e,i is the emission of the transient species i.

Conversely, the reduced contribution of the ground state is:

l t c t A a    ) ( ) ( ) , ( 0 0    (15b)
where ε a (λ) is the molar extinction coefficient, l is the optical path length, and c 0 is the concentration of the ground state population.

The total absorbance of the pumped sample now reads:

  l t c t c t A t A A N i i i e i u a N i i p                   1 , , 0 1 0 ) ( ) ( ) ( ) ( ) ( ) , ( ) , ( ) (         
Given that the total concentration of all species present (c) is constant and equal to the sum of the concentration of the ground state species (c 0 ) and all the transient species (c i ), we have:

   i c c
         (16)
also possible. However, this is more costly in terms of computation time and does not benefit from noise reduction due to the truncation of small singular values.

The second part of the kinetic analysis consists of the formulation of a kinetic model that describes the system. For a system of coupled first order unimolecular reactions with N transition states, the kinetic equations can be written in matrix form as follows:

) ( ) ( t C MC t C dt d   ( 22
)
where C(t) is a vector of dimension N containing the concentrations of the N transient species at time t and MC is a matrix containing the first order rate constants that describe the relation between the transient species.

At t < 0 s, C(t) = 0, and t =0s, the pimp pulse creates the initial population C(0). In practice, the initial concentrations of the excited states are not precisely known hence C(0) cannot be precisely introduced. A normalised solution such that the sum of the initial concentrations of the transient species is equal to 1 is sought. This is noted as K(t). The real solution C(t), corresponding to the initial condition C(0), is proportional to K(t):

) ( ) ( * 0 t K c t C   (23)
where c 0 * is the sum of the concentrations of transient species at t = 0 s.

Equation 22 becomes:

) ( ) ( t K MC t K dt d   (24)
This differential matrix can be easily solved if the matrix MC is diagonal and the solution is an exponential function for each concentration. If the matrix is not diagonal but can however be diagonalised, the relationship between the matrix MC and its diagonal MD exists as follows:

1     P MD P MC (25) where P is the transition matrix where the columns contain the eigenvectors of MC and MD and the diagonal contains the eigenvalues of MC.D where:

) ( ) ( 1 t K P MD P t K dt d      ( 26
) ) ( ) ( t Y MD t Y dt d   
where:

) ( ) ( 1 t K P t Y    (27)
The solution of such a system, in general, leads to an exponential function for each element of the vector Y and hence to the sum of N exponential functions for each concentration:

) ( ) ( t E MF t K   (28)
where E(t) is a vector of dimension N containing the values of the exponential functions at time t and MF is an N  N matrix containing the pre-exponential factors. The vector E(t) measured at q delays in the buffer at pH 8 with 0.1 M NaCl and 25% glycerol. During protein handling, care has been taken to ensure that the protein was kept at 5°C at all times. The protein was placed in a bespoke 60 µL cuvette with an optical path length of 0.1 cm (115B-QS, Hellma Analytics).

Nanosecond real-time transient absorption spectroscopy

A 70 µL solution of 40 µM oxidised X. laevis (6-4) photolyase was prepared. The appropriate volume (~30 µL 100 µM) of stock X. laevis photolyase to make the solution was first passed through a gel-filtration column (Micro Bio-Spin6, Biorad) to remove any unbound flavin. The protein was then bulked to 150 µL with 0.1 M TRIS buffer at pH 8 with 0.1 M NaCl and 25% glycerol. The sample was then placed in a 1 cm path length cuvette (Starna). The cuvette has a capacity of 70 µL and has four clear windows to allow both excitation and detection. The path length of the optical detection is 1 cm while that for excitation is 0.1 cm. The sample and cuvette were maintained at 5°C at all times.

Homology Modelling

Since the crystal structure of X. laevis (6-4) photolyase is not available in literature, a model of the protein structure was needed as basis for the structural analysis of the protein. The model was obtained using a method called homology modelling. The method used in this work is described below. This was done with the help of Raphaël Guerois of the Laboratoire de Biologie Structurale et Radiobiologie, iBiTec-S, CEA Saclay.

The principle of homology modelling is based on two major observations from Krieger, et al.

[13]:

1. The structure of a protein is uniquely determined by its amino acid sequence [14]. In theory, the sequence is enough be sufficient to obtain the structure;

2. During evolution, the structure of the protein is more stable and changes more slowly than the sequence associated with it. This means that similar sequences adopt practically identical structures and distantly related sequences fold into similar structures [15,16]. The growth of the Protein Data Bank over the years enabled the quantification of a limit for this rule [17]. In general, if two sequences have an identity of at least 35% (safe zone limit for homology modelling), the sequences are practically guaranteed to adopt a similar structure.

The limit of homology modelling also comes from these observations. If fragments of the sequence do not form secondary structures (i.e. they form flexible loops), their positions in the model cannot be easily determined. Hence the regions of the flexible loops are the regions where the greatest uncertainty in the model lies.

There are two structures involved in homology modelling. The first one is a known structure called the template. Using the template as guide, the model of our desired protein will be obtained using the protein's primary sequence (called the target). To create our homology model of X. laevis photolyase we used the structure of D. melanogaster (6-4) photolyase (PDB: 3CVU) [18] as the template.

The model was created using the online SWISS-MODEL platform (swissmodel.expasy.org) [19][20][21][22]. The first part of the modelling consisted of the alignment of the sequences of X. laevis photolyase (NCBI ref: NP_001081421.1) and D. melanogaster (6-4) photolyase. The alignment has been done using ClustalW2 (www.ebi.ac.uk/Tools/msa/clustallw2). From the alignment we saw that the two sequences share a 57% identity and 78 % similarity. The high sequence identity and similarity is the reason why 3CVU was chosen as the template. The high percentage identity also indicates that we are well on the safe zone for homology modelling. Alternatively, SWISS-MODEL can automatically search for templates when given a particular sequence. This option in the platform is called HHpred. Using this method, the suitable template found was A. thaliana (6-4) photolyase (PDB: 3FY4 [23]) that also shares a

In the introductory section, it was pointed out that the main objective of my thesis was to study the two major reactions of the (6-4) photolyase: (i) photoactivation and (ii) photorepair of the photoproduct.

In the case of photoactivation, we wished to determine the exact pathway of electrons during the reduction of the FAD ox cofactor of (6-4) photolyase. We have studied the photoreduction of the oxidised cofactor of X. laevis photolyase by broadband femtosecond transient absorption spectroscopy up to a pump-probe delay of 1500 ps after the initial excitation. We have observed a sub-ps electron transfer after excitation of the flavin cofactor. From the obtained transient absorption spectra, it has been observed that the species created after excitation is the FAD •-/WH •+ radical pair. This radical pair is also the dominant contribution observed until the last transient spectrum of our experimental time window. Global analysis of the transient spectra generally resulted to three time constants (plus a plateau).

From the three time constants found in the global analysis of our obtained data, we asked the question whether or not these constants are attributed to the transfer of an electron from one tryptophan residue to another. The logic behind this hypothesis is the existence of the conserved tryptophan triad among photolyases that are previously shown to be involved in the cofactor reduction. To answer this question, we calculated the anisotropy spectra from our polarised measurements. Our results show that the mechanism of photoactivation in X. laevis suggests that the final donor is W C . We have suggested that the simplest explanation of our observed anisotropy would be the presence of back reactions. Back reactions complicate the electron transfer dynamics and may explain our anisotropy observations. The measurement of these back transfers is therefore a possible future study using the redox-inactive phenylalanine mutants of the tryptophan residues.

However it is possible that real picture of photoactivation may be even more complex. That is to say, the mechanism involves other residues apart from W A , W B , and W C . In our structural analysis of Xl64, were able to identify a tryptophan residue that is of sufficient distance from W C to engage in electron transfer.We call this tryptophan, W D 1 . W D has, up to the time of this writing, never been mentioned in literature. It is important to note that this residue is present in the (6-4) photolyases and 1 W D was identified by Pavel Müller.

Figure 1 .

 1 Figure 1. The pyrimidine nucleobase ring showing the numbering of positions. Thymine: R = CH 3 , X = O; Cytosine (keto form): R = H, X = NH

Figure 2 .

 2 Figure 2. Formation of the CPD photoproduct between two thymine bases

Figure 4 .

 4 Figure 4. Dewar photoproduct

Figure 6 .

 6 Figure 6. EcCPD crystal structure showing FAD and MTHF cofactors (left). Surface rendering of EcCPD showing the FAD binding pocket (right). Figure from Sancar Chem Rev 2003 [36].

Figure 7 .

 7 Figure 7. Crystal structure of Dm64 photolyase bound to DNA containing a (6-4)PP lesion. The FAD cofactor is shown in yellow.

Figure 8 .

 8 Figure 8. Structures of the different chromophores of photolyase.

  -dipole coupling (i.e.

Figure 9 .

 9 Figure 9. Classical overall repair mechanism of the CPD lesion by CPD photolyase. Figure reproduced from Sancar [36].

Figure 10 .

 10 Figure 10. Detailed mechanism of CPD photorepair by CPD photolyase according to MacFarlane et al. [85].

Figure 11 .

 11 Figure 11. Detailed mechanism of CPD photorepair by CPD photolyase according to Thiagarajan et al. [88].

Figure 12 .

 12 Figure 12. Detailed mechanism of CPD photorepair by CPD photolyase according to Liu et al. [92].

Figure 13 .

 13 Figure 13. Classical repair mechanism of the (6-4) photoproduct by (6-4) photolyase. Figure reproduced from Sancar [36].

  photolyase of X. laevis (Xl64) involves two histidine residues: His358 and His354[START_REF] Hitomi | Role of two histidines in the (6-4) photolyase reaction[END_REF]. Both the H354A and H358A mutations of Xl64 photolyase indeed resulted in an almost complete loss of the repair activity. The proposed role of these two histidines in the formation of the oxetane intermediate is presented in Figure14.

Figure 14 .

 14 Figure 14. Role of the two histidines in (6-4) photolyase. Figure taken from Hitomi et al. [94].

  mechanism is dependent on the assumption that HisA is unprotonated while HisB is protonated in the active form of the protein. In 2007, Schleicher et al. claimed that, at pH 9.5 (optimum repair pH according to Hitomi et al.[START_REF] Hitomi | Role of two histidines in the (6-4) photolyase reaction[END_REF]), HisA is unprotonated while HisB is protonated thereby effectively supporting Hitomi's model[START_REF] Schleicher | Electron nuclear double resonance differentiates complementary roles for active site histidines in (6-4) photolyase[END_REF] (Figure15).

Figure 15 .

 15 Figure 15. Changes in the microenvironment of H 8 and H 1' protons of the isoalloxazine ring introduced by pH changes according to Schleicher et al. [98]. This classic notion of (6-4) photoproduct repair has been challenged in 2008. The crystallographic structure of the D. melanogaster (6-4) photolyase binding a T(6-4)T substrate was published by Maul et al. (cf.Figure7)[41]. In their report, they showed that the T(6-4)T photoproduct was present in the active

Figure 16 .

 16 Figure 16. According to this mechanism, after injection of an electron by the flavin to the (6-4) lesion, a protonated HisB donates a proton to the hydroxyl group on the 5' base. This transiently forms a water molecule that attacks the 3' base. This step effectuates the hydroxyl group transfer. The created species after the attack of water readily fragments and rearranges to give the repaired bases. This step is accompanied by the return of the proton to the HisB and the regeneration of FADH -.

Figure 16 .

 16 Figure 16. Transient water mechanism of (6-4) photolyase. Figure taken from Maul et al. [41].

Figure 17 .

 17 Figure 17. A: Oxetane-free repair mechanism of (6-4) photolyase. B: Energy profile of the hydroxyl transfer repair pathway. Figures taken from Domratchevaet al. [99].

Figure 18 .

 18 Figure 18. Transient oxetane repair mechanism of (6-4) photolyase. Figure taken from Li et al. [103].

AFigure 19 .BFigure 20 .

 1920 photoexcitation is not enough for photorepair to proceed [104]. They showed using QM/MM calculations that two photoexcitations are needed for the photorepair. Their mechanism is shown in Figure19. His365 is HisB in Dm64 photolyase.

Figure 22 .

 22 Figure 22. Brettel-Vos mechanism of electron hopping shown here in EcCPD photolyase (PDB: 1DNP). The FADH • is shown in yellow. The pathway of electron transfer is shown with the blue arrows with their corresponding time constants. In 2003, Byrdin et al. performed a set of transient absorption measurements on EcCPD and its W382F mutant (W A is replaced by phenylalanine) [122]. They observed that the decay of the excited FADH • is slower in the mutant (at 80 ps) than in the wild type protein (at 26 ps). Furthermore, neither FADH -nor WH •+ 's absorptions were detected. This led to the conclusion that W A is the primary electron donor in the photoactivation reaction and the 80 ps lifetime observed is the intrinsic lifetime of the excited FADH • . In 2004, Byrdin et al. updated their previous studies on EcCPD photoactivation by studying the charge recombination between WH C •+ and FADH -[123]. In their model, they proposed fast protonation equilibrium between WH C •+ and W C • . Depending on the pH, two parallel recombination pathways contribute differently: (i) reduction of WH C •+ by FADH -to give FADH • and WH C (τ ET = 10-50 µs) and (ii) reduction of W C • by FADH -to give W C -and FADH • (τ ET = 25 ms). W C -will then be protonated yielding FADH • and WH C . The updated mechanism is shown in Figure 23.

Figure 23 .

 23 Figure 23. Updated picture of EcCPD photoactivation as reported by Byrdin et al. [123]. In 2006, Luckas et al. studied the phenylalanine mutant of W B in EcCPD (W359F) by transient absorption spectroscopy [124]. The authors reported that the decay of the FADH •* excited state in ~30 ps and did not observe the accumulation of FADH -and WH •+ photoproducts at a detectable concentration. To explain why the photoproducts are conversely detected in the wild-type protein, the authors deduced that the electron transfer from WH B to WH A •+ takes place in less than 4 ps. Brettel et al. in 2008 performed polarised nanosecond transient absorption experiments on the phenylalanine mutant of W C (W306F) and the wild type protein [125]. Their experiment was based on the fact that W B and W C are at distinct orientations from each other with respect to the flavin. The authors took advantage of this fact in order to differentiate between the two residues on the basis of the anisotropy of the absorption transition of the corresponding protonated tryptophanyl radicals. They observed the oxidation of W C for the wild type protein and the oxidation of W B for the W306F mutant. Their results confirmed the electron transfer from W C to WH B •+ takes places in less than 10 ns. They further performed femtosecond transient absorption experiments on the same proteins by collaborating with the group of Vos [126]. Their results showed that after 30 ps, the electron hole (i.e. WH •+ ) is on W C . These results demonstrate that there is no accumulation of WH B •+ and that the electron transfer from W C to WH B •+

Figure 24 .

 24 Figure 24. Brettel-Vos mechanism of photoactivation: kinetic and energetic scheme of the triple tryptophan electron transfer chain in semi-reduced EcCPD, according to Byrdin et al. [127].

  performed femtosecond transient spectroscopy on the wild-type protein and all the mutants in order to characterise each electron transfer step.

Figure 25 .

 25 Figure 25. EcCPD photolyase showing the new donors W A' (W384) and W B' (W316). Figure taken from Liu et al. PNAS 2013 [136].

Figure 26 .

 26 Figure 26. Photoactivation of oxidised EcCPD photolyase. Figure taken from Liu et al. [136].

Figure 27 .

 27 Figure 27. Photoactivation of semireduced EcCPD photolyase. Image taken from Liu et al. [137]. Gray arrows and time constants correspond to the case of oxidised EcCPD, detailed in Figure 25.

  4 µs after the excitation pulse are FADH • and a neutral tyrosyl (Y • ). Since the shape of the tr-EPR spectrum does not significantly change for observation times larger than 100 ns, they propose FAD •-is protonated within 100 ns of its formation. The absence of a tryptophan in their experiment's observation window prompted the proposition of Y • being the final electron donor. In 2010, Brazard et al. studied the photoreduction of the oxidised (6-4) photolyase of Ostreococcus tauri (Ot64), a species of unicellular marine alga, by femtosecond transient absorption spectroscopy. They observed that a primary electron transfer, from a tryptophan residue to the excited FAD ox , occurs in 390 ps [135]. Subsequent kinetic steps, namely partial decays of the FAD •-/ WH •+ pair of radicals with time constants of 9 and 81 ps, where detected. No protonation of FAD •-or deprotonation of WH •+ was observed in the 1.4 ns time window of the study. These observations were interpreted within the framework of the Bretttel-Vos mechanism. To each kinetic step was associated one elementary electron hopping along the chain of conserved tryptophans, accompanied by competitive charge recombination, as shown in Figure 28. The initial donor was assumed to be the closest tryptophan, W A .

Figure 28 .

 28 Figure 28. Kinetic model proposed to describe the photoreduction of FAD ox in Ot64, according to Brazard et al. [135].

Figure 29 .

 29 Figure 29. Electron transfer pathways in WT and mutant X. laevis cryptochrome DASH proposed by Biskup et al. [152].

Figure 30 .

 30 Figure 30. The different forms of flavin.

Figure 31 .Figure 32 .

 3132 Figure 31. Absorption and emission spectrum of FAD ox (A, red) and FADH -(C, blue). Figure taken from Kao et al. [157].

  tokodaii (St, archaeon), Xenopus laevis (Xl, frog), Drosophila melanogaster (Dm, fruit fly), Arabidopsis thaliana (At, plant), Synechocystis sp. (Ss, cyanobacterium), Vibrio cholera (Vc, bacterium), Danio rerio (Dr, zebrafish), Chlamydomonas reinhardtii (Cr, alga), Erithacus rubecula (Er, robin), Sylvia borin (Sb, garden warbler), Mus musculus (Mm, mouse), Homo sapiens (Hs, human), and Danaus plexippus (Dp, monarch butterfly).

(

  homologous to W382, W359 and W306 in EcCDP; respectively noted W A , W B and W C in generic terms) across all proteins in the sampled population. Only W B and W B' are conserved across the entire population. The cryptochrome 2 of the bacteria V. cholera (VcCry2) only has W B and W B' . Its DASH cryptochrome on the other hand (VcCryD) contains only W A , W B , and W B' . For Xl64 it is W396, W373

Figure 1

 1 Figure1shows the active site of Xl64 for the photoactivation process (redox active residues in the vicinity of the flavin and the canonical triad), taken from our homology model. In yellow is shown the flavin cofactor and in blue the "canonical" tryptophan triad (W398, W373 and W319). Also shown is the amino acid closest to the N 5 of the flavin isoalloxazine ring which we have seen in Chapter 1 to play a role in the stabilisation of the flavin radical. The asparagine residue (N392) playing this role in Xl64 is shown in red[18].

Figure 1 .

 1 Figure 1. The photoactivation active site in X. laevis (6-4) photolyase. Intermolecular distances between the residues and FAD ox are shown in angstroms, Å.

  Figure 1 are three tyrosine residues in the vicinity of the photoactivation active site: Y 295 , Y 431 , and Y 487 . The Y 295 residue lies 7.2 Å away from the flavin. It could be a potential electronor proton donor of the flavin. However due to its distance from the flavin, the probability of its involvement could be low. Also shown are Y431 and Y487. These two residues are found lying closely to W C and W D at a distance of about 8 Å. These tyrosines could potentially play the role of a final donor in the electron transfer chain, for instance by reducing oxidised W C . Either of these residues could be the one claimed by Weber et al.[6] to be the final electron donor of Xl64 at 1.4 µs. Another observation of a tyrosine radical species has been reported by Yamada et al.[7] (cf. Chapter 1) in the reduction of Xl64, from FAD ox to FAD •-, at 200 K. It is important to note that Yamada et al., in contrast to Weber et al., do not claim explicitly that a tyrosine is the final donor in the electron relay. Rather it is somehow involved in the formation of FAD •-either in lieu of or in together with a tryptophan.

Figure 2 .

 2 Figure 2. Protein surface showing W C (blue) and W D (orange) of Xl64 photolyase

  Xl64 photolyase has, to the best of our knowledge, not yet been studied by transient absorption spectroscopy. The only spectroscopic studies available on the photoactivation of Xl64 photolyase are the EPR study by Weber et al. [6] and the FTIR studies by Yamada et al. [7]and Zhang et al. [20] (see Chapter 1). Femtosecond pump-probe transient absorption spectroscopy experiments were performed in the Group of Ultrafast Photochemistry at the Département de Chimie of Ecole Normale Supérieure in Paris. Real-time nanosecond transient absorption spectroscopy was performed in the laboratory of Klaus Brettel in the Service de Bioénergetique, Biologie Structurale et Mécanismes, iBiTecS at CEA Saclay. The Xl64 samples that we used in our experiments were prepared by Junpei Yamamoto of the Osaka University, Japan.

Figure 3 Figure 3 .

 33 Figure3shows these normalised spectra in a buffer solution of 100 mM TRIS at pH 8 containing 100 mM NaCl, and 25 % (v/v) glycerol.

BFigure 4 .

 4 Figure 4. A) Absorption spectra of FAD ox (black), FADH • (red), and FADH -(green) in Xl64 reproduced from Schleicher et al.B) Absorption spectrum of (6-4) photolyase from X. laevis (cyan) with fit line (blue). The sample

Figure 5 .

 5 Figure 5. Transient absorption spectrum of Xl64 photolyase at 100 ps pump-probe delay (3z kin recording).

Figure 7 .

 7 Figure 7. A: Decay-associated difference spectra (DADS) of the global fit of the isotropic 3z kin data, with 3 exponentials plus a plateau. B: Corresponding evolution-associated difference spectra (EADS).

τ 1 =

 1 0.44 ± 0.01 ps, τ 2 = 8 ± 1 ps, τ 3 = 51 ± 2 ps, τ 3 = 229 ± 17 ps, and τ 5 = ∞, with a mean residue of 1.5 × 10 -5 . The corresponding DADS and EADS are shown in Figure 8.

Figure 8 .

 8 Figure 8. A): Decay-associated difference spectra (DADS) of the global fit of the isotropic 3z kin data, with 4 exponentials plus a plateau. B): Corresponding evolution-associated difference spectra (EADS).

10 ( 8 (Figure 9 .

 1089 Figure9. Simulated difference spectra of the putative products of photoactivation, according to Brazard et al.[5]. In frame A the reduced flavin is FAD •-and in frame B it is FADH • . Frame C recalls the reference spectra used to calculate the simulated difference spectra.

Figure 10 .

 10 Figure 10. Superposition of EADS 2 to 4 (3-exponential fit of the 3z kin recording) and simulated spectrum, all normalised at 447 nm.

Figure 11 .

 11 Figure 11. Difference spectrum of Xl64 photolyase upon formation of FAD •-at 200 K. Reproduced from Yamada et al. [7].

Figure 12 .

 12 Figure 12. A: Cascading scheme involving 4 transient species, no back reaction but ground-state recovery channels at each step of the conversion. B: Same with 5 transient species.

Figure 13 .

 13 Figure 13. A: Isotropic species-associated difference spectra (SADS) in the context of the 4-species cascading model with population losses of Figure 12. B: Corresponding deconvoluted population dynamics.

Figure 14 .

 14 Figure 14. A: Isotropic species-associated difference spectra (SADS) in the context of the 5-species cascading model with population losses of Figure 12. B: Corresponding deconvoluted population dynamics.

τ 1 =

 1 0.44 ± 0.01 ps, τ 2 = 7 ± 1 ps, τ 3 = 54 ± 2 ps, τ 3 = 274 ± 21 ps, and τ 5 = ∞, with a mean fitting residue of 1.7 × 10 -5 . One can verify that these values are close to those obtained with the isotropic data alone. The corresponding EADS, in both polarisations, are shown in Figure16.

Figure 16 .

 16 Figure 16. Polarisation-dependent EADS associated to the global analysis of the 3z kin measurement with a sum of 4 exponentials and a plateau: parallel on the left, perpendicular on the right.

  1.2) to the flavin, then the middle time (36 ps) would correspond to the transfer from W B to W A and the slowest time constant (114 ps) to the transfer from W C to W B , as depicted in Figure 17. In other words species S2 would correspond to WH •+ sitting in the W A position (W A H •+ ), S3 would correspond to W B H •+ and S4 to W C H •+ . The meaning of the ground-state recovery process at each kinetic step in such a view would be charge recombination between the radicals, in competition with electron transfer, which would explain the non-unit conversion yields (essentially ϕ 2 and ϕ 3 ) estimated in §3.2.2.4. The fact that the recombination rate constant slows down as the reaction proceeds along the Trp chain (because the time constants increase) would be consistent with the increasing distance between the FAD •-and WH •+ radicals.

Figure 17 .

 17 Figure 17. Schematic view of the hypothetical electron transfer flow considered to explain the time-resolved spectroscopic data. The charge recombination channels in competition with electron transfer are not represented.

Figure 18 .

 18 Figure 18.Epsilon ratio between WH •+ (Solar et al.[21]) and FAD •-(Berndt et al.[23]) . Insert: Absorption spectra of WH •+ (red) and FAD •-(black).

Figure 20 displays

 20 Figure 20 displays the same raw anisotropy data as a function of time at a few of selected wavelengths chosen in the blue transient absorption band (375 and 400 nm; blue crosses and cyan squares, respectively), in the bleaching band (447 nm; green triangles) and the red transient absorption band of the products (580 nm; red circles). The data have here been averaged over about 2 nm (5 pixels) in order to smooth out minor spectral fluctuations. The continuous lines are the corresponding anisotropy traces deduced from a global fit of the polarised transient absorption data, with four exponentials plus a plateau (presented in §3.2.3). Section 3.2.4.4 below will give some more details on these traces; they here essentially serve as a guide to the eye.

Figure 20 .

 20 Figure 20. Kinetic traces of the raw transient anisotropy of Xl64 (3z kin measurement) at selected wavelengths.

Figure 21 .

 21 Figure 21. Evolution-associated anisotropy spectra" (EAAS) associated to the global analysis of the 3z kin raw anisotropy spectra with a sum of 3 exponentials and a plateau.

  This objective can easily be reached by recurring to a global simultaneous fit of the polarised transient absorption data, both parallel and perpendicular, as presented above in §3.2.4. This approach first provides polarised EADS, associated to a purely formal sequential scheme with 100% conversion yields between species. At this stage we can use the EADS to construct what could loosely be called "fitassociated anisotropy spectra" (FAAS), according to Equation (3.7):These FAAS are thus defined as the intrinsic anisotropy spectra associated to the EADS.Interestingly let us recall from §3.2.2.5 and §3.2.3 that the species-associated difference spectra (SADS) of the cascading models of Figure12are proportional to the EADS. This was established in the simplified case of the isotropic transient absorption spectra and remains true for the polarised data, the proportionality factor being identical for parallel and perpendicular polarisations. It is then straightforward to now define "species-associated anisotropy spectra" (SAAS), according to Equation(3.8). Within the models of Figure12, SAAS are just equal to FAAS.

Figure 22 .

 22 Figure 22.A: "Species-associated anisotropy spectra" (SAAS) relative to the kinetic model of Figure12A(and global analysis of the 3z kin measurement with a sum of 3 exponentials and a plateau). B: SAAS associated to the kinetic model of Figure12B(and fit with 4 exponentials and a plateau).

Figure 23 .

 23 Figure 23. Anisotropy kinetic trace at 580 nm (3z kin recording). Open circles: raw anisotropy. Green line: anisotropy reconstituted from global simultaneous fit with 3 exponentials and a plateau. Blue line: 4-exponential simultaneous fit. Red line: direct global fit of the raw anisotropy data with 3 exponentials and a plateau.

  our best anisotropy kinetics at 580 nm, i.e. of the 3z kin measurement, along with error bars at 95% confidence. The data have been in this figure averaged over about 2 nm (5 pixels) in order to smooth out minor spectral fluctuations. The error bars were calculated as follows: (i) at each time (and each wavelength), we collected the list of 80 polarised transient absorption recordings (corresponding to 80 consecutive kinetic scans) that we averaged to produce the final value of this experiment, (ii) a list of 80 individual anisotropies were calculated and statistically analysed, (iii) the standard deviation of this last distribution was computed and the standard error of the mean (SEM) was deduced, (iv) the SEM was multiplied by two to yield the half width of the 95% confidence interval.

Figure 24 .

 24 Figure 24. A: Kinetics of the raw anisotropy at 580 nm (blue circles) for the 3z kin data with error bars at 95% confidence. In continuous blue line is recalled the anisotropy reconstituted from the 4-exponential fit. B: Corrected anisotropy kinetics (translated by +0.02), both raw (red circles) and fit (red line).

Figure 25 .

 25 Figure 25. Kinetics of anisotropy at 580 nm. Open green circles: corrected 3z kin recording (fit in continuous line). Open orange squares: 2z kin2 recording (fit in continuous line). Filled blue circles: corrected 3z 3T recording at 3 times only. Filled red squares: 2z 3T recording. Error bars at 95% confidence are represented.

Figure 26 .

 26 Figure 26. Kinetics of anisotropy at 580 nm. Blue line: fit of corrected 3z kin recording. Red line: fit of 2z kin2 . Green dotted line: fit of 2z kin1 . Cyan circles: 3z 3T . Orange squares: 2z 3T .

  scheme of Figure12A. As a first step we here calculate the intrinsic anisotropies expected from each of the tryptophans in the active site which we expect to take part in the photoreduction mechanism of the flavin. To do this, we must calculate the angle (β) between the transition dipole moment the protonated tryptophanyl radical (lowest transition probed at 580 nm) and that of the oxidised flavin (lowest transition excited at 475 nm). The orientations of these dipole moments were determined using the homology model and published transition moments for the flavin and the tryptophanyl radical. After having calculated β, calculating the expected anisotropy becomes straightforward by using Equation (3.5).The experimental determination of the orientation of the S 0 →S 1 transition dipole moment of the isoalloxazine ring of the flavin has been published in at least three reports. The first one byEaton et al. reported the angle between the transition moment (lowest transition) and the short (N5-N10) axis of the molecule (x axis in Figure27) to be 75° ± 4° (counted counterclockwise)[39]. This result was obtained from the polarised absorption of a single crystal of oxidised flavodoxin, a flavoprotein noncovalently binding flavin mononucleotide (FMN); it was internally supported by molecular orbital calculations performed by the authors themselves. This work is supported by a relatively recent computational study by Climent et al. by giving the same value of 75°[40]. This latter value has been recently used by Immeln et al. for the same purpose we are now pursuing, in a plant cryptochrome[17].On the other hand, Johansson et al. reported an angle of 58° ± 4° for FMN in model membranes[41].Matsuoka et al. and claims this angle to be 45 ± 4° for riboflavin in stretched polyvinyl alcohol films

Figure 27 .

 27 Figure 27. A: Orientation of the transition dipole moment for the lowest transition of the isoalloxazine ring of FAD (75° counted counterclockwise from x axis). B: Orientation of the transition dipole moment for the lowest transition of the indole ring of the WH •+ radical (110° from x axis)

Figure 28 compares

 28 the experimental kinetic trace of Liu et al. to our simulation No particular attention should be paid to times below 2 ps.

Figure 28 .

 28 Figure 28.Comparison of the experimental anisotropy kinetics at 580 nm (red line) of Liu et al.[16] relative to the photoreduction of oxidised EcCPD, and our simulation (blue line) of the same based on the crystal structure of EcCPD[44].

Figure 29 .

 29 Figure 29. Anisotropy at 580 nm of the 2z and 3z data overlain with the predicted anisotropy based on the working cascade model.

Figure 30 .

 30 Figure 30. Isotropic transient absorption spectrum of Xl64 photolyase at 3 ms.

Figure 31 showsFigure 31 .Figure 32 .

 313132 Figure31shows the absorbance change of the system at 457 nm. The signals are negative. This is expected from the results of the ultrafast experiments. The negative signal is due to the bleaching of the ground state of the flavin. Polarisation effects are observed until 5 µs. After this time, the signals of both the parallel and perpendicular polarisations are constant at 3 mOD. The same constant bleach was observed in the ultrafast experiments starting from 500 ps until 1500 ps.

Figure 33 .

 33 Figure 33. Anisotropy decay at 457 nm

Figure 34 .

 34 Anisotropy decay monitored at 562 nm.

Figure 35 .

 35 Figure 35. Isotropic transient absorption kinetics monitored at 457 nm.

Figure 36 .

 36 Isotropic transient absorption decay monitored at 562 nm.

Figure 1 .

 1 Figure 1. Absorption spectra of the T(6-4)T photoproduct (green) and the intact TT nucleobases (black).

Figure 2 .

 2 Figure 2. Absorption spectrum of fully reduced X. laevis (6-4) photolyase.

Figure 3 .

 3 Figure 3. Spectra of the photolyase-photoproduct solution over successive periods of blue light illumination.Arrows show the evolution of the spectra with increasing illumination time. The sample contained 60 µM of the DHT substrate and 1.7 µM of X. laevis photolyase. The arrows show the evolution of the spectra with every illumination period.

  linear. This corresponds to the conversion of the T(6-4)T to the intermediate. After the formation of the intermediate, further illumination would bring about the restoration of the intact TT. Only then would absorbance increase be observed at 265 nm. These predictions are based on the assumption that the intermediate, if present, does not absorb at the observation wavelengths. The plots of absorption change as a function of illumination time of the spectra in Figure 3 are shown in Figure 4.

Figure 4 .

 4 Figure 4. Absorption changes at 265 and 325 nm plotted against illumination time.

Figure 5 .

 5 Figure5. Photorepair spectra for the normal 8-mer substrate. Absorption spectra were recorded after successive periods of irradiation with 384 nm light. Insert: Absorption change at 325 nm as a function of total irradiation time (symbols, using the same colors as the corresponding spectra) and linear regression line through the origin. The sample initially contained 1.13 µM Xl (6-4) photolyase with fully reduced flavin cofactor and 36.4 µM substrate.

Figure 7 .

 7 Figure 7. Control signals for the flash excitation data. The signals were arbitrarily offset for clarity.

Figure 8 .

 8 Figure 8. a) Averaged absorption changes induced by eight successive laser flashes observed on a sample having been exposed to eight such flashes immediately prior. Average of three measures. b) Absorption changes induced by a single such flash on a sample kept in the dark for at least 20 min. Average of seven measures. The sample contained 4.9 µM of the photoreduced X. laevis (6-4) photolyase and 20 µM of DHT (6-4) photoproduct 10-mer.

Figure 9 .

 9 Figure 9. Stairs measurement. Arrows indicate the moment the light flash is introduced.A line is drawn arbitrarily to show the quadratic trend. The sample contained 3.9 µM of the reduced X. laevis photolyase and 20 µM of the DHT 10-mer substrate.

Figure 10 .

 10 Figure 10. Stairs measurements performed at different dark adaptation times (Dark time in figure) after a preceding set of 20 flashes. The "repair lag" is the time necessary to achieve a constant step height within a flash sequence. It was obtained as illustrated by the straight lines in the figure.

Figure 11 .

 11 Figure 11. Repair lag as a function of the dark adaptation time fitted with a single exponential function with a time constant of 1.7 min.

Figure 12 .

 12 Figure 12. Long-term absorption changes at individual wavelengths of a dark-adapted sample (8.8 µM reduced X. laevis (6-4) photolyase and 33 µM of the DHT 10-mer substrate). The signals were globally fitted with a single exponential function with a time constant of 1.8 min.

Figure 13 .

 13 Figure 13. Normalised amplitudes associated with the decay curves in Figure 13 superimposed with the normalised absorption spectrum of the T(6-4)T photoproduct. The error bars presented here represent the standard amplitude errors of the global fit.

Figure 14 .

 14 Figure 14. FADH • -FADH -difference spectrum obtained from Thiagarajan et al. PNAS 2011.

1

 1 Analysis of the suggested repair reaction scheme3.1.5.1.1 General assumptionsUnder the experimental conditions, the following simplifying assumptions appear justified:1.Substrate binding and product release are much faster than the overall repair rate (<0.01 s -

Figure 15 .

 15 Figure 15. Simulation of the kinetics of the 265 nm absorption changes under continuous light excitation using Equation (6) and relation  r A s r 265 265 ) (     

Figure 16 .

 16 Figure 16. Corrections of the first flash signal at 325 nm. Black trace, raw signal taken from Figure 2. Blue trace, raw minus S only signal. Red, raw minus S only minus E only signal. The subtracted S only signal was measured under repetitive flash excitation to avoid unnecessary degradation of the signal-to-noise ratio.

Figure 17 .

 17 Figure 17.Confidence regions for the ( 1 ,  2 ) pair. The zone between the blue curves corresponds to limiting values of  SS (cf. Section 3.1.5.3.1). The zone delimited by the green dashed lines (and of course  2 ≤ 1) is deduced from the flash (first vs. successive) excitation experiment (cf. Section 3.1.5.3.2). The zones limited by red curves are deduced from the fit of the "stairs" experiment, and our simulations of it (cf. Section 3.1.5.3.3). The red dotted curve corresponds to  = 0.191 and p 0 = 3.9 µM. The (internal) thin red solid curve corresponds to  = 0.191 and p 0 varying between 3.7 and 4.1 µM. The (external) thick red solid curve corresponds to  varying between 0.156 and 0.225, and p 0 varying between 3.7 and 4.1 µM. The quantum yields obtained from the best fit of the "stairs" experiment ( 1 = 6.7% and  2 = 83%) are recalled by the red cross.

. 1 * , η 2 *) 1 * , η 2 *. 1 *

 12121 Let us first analyse the meaning of the data collected by the above series of simulations. By making histograms of the pool of optimal A and B data (we ignore f x0 as this parameter is not relevant to our problem), one can readily estimate the probability density function p(A, B| η that a fit provides parameters A and B, given that the generators, playing the role of underlying reality, are η It is implicitly assumed that α * = 0.191. The quantity that we actually need is the probability p(η the fit parameters are A and BApplication of the wellknown Bayes theorem[18] leads to the following relation:

, 1 * , η 2 *)

 12   ) generators, chosen in the vicinity of the values deduced from the experimental fit, that the following functional form approximately applies: p(x, y| η = n x (x; x * ,  x ) n y (y; y * ,  y )

2 Figure 18 .

 218 Figure18. Spectra used to estimate the excitation rate of FADH -in photolyase exposed to sunlight. (a) Solar irradiance reference spectrum, representing the annual averaged daylight terrestrial hemispherical irradiance incident on a sun-facing, 37° tilted surface at clear sky conditions for the 48 contiguous United States. The original data[19] were converted from W m -2 nm -1 to photons s -1 cm -1 nm -1 . (b) Absorption spectrum of Xl (6-4) photolyase with fully reduced flavin cofactor (FADH -; red line) obtained by photoreduction (see Section 1) of the enzyme containing fully oxidized flavin (FAD ox ; black line). The absorption cross sections σ (right hand scale) were obtained by scaling the spectrum before photoreduction to ε = 11.2 mM -1 cm -1 in the 450 nm peak of FAD ox which corresponds to σ = 4.28 × 10 -17 cm².

Figure 19 .

 19 Figure 19. T(6-4)C photoproduct

Figure 20 .

 20 Figure 20. Absorption spectrum of the T(6-4)C photoproduct phosphoramidite § § .

Figure 22 .

 22 Figure 22. Repair of the T(6-4)C DHT substrate by X. laevis (6-4) photolyase

Figure 23 .

 23 Figure 23. Differential spectra of the photorepair of the DHT substrate of the T(6-4)C photoproduct.

  photolyase. Bearing in mind the landmark ultrafast experiments Li et al.[1] performed on T(6-4)T photorepair by A. thaliana photolyase (see Chapter 1, §1.3.1.2), which is the only one of this type published to date, our motivations were at least threefold. A first obvious interest of our work is to independently reproduce this type of time-resolved experiment on a different photolyase, thereby expanding the number of available studies to at least two. More specifically the advantage of our experimental technique is to provide full transient absorption spectra on a wide spectral interval, which is then expected to allow more precise identification of the transient states than with a limited number of independently measured kinetics. Thirdly our study will be the first one taking the 2-photon repair mechanism into consideration. Li et al.[1] indeed fully interpreted their results within the framework of a 1-photon process.It has to be noted that, since the spectral observation window of our setup does not extend below about 350 nm, the experiments presented in this chapter essentially reveal photoinduced changes occurring at the level of the flavin cofactor. Depletion of the substrate absorption (visible at 325 nm) and restoration of intact T-T absorption (at 265 nm) are not observable in those measurements. Let just mention that several follow-up studies at scheduled in the near future. Extension of the continuum probe below 350 nm may be possible by replacing some optical components (filter, lenses) that presently absorb light in this region (transient absorption spectra of riboflavin extending down to ~280 nm were for example obtained by Weigel et al.[31]. It could even be considered to produce a specific probe at 265 nm, for instance by frequency doubling of a beam at 530 nm. It is also planned to monitor the photorepair reaction in the ns-µs-ms regime, including by probing at 265 nm, by real-time ns spectroscopy in the laboratory of a K. Brettel at CEA-Saclay.Before presently the results is worth at this asking what our femtosecond transient absorption experiment is going to probe. Will it be the mostly the first step of the photorepair reaction (i.e. the conversion of the T(6-4)T photoproduct to intermediate X) or will the second step of the photorepair (conversion of intermediate X to intact T-T pair) also contribute to the data -and to what extent? The question in fact arises because a very large number of excitation pulses (several hundred thousand to millions) is necessary to record good transient absorption spectra and kinetics with such our set-up. This

Figure 24 .

 24 Figure 24. Fluorescence spectra of FADH-when in solution (black) and when bound to EcCPD photolyase (red) excited at 360 nm extracted from Kao et al.[32].

Figure 26 .

 26 Figure 26.Transient isotropic absorption spectrum of the fully reduced X. laevis photolyase at a pump-probe delay of 300 fs, after femtosecond excitation at 387.5 nm.

Figure 28 .

 28 Figure 28. DADS (A) and EADS (B) spectra obtained from the global fit of the transient absorption spectra of fully reduced Xl64.

Figure 29 .

 29 Figure 29. Normalised EADS (from Figure 28B) at 575 nm showing the similarity of all four spectra.

Figure 31 .

 31 Figure 31. DADS (A) and EADS (B) spectra obtained from the global fit of the transient absorption spectra of fully reduced Xl64 in the presence of T(6-4)T substrate.

Figure 32 .

 32 Figure32. A) Comparison of EADS5 and the FADH • -FADH -difference spectrum. The spectra were normalised at 580 nm. B) Spectra of FADH -and FADH • from Xl64 digitised from Schleicher et al.[34].

Figure 34 .

 34 Figure 34. Hypothetical equilibrium between FADH -* /T(6-4)T and FADH • /T(6-4)T •-.

Figure 1 .

 1 Figure 1. Thymine and Dihydrothymine (DHT).

Figure 3 .

 3 Figure 3. Set-up for femtosecond transient absorption spectroscopy. The additional path in the 3z configuration is shown as broken lines.

  the intensity of probe beam transmitted by the sample and I r that of the reference beam. Depending on their presence or absence of the two beams four different configurations can be generated, as presented below:  pump on (p) and probe on (pr): probes the absorption of the excited sample  pump off (np) and probe on (pr): probes the absorption of the ground state  pump on (p) and probe off (npr): measures pump scattering and sample fluorescence  pump off (np) and probe off (npr): detects the background noise

4. 3 Artefacts 4 . 3 . 1

 3431 Pump energy fluctuationDuring the course of an ultrafast spectroscopy experiment, the pump energy undergoes fluctuations to the order of about 2%. We have pointed out that the intensity of the pump affects the amplitudes of the signals we measure (cf. Section 4.2.2.2.3). Since the intensity of the pump beam is proportional to the intensity of the signal, we also expect the measured signals to have fluctuating amplitudes. These fluctuations should, in principle, be correctable by normalising the measured signal intensities with those of the pump. However for small pump fluctuations as those that we have in our experiments, we observed that the fluctuations of the transient absorption signal and the fluctuations of the pump intensity (as measured by the photodiode) are ill-correlated. As a result normalisation gives almost no effect. We therefore are unable to correct for the pump fluctuations. The transient absorption fluctuations give a large effect in anisotropy measurements since the anisotropy depends on the difference between the intensities of the parallel and perpendicular signals. This effect is most evident when parallel and perpendicular signals are measured separately. It is for this reason that we implemented the simultaneous detection of parallel and perpendicular signals in our 3z setup.

Figure 4 .

 4 Figure 4.

Figure 5 .

 5 Figure 5. Typical chirp curve where t 0 (ps) varies with the wavelength due to group velocity dispersion.

Figure 7 .

 7 Figure 7. Comparison of the anisotropy of coumarin 6 at 1500 ps using our 2z and 3z set-ups.

Figure 8 .

 8 Figure 8. Average spectrum of the negative time delays showing a non-zero signal.

Figure 9 .

 9 Figure 9. Smoothing of the "ms constant" using the Savitzky-Golay algorithm.

Figure 10 .

 10 Figure 10. A: Evolution of parallel transient absorption spectra at 2.2 ps B: the same spectra normalised at 450 nm.

Figure 11 .

 11 Figure 11. Close view of the 500 nm < λ < 650 nm region of the normalised transient absorption spectra at 2.2 ps.

Figure 12 .

 12 Figure 12. ∆∆A spectrum of spectra 7 and 1 (with k=0.9) superimposed with the 2.2 ps transient absorption spectrum of FAD ox in solution.

Figure 13 .

 13 Figure 13. Amplitude decrease at different wavelengths of all 53 parallel spectra at 2.2 ps.

4. 4 . 1

 41 Spectral analysis of transient absorption signalsThe different spectral features observed in a differential absorption spectrum are due to the sum of three contributions: transient absorption, bleaching, and stimulated emission. The absorption of transient species (ε u : S 1 →S n , T 1 →T n , etc.) corresponds to the positive contributions. Two processes account for the negative contributions. The first of these is the loss of ground state absorption (ε a : S 0 →S 1 ) due to excitation. Stimulated emission also contributes negatively. It corresponds to the amplification of the probe photons by stimulated emission from the excited state (ε e : S 1 →S 0 ).

  photolyase is not fully compatible with the simple picture of an electron hole travelling from W A to W B and to W C . The anisotropy kinetics identifies the first reducing agent to be most probably W A . Data further suggests that the population of W B is less important than what is predicted by the hypothetical model. This may indicate that a simply transient role for W B . The anisotropy value at the end of our time window of 1500 ps in the femtosecond experiments

  Maul et al.'s work clearly showed the absence of the oxetane species in the enzyme-substrate complex prior to photorepair which refutes the previous notion by Sancar that states that the oxetane is thermally formed upon the binding of photolyase to the (6-4) lesion[36]. This landmark contribution has sparked new studies of the (6-4) photolyase repair mechanism; mostly theoretical and exclude the oxetane intermediate[START_REF] Domratcheva | Electronic structure of (6-4) DNA photoproduct repair involving a non-oxetane pathway[END_REF][100][101][102]. Maul et al. themselves proposed a mechanism, called the transient water mechanism, shown in

  Heelis and A. Sancar while working with the FADH • to FADH 2 (i.e. what was believed to be FADH 2 at that time) reduction in E. coli CPD photolyase (EcCPD contains FADH • after purification ) observed that the reoxidation of FADH 2 to FADH • is slower in the presence of an external reducing agent such as DTT[113]. They deduced that the protein contains an intrinsic photoreductant that is probably an amino acid residue. A few years later in 1990, they identified this to be a tryptophan[114].In 1991, Li et al. prepared phenylalanine mutants of the 15 tryptophan residues present in EcCPD.

1.3.2.3 Cryptochrome photoreduction

	case, protonation is believed come from an occluded water molecule [140]. The presence of cysteine on
	this position stabilises FAD •-as had been observed by Iwata et al. [144]. Liu et al. could even stabilize
	FAD •-for in a double mutant of EcCPD [145].They replaced the N378 asparagine (facing N5) by a
	cysteine and the surface-exposed E363 glutamate by a leucine.
	Cryptochromes share high sequence homology with photolyases. In particular, (6-4) photolyases
	show very high homology with mammalian circadian clock cryptochromes [42]. It has been suggested
	that mammalian cryptochromes exhibit light-response mechanisms akin to the photoactivation
	mechanism of photolyases [146]. Plant cryptochrome on the other hand appear to be capable of
	mechanisms distinct from the tryptophan triad based mechanism of photolyases. These mechanisms are
	presented in the following.
	In 2005, Zeugner et al. showed that the photoreduction of Arabidopsis cryptochrome 1 is
	dependent on the tryptophan triad using phenylalanine mutants of the W A and W C [147]. They observed
	that the phenylalanine mutants showed a marked reduction of the light-activated autophosphorylation in
	vitro and in vivo photoreceptor function. In 2010, Brazard et al. showed in the DASH cryptochrome of
	O. tauri that the photoactivation follows the Brettel-Vos mechanism of photolyases [135].
	In 2011, Li et al. performed photoreduction experiments on the phenylalanine mutants of the
	tryptophan triad in the highly homologous Arabidopsis cryptochrome 2 [92]. They found that while the
	phenylalanine mutants lost photoreduction activity in vitro, the in vivo biological functions were not
	hampered as a consequence of the mutation. Their results led them to conclude that cryptochromes
	function with a photoreduction mechanism independent of the tryptophan triad established in photolyases.
	They further stated that the tryptophan triad was conserved in cryptochromes most likely for the
	preservation of structural integrity over its photochemistry. Other studies in which single Trp residues of
	the conserved Trp triad were mutated showed no physiological effect in vivo and/or only minute effects
	in vitro [75, 148-150]. Biskup et al. "identified a partly conserved Trp nearby the Trp triad as an
	alternative terminal electron donor" in Synechocystis Cry-DASH [151]. More recently Biskup et al.
	demonstrated that "electron abstraction from a nearby amino acid by the excited FAD triggers further
	electron transfer steps even if the conserved chain of three tryptophans is blocked" in X. laevis
	cryptochrome DASH. Secondary electron transfer pathways were identified as shown in Figure 29 [152].
	•-is unstable and is easily protonated to FADH • . It has been shown by
	Kottke et al. in A. thaliana cryptochrome 1(AtCRY1) that the aspartic acid residue itself is the proton
	donor [141]. This is not the case for asparagine due to its very high pK a of 17 [142]. Rather, it is believed
	that asparagine creates a network of hydrogen bonds that facilitate the protonation of FAD •-[143]. In this

  •-/W379 • radical pair. The authors state that this fast deprotonation is hidden behind a vibrational cooling process seen with a timescale of 31 ps.

	The idea of radical pair stabilisation via proton transfer was supported later in 2012 by a
	theoretical study by Solov'yov et al. on another plant cryptochrome, Cry1from A. thaliana [154]. Using
	quantum chemical calculations and molecular dynamics simulations, the authors found that upon
	formation of the FAD •-/WH A •+ radical pair (as a consequence of photoexcitation), FAD •-is protonated
	by an aspartic acid residue (D396, not present in animal cryptochromes). The stable FADH • /W A • pair is
	then formed by the protonation of D396 -by WH A •+ . Alternatively, WH A •+ can be reduced by W B thus
	providing two pathways for photoreduction. The two pathways were said to be interconvertible to the
	other via an electron transfer between W A and W B .
	Recently, the effect of ATP binding on the photoactivation of cryptochromes has been reported by
	Müller et al. [155]. The authors studied the effects of ATP, bound by A. thaliana cryptochrome
	(containing FAD ox ), on the dynamics and yield of FADH • formation. They found that ATP binding
	enhances the yield of FADH • from the photoreduction. They claim that this is explained by the fact that
	ATP effectively favours the protonation of a nearby aspartic acid residue and a productive electron
	transfer from the tryptophan triad.

462(1-3): p

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	Zabulon, and C. Bowler, Spectro-temporal characterization of the photoactivation mechanism of

  •-in DmCry, not in Xl64. The red structured band of EADS3 and EADS4 around 590 nm, with local maxima around 575 and 612 nm, looks very similar to a homologous band of the simulated spectrum, located around 570 nm, arising from the WH •+ contribution. It is however significantly red-shifted by about 20 nm. One could argue that this shift is due to the specific environment of WH •+ in Xl64, which is different from that in Solar et al.[21][22][23] (radiochemically produced species in solution). Local polarity, hydrogen bond interactions or electrostatic interaction with FAD •-or other charged residues could be invoked. It cannot be excluded that the difference might also arise from the presence of additional contributions, due to other species excited by the pump beam. One can, in particular, think of FADH -, or FADH • possibly present in small proportions and escaping from the analysis of the steady-state spectrum ( §3.2.1). We have checked that excited FADH -in Xl64 (see Chapter 3) has a long lifetime (3.8 ns) and exhibits a broad positive band extending far in the red (in agreement

  •-and WH •+ radicals. The particularly broad and structureless red absorption of EADS2 is tentatively assigned to a hot WH •+ radical. EADS 3

and EADS3 would correspond to vibrationally relaxed WH •+ radicals. The apparently red shifted spectrum of WH •+ , as compared to solution measurements, is proposed to be due to specific interactions inside the protein. The data do not show any trace of protonation of FAD •-. The contribution of small amounts Y • and/or W • in EADS2 is not completely ruled out.

  2.2.3). The corresponding DADS are given in Annex D. As a compact substitute for the raw spectra Figure15displays the polarisation-dependent EADS. Polarisation-dependent EADS associated to the global analysis of the 3z kin measurement with a sum of 3 exponentials and a plateau: parallel on the left, perpendicular on the right.

										0.004
	Spectra	0.008	Parallel			EADS1 EADS2	Spectra		Perpendicular		EADS2 EADS1
	Difference	0.004					EADS3 EADS4	Difference	0.002					EADS3 EADS4
	Species-Associated	-0.004 0.000							Species-Associated	0.000
										-0.002
		400	450	500	550	600	650	700	750	400	450	500	550	600	650	700	750
				Wavelength / nm						Wavelength / nm
	Figure 15.

Table 2 .

 2 Results of the global simultaneous fits of both the parallel and perpendicular transient absorption spectra of Xl64. Two different measurements (2z kin2 and 3z kin ) were analysed, with either 3 exponentials plus a plateau or 4 exponentials plus a plateau. The time constants are given in lines τ i and the average residue of the fit is indicated in line <res>.

		2z kin2		3z kin
		3 exp. + plateau	4 exp. + plateau	3 exp. + plateau	4 exp. + plateau
	τ 1	0.47 ± 0.02 ps	0.46 ± 0.01 ps	0.44 ± 0.01 ps	0.44 ± 0.01 ps
	τ 2	26 ± 3 ps	14 ± 4ps	39 ± 4 ps	7 ± 1 ps
	τ 3	113 ± 11 ps	56 ± 8 ps	120 ± 14 ps	54 ± 2 ps
	τ 4	∞	319 ± 108 ps	∞	274 ± 21 ps
	τ 5	-	∞	-	∞
	<res>	3.9 × 10 -5	3.7 × 10 -5	2.5 × 10 -5	1.7 × 10 -5

  Raw transient anisotropy of Xl64 (3z kin measurement) at selected pump-probe delays between A) 0.1 to 2.2 ps, B) 2.2 to 221.8 ps, and C) 221.8 and 1500 ps.

		0.8	A					0.1 ps
								0.3 ps
								0.5 ps
								0.7 ps
		0.4						2.2 ps
	r	0.0
		-0.4
		0.8
		0.4	B					27.9 ps 2.2 ps
		0.2						111.1 ps
	r	0.0						221.8 ps
		-0.2
		0.4						221.8 ps
								701.5 ps
		0.2						1500 ps
	r	0.0
		-0.2
			400	450	500	550	600	650	700
					Wavelength / nm

C Figure 19.

  •-. Figures 19B and 19C displays the evolution of the anisotropy on a longer timescale. Between 2.2 and 221.8 ps the changes are quite subtle. A spectral evolution is observed in the spectral region below 375 nm; the anisotropy decreases from 0.34 to 0.24 at 360 nm. At 580 nm the anisotropy is seen to slightly rise, from -0.07 to -0.04, then return back around -0.07. At 447 nm the anisotropy remains nearly constant, which further the assumptions of §3.2.2.4. Between 221.8 ps and 1500 ps, no discernible change is observed in the anisotropy spectra.

Table 3 .

 3 Anisotropy values at 580 nm of the SAAS (FAAS) corresponding to the kinetic models of Figure9and the global simultaneous fits of the polarised transient absorption spectra. Two different measurements (2z kin2 and 3z kin ) were analysed, with either 3 exponentials plus a plateau or 4 exponentials plus a plateau. The corresponding time constants are given in Table2.

		2z kin2		3z kin
		3 exp. + plateau	4 exp. + plateau	3 exp. + plateau	4 exp. + plateau
	r 1	0.56	0.53	1.55	1.47
	r 2	-0.067	-0.078	-0.062	-0.077
	r 3	-0.008	-0.003	-0.013	-0.029
	r 4	-0.040	-0.073	-0.077	-0.064
	r 5	-	-0.012	-	-0.069

Table 4 .

 4 Angles between WH •+ and FAD ox transition moments and corresponding anisotropies, associated to the different tryptophan residues potentially involved in the photoactivation of Xl64 photolyase. The calculations were based on our homology model of the protein. Errors corresponding to variations of the orientations of both transitions moments within an interval of ±5° are indicated.

		Angle between WH •+ and FAD ox	Theoretical Anisotropy
		transition moments
	W A	55.3° ± 6.5°	-0.006 ± 0.07
	W A'	88.4° ± 8.5°	-0.20 ± 0.02
	W B	149.7° ± 5°	+0.25 ± 0.05
	W B'	145° ± 7.5°	+0.20 ± 0.07
	W C	60.0° ± 9°	-0.05 ± 0.08
	W D	76.8° ± 4.5°	-0.17 ± 0.02

  we could simulate the anisotropy kinetics one would expect at 580 nm for the model of Figures12A and 17. Equation (3.7) was used for that purpose. Here the index i refers to all molecular species supposed to contribute to the transient absorption at 580 nm: i.e.

	FAD ox * , W A H •+ , W B H •+ and W C H •+ (assuming FAD •-does not contribute at 580 nm after the decay of
	FAD ox

*

).

  13 s -1 .

		0.10		-2.3
			nm	-2.4
		0.08	A 378	-2.5
	Absorbance	0.06	ln	-2.7 -2.6	0.0 0.2 0.4 0.6 0.8 1.0 1.2 Illumination time / s
		0.04
		0.02
		0.00
		350	400	450	500
			Wavelength / nm

Figure 6

. DMAD actinometry. Absorption spectra were recorded after successive periods of irradiation with 384 nm light. Insert: Natural logarithm of the absorbance at 378 nm as a function of total irradiation time (symbols, using the same colors as the corresponding spectra) and linear regression line. The sample contained 2.8 µM DMAD in 50 mM H 2 SO 4 . The slope obtained in this experiment is -0.367.

Table 1 .

 1 Table of repair lag values corresponding to different dark adaptation times.

	Dark adaptation time	Repair lag
	35 s	1.85 s
	2 min	4.59 s
	5 min	6.17 s
	20 min	6.56 s

  Evolution of the transient absorption spectrum of fully reduced Xl64 in the presence of T(6-4)T substrate for increasing pump-probe delays between 0.3 to 3200 ps. The sample was excited at 387.5 nm.

		0.0014
		0.0012							875 ps 1348 ps
									2077 ps
		0.0010							3200 ps
		0.005 0.0008
	A	A
		0.004 0.0006							0.3 ps 0.7 ps
									3 ps
		0.0004
		0.003
	A	0.0002
		0.002 0.0000
		350	400	450	500	550	600	650	700	750
					Wavelength / nm
		0.001
		0.000
			400	450	500	550	600	650	700	750
					Wavelength / nm
		0.005
									3 ps
									9 ps
		0.004	B						28 ps 14 ps
									43 ps
		0.003							66 ps
	A
		0.002
		0.001
		0.000
		350	400	450	500	550	600	650	700	750
					Wavelength / nm
		0.0040							66 ps
		0.0035	C						239 ps 125 ps
		0.0030							369 ps 568 ps
		0.0025							875 ps
	A	0.0020
		0.0015
		0.0010
		0.0005
		0.0000
		350	400	450	500	550	600	650	700	750
					Wavelength / nm

D Figure 30.

  3.3.2 on DADS, it can be said that EADS4 is in fact a mixed spectrum containing contributions of both excited FADH -(possibly not the same one as that observed at early time; see EADS1 to EADS3) and FADH • . The 2.9-ns component corresponds to the decay of this FADH -* contribution. Comparison of EADS4 and the FADH • -FADH -difference spectrum. The spectra were normalised at 580 nm.

		1.25
			EADS4
			FADH° -FADH-
		1.00
	amplitude	0.75
	Normalised	0.50
		0.25
		0.00
		350	400	450	500	550	600	650	700	750
					Wavelength / nm
	Figure 33.

  is the absorbance of the active photolyase at the excitation

	wavelength.	t A PL  	and k AM are obtained directly from the photorepair and actinometry experiments,
	respectively.

  c 0 Hence, the differential absorption is given by:

		A (	,	t	)		A	p	(	,	t	)		np A	(	,	t	)		N i 1  		u	,	i	(	)		e	,	i	(	)		a	(	)	 	c	i	t (	)		l

Literary Review

Photoactivation

Photorepair

Experimental

The fourth and final chapter is dedicated to the presentation of all experimental methods that we employed in our studies. All spectroscopic methods, photorepair assays, data analysis, sample preparation and protein structure modelling are discussed in detail.

However, dues to the large uncertainties involved in the prediction of anisotropy values, we cannot conclude with absolute certainty on the role W D . Further experimentation is required and hence this is question is left open. The slope of the linear fit obtained for the curve in the insert of Figure 3 is -6×10 -5 s -1 . This slope is similar to the values found in the assays for T(6-4)T photorepair. For instance, for the experiment found in Figure 5 has a value of -5×10 -5 s -1 . As the T(6-4)T and T(6-4)C photorepair experiments were performed under the same irradiation conditions, the similarity of the slopes suggests that the two lesions are repaired with similar (steady-state) yields.

We also performed measurements on the DHT substrate of the T(6-4)C photoproduct. Using the DHT substrate will allow us to monitor the restoration of the repaired T-C bases at 266 nm. The DHT 10mer substrate we used has the following sequence: d(HHHHT(6-4)CTHHH). The result of this repair experiment is shown in Figure 22. Transient absorption spectrum of free reduced FAD in aqueous solution, as reported by Brazard et al. [33]; between 220 fs and 1.2 ps in (A), between 1.2 and 20 ps in (B), and between 20 and 1400 ps in (C).

The transient spectrum contains two positive peaks: one at about 390 nm and the other at 520 nm.

The authors described the multi-exponential decay of excited FADH -with the following time constants: τ 1 = 1 ± 0.2 ps, τ 2 = 5.5 ± 0.6 ps, and τ 3 = 32 ± 1 ps. The authors attribute the decay to the relaxation of the molecule along some internal degree of freedom. The decay is interpreted by Kao et al. as the crossing of conical intersections through a butterfly-like bending motion of the flavin ring. This butterflylike bending is the bending of the isoalloxazine ring of FADH -along the N5-N10 axis of the molecule.

Transient absorption spectroscopy of fully reduced Xl64, in the absence of substrate

Here we present our studies on the fully reduced Xl64. Reduction was achieved by the illumination of a sample, contained in a 10 mM phosphate buffer at pH 7 containing 100 mM NaCl, 6

Conclusion

In the first part of this chapter, we were able to demonstrate that the photorepair of the T(6-4)T photolesion by X. laevis photolyase proceeds via a successive two-photon mechanism. We have accomplished this by observing the absorption changes at 265 nm and 325 nm. The observation of the absorption changes at 265 nm were made possible by the use of the DHT substrate in order to avoid saturating the absorption by normal nucleobases. The absorbance changes at these two wavelengths as a consequence of single turnover flashes allowed us to better monitor the repair process. This is important since to prove that the repair has taken place, evidence of both the loss of the T(6-4)T photoproduct and the restoration of the intact TT bases is essential. We have estimated that the yields of the first and second photoreactions are 6.7% and 83% respectively. The low yield of the first photoreaction, corresponding to the formation of a reaction intermediate X, explains the low overall repair yield of the T(6-4)T photoproduct that we observed in the steady-state experiments to be in the order of 6%. Our results are not in contradiction with the intermediate X being the oxetane linked thymine dimer T(ox)T, as proposed by Sadeghian et al. [3], but do not constitute a proof of it. Our results should give way to future studies of the mechanism of the (6-4) photoproduct repair. Most notable would be the separate characterisation of the two photoreactions.

In the second part of this chapter, we have shown the repair of the T(6-4)C photoproduct by photolyase. We have seen using the normal T(6-4)C substrate that under the same conditions, the T(6-4)C lesion appears to be repaired similarly as the T(6-4)T photoproduct. We have done this by comparing the slopes of the repair curves of the two lesions. The quantification of the steady-state repair yield has not been performed due to the absence of the ∆ε value for the repair T(6-4)C substrate. We have also performed the repair experiment using the DHT substrate of the T(6-4)C photoproduct. This substrate allowed us to observe the restoration of the repaired TC bases at 265 nm. We have observed a non-linear repair trend in the repair curve of the DHT substrate. This was most likely due to some impurity in the substrate used. Due to insufficient material, we were unable to repeat the experiment.

We have also presented our preliminary work in the investigation of the

Photorepair

With the exception of the ultrafast measurements of the photorepair of (6-4) photolyase, all photorepair experiments were performed in the laboratory of Klaus Brettel at the Service de Bioenergetique, Biologie Structurale et Mécanismes, iBiTecS, CEA Saclay. These experiments were the steady-state photorepair of the T(6-4)T and T(6-4)C photoproducts and the flash experiments on the photorepair of the T(6-4)T photoproduct.

Substrate

We have mentioned in Chapter II that the sole criterion to really demonstrate that the substrate is actually photorepaired is to be able to detect the repaired product. We want to show that a short strand of DNA containing a (6-4) pyrimidine dimer is repaired. We therefore wish to see the pyrimidines in their restored forms, that is, as TT or TC nucleobases. Our way of viewing this restored form is by looking at their absorption bands: at 265 nm. In an oligonucleotide strand containing a single pyrimidine dimer, many nucleobases are flanking the dimer. The nucleobases flanking the lesion ensures the proper binding of the substrate to the photolyase. In our case, an 8-mer was used and we refer to this substrate as the normal substrate (see Figure 5, Chapter II). DNA nucleobases are known to absorb very strongly in the UV region. With all the nucleobases on the DNA oligomer, the absorbance increase due to the absorption of an extra TT/TC (when they are restored after photorepair) is difficult to quantify, let alone detect. For instance, in a DNA 15-mer containing one T(6-4)C photolesion * , the ratio between the absorption band of the nucleobases at 260 nm and the lesion at 315 nm is about 18. A concentration that would be enough to visualise the absorption band of the (6-4) lesion would be too much to see the 265 nm band as the detection at 265 nm is saturated. On the other hand, diluting the solution to see the 265 nm would render the absorption of the lesion too weak. We hence need to ensure that allows us to see the absorption changes on both bands clearly.

To remove the background absorption of the bases flanking the (6-4) lesion in our oligonucleotide, Junpei Yamamoto replaced these bases with dihydrothymine (DHT, H). DHT is a nonaromatic form of thymine as shown in * Used in our T(6-4)C photorepair experiment.

thymines was also available at 265 nm;  265 for repair of the DHT substrate was obtained as follows.

The absorption coefficient at 260 nm of a repaired substrate that contains a TTT sequence was calculated (according to Ref. [3]) to be 15.0 mM -1 cm -1 larger than that of the unrepaired DHT substrate, hence  260 = 15.0 mM -1 cm -1 . This value was used to scale the absorption difference spectrum for photorepair of the DHT substrate extracted from experiments as those shown in Figure 3 of Chapter III. The scaled difference spectrum yielded  265 = 16.2 mM -1 cm -1 .

Photorepair under steady-state excitation

The measurement sample consisted of 1-2 µM reduced enzyme, 35-60 µM substrate, 10 mM phosphate (pH 7.0), 100 mM NaCl, 6 mM cysteine, and 5% glycerol in the anaerobic cell. Working in the dark, the sample (maintained at 10°C) was illuminated through the 108 mm window for defined time intervals by continuous light from a LQX1800 xenon lamp passing through an optical fiber then filtered with a neutral density filter (2 mm NG11, Schott), a 3 mm colored glass (B390, Hoya), and an interference filter (IF380, Pomfret), yielding a spectrum centered at 384 nm with a bandwidth of 10 nm (FWHM). A mechanical shutter integrated in the lamp was coupled to a DG535 function generator (Stanford Research) to control irradiation times (170 ms for actinometry and up to 300 s for photorepair).

The absorption spectrum along the 10 mm path was recorded after each irradiation period with a Uvikon XS spectrophotometer (Secomam), in which the sample holder was cooled at 10°C, in a spectral range of 240-500 nm or 300-500 nm for DHT containing and normal substrates, respectively.

Photorepair under flash excitation

Samples were excited through the 10×8 mm window of the cuvette by 100 ps flashes of ~4 mJ at 355 nm provided by a Leopard SS-10 Nd:YAG laser (Continuum). The flash energy was measured using an OE25SP-H-MB energy meter (Gentec) placed behind the 108 mm output window of a cuvette identical to the one used for photorepair experiments, but filled with water. For calculating the mean irradiation flux per flash in the sample, corrections were made for reflections at the windows and absorption by the sample.

Continuous monitoring light was provided by a T9F26 light-emitting diode (Seoul Semiconductor) at 265 ± 7 nm (half height of the emission band) or an UVTOP320 light-emitting diode (Roithner Laser Technik) at 325 ± 6 nm. In the former case, the specific wavelength of 265 nm was selected by passing the light through a slightly tilted narrow band (2 nm FWHM) interference filter (LL01-266, Semrock), placed between the light emitting diode and the sample. Detection was made by directing the light onto a solar-blind R6834 photomultiplier tube (Hamamatsu), protected against stray light from the excitation flash by interference filters with peak transmission at 265 and 325 nm, respectively. The output of the photomultiplier was fed to the 11A33 plug-in of a DSA 602A digital oscilloscope (Tektronix) where a 100 nF capacitor was set in parallel to the input of 1 M impedance.

The set-up is shown schematically in Figure 2. The intensity of the 265 nm monitoring light beam at the position of the cuvette was measured using a calibrated fibred spectrometer (HR4000 from Ocean Optics, with a 200 µm core diameter fibre) to be about 60 µW cm -² in the center of the beam. Taking into account reflection losses and the absorption by the sample, we estimate that the mean irradiance along the beam in the sample was about 25 µW cm -2 .

As FADH -absorbs very strongly at 265 nm (ε ≈ 60 mM -1 cm -1 [4]), the actinic effect of our 265 nm monitoring light was not completely negligible. It should have excited ~0.8% of the photolyases present in the beam per second and converted ~0.05% of the enzyme:substrate complexes per second from PL-T(6-4)T to the intermediate PL-X (based on η 1 ≈ 6.7%). After sample insertion into the cell holder, an average delay of ~30 s before the first flash was needed to make the necessary adjustments. After this time, ~1.5% of the enzyme:substrate complex in the beam path would be in the intermediate state PL-X immediately before flash irradiation.

The "repair lag" for the 20-flashes repair experiments ("stairs") in Figure 4a in the main text was obtained as follows. For each of the last 10 flashes of a series, the signal amplitude at the time of arrival of the flash was approximated by averaging, for the purpose of noise reduction, the amplitudes of the preceding 101 data points (i.e. over 0.5 s). For the thus obtained 10 time-amplitude points, a linear regression line was calculated and its zero crossing (intersection with the average level of all data points before the first flash) was determined. The time interval from the first flash to this intersection was named "repair lag". It would be zero for stairs with constant repair per flash from the first flash on. It increases with decreasing repair on the first flashes compared to later flashes (see Figure 4a in the main text). Error bars of the repair lag were determined from the errors of the parameters of the linear regression.

Second harmonic generation

The second harmonic (noted 2ω) of the 775 nm beam is obtained by passing the beam through an optical nonlinear crystal of barium borate (Ba(BO 2 ) 2 , BBO). A beam with a wavelength of 387.5 nm is thus obtained. It was used for our experiments with reduced photolyase (i.e. FADH -).

Non-collinear optical parametric amplifier

For exciting FAD ox , pump wavelengths that are not accessible by second harmonic generation of the 775 nm laser, optical parametric amplification (OPA) was used. We in fact use a non-collinear optical parametric amplifier (NOPA). The NOPA is an OPA with a non-collinear geometry that allows obtaining short pulses. The principle of OPA is to convert a beam (also called a "pump") having frequency ω into two beams of lower frequency (ω 1 and ω 2 called the signal and idler) using non-linear optics. This conversion follows the relation that the sum of the signal and idler frequencies is equal to the frequency of the original beam: ω = ω 1 + ω 2 (the beams at ω 1 and ω 2 are termed as the signal and idler beams).

In the NOPA, the beam called the pump at 387.5 nm is generated by second harmonic generation of a fraction of the 775 nm beam through a BBO crystal. The rest of the beam is used to generate a white light continuum called a seed. The seed beam is the source of the signal beam. The generation of the white light continuum is achieved by passage through a sapphire disk. The pump and seed beams are then overlapped in a second BBO crystal. The result is the amplification of the signal wavelength (475 nm)

and the generation of a residual beam called the idler (infrared). The orientation of the crystal is adjusted in such a way such that phase matching conditions are satisfied.

After passing through the NOPA, it should be noted that the pulse duration of the beam is temporally stretched due to group velocity dispersion from the different optical elements in the NOPA.

The temporally stretched beam is said to be chirped. To reverse the chirp, the beam is recompressed by several passages between two prisms. The pump wavelength of 475 nm used in our experiments with oxidised photolyase (i.e. FAD ox ) has been generated with the NOPA.

We will see in Section 4.2.2.6 that in order to calculate our transient absorption spectra, we need measurements where the pump is either present (on) or absent (off). To create this distinction, we need to measure the intensity of the pump beam. The measurement of this intensity is done with the use of a photodiode. A small portion (~4%) of the beam, coming from second harmonic generation or NOPA, is sent to the photodiode for this function. The pump beam is polarised at the magic angle (54.7°) with respect to the probe for experiments done at the magic angle (i.e. FADH -) and at 45° for polarised experiments (i.e. FAD ox ).

Saturation

It is most desirable to work at conditions where increasing the pump energy would lead to a proportional increase in the population of the excited state we wish to investigate. A large excited population would of course give a large transient absorption signal. However, if we send too much energy to the sample, undesirable effects may take place. One of these is multiphotonic excitation. In particular, we do not want to excite the excited state with a second photon. The excitation of higher excited state could potentially lead to a variety of transient populations and dynamics thereby obscuring the measurements. Another reason is the photodegradation of the sample due to irreversible photoinduced effects caused by excessive pumping. To aid us in choosing the pump energy, we are guided by the saturation fluence (F s ) § . In a simple ballistic model, the saturation is defined as the fluence wherein each absorbing centre gets a photon; all molecules are excited. Above the saturation fluence, some molecules may absorb more than one photon and new excited states are formed as a result. Below this the saturation fluence, the number of molecules getting a photon increases proportionally with the pump energy. The range of pump energies that creates this linear response is referred to as the linear regime. Beyond this region, the deviation from linearity first increases slowly then more rapidly with increasing pump energy.

Hence it is best to work at the linear regime at a pump energy close to the saturation fluence of our molecule of interest in order to maximise the excited state population without the risk of multi-photon excitation.

Probe beam

The absorbance of the pump-excited sample is measured with the use of a beam called the probe.

About 3 mW of the amplified beam not used for pump generation is used to generate the probe. To obtain an absorbance spectrum, we need to have many wavelengths and not just that of the laser. To obtain such polychromatic light, the probe beam is passed through a Kerr medium (in our case CaF 2 ). Self-phase modulation produces a white light continuum that is used to measure the absorbance of the sample over a spectral range of about 350-750 nm. To generate the dynamic image that we want of our excited system, we need to measure the absorption spectrum at different times after the excitation. These times are obtained by introducing a variable delay between the pump and the probe pulses. The pump-probe delay is achieved by sending the probe beam through a variable delay line. The variable delay line increases the trajectory of the probe beam (before arriving at the sample) and hence the time of arrival of the probe to the sample can be manipulated. The probe beam is further split into two identical beams with a beam splitter. One beam is focused on the sample while the other serves as the reference, as in a standard double-beam spectrophotometer.

4.2.2.4

The sample § Fluence refers to the energy per unit area. The saturation fluence is related to the absorption cross-section by . The corresponding saturation intensity, I s, at wavelength λ is given by , where h is Planck's constant and c is the speed of light.

) 0 (

To measure polarised signals, the polarisation of the monitoring light is adjusted (parallel, perpendicular, or at magic angle with respect to the pump) before the sample. The adjustment of the polarisation is achieved with the use of a polariser. This is seen in Figure 3 as "p". situation, where the pump plays the role of excitation beam for the Raman effect and the continuum probe plays the role of stimulation beam (hence is amplified). These bands have a vibrational frequency of about 3400 cm -1 corresponding to the O-H stretching of the water molecules of the solvent. This artefact is taken into account during data treatment using an empirical function which is the sum of a Gaussian and its first few time derivatives as has been described by Kovalenko et al. [5]. In our case, we use the sum of up to the fourth time derivative as we have observed that this described this effect better. This function M is presented as follows:

3z artefact

We tested our 3z set-up on a sample of coumarin 6. This molecule has a short rotation correlation time of 69.4 ps in methanol at 25°C [6]. We verified this using our set-up and while we were able to measure essentially 0 for the anisotropy using our 2z set-up, we found non-zero values using our 3z setup. We see in Figure 7 that while the global forms of the anisotropy spectra are similar, the 3z anisotropy spectrum appears to be more negative with respect to the 2z set up as shown on Figure 11. Furthermore, it appears to be more negative in some regions than others. We particularly see this in the region between 500 and 650 nm. When we measured the anisotropies of other dyes, we observed the same effect as with coumarin 6 where the supposed-to-be-zero anisotropy was shifted negatively by approximately the same A net positive contribution of species i is observed when ε u,i > ε a + ε e,i . In this case, the transient absorption is said to be dominant. A negative contribution is expected when ε u,i < ε a + ε e,i . In this case, either of two contributions is dominant: i) when ε a < ε e,i , one observes a stimulated emission or gain; ii)

When ε a > ε e , bleaching is observed. This three level model is illustrated in Figure 14.

Kinetic analysis

A kinetic analysis of the transient absorption data is necessary to be able to quantify the dynamics of the photoinduced phenomena. After measuring the transient absorption spectra (ΔA(λ,t)) at different pump-probe delays, we obtain data in the form of a matrix that we will call A. A has been corrected of the artefacts described previously and is of the dimension p  q. The p rows of A are the kinetics (typically tens of time delays) and the q columns are the spectra (typically hundreds of wavelengths).

The kinetics are parametrically fitted to design a kinetic model of the observed reactions. The kinetics are often fitted with a sum of exponentials with decreasing lifetime τ i (e -t/τi ). The choice of this solution is due to the fact that a sum of exponentials is the most general solution for a system of coupled unimolecular reactions of the first order describing the evolution of the concentrations of the different At a given wavelength, the fit of the kinetic parameters is performed with the following function:

where the fitting parameters are the pre-exponential factors a i , the lifetimes τ i , the internal parameters of M(b i ) (Equation 13), and dt.

The global analysis of matrix A is aided by singular value decomposition (SVD) of the matrix [10]. SVD greatly reduces the number of kinetics that are to be simultaneously fitted. The decomposition is a generalisation of the diagonalisation of square matrices. This function is readily available in the computational software Mathematica (Wolfram Research) that we use. The matrix A (of dimension p  q) is decomposed in the following way according to Equation 18:

U is a matrix (dimension p  p) of the normalised "orthogonal spectra". S is a pseudo-diagonal matrix (dimension p  q) containing the singular values (in decreasing order) on its diagonal. V is a matrix (dimension q  q) of the normalised "orthogonal kinetics". The orthogonal kinetics are the rows of the transpose of V, t V.

Having decomposed A into matrices U, S, and V, the truncation of these three matrices becomes possible. This is done by taking only the m largest singular values. The number m is chosen in such a way that the part of the matrix ignored due to the truncation does not contain any significant signal or is essentially noise. Thus the matrix U is reduced to a matrix of dimension p  m (m orthogonal spectra), S is reduced to a diagonal matrix of dimension m, and V is reduced to a matrix of dimension q  m. We will note these reduced matrices with primed symbols as shown in Equation 19:

where A' is related to A by the following A = A' + noise (19) We then perform a global analysis of the m orthogonal kinetics weighted by their singular values, meaning S'  t V', by a sum of N exponentials. By doing this, the different lifetimes and their corresponding pre-exponential factors are obtained. This is written as:

where F is a matrix of the pre-exponential factors of dimension m  N and T is a matrix of the N exponentials of the q time delays in the experiment with dimension N  q. The global fit cannot exactly reproduce A'. What we obtain is the matrix A" defined as:

where A' = A" + noise. F U B   ' . B (dimension p  N) is the matrix of the pre-exponential factors associated to the N exponentials taken at p wavelengths of the experiment. The spectra obtained, the columns of the matrix B, are called the decay associated difference spectra (DADS). A" should also be an approximation of A. We verify at this point that the matrix obtained by A-A" still essentially contains noise.

The advantage of global analysis by SVD is that all spectral information are summarised to a reduced number of orthogonal kinetics which are easily fitted. Alternatively, a direct global fit of A is experiment becomes the matrix T with dimensions N  q (see Equation 20). The concentration vectors, K(t), measured at q delays in the experiment form the matrix κ related to T as follows:

The matrix of the model-predicted values A" can be rewritten as follows: The parameters of the model MC must be fitted numerically to obtain the N experimentally measured lifetimes, τ i , and to satisfy the eventual constraints on the SADS. In the case where the kinetic model is a simple sequential cascade where each state decays to the next with a yield of 1, the SADS are called the evolution associated difference spectra (EADS).

Sample preparation 4.5.1 Xenopus laevis (6-4) photolyase

Xenopus laevis photolyase was prepared according to the following procedure [11]. A construct containing the cDNA of Xenopus laevis photolyase inserted in a pGEX 4T-2 vector at the BamHI/EcoRI site was kindly gifted from Dr. Takeshi Todo (Osaka University, Japan). The construct was transfected into MAX Effiency® DH10B™ E. coli competent cells (Invitrogen). The transfected cells were cultured in LB medium with 100 µg mL -1 of ampicillin at 25°C, and isopropyl-β-D-1thiogalactopyranoside was then added at the final concentration of 0.4 mM. After 12 hours, cells were harvested and disrupted by sonication on ice. The supernatant was loaded onto a column containing Glutathione-sepharose 4B resin (GE Healthcare UK Ltd., Buckinghamshire, UK) in order to trap the GST-tagged enzyme, and elution was performed according to the manufacture instruction. After the buffer-exchange into PBS, 100 µL of thrombin (100 units µL -1 ) was added to cleave the GST tag; the mixture was incubated at 20°C for 12 hours. Then the solution was passed through the Glutathionesepharose 4B column again to remove the remaining GST-tagged enzyme. The flow-through fractions were further purified by an Äkta purifier (GE Healthcare UK Ltd.) equipped with a HiTrap Heparin HP column (GE Healthcare UK Ltd.) to obtain tag-free photolyase in an oxidized form. The purification buffer was finally replaced with a buffer containing 50 mM Tris-HCl, 50 mM NaCl, and 5% glycerol (pH 8.0) by repeated concentration/dilution processes.

Reduced photolyase

Ultrafast experiments

50 µL of oxidised Xenopus laevis (6-4) photolyase with a concentration of about 100 µM was used in the femtosecond pump-probe experiments. Prior to the experiments, the protein solution was passed through a gel-filtration column (Micro Bio-Spin6, BioRad) to eliminate any unbound FAD and to exchange the buffer solution. For experiments on reduced photolyase the protein was placed in a 10 mM phosphate buffer at pH 7 containing 100 mM NaCl, 6 mM cysteine and 25% glycerol. The protein was placed in a custom 60 µL anaerobic cuvette with an optical path length of 0.1 cm (117.194B-QS, Hellma Analytics). After sealing the cuvette with the use of rubber septa, the inside was rid of oxygen by repeated cycles of vacuum and argon. This operation is done via a needle inserted through the rubber septum. After purging, the rubber septa were replaced inside a glovebox (<3 ppm O 2 , Campus by Jacomex) to avoid any seepage of oxygen that would re-oxidise FADH -. The cuvette containing the protein was then illuminated with continuous-wave light: ~430-800 nm. This was obtained by filtering the light emitted by a xenon lamp (LQX1800, Linos) with a 2-mm GG435 colour filter (Schott). The sample was illuminated until its steady-state spectrum is converted to that of fully reduced photolyase. The sample was kept on ice at all times.

For ultrafast experiments involving the substrate, the normal T(6-4)T substrate (8-mer) was added after photoreduction. The ratio of the substrate to protein concentrations was 3:1. The 60 µL cuvettes used for pump-probe spectroscopy are too narrow to allow for homogenisation of the sample by shaking after substrate addition. The sample was prepared in the larger 1 cm 250 µL anaerobic cuvettes used for steady-state photorepair (16.160-F/4/Q/10 GL 14/S/Z15, Starna) and then transferred to the 0.1 cm 60 µL cuvette used for pump-probe spectroscopy after substrate addition. The protein was maintained at 5°C at all times in both cases.

Steady-state photorepair experiments

The steady-state photorepair sample (250 µL), consisting of 10-60 M substrate, 1-9 M reduced enzyme, 10 mM phosphate buffer (pH 7), 100 mM NaCl, 6 mM cysteine, and 5% glycerol, was prepared as follows. An aliquot of stock solution of the enzyme was diluted with a buffer containing 10 mM phosphate, 100 mM NaCl, and 5% glycerol (pH 7), and the mixture was passed through a gel-filtration column (Micro Bio-Spin 6, BioRad), equilibrated with the buffer, in order to remove unbound FAD. The eluate (60 µL) was transferred to an anaerobic 1028 mm (lengthwidth height) inner volume quartz cuvette with self-masking solid black walls and four clear windows (Starna, 16.160-F/4/Q/10 GL 14/S/Z15), and the solution was purged with argon through a septum. The cuvette was then put into a glove box in which the oxygen level was kept below a few ppm, and the sample without the substrate (typically 240 µL) was prepared in the glove box by addition of cysteine dissolved in the buffer. Prior to irradiation by light, an absorption spectrum of the solution was measured to determine the enzyme concentration from the absorption band of oxidized FAD (molar decadic absorption coefficient ε = 11.2 mM -1 cm -1 at 450 nm [12]). The sample was illuminated through the 10×8 mm window with continuous light (~430-800 nm) from a LQX1800 xenon lamp (Linos), through an optical fiber and colored glass filters (2 mm GG435 and 2 mm KG3 from Schott), until the absorption spectrum was completely converted into that of fully-reduced FADH -. Then, the cuvette was put into the glove box again, and aliquots of the substrate were added into the solution containing the reduced FADH -to prepare the sample for measurements. The sample was maintained at a temperature of 4°C at all times. to the experiments, the protein solution was passed through a gel-filtration column (Micro Bio-Spin6, BioRad), equilibrated with a buffer, to eliminate any unbound FAD. The protein was put in a 0.1 M TRIS 57% identity with X. laevis photolyase. We ultimately chose to stay with the (6-4) photolyase from D. melanogaster as both are animal photolyases and hence are more likely related.

The next step is to create the model. This is done in a program called SWISS-PDB viewer. This is the companion program of the online SWISS-MODEL platform. To generate the model, the target sequence is threaded (i.e. wound) around the structure of the template. The primary sequence of the target is "physically" traced on the template structure. This was done by aligning the protein sequences using the alignment previously done in the preceding step as a guide. The alignment may also be optimised in

Conclusion

animal cryptochromes that were sampled in our structure analysis. This fact might suggest the involvement of the residue in a mechanism that is unique to animal CPF proteins. The position of W D in the protein makes it an ideal candidate as the final electron donor. A study of the photoactivation at longer timescales should also be undertaken. Our initial measurements of the anisotropy in the nanosecond regime showed a similar value of anisotropie that was found in the femtosecond experiments. In order to confirm the involvement of W D , the phenylalanine mutant of this residue must be studied. Furthermore at long timescales, what would be of great interest would be the observation of FAD •-protonation. The proton donor to the flavin has also not yet been identified in the photolyase. The possibility of residues other than tryptophan is also an interesting subject to investigate. The hypothetical involvement of a tyrosine as the final electron donor of Xl64 photoctivation has up to this point not yet been followedup in literature. The challenge in the study of the tyrosine is the limited wavelength range over which it can be observed as can be seen in the absorption spectrum of the tyrosyl radical.

On the subject of the photorepair of Xl64 photolyase, the main question that we sought to answer is the number of photons required for repair of the (6-4) photoproduct. Save for a theoretical study, literature on the photorepair by (6-4) photolyase is at a consensus that the mechanism involves a single photon. Using series of single turnover flashes, we have found that the repair reaction of Xl64 proceeds by a successive two-photon mechanism. The first photon converts the (6-4) lesion into a metastable intermediate that we call X. Absorption of a second photon within the lifetime of X gives rise to the restoration of the intact TT bases. We estimated the quantum yields of the first and second photoreactions to be 6.7 % and 83 % respectively.

In light of our results, a new pathway for future studies the repair mechanism of the photoproduct has been opened. The first of these studies is the separate characterisation of the two photoreactions. To unravel the mechanism, the reaction induced by each of the photons must be given.

The separate study of the photoreactions is, however, challenging. We have performed preliminary ultrafast experiments of the photorepair reaction where we observed the dynamics of the flavin chromophore in the protein during repair. In our experiments, we were able to deduce that what we measured was essentially the first photoreaction. We were able to observe that the electron transfer from the excited FADH -cofactor occurs in 247 ps. In our time window of 3 ns, the reaction is not yet finished.

The expected regeneration of FADH -was not observed. There is hence a great deal of interest in following the reaction to longer timescales. For future experiments, it can be conceivable to further minimise the contribution of the second photoreaction. Our broadband studies contribute to the sole existing ultrafast study of the (6-4) photoproduct repair. In our experiments, we were unable to see the spectral region <350 nma region of large interest as this is where intact base restoration can be directly observed. We are currently improving on our transient absorption set-up to surpass this limitation.

Consequential to the two-photon repair mechanism is the existence of the repair intermediate, X.

In our characterisation of the intermediate, we have estimated its to be ~2 min. We have shown that X decays back to the original T(6-4)T lesion in the absence of a second photon. The question at the moment would be the identity of X. Our data is not in contradiction to X being the oxetane-bridged pyrimidine dimer as in the theoretical two-photon model. However, neither does it give proof that X is indeed the oxetane. Hence of great interest is the identification of X. In the spectral identification of the intermediate, the advantage of our technique of ultrafast broadband spectroscopy becomes evident. The characterisation of the reaction intermediate would be more complete with transient absorption spectra.

Another issue of interest is the characterisation of the repair of the T(6-4)C photoproduct. We have performed photorepair assays on the T(6-4)C photoproduct. We qualitatively observed that the T(6-4)C is repaired similarly as the T(6-4)T based on our repair curves. Our observation indicates that the lesion is repaired by the (6-4) photolyase. It does not give information on how the lesion is repaired. It is of great interest to know if the T(6-4)C lesion is repaired with a similar mechanism as the T(6-4)T. If it is, the elementary steps will also need characterisation as it might not be the same the T(6-4)T. If not, then it would further open a new field of investigation as it will also put into question the repair mechanisms of the other forms of the (6-4) photoproduct.

From the results obtained, it can be said in closing that the main objectives of this thesis have been attained and new insights have been provided both in the case of photoactivation and photorepair of photolyase.

Annexes

Annex A : Phylogenetic analysis of the proteins mentioned in Chapter 2

The colour coding in the figure represents the different protein subfamilies. Starting from a sequence alignment of selected proteins (using COBALT, NCBI), a phylogenetic tree was constructed using the Phylogeny tool of ClustalW2 (EMBL-EBI).

Annex B : Steady-state absorption spectum of FAD ox in solution

The steady state absorption spectrum of 200 µM FAD ox in a 100 mM Tris buffered at pH 8 containing 100 mM NaCl, and 5% glycerol is shown below.