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

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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

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.
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 [32-35]. There is a limited degree of homology between plant/animal photolyases and microbial 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]), (6-4) 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 [45]), Cry2 from Mus musculus (416E [46]), Cry3 from Arabidopsis thaliana (2J4D [47]), CryDASH from Synechocystis sp. (1NP7 [48]) and Arabidopsis thaliana (2IJG [49]). We will start describing the well-known structure of EcCPD in some details. It is represented in Figure 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].
EcCPD is a 471-residue long globular protein. EcCPD contains two distinct regions: the N-terminal 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.
Figure 7. Crystal structure of Dm64 photolyase bound to DNA containing a (6-4)PP lesion. The FAD cofactor is shown in yellow.

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 [50, 51]. 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 enzyme-substrate 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 F0), 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 [52]. The MTHF chromophore also dissociates readily from photolyase and is present in substoichiometric amounts in purified samples [53-55]. In purified EcCPD samples, MTHF is present only in 20-30 % of the total photolyase present [55]. However, MTHF can be reconstituted by incubating the apoenzyme in solutions with high MTHF concentrations [56]. The antenna chromophore in (6-4) photolyase has not yet been found in purified samples although MTHF was observed in a semi-purified sample [57]. A binding pocket that strongly binds F0 has been identified in D. melanogaster (6-4) photolyase [58].
In 2013, the crystal structure of the (6-4) 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 [59]. 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 [60]. 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.[61]. Though the existence of the (6-4) photoproduct was known before the discovery of the (6-4) photolyase, the only substrate known to photorepaired was the CPD [61]. As far as DNA repair is concerned the (6-4) 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 (6-4) photoproduct form thus promoting repair [62, 63].
The (6-4) photolyase has been believed to only exist in eukaryotes until the identification of the A. tumefaciens (6-4) photolyase was reported by Zhang et al. in 2013 [64]. The most recent phylogenetic analyses of the CPF proteins have not yet revealed subfamilies among the (6-4) photolyases [24, 52, 65].

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 [66]. We have seen in Figure 5 that cryptochromes are classified into several subfamilies: plant cryptochromes, animal cryptochromes, DASH cryptochromes, and CryPro [23, 65]. 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. [52], Oliveri et al. [65] 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 [67]. More specifically, it inhibits hipocotyl elongation [15]. AtCry1also acts as a regulator of gene expression during plant photomorphogenesis [68, 69].

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 [70, 71]. 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 [72]. 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 [73].
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 [74, 75].

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) [48, 76]. 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 [48]. Like photolyases, the Synechocystis CryDASH has also been reported to bind to DNA and possesses a weak DNA photorepair activity [29, 76]. 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 [77, 78]. 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 [79].

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 (6-4) 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.[64].

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 (KS) 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 (KNS) is ~10-4 M. This translates to a selectivity factor (KNS/KS) of ~105 [80]. The rate of substrate binding (kon) has been reported to be in the range of 107 M-1 s-1 for CPD photolyase [36] while the dissociation constant (koff) 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 light-dependent. 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 (FADox 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 5109 s via dipole-dipole coupling (i.e. FRET). The now excited FADH– transfers an electron to the CPD lesion at a rate of 7 109 to 2 1010 s-1 over 5-10 Å [81-83]. 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% [84]. The mechanism is illustrated in Figure 9.
Figure 9. Classical overall repair mechanism of the CPD lesion by CPD photolyase. Figure reproduced from Sancar [36].
In the same year, a real-time transient absorption spectroscopy study of the splitting of the CPD dimer has been reported by MacFarlane and Stanley [85]. The authors reported that the time constant of electron transfer from the flavin to the substrate is 32 ± 20 ps. The asynchronous splitting of the two C–C bonds was claimed to be initiated in ~60 ps and is completed by 1500 ps. Furthermore, the authors did not observe the regeneration of FADH– after full bond splitting in their experiments.
Figure 10. Detailed mechanism of CPD photorepair by CPD photolyase according to MacFarlane et al. [85].
In 2011, Thiagarajan et al. reported a study of the kinetics of CPD dimer splitting [86]. 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% [87]. 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.
Figure 11. Detailed mechanism of CPD photorepair by CPD photolyase according to Thiagarajan et al. [88].
Shortly after the publication of the report by Thiagarajan et al., Liu et al. reported another study on the CPD repair [89]. 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 [90, 91].

Table of contents :

General Introduction
1. Literary Review
Introduction
1.1 UV-Induced DNA Lesions
1.1.1 Cyclobutane pyrimidine dimer, CPD
1.1.2 Pyrimidine (6-4) pyrimidone photoproduct, (6-4) photoproduct
1.2 Photolyases and Cryptochromes
1.2.1 Structure of the cryptochrome and photolyase family
1.2.1.1 Phylogenetics
1.2.1.2 Primary and secondary structure
1.2.1.3 Chromophores
1.2.2 Photolyases
1.2.2.1 CPD photolyase
1.2.2.2 (6-4) photolyase
1.2.3 Cryptochromes
1.2.3.1 Plant cryptochromes
1.2.3.2 Animal cryptochromes
1.2.3.3 DASH cryptochromes
1.2.3.5 Dual-function cryptochromes
1.2.3.4 CryPro cryptochromes
1.3 The reaction mechanisms of photolyase
1.3.1 Photorepair
1.3.1.1 Photorepair by CPD photolyase
1.3.1.2 Photorepair by (6-4) photolyase
1.3.1.3 Photorepair of the T(6-4)C photoproduct
1.3.2 Photoactivation
1.3.2.1 Photoactivation of CPD photolyase
1.3.2.2 Photoactivation of (6-4) photolyase
1.3.2.3 Cryptochrome photoreduction
1.4 The flavin cofactor of the (6-4) photolyase
1.4.1 Flavins in solution
1.4.2 Flavins in photolyase
Conclusion
References
2. Photoactivation
Introduction
3.1 Structural analysis of the X. laevis (6-4) photolyase
3.1.1 Sequence alignment
3.1.2 Homology model
3.2 Spectroscopic study of the wild type X. laevis (6-4) photolyase
3.2.1 Steady-state spectra
3.2.2 Isotropic femtosecond transient absorption spectroscopy
3.2.2.1 Initial transient absorption spectrum
3.2.2.2 Spectral dynamics of the isotropic transient absorption
3.2.2.3 Global kinetic analysis of the isotropic transient absorption spectra
3.2.2.4 Nature of the photoproducts
3.2.2.5 Elementary kinetic model
3.2.3 Polarised femtosecond transient absorption spectroscopy
3.2.4 Femtosecond transient absorption anisotropy
3.2.4.1 Sensing the orientation of the WH●+ radical
3.2.4.2 Raw transient anisotropy
3.2.4.3 Global kinetic analysis of the raw anisotropy spectra
3.2.4.4 Species-associated anisotropy spectra
3.2.4.5 Errors on the anisotropy
3.2.4.6 Expected anisotropies based on the homology model
3.2.4.7 Comparison of experiment with simulation
3.2.4.8 “Millisecond constant”
3.2.5 Nanosecond transient absorption spectroscopy
3.2.5.1 Polarised transient absorption decays
3.2.5.2 Anisotropy in the ns timescale
3.2.5.3 Isotropic transient absorption in the ns-μs timescale
Conclusion
References
3. Photorepair
Introduction
3.1 Photorepair of the T(6-4)T photoproduct by (6-4) photolyase
3.1.1 Absorption spectra
3.1.1.1 T(6-4)T photoproduct
3.1.1.2 Fully reduced X. laevis (6-4) photolyase
3.1.2 Photorepair under steady-state irradiation
3.1.2.1 Observation of T(6-4)T and TT restoration
3.1.2.2 Photorepair of the normal substrate
3.1.2.3 DMAD actinometry
3.1.3 Photorepair under single-flash excitation
3.1.3.1 Single-flash excitation
3.1.3.2 “Stairs” experiments
3.1.4 Characterisation of X
3.1.4.1 The observation of X
3.1.4.2 The decay of X
3.1.5 Quantitative analysis of the proposed reaction scheme
3.1.5.1 Analysis of the suggested repair reaction scheme
3.1.5.1.1 General assumptions
3.1.5.1.2 Continuous light excitation
3.1.5.1.3 Single flash excitation
3.1.5.1.4 Flash sequence (“stairs”)
3.1.5.2 Estimation of the repair quantum yields η1 and η2
3.1.5.2.1 Equations for the absorption changes
3.1.5.2.2 Determination of η1 and η2 from flash (first vs. successive) excitation
3.1.5.2.3 Determination of η1 and η2 from flash sequence (“stairs”)
3.1.5.3 Quantum yield uncertainty estimation
3.1.5.3.1 From continuous light excitation
3.1.5.3.2 From flash (first vs. successive) excitation
3.1.5.3.3 From flash sequence (“stairs”)
3.1.6 Photorepair under solar irradiation
3.2 Preliminary study of the steady-state photorepair of the T(6-4)C photoproduct
3.2.1 The T(6-4)C photoproduct
3.2.2 Steady state photorepair of the T(6-4)C photoproduct by X. laevis (6-4) photolyase
3.3 Preliminary studies of T(6-4)T photorepair by femtosecond pump-probe spectroscopy
3.3.1 Reference spectra
3.3.1.1 Steady-state fluorescence spectra
3.3.1.2 Transient absorption spectra of FADH– in solution
3.3.2 Transient absorption spectroscopy of fully reduced Xl64, in the absence of substrate
3.3.2.1 Initial transient absorption spectrum
3.3.2.2 Spectral dynamics of the transient absorption spectrum
3.3.2.3 Global kinetic analysis
3.3.3 Transient absorption spectroscopy of fully reduced Xl64 in the presence of T(6-4)T
substrate
3.3.3.1 Spectral dynamics of the transient absorption spectrum
3.3.3.2 Global kinetic analysis
3.3.3.3 Interpretation
Conclusion
References
4. Experimental
Introduction
4.1 Photorepair
4.1.1 Substrate
4.1.2 Quantum yield determination
4.1.3 Photorepair under steady-state excitation
4.1.4 Photorepair under flash excitation
4.2 Spectroscopic methods
4.2.1 Steady-state spectroscopy
4.2.2 Femtosecond transient absorption spectroscopy
4.2.2.1 Femtosecond laser source
4.2.2.1.1 Ti:Sapphire laser
4.2.2.1.2 Regenerative amplifier
4.2.2.2 Pump beam
4.2.2.2.1 Second harmonic generation
4.2.2.2.2 Non-collinear optical parametric amplifier
4.2.2.2.3 Saturation
4.2.2.3 Probe beam
4.2.2.4 The sample
4.2.2.5 Data acquisition
4.2.2.6 Spectral acquisition
4.2.3 Nanosecond real-time transient absorption spectroscopy
4.2.3.1 Laser sources
4.2.3.1.1 Excitation pulse
4.2.3.1.2 Monitoring light
4.2.3.2 The sample
4.2.3.3 Detection system and data acquisition
4.3 Artefacts
4.3.1 Pump energy fluctuation
4.3.2 White light chirp and t0
4.3.3 Cross-phase modulation and stimulated Raman scattering
4.3.4 3z artefact
4.3.5 “Millisecond constant”
4.3.6 Free flavin
4.4 Data analysis
4.4.1 Spectral analysis of transient absorption signals
4.4.2 Kinetic analysis
4.5 Sample preparation
4.5.1 Xenopus laevis (6-4) photolyase
4.5.2 Reduced photolyase
4.5.2.1 Ultrafast experiments
4.5.2.2 Steady-state photorepair experiments
4.5.3 Oxidised photolyase
4.5.3.1 Femtosecond pump-probe spectroscopy
4.5.3.1 Nanosecond real-time transient absorption spectroscopy
4.6 Homology Modelling
References
Conclusion
Annexes

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