Ultrafast Dynamics of Flavin Cofactor in
Photolyase/Cryptochrome Family
Dissertation
Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University
By
Chuang Tan, M.S.
Graduate Program in Chemical Physics
The Ohio State University
2013
Dissertation Committee:
Professor Dongping Zhong, Advisor
Professor Heather C. Allen
Professor Chenglong Li Copyright by
Chuang Tan
2013
ABSTRACT
Due to the essential role of flavoproteins in light-driven biological activities, such as photoinduced DNA repair and signal transduction, it is of great interest to study the photochemistry and photophysics of flavin cofactor. This dissertation presents a systematic investigation on the ultrafast dynamics of flavin cofactor in four redox states in photolyase/cryptochrome family proteins. In photolyase, the antenna molecule MTHF absorbs a photon and transfer the excitation energy to the catalytic cofactor FADHˉ to enhance the DNA-repair efficiency. With the femtosecond-resolved fluorescence and transient absorption spectroscopy, we examined ultrafast dynamics of the resonance energy transfer from MTHF to three flavin redox states in photolyase and accurately determine the rates. The energy transfer from MTHF to the fully reduced hydroquinone FADHˉ occurs in 170 ps, but it takes 20 and 18 ps to the oxidized FAD and neutral semiquinone FADH, respectively. The orientation factors were estimated from the acquired energy transfer rates and compared with values from the theoretical study. Both results demonstrate that MTHF-FADHˉ has the largest κ2 value, indicating the best orientation alignment for the physiologically functional state.
Photolyase uses blue light to repair the UV-induced pyrimidine dimer through a
ii
cylic electron transfer radical mechanism. The significant loss of repair efficiency by the mutation of the active-site residues indicates that those residues play a critical role in the DNA repair by photolyase. To understand how the protein active site modulates the repair and achieve the high efficiency, we mapped out the complete evolution of functional dynamics with 6 active-site mutated photolyases in real time, and analyzed the individual electron transfer processes in the catalytic reaction with Sumi-Marcus model. The results suggest that photolyase controls the critical electron transfer and the ring-splitting of pyrimidine dimer through modulation of the redox potentials and reorganization energies, and stabilization of the anionic intermediates, maintaining the dedicated balance of all the reaction steps and achieving the maximum function activity.
The flavin cofactor in photolyase is converted to oxidized FAD or neutral semiquinone FADH and loses the enzyme activity. Upon light excitation, both FAD and FADH can be photoreduced to catalytic form FADHˉ via electron transfer mainly through the neighboring conserved tryptophan triad. The ultrafast photoreduction dynamics of flavin cofactors in (6-4) photolyase was revealed by the femtosecond-resolved laser spectroscopy. Significantly, we found that the tryptophan triad has a reduction potential gradient modulated by the local protein environment, which promotes the electron hopping process with a distinctive directionality and enhances the photoreduction efficiency.
In Arabidopsis thaliana cryptochrome 2, we observed the ultrafast photoreduction
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of oxidized FAD in a few picoseconds and of neutral radical semiquinone FADH in tens of picoseconds through intraprotein electron transfer. Such ultrafast dynamics exclude their potential roles as the functional states. In contrast, we found that the anionic hydroquinone FADHˉ and semiquinone FAD− have complex deactivation dynamics on the time scale from a few picoseconds to a few nanoseconds, which is believed to occur through conical intersections with a flexible bending motion of the isoalloxazine ring. These results imply that the anionic hydroquinone FADHˉ is more likely to be the functional state rather than the two neutral states.
iv
Dedicated to my family.
v
ACKNOWLEDGMENTS
I would like to express my sincere gratitude to my advisor, Dr. Dongping Zhong, for his intellectual guidance, professional support, and unique insights. It is my great honor to study under his direction. His enthusiasm, intellect and diligence have excited me to explore the field of biophysics. I have learned many virtues such as honesty, strictness and persistence which are essential for doing good research. His guidance and insight have been instrumental throughout my graduate education.
I am grateful to Dr. Heather C. Allen and Dr. Chenglong Li for serving on my dissertation examination committee and all the help and assistance they have given to me over the years. I would appreciate very much their patience and kindness.
I would also like to thank the former and present lab members for their friendship and support. In particular, I am thankful to Lijuan Wang, Dr. Ya-Ting Kao, Dr. Jiang
Li, Dr. Xunmin Guo and Zheyun Liu for their help and insight in performing experiments. My special thanks are due to Dr. Suxia Yan for teaching me the laser skills and the kindly help in the daily life. My thanks also need to go to my old friends,
Dr. Weihong Qiu and Dr. Yi Yang, for the job search.
I am immensely thankful to my parents and brother. Their love and expectation encourage me to continue with rolling my dream to be a scientist.
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I sincerely thank my wife Xuetao Huang for her long love and care.
I would also like to thank Department of Chemistry, the Ohio State University and National Institute of Health for funding.
vii
VITA
November 1981…………………...Born in Guangxi, China
2004……………………………….B. S. Chemical Physics, University of Science and
Technology of China, China
2008……………………………….M.S. Chemical Physics, The Ohio State University
2008 to present…………………Graduate Research Associate, Department of
Chemistry, The Ohio State University
PUBLICATIONS
7. Liu Z, Guo X, Tan C, Li J, Kao Y-T, Wang L, Sancar A, Zhong D (2012) Electron tunneling pathways and role of adenine in repair of cyclobutane pyrimidine dimer by DNA photolyase. J. Am. Chem. Soc. 134: 8104-8114. 6. Li X, Wang Q, Yu X, Liu H, Yang H, Zhao C, Liu X, Tan C, Klejnot J, Zhong D, Lin C (2011) Arabidopsis cryptochrome 2 (CRY2) functions by the photoactivation mechanism distinct from the tryptophan (trp) triad-dependent photoreduction. Proc. Natl. Acad. Sci. U.S.A. 108, 20844-20849. 5. Liu Z, Tan C, Guo X, Kao Y-T, Li J, Wang L, Sancar A, Zhong, D (2011). Dynamics and mechanism of cyclobutane pyrimidine dimer repair by DNA photolyase. Proc. Natl. Acad. Sci. USA 108: 14831-14836. 4. Li J, Liu Z, Tan C, Guo X, Wang L, Sancar A, Zhong D (2010). Dynamics and mechanism of repair of ultraviolet-induced (6-4) photoproduct by photolyase. Nature 466: 887-890.
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3. Chang C-W, Guo L, Kao Y-T, Li J, Tan C, Li T, Saxena C, Liu Z, Wang L, Sancar A, Zhong D (2010) Ultrafast solvation dynamics at binding and active sites of photolyases. Proc. Natl. Acad. Sci. USA 107: 2914 -2919. 2. Öztürk N, Selby C, Song S-H, Ye R, Tan C, Kao Y-T, Zhong D, Sancar A (2009) Comparative photochemistry of animal type 1 and type 4 cryptochromes. Biochemistry 48: 8585-8593. 1. Kao Y-T, Tan C, Song S-H, Öztürk N, Li J, Wang L, Sancar A, Zhong D (2008) Ultrafast dynamics and anionic active states of the flavin cofactor in cryptochrome and photolyase. J. Am. Chem. Soc. 130: 7695-7701.
FIELDS OF STUDY
Major Field: Chemical Physics
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TABLE OF CONTENTS
ABSTRACT …………………………………………………………………………...ii ACKNOWLEDGMENTS ...... vi VITA………………………………………………………………………………...viii TABLE OF CONTENTS ...... x LIST OF TABLES ...... xiii LIST OF FIGURES ...... xiv CHAPTER 1 INTRODUCTION ...... 1 CHAPTER 2 ULTRAFAST DYNAMICS OF RESONANCE ENERGY TRANSFER WITH THREE FLAVIN REDOX STATES IN PHOTOLYASE...... 18 2.1 Introduction ...... 19 2.2 Materials and Methods ...... 20 2.2.1 Preparation of CPD Photolyase and Mutants...... 20 2.2.2 Fluorescence Quantum Yield of MTHF...... 21 2.2.3 Femtosecond-resolved Spectroscopy...... 21 2.2.4 Theoretical study...... 22 2.3 Results and Discussion ...... 22 2.3.1 Quenching dynamics of excited MTHF...... 22 2.3.2 Capture of the FADH¯* intermediate...... 23 2.3.3 FRET from MTHF to FAD and the subsequent reduction...... 24 2.3.4 FRET from MTHF to FADH and the subsequent reduction...... 25 2.3.5 Calculation of the orientation factor...... 26 2.3.6 Theoretical study of the orientation factors...... 28 2.4 Conclusion ...... 28 CHAPTER 3 MOLECULAR UNDERSTANDING OF EFFICIENT DNA REPAIR MACHINERY OF PHOTOLYASE ...... 41 3.1 Introduction ...... 42 3.2 Materials and Methods ...... 44 3.2.1 Preparation of CPD Photolyase and Mutants...... 44 3.2.2 Preparation of CPD Substrate...... 45 3.2.3 Enzyme Activities...... 45 3.2.4 Femtosecond-resolved Spectroscopy...... 46
x
3.2.5 Electron Transfer Fitting Model...... 46 3.3 Results and Discussion ...... 49 3.3.1 Dissociation Constants and Relative Quantum Yields...... 49 3.3.2 Reaction Times of Each Elementary Step...... 49 3.3.3 Analysis of Forward ET and Electron Return by Sumi-Marcus Model...... 51 3.3.4 Modulation of Driving Force and Solvent Reorganization by Active-site Residues...... 57 3.3.5 Ring Splitting and Back Electron Transfer...... 59 3.3.6 Maximization of The Quantum Yield...... 60 3.4 Conclusions ...... 61 CHAPTER 4 ULTRAFAST PHOTOREDUCTION OF FLAVIN COFACTORS IN (6-4) PHOTOLYASE ...... 73 4.1 Introduction ...... 74 4.2 Materials and Methods ...... 76 4.2.1 Preparation of (6-4) Photolyase and Mutants...... 76 4.2.2 Femtosecond-resolved Spectroscopy...... 77 4.3 Results and Discussion ...... 77 4.3.1 Direct electron transfer between adenine and flavin moiety in oxidized FAD...... 77 4.3.2 Electron flow from the nearby tryptophan...... 80 4.3.3 Electron hopping among the conserved tryptophan triad...... 81 4.3.4 Ultrafast photoreduction of FADH...... 83 4.3.5 Analysis of the ET networking by Marcus model...... 84 4.4 Conclusions ...... 87 CHAPTER 5 ULTRAFAST DYNAMICS OF FLAVIN COFACTOR IN BLUE-LIGHT RECEPTOR CRYPTOCHROME ...... 101 5.1 Introduction ...... 102 5.2 Materials and Methods ...... 103 5.2.1 Preparation of AtCRY2 and Mutants...... 103 5.2.2 Photoreduction of AtCRY2 and Trp-triad mutants...... 104 5.2.3 Femtosecond-resolved Spectroscopy...... 104 5.3 Results and Discussion...... 104 5.3.1 Steady-state spectroscopic properties of AtCRY2...... 104 5.3.2 Ultrafast photoreduction dynamics of flavin cofactor in AtCRY2...... 106 xi
5.3.3 Deactivation dynamics of anionic flavin cofactors in AtCRY2...... 108 5.4 Conclusions ...... 110 Bibliography ...... 121 APPENDIX: PROTOCOLS ...... 129
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LIST OF TABLES
Table 2.1 The calculation of the orientation factor κ2 ...... 30
Table 2.2 The optimized structures of MTHF and flavin molecules ...... 31
Table 3.1 Results of reaction times, efficiencies of the elementary steps and overall repair quantum yields of wild-type and mutant photolyases ...... 62
Table 3.2 The energies in electron transfer reactions ...... 63
Table 4.1 Results of reaction times, free energies and reorganization energies of the individual electron transfer steps in the photoreduction of FAD ...... 89
Table 4.2 Results of reaction times, free energies and reorganization energies of the individual electron transfer steps in the photoreduction of FADH ...... 90
Table 5.1 The time scales of the individual electron transfer steps in the photoreduction of FAD in A. thaliana cryptochrome 2 ...... 112
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LIST OF FIGURES
Figure 1.1 Structure of flavin in oxidized state...... 10
Figure 1.2 Different redox and ionic states of flavin (FAD) under physiological conditions...... 11
Figure 1.3 Formation of CPD and (6-4) photoproducts under UV-light irradiation... 12
Figure 1.4 Crystal structures of two types of photolyases...... 13
Figure 1.5 Molecular structures of chromophore cofactors in the photolyase family. 14
Figure 1.6 Sketch of femtosecond pump-probe methods...... 15
Figure 1.7 Principle of the up-conversion technique...... 16
Figure 1.8 Schematic representation of the experiment setup with both fluorescence up-conversion and transient absorption configurations ...... 17
Figure 2.1 Crystal structure of photolyase...... 32
Figure 2.2 Steady-state absorption and emission spectra of flavin cofactor and MTHF.
...... 33
Figure 2.3 Determination of fluorescence quantum yield of MTHF in photolyase. 34
Figure 2.4 Computational models for MTHF and flavin molecules...... 35
Figure 2.5 Femtosecond-resolved fluorescence dynamics of FRET from MTHF to flavin cofactors...... 36
Figure 2.6 Femtosecond-resolved transient absorption dynamics of FRET from
MTHF to FADHˉ...... 37
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Figure 2.7 Femtosecond-resolved transient absorption dynamics of FRET from
MTHF to FAD...... 38
Figure 2.8 Femtosecond-resolved transient absorption dynamics of FRET from
MTHF to FADH ...... 39
Figure 2.9 Optimized structures of MTHF and three states of flavin cofactors and calculated transition dipoles...... 40
Figure 3.1 Enzyme-substrate complex structure of photolyase...... 64
Figure 3.2 Schematic representation of the hypothetical dynamic process in DNA photorepair...... 65
Figure 3.3 Determination of dissociation constants and CPD repair quantum yields by mutant photolyase...... 66
Figure 3.4 Femtosecond-resolved transient absorption dynamics of CPD repair by mutant photolyases of R226A, N378S and M345A...... 67
Figure 3.5 Femtosecond-resolved transient absorption dynamics of CPD repair by mutant photolyases of E274A, R342A and N378C...... 68
Figure 3.6 Schematic representation of ET coupled with solvation susbtrates...... 69
Figure 3.7 Analysis of the forward ET and the electron return by Sumi-Marcus model.
...... 70
Figure 3.8 The energies and rates of ET reactions in wild-type and mutated photolyases ...... 71
Figure 3.9 Quantum yields, reaction times and free energy diagram of the elementary
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steps in the repair of CPD...... 72
Figure 4.1 Electron transfer network of FAD in A. thaliana (6-4) photolyase ...... 91
Figure 4.2 Femtosecond-resolved transient absorption dynamics of photoreduction of
FAD in mutant and wild-type (6-4) photolyases...... 92
Figure 4.3 Femtosecond-resolved dynamics of direct electron transfer to FAD in (6-4) photolyase ...... 93
Figure 4.4 Femtosecond-resolved dynamics of FAD photoreduction involving W383 in (6-4) photolyase ...... 94
Figure 4.5 Femtosecond-resolved dynamics of FAD photoreduction involving W329 in (6-4) photolyase ...... 95
Figure 4.6 Femtosecond-resolved transient absorption dynamics of photoreduction of
FADH in mutant and wild-type (6-4) photolyases ...... 96
Figure 4.7 Femtosecond-resolved dynamics of direct electron transfer to FADH in
(6-4) photolyase...... 97
Figure 4.8 Femtosecond-resolved dynamics of FADH photoreduction involving
W383 in (6-4) photolyase ...... 98
Figure 4.9 Femtosecond-resolved dynamics of FADH photoreduction involving
W329 in (6-4) photolyase ...... 99
Figure 4.10 Kinetic and energetic scheme of the photoreduction of flavin cofactor in A. thaliana (6-4) photolyase ...... 100
Figure 5.1 Sequence alignment and local X-ray structure of A. thaliana cryptochrome.
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...... 113
Figure 5.2 Steady-state spectra of A. thaliana cryptochrome 2 ...... 114
Figure 5.3 Photoreduction of A. thaliana cryptochrome 2 ...... 115
Figure 5.4 Femtosecond-resolved oxidized flavin dynamics in A. thaliana cryptochrome 2 ...... 116
Figure 5.5 Femtosecond-resolved dynamics of direct electron transfer to FAD in A. thaliana cryptochrome 2 ...... 117
Figure 5.6 Femtosecond-resolved dynamics of FAD photoreduction involving W374 in A. thaliana cryptochrome 2 ...... 118
Figure 5.7 Femtosecond-resolved dynamics of photoreduction of FAD and FADH in
A. thaliana cryptochrome 2 ...... 119
Figure 5.8 Femtosecond-resolved dynamics of anionic flavin cofactors (FADH¯ and
FAD¯) in A. thaliana cryptochrome 2 ...... 120
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CHAPTER 1 INTRODUCTION
1
CHAPTER 1
INTRODUCTION
Flavin is widely distributed in nature and plays a very important role in many enzymatic functions.1-5 The most common flavins occurring in flavoproteins are riboflavin, flavin mononucleotide (FMN) and flavin adenine dinucleotide (FAD), which consist of a heterocyclic isoalloxazine ring with ribityl, ribityl phosphate and ribityl adenine diphosphate, respectively (Figure 1.1). Although the chemical entity responsible for the diverse biological activity of flavoproteins is the isoalloxazine moiety, the side chains are also important for the selective binding to a particular flavoprotein.3 Flavins exist in three different redox states: the oxidized quinone, one-electron reduced semiquinone (radical), and two-electron reduced hydroquinone (Figure 1.2). Under physiological conditions, semiquinone and hydroquinone can be present in their neutral
3 and anionic forms based on their pKa values of 8.3 and 6.7, respectively. Among these redox and ionic states, two redox pairs, oxidized quinone/anionic semiquinone
(FAD/FAD−) and neutral semiquinone/anionic hydroquinone (FADH/FADHˉ), can convert between by accepting or donating electrons. The chemical versatility of flavin makes flavoproteins ubiquitous and involved in a wide array of biological process, such as cell apoptosis,6,7 photosynthesis,8 oxygen activation,9 light-driven activity,10 metabolism and so on.11
2
Photolyase and cryptochrome comprise a family of structurally related flavoproteins, which play a vital role in the light-driven biological activities,10 such as photoinduced
DNA repair and signal transduction.10,12-15 The photochemistry and photophysics of this protein family has drawn considerable attention.
Photolyase, one of photoenzymes, can use solar energy to reverse the harmful effects of ultraviolet (UV) radiation.10 Exposure to the UV-light can cause severe DNA damage.
The two major lesions in UV-irradiated DNA are the cyclobutane pyrimidine dimer (CPD) and the pyrimidine (6-4) pyrimidone photoproduct (6-4PP) that are formed between two adjacent pyrimidine bases (Figure 1.3).16,17 These photoproducts result in the mutagenesis, block the replication and transcription, and finally lead to be the main causative agent for skin cancer.16,18-21 These two lesions can be repaired by photolyases with blue light as energy source.10 Photolyases are monomeric proteins with molecular masses in the 53-66 kDa range (454-614 amino acid residues) depending on the organism.22 They can be distinguished by their different substrate specificity: CPD photolyase binds and repairs
CPD lesions in single strand or double helical DNA, whereas (6-4) photolyase reverses
6-4PP-damage in DNA. These two types of photolyases share a high sequence and structure homology and contain a fully reduced flavin adenine dinucleotide (FADHˉ) as the catalytic cofactor (Figure 1.4).10 Besides the flavin cofactor, CPD photolyase has another noncovalently bound chromophore as the light-harvest antenna, either methenyltetrahydrofolate (MTHF) or 8-hydroxy-7,8-didemethyl-5-deazariboflavin
(8-HDF) (Figure 1.5).23 During the catalysis, the photo antenna absorbs a photon and transfers the excitation energy to the catalytic cofactor FADHˉ to enhance the
3
DNA-repair efficiency.24
In both CPD photolyase and (6-4) photolyase, the FADHˉ cofactor is deeply buried within the α-helical domain and has an unusual U-shaped conformation with the isoalloxazine ring and adenine moieties in close proximity.25,26 The FADHˉ is held tightly in place by contact with 14 amino acids, and is accessible to the flat surface of the
α-helical domain through a hole in the middle of this domain. Of special significance, this hole has the right dimension and polarity to allow the entry of a CPD or 6-4PP dimer within van der Waals contact distance to the isoalloxazine ring of the flavin cofactor.10,25
Both CPD photolyase and (6-4) photolyase bind their substrates (CPD to CPD photolyase
-9 10 and 6-4PP to (6-4) photolyase, respectively) in DNA with high affinity (KD ~10 M).
By mixing with appropriate concentrations of photolyases and their substrates, the enzyme-substrate complex can be prepared in dark for the repair study in real time.
The blue-light photoreceptor cryptochromes share the high sequence and structure homology with photolyase, but lack the ability to repair DNA.10 Instead, cryptochromes can mediate a variety of blue-light responses through a series of signal transduction processes, including regulation of the growth and development in plants and photoentrainment of the circadian rhythms in animals.13,14,27,28 Cryptochrome also contains flavin adenine dinucleotide as the key cofactor for its function. The fully reduced FADHˉ has been identified as the catalytic state in photolyase,29 whereas the functional state of flavin in cryptochrome is still under debate. Therefore, it requires more in-depth study to answer this question and more experimental insight need to be provided. 4
Due to the essential role of FAD in photolyase and cryptochrome, the photochemistry and photophysics of this flavin cofactor become the key to understand the behavior and function of these proteins. To address this issue, we conducted a systematic investigation on the dynamics of flavin cofactors in photolyase and cryptochrome in several aspects. In chapter 2, we examined the ultrafast dynamics of the resonance energy transfer (RET) from MTHF to three flavin redox states of FAD, FADH and FADHˉ, respectively, in E. coli CPD photolyase. With the measured dynamics and the knowledge of the spectral overlap integral between MTHF and each flavin state, the orientation factors were obtained. The highest orientation factor for FADHˉ suggests the excellent structure alignment for the functional state to maximize the enzyme efficiency.
Photolyase uses blue light to repair the major ultraviolet (UV)-induced DNA damage, the cyclobutane pyrimidine dimer (CPD), through a cylic electron transfer radical mechanism. The repair efficiency (0.82) is much higher than those (0.004-0.41) of all chemical model systems synthesized so far.30-33 Although the complete photocycle of
CPD repair by photolyase has been revealed in our recent studies,12,34 the central questions of what the role of the protein environment is and how the repair machinery functions in photolyase still need to be resolved. In chapter 3, we mutated a series of key residues at the active-site of E. coli photolyase, and found out those residues play a critical role in the DNA repair. To examine how the protein active-site residues mediate the repair efficiency, we also mapped out the complete evolution dynamics of flavin and
CPD-related intermediates within 6 active-site mutated photolyases in real time, and analyzed the electron transfer processes in the catalytic reaction with Sumi-Marcus model.
5
The results provide some insights of how the active-site residues modulate the flavin cofactor and the bound CPD to maximize the enzyme activity during the DNA repair.
The flavin cofactor in photolyase is oxidized and loses enzyme activity in vitro.
However, the oxidized flavin can be photoreduced to the active state FADHˉ upon blue light excitation via electron transfer from neighboring aromatic residues. The electron transfer pathway was proposed to go through a chain of three conserved tryptophan residues (Trp-triad).25 In chapter 4, with site-directed mutagenesis, we systematically investigated the dynamics of the electron flow among adenine moiety and the conserved three tryptophan residues to oxidized flavin (FAD) and neutral semiquinone (FADH) in
Arabidopsis thaliana (6-4) photolyase. With femtosecond-resolved spectroscopy, we are able to determine the time scales of the ultrafast photoreduction of the flavin and the electron relay through the Trp-triad, and provide our understanding of intraprotein electron transfer.
To unravel the physiological relevant redox state of flavin in cryptochrome, in the chapter 5, we systematically studied the dynamics of flavin cofactor in four different redox states and two redox pairs (FAD/FAD− and FADH/FADHˉ) in Arabidopsis thaliana cryptochrome 2. With femtosecond resolution, we observed the ultrafast photoreduction of oxidized FAD and neutral semiquinone FADH in picoseconds and tens of picoseconds, and multiple deactivation timescales of anionic flavins. This observed behavior of flavin provides some insight into the funcitional active state of flavin cofactor in cryptochrome.
6
To follow the ultrafast photophysical processes and photochemistry reactions of flavin in photolyase and cryptochrome, femtosecond fluorescence up-conversion and transient absorption techniques were used to carry out all the experiments. Currently, most ultrafast experiments use the well-established pump-probe technique, in which a femtosecond laser pulse initiates some process of interest and another femtosecond pulse delayed in time monitors the response of the changes in the sample. The dynamic response of the photo-process can be reconstructed by repeating the experiment at series of pump-probe delay times between the pump and probe pulses through mechanically changing the optical path length of the pump pulse. The principle of this femtosecond pump-probe spectroscopy is clearly illustrated in Figure 1.6 and Figure 1.7. Briefly, for femtosecond up-conversion method, the sample emits fluorescence upon excitation by the pump pulse. The emitted fluorescence is collected by using a pair of parabolic mirrors and focused on a small spot of the non-linear crystal (BBO crystal). The other femtosecond pulse (gating pulse) is also focused on the same spot of the BBO crystal.
When the gaiting pulse and the fluorescence emission are present simultaneously in the
BBO crystal, through a non-linear process called sum-frequency generation, an up-converted signal is generated. After passing through a double-gating monochromator, the signals are detected by a computer-controlled photomultiplier tube (PMT). For the transient absorption method, the main measurement is the difference in absorption in the absence and presence of the pump beam. After the pump beam initiates the reaction, the probe beam captures the transient absorption of the molecular species as a function of the delay time.
7
The experimental setup is schematically shown in Figure 1.8. Specifically, the femtosecond pulse after the two-stage amplifier (Spitfire, Spectra-Physics) has a temporal width of 110 fs with energy of more than 2 mJ and a repetition rate of 1 kHz. The laser beam is then split into two equal parts to pump two optical parametric amplifiers
(OPA-800C, Spectra-Physics).12,35
For fluorescence up-conversion experiments, we used the pump wavelength at 400 nm by direct doubling of the 800-nm fundamental beam from the first optical parametric amplifier (OPA) through a 0.2-mm-thick BBO crystal. The pump wavelength 480 nm was generated through a mixing of the fundamental (800 nm) and the signal (1200 nm).
Before entering the sample cell, the pump pulse energy was attenuated to 70-200 nJ. The fluorescence emission was collected by a pair of parabolic focus mirrors and mixed with the gating pulse (800 nm) in a 0.2-mm-thick BBO crystal through a noncollinear configuration. The up-converted signals were detected by a photomultiplier tube (PMT) after passing through a double-gating monochromator.
For our transient absorption measurements, we used the pump wavelength at 400 nm by direct doubling of the 800-nm fundamental from the first optical parametric amplifier (OPA) through a 0.2-mm-thick BBO crystal. We also used some other pump wavelengths, such as 480, 500, and 640 nm. Pump wavelengths from 450 nm to 650 nm can be generated through a mixing of the fundamental (800 nm) with the signal or idler.
By translating the movable mirror (MM in Figure 1.8) out of the laser path, the half-intensity fundamental beam (1 mJ) pumps the second OPA, and various probe wavelengths form 400 to 720 nm were generated by a mixing of the idler or signal with
8
the fundamental. Both the pump and probe pulses were compressed through a pair of prisms with double paths to reach a temporal resolution of 60 fs. The sensitivity of the transient absorption method can reach 10-4—10-5 of the absorbance change. All signals were digitized and processed by computers.
In our experiment, the pump beam polarization for up-conversion was set at a magic angle (54.7o) with respect to the acceptance axis of the up-conversion crystal
(vertical), and the gating beam polarization was set parallel to this axis by using a half-wave plate. For the transient absorption experiments, the pump beam polarization was set at a magic angle directly with respect to the probe beam, which is vertical.
9
NH2 Vitamin B (Riboflavin) 2 N 5 7 6 N OH OH O O 1 8 2 O O O 9 3 P P O N 4 OH N O O 9a N 10a N 9 10 1 O Adenine 8 2 7 3 OH OH 6 5 4 NH 5a N 4a Adenosine Isoalloxazine O Flavin Mononucleotide (FMN) Flavin Adenine Dinucleotide (FAD)
Figure 1.1 Structure of flavin in oxidized state.
Riboflavin, flavin mononucleotide (FMN) and flavin adenine dinucleotide (FAD) consist of a heterocyclic isoalloxazine ring with ribityl, ribityl phosphate and ribityl adenine diphosphate, respectively.
10
Figure 1.2 Different redox and ionic states of flavin (FAD) under physiological conditions.
Three different redox states of flavins: oxidized quinone, one-electron reduced semiquinone and two-electron fully-reduced hydroquinone. Under physiological conditions, semiquinone and hydroquinone can be present in their neutral and anionic forms based on their pKa values of 8.3 and 6.7, respectively. Among these five redox and ionic states, there are two redox pairs: oxidized flavin/anionic semiquinone (FAD/FAD−) and neutral semiquinone/ anionic hydroquinone (FADH/FADH−).
11
Figure 1.3 Formation of CPD and (6-4) photoproducts under UV-light irradiation.
UV irradiation on cell can induce severe DNA damage. The cyclobutane pyrimidine dimer (CPD) and the pyrimidine (6-4) pyrimidone photoproduct (6-4PP) are the two major classes of cytotoxic and carcinogenic photolesions in damaged DNA.
12
Figure 1.4 Crystal structures of two types of photolyases.
Ribbon diagram representation of Escherichia coli CPD photolyase (A) and Arabidopsis thaliana (6-4) photolyase (B). Note that the crystal structure of CPD photolyase contains both FAD (green) and folate (red), whereas that of (6-4) photolyase contains only FAD.
13
NH2
N N
O O N N CH (CHOH) CH O 2 3 2 P O P O CH2 O
H3C N N O O O
NH OH OH H C N - 3 H FADH O CH OH O 2 (CHOH) O N C (Glu)n 3
N CH2 HN OH N N O NH 2 N N H NH
5,10-MTHF 8-HDF O Figure 1.5 Molecular structures of chromophore cofactors in the photolyase family.
Molecular structures of the chromophore cofactors in the photolyase family. Note that the FADHˉ is the essential catalytic cofactor.
14
Figure 1.6 Sketch of femtosecond pump-probe methods.
The time delay between pump and probe pulses is determined by the optical path length differences between the two pulses. Variety of the optical path length differences can be achieved by changing the optical path length of the pump pulse with a translation stage in pump line. (A) Setup for up-conversion method. (B) Setup for transient absorption method.
15
Figure 1.7 Principle of the up-conversion technique.
(A) Principle of spatial and temporal overlap. When spatial and temporal overlap of the two beams is achieved in the non-linear crystal, the phase-matching condition ku=kg+kf must be fulfilled for sum-frequency generation ωu=ωg+ωf. (B) By repeating the experiment at various delay times (), a series of up-converted signal would be collected to form final fluorescence transient.
16
Figure 1.8 Schematic representation of the experiment setup with both fluorescence up-conversion and transient absorption configurations
Blue line: pump-pulse pathway. Red line: gate-pulse pathway of up-conversion detection. Purple line: up-converted signal. Green line: probe-pulse pathway of transient absorption detection. F: filter. MM: movable Mirror. MP: movable parabolic mirror. P: parabolic mirror. TS: translation stage. BBO: -barium borate crystal. /2: half-wave plate. PD: photodiode. PMT: photomultiplier tube.
17
CHAPTER 2
ULTRAFAST DYNAMICS OF RESONANCE ENERGY
TRANSFER WITH THREE FLAVIN REDOX STATES IN
PHOTOLYASE
18
CHAPTER 2
ULTRAFAST DYNAMICS OF RESONANCE ENERGY
TRANSFER WITH THREE FLAVIN REDOX STATES IN
PHOTOLYASE
2.1 Introduction One of the detrimental effects of Ultraviolet (UV) irradiation on the biosphere is the formation of cyclobutane pyrimidine dimer (CPD) between two adjacent pyrimidine bases in DNA.36 The CPD photolesion blocks the replication and transcription, and is implicated as the main causative agent for skin cancer.18,37
Photolyase, a photoenzyme, reverses the CPD back to normal DNA bases using blue light as energy source, and therefore prevents the harmful effects of UV irradiation.10,38
Photolyase is a flavoprotein and contains two noncovalently bound chromophores.
One is the fully reduced flavin adenine dinucleotide (FADHˉ), which is the catalytic cofactor to carry out the repair function upon excitation. The second chromophore is an antenna pigment, either methenyltetrahydrofolate (MTHF) or 8-hydroxy-
7,8-didemethyl-5-deazariboflavin (8-HDF).10 In E. coli photolyase, the flavin cofactor
FADHˉ is deeply buried within the α-helical domain and has an unusual U-shaped conformation with the isoalloxazine ring and adenine moieties in close proximity, while the photoantenna MTHF is located in a shallow cleft between the α-helical and
α/β domains and partially sticks out from the surface of the enzyme (Figure 2.1).25 19
During catalysis, the antenna MTHF absorbs a photon and transfers the excitation energy to the catalytic cofactor FADHˉ to yield the excited singlet state FADHˉ*.24,39
The mechanism of energy transfer is believed to be of Förster type via long-range dipole-dipole interaction between MTHF and FADHˉ. Since MTHF has a higher extinction coefficient than FADHˉ and an absorption maximum at longer wavelength relative to that of catalytic cofactor,10 it can enhance the DNA-repair efficiency. The energy transfers from MTHF to FADHˉ and FADH have been studied by measuring the quenching dynamics of MTHF,35,40 but the actual time scale have not yet been determined. Here, we report our systematic study on the Förster resonance energy transfers (FRET) from MTHF to three flavin redox states of FAD, FADH and FADHˉ, respectively, with both femtosecond up-conversion and transient absorption methods.
We not only measured the fluorescence quenching from MTHF, but also captured the formation of the excited state of flavin species by FRET, and finally resolved the energy transfer dynamics precisely. Moreover, we also examined the ultrafast photoreduction dynamics of the excited FAD and FADH led by the energy transfer.
2.2 Materials and Methods
2.2.1 Preparation of CPD Photolyase and Mutants. The CPD photolyase from E. coli was prepared as described previously with modification.41 The protein purification procedure is shown in Appendix I. After purification, the flavin cofactors in CPD photolyase are mostly in the neutral semiquinone form FADH. To obtain the photolyase containing pure FAD or FADHˉ, the sample need the further manipulation of oxidization and photoreduction (details in
Appendix II). The purification of mutant N341A is different from the wild-type
20
photolyase. Instead of going through blue sepharose column and gel-filtration Bio-p
100 column, mutant N341A is purified using Hi Trap Heparin HP column twice.
2.2.2 Fluorescence Quantum Yield of MTHF. Due to the lack of flavin cofactor, the mutant N341A is a good candidate to measure the fluorescence quantum yield (ΦF) of MTHF in photolyase. The most
42 reliable method for recording ΦF is the comparative method of Williams, which involves the use of a well characteristic standard. The Coumarin 1 in ethanol was selected as the standard with the known ΦF value of 0.73 at the excitation wavelength of 360 nm.43 A set of standard samples and a set of N341A samples with different concentrations were prepared. Both samples should be very dilute with the absorbance of less than 0.05 in 5-mm quart cuvette to avoid the stimulated emission and other side effects. The UV-vis absorption spectra and the fluorescence spectra excited at 360 nm of both standard and N341A were recorded from the lower concentration to the higher concentration. Then the integrated fluorescence intensities were plotted via the absorbance at the excitation wavelength (Figure 2.3). Since the standard and the test samples with identical absorbance at the same excitation wavelength can be assumed to be absorbing the same number of photons, the ratio of the slopes of the two plots is equal to the ratio of the quantum yields of the two samples. With known ΦF value of the standard, the quantum yield of N341A was calculated. The experiments were repeated 3 times, the three fluorescence quantum yield of N341A were taken an average.
2.2.3 Femtosecond-resolved Spectroscopy. All the femtosecond-resolved measurements were carried out using up-conversion
21
fluorescence and transient absorption methods. The laser experimental layout and procedure have been detailed elsewhere.12,35 The excitation wavelengths were 400 nm
for all three photolyase enzyme complexes of EPL-MTHF-FAD, EPL-MTHF-FADH and EPL-MTHF-FADHˉ. The instrument response time is about 250 fs and the experiments were done at the magic angle (54.7º). 5-mm quartz cuvettes (Starna) were used as the sample cell in experiments and samples were kept stirring during irradiation to avoid heating and photobleaching. The experiments in the femtosecond-resolved measurements were carried out under anaerobic conditions for
EPL-MTHF- FADHˉ and under aerobic conditions for EPL-MTHF-FAD and
EPL-MTHF-FADH , respectively.
2.2.4 Theoretical study. The conventional models for MTHF and flavin molecules are used for computational studies, as shown in Figure 2.4. All the structural optimizations of these models were carried out by density functional theory (DFT) with the hybrid functional B3LYP and a large basis set 6-311++G (2d,2p). Transition dipole moments and excitation energies were then calculated on the optimized structures with time-dependent density functional theory (TDDFT) at the same computational level.
In order to mimic the protein environment, a polarizable continuum model (IEF-PCM) in the equilibrium (eq) time regime in combination with a static dielectric constant of
4 was adopted throughout the calculations. All the calculations were performed with the Gaussian 09 package.
2.3 Results and Discussion
2.3.1 Quenching dynamics of excited MTHF. The typical femtosecond-resolved fluorescence transients of the enzyme 22
complexes of EPL-MTHF-FAD, EPL-MTHF-FADH and EPL-MTHF- FADHˉ at
400-nm excitation are shown in Figure 2.5. All transients were well fitted by a single-exponential decay with the time constants of 20 ps for EPL-MTHF-FAD, 18 ps
for EPL-MTHF-FADH and 160 ps for EPL-MTHF- FADHˉ. At 400-nm excitation, the absorption of MTHF is dominant due to its relatively higher absorption coefficient.
For EPL-MTHF-FAD, the excited FAD* would be quenched by the ultrafast electron transfer from the neighboring aromatic residues in less than 1 ps,44 and the 20-ps
dynamics should not come from the excited FAD*. For EPL-MTHF-FADH , the excited FADH* has no emission at the gated wavelength of 480 nm because its absorption extends to700 nm. For EPL-MTHF- FADHˉ, the excited FADHˉ* has emission peaks at 510 and 540 nm, and its lifetime (1.3 ns) is much longer than 160 ps.12 Therefore, the observed fluorescence dynamics of the three complexes are purely from the excited MTHF*. Moreover, the fluorescence lifetime of MTHF* in mutant
N341A is 2.6 ns, which is consistent with the recently reported lifetime (2.55 ns) of
45 EPL-MTHF complex without flavin cofactor in Caulobacter crescentus photolyase.
Thus, the recorded dynamics of MTHF* probably represent the excitation energy transfer from MTHF* to the three flavin states.
2.3.2 Capture of the FADH¯* intermediate.
To confirm that the quenching of EPL- MTHF*-FADHˉ occurs by the resonance energy transfer, we searched for the proposed intermediate FADHˉ* with femtosecond-resolved transient absorption spectroscopy. Strikingly, the absorption transients show a distinct rise-decay pattern besides the quenching of MTHF* at all wavelengths longer than 500 nm (Figure 2.6), indicating the formation of an
23
intermediate. Since ground state MTHF and FADHˉ do not have absorption at this region, the intermediate was ambiguously identified as FADHˉ*. Thus, the quenching of MTHF* is caused by the resonance energy transfer. Taking the radiative lifetime of
MTHF* into account, the time constant of resonance energy transfer from MTHF* to
FADHˉ is 170 ps.
Due to the loss in the purification, the purified E. coli photolyase contains only
46-48 substoichoimetric (20-50%) of MTHF. In our sample EPL-MTHF*-FADHˉ, the ratio of MTHF to FADHˉ is 0.34. According to the extinction coefficients of MTHF and FADHˉ at 400 nm (21675 to 2861), the excited population of MTHF and FADHˉ
(0.7 to 0.3) was obtained. Thus, the absorption transients should contain the signals from two parts of excited FADHˉ: one is directly excited by photon (dashed pink in
Figure 2.6) while the other is excited by FRET from MTHF (dashed dark goldenrod).
With the known lifetime of 1.3 ns for FADHˉ*, the time scale of 170 ps for the formation of excited FADHˉ intermediate by FRET was obtained, which is consistent with the result from the above fluorescence study.
2.3.3 FRET from MTHF to FAD and the subsequent reduction.
The dynamics of excited EPL-MTHF-FAD were recorded by the femtosecond-resolved transient absorption (Figure 2.7). The absorption transients probed at 580-640 nm clear show the strong signal of a 20-ps rise. Since the excited
44 EPL-FAD without MTHF does not have such dynamics, the 20-ps rise was unambiguously identified as the formation of intermediate FAD* by FRET from
MTHF. Due to the loss of MTHF, the absorption transients should contain the FAD* signals from both directly excited FAD and excited FAD by FRET. The ratio of 1:1
24
was obtained based on the ground state population of FAD and MTHF and their extinction coefficients at 400 nm.
The FAD* is readily to be reduced via electron transfer (ET) from the neighboring aromatic residues. The ET was proposed to mainly go through a conserved triple tryptophan chain (W382-W359-W306 tryptophan triad).25 In our previous study, the photoreduction dynamics of EPL-FAD in CPD photolyase have been resolved (the similar work for (6-4) photolyase will be described in detail in chapter 4). The excited
FAD is reduced by the electron transfer from the nearby W382 in 0.7 ps and the charge recombination takes 65 ps. Subsequently, the conserved tryptophan triad delivers an electron via multistep hopping mechanism in 75 ps from W359 to W382+ and in 150 ps from W306 to W359+. With this reduction dynamics and the excited population ratio of MTHF and FAD, all the absorption transients of EPL-MTHF-FAD can be fit very well. Four typical transients with distinct patterns are deconvoluted by the evolution of excited MTHF, the photoreduction of directly excited FAD and excited FAD by FRET with tryptophan triad (insets of Figure 2.7). The reduction of
FAD* from FRET shows a typical 20-ps rise at the initial part and a long plateau overlapped with that of directly excited FAD at the long time window due to the 1:1 ratio.
2.3.4 FRET from MTHF to FADH and the subsequent reduction. The dynamics of excited EPL-MTHF-FADH were recorded by the femtosecond-resolved transient absorption (Figure 2.8). Similar to EPL-MTHF-FAD,
the absorption transients of excited EPL-MTHF-FADH at wavelengths of longer than
620 nm show a clear rise signal of about 20 ps. This rise was assigned as the
25
formation of the intermediate FADH * by FRET from MTHF, because the EPL-
FADH alone does not have such rise dynamics of about 20 ps and MTHF* has the decay dynamics at the wavelengths of 700 and 660 nm. Since the FADH in the enzyme is partially reduced to FADHˉ upon 400-nm excitation, the absorption transients contain minor signal from excited EPL-MTHF-FADHˉ. Moreover, the transients also include the signal from directly excited FADH* due to the loss of
MTHF. The previous photoreduction dynamics study of EPL-MTHF-FADH revealed that the FADH* is ready to be reduced by the ET from the nearby W382 in 40 ps and the charge recombination takes 4 ps. The following electron hopping processes occur in 2 ps from W359 to W382+ and in 120 ps from W306 to W359+. With the knowledge of the dynamics of excited EPL-MTHF-FADHˉ and reduction of
EPL-FADH * as well as the excited state population ratio (0.7:0.3) of MTHF and
FADH , all the absorption transients of EPL-MTHF-FAD can be fit systematically. The time scale of 18 ps for the formation of excited FADH intermediate by FRET was accurately determined, which matches the fluorescence study very well.
2.3.5 Calculation of the orientation factor. According to Föster energy transfer theory, the rate of FRET from MTHF* to flavin cofactor depends on the relative position (r) and orientations of donor (MTHF) and acceptor (flavin) and can be expressed as follows:49
6 1 R0 k RET (2.1) D r
2 2 4 1/ 6 R0 9.7810 n Q D J (2.2)
26
where R0 , the Föster distance, is defined as the donor-acceptor distance at which the
transfer efficiency is 50%. τD and QD are the donor’s fluorescence quantum yield and radiative lifetime in the absence of acceptor, respectively, r is the center-to-center distance between the donor and acceptor, κ2 is the orientation factor, n is the refractive index of the medium (1.39), and J is the spectral overlap integral between the donor’s emission and acceptor’s absorption. The X-ray structure reported a distance (r) of 16.8
Å between MTHF and the flavin cofactor.25 Given the FRET rates from MTHF to the three flavin states and the radiative lifetime of MTHF, the derived values of R0 are
37.81 Å for MTHF-FAD, 38.48 Å for MTHF- FADH and 26.47 Å for MTHF-FADHˉ.
From the study of N341A mutant, the fluorescence quantum yield of 0.408 for
MTHF* was obtained. The spectral overlap integral is generally expressed as follows:49
F() ()4d J (2.3) F()d
Where F(λ) represents the emission spectrum of donor in the absence of the acceptor and ε(λ) is the absorbance molar extinction coefficients of acceptor in units of cm-1•M-1. Typically, the steady-state fluorescence and absorption spectra (Figure 2.2) are used to calculate the J value. The obtained three J values are listed in the Table 2.1.
Through the equation 2.2, the orientation factors were found to be 1.53 for
MTHF-FAD, 1.26 for MTHF-FADH and 2.84 for MTHF-FADHˉ. The largest orientation factor for MTHF-FADHˉ indicates the best orientation for the functional state.
27
2.3.6 Theoretical study of the orientation factors. With the theoretical calculation using the initial X-ray structures of MTHF and flavin molecules, the κ2 values were estimated though the following equation.49
2 2 cosT 3cos D cos A (2.4)
Where θT is the angle between the emission transition dipole of the donor and the absorption transition dipole of acceptor, θD and θA are the angles between these dipoles and the vector joining the centers of the donor and acceptor. From the optimized structures (Table 2.2), the center-to-center vectors were found to be (-3.76, 9.71, 12.91) for MTHF-FAD, (-3.77, 9.71, 12.92) for MTHF-FADH and (-3.79, 9.70, 12.96) for
MTHF-FADHˉ. Based on the optimized structures, the transition dipoles were calculated with time-dependent density functional theory. The emission transition dipole of MTHF (2.39, -1.06, -1.16), and the absorption transition dipoles of FAD
(1.37, 0.34, 1.23), FADH (1.25, 0.097, 1.05) and FADHˉ (0.14, 0.23, 0.18) are shown in Figure 2.9. The transition dipoles of FAD and FADH are in the flavin plane and with the angles of 77.6˚ and 84.9˚ relative to the N5-N10 axis, respectively, which are consistent with previous study.50,51 With the directions of these transition dipoles and the center-to-center vector, the orientation factors were determined to be 1.57 for
MTHF-FAD, 1.29 for MTHF-FADH and 2.23 for MTHF-FADHˉ. These results match the experimental values very well.
2.4 Conclusion The Förster resonance energy transfers from MTHF to three flavin redox states in photolyase were systematically studied using femtosecond-resolved fluorescence up-conversion and transient absorption methods. The excitation energy transfer from 28
the antenna molecule MTHF to the fully reduced hydroquinone FADHˉ occurs in 170 ps, but it takes 20 and 18 ps to the oxidized FAD and neutral semiquinone FADH, respectively. The orientation factors were estimated from experimental data as well as the theoretical study. Both results show that MTHF-FADHˉ has the largest κ2 value, which indicates the best orientation alignment for the functional state.
29
a 6b 3 -1 2 τFRET (ps) τD (ps) (R0/r) r (Å) R0 (Å) QD n J (cm •M ) κ MTHF-FAD 20 2600 130.00 16.8 37.81 0.408 1.39 1.99x10-14 1.53 MTHF-FADH 18 2600 144.44 16.8 38.48 0.408 1.39 2.68x10-14 1.26 MTHF-FADHˉ 170 2600 15.29 16.8 26.47 0.408 1.39 1.26x10-15 2.84
Table 2.1 The calculation of the orientation factor κ2 a b 6 τD is the radiative lifetime of donor MTHF in photolyase without any quenching channel. (R0/r) is calculated by (τD/τFRET).
30
MTHF X Y Z FAD X Y Z FADH X Y Z FADHˉ X Y Z
N 29.67 63.55 17.90 N 27.13 73.38 27.69 N 27.11 73.38 27.68 N 27.15 73.39 27.69
C 29.13 64.18 16.88 C 26.27 72.79 26.80 C 26.25 72.80 26.79 C 26.48 72.89 26.62
N 28.01 64.91 17.07 O 25.36 73.38 26.25 O 25.33 73.39 26.24 O 25.72 73.56 25.90
N 29.68 64.17 15.63 N 26.46 71.42 26.51 N 26.42 71.43 26.51 N 26.66 71.54 26.32
C 30.86 63.47 15.31 C 27.42 70.57 27.02 C 27.39 70.60 27.03 C 27.47 70.63 27.01
O 31.31 63.50 14.16 O 27.49 69.40 26.70 O 27.49 69.40 26.74 O 27.56 69.44 26.64
C 31.42 62.78 16.42 C 28.33 71.25 27.98 C 28.26 71.27 27.95 C 28.11 71.20 28.13
N 32.57 61.99 16.28 N 29.28 70.58 28.55 N 29.27 70.58 28.56 N 28.91 70.40 29.00
C 32.89 61.00 17.33 C 30.11 71.21 29.43 C 30.13 71.18 29.45 C 29.92 71.08 29.70
C 32.70 61.66 18.69 C 31.16 70.50 30.05 C 31.17 70.48 30.08 C 31.05 70.44 30.20
N 31.36 62.23 18.74 C 32.01 71.10 30.95 C 32.02 71.10 30.97 C 32.01 71.11 30.97
C 30.82 62.87 17.68 C 33.11 70.32 31.60 C 33.13 70.32 31.63 C 33.22 70.36 31.48
C 34.32 60.57 16.95 C 31.83 72.49 31.25 C 31.84 72.48 31.26 C 31.83 72.47 31.24
N 34.47 61.11 15.58 C 32.74 73.18 32.22 C 32.75 73.19 32.22 C 32.83 73.24 32.07
C 33.47 61.96 15.32 C 30.81 73.20 30.64 C 30.82 73.17 30.63 C 30.70 73.12 30.73
C 37.77 60.31 13.10 C 29.95 72.58 29.73 C 29.96 72.55 29.73 C 29.74 72.44 29.98
C 37.85 60.10 14.47 N 28.92 73.25 29.11 N 28.92 73.23 29.10 N 28.58 73.08 29.50
C 36.76 60.36 15.30 C 28.07 72.66 28.23 C 28.06 72.65 28.22 C 27.92 72.55 28.39
C 35.57 60.84 14.74
C 35.48 61.03 13.36
C 36.58 60.78 12.55
Table 2.2 The optimized structures of MTHF and flavin moleculesa aAll coordinates are in unit of Å.
31
Figure 2.1 Crystal structure of photolyase
X-ray structure of E. coli photolyase (orange ribbon) containing the antenna MTHF, the catalytic cofactor FADHˉ, and the tryptophan triad (red stick) for the photoreduction of the flavin cofactor. FRET, förster resonance energy transfer.
32
Figure 2.2 Steady-state absorption and emission spectra of flavin cofactor and MTHF
Shown are the absorption spectra (thick line) of MTHF (black), FAD (blue), FADH (red) and FADHˉ (green), and the emission spectra (thin line) of MTHF (gray), respectively. The excitation wavelength was 360 nm for MTHF. The spectrum was taken under anaerobic conditions for FADHˉ, whereas the spectra were taken under aerobic conditions for FAD, FADH and MTHF.
33
Figure 2.3 Determination of fluorescence quantum yield of MTHF in photolyase
N341A is a good candidate to measure the fluorescence quantum yield of MTHF in photolyase because it has MTHF but lacks the flavin cofactors. The coumarin 1 in ethanol was selected as the standard sample. The integrated fluorescence intensities of coumarin 1 (standard) and N341A were plotted via the absorbance at 360 nm (excitation wavelength). With the known fluorescence quantum yield of 0.73 for the standard, the fluorescence quantum yield of N341A was calculated by the ratio of the slopes of the two plots.
34
Figure 2.4 Computational models for MTHF and flavin molecules
The initial structures of MTHF and flavin molecules were extracted from the PDB file (1DNP). Then the structural optimizations of these models were carried out by density functional theory (DFT) with the hybrid functional B3LYP and a large basis set 6-311++G (2d,2p).
35
Figure 2.5 Femtosecond-resolved fluorescence dynamics of FRET from MTHF to flavin cofactors
Shown are the normalized fluorescence transients of FRET from MTHF to FAD (blue), FADH (red) and FADHˉ (green), respectively. The gated wavelengths were 490 nm for FADHˉ, and 480 nm for FAD and FADH.
36
Figure 2.6 Femtosecond-resolved transient absorption dynamics of FRET from MTHF to FADHˉ
The FRET dynamics are probed systematically from 700 to 500 nm and four typical transients with distinct patterns are deconvoluted by the evolution of excited MTHF (dashed blue), the excited FADHˉ from FRET (dashed dark goldenrod), and the directly excited FADHˉ with tryptophan triad (dashed pink).
37
Figure 2.7 Femtosecond-resolved transient absorption dynamics of FRET from MTHF to FAD
The FRET dynamics are probed systematically from 700 to 500 nm and four typical transients with distinct patterns are deconvoluted by the evolution of excited MTHF (dashed blue), the photoreduction of excited FAD from FRET with tryptophan triad (dashed dark goldenrod), and the photoreduction of directly excited FAD with tryptophan triad (dashed pink).
38
Figure 2.8 Femtosecond-resolved transient absorption dynamics of FRET from MTHF to FADH
The FRET dynamics are probed systematically from 700 to 500 nm and three typical transients with distinct patterns are deconvoluted by the evolution of excited MTHF (dashed blue), the photoreduction of excited FADHfrom FRET with tryptophan triad (dashed dark goldenrod), the photoreduction of directly excited FADH with tryptophan triad (dashed pink), and the signal of some residue of (MTHF-FADHˉ).
39
Figure 2.9 Optimized structures of MTHF and three states of flavin cofactors and calculated transition dipoles.
Optimized structures of three donor-acceptor pairs of MTHF-FAD, MTHF-FADH and MTHF-FADHˉ with vectors from mass-center of donor to mass-center of acceptor (black arrow) and calculated transition dipole moments of MTHF (violet arrow) and flavin cofactors (red arrow). The insets show the overlap between the emission of MTHF and the absorption of three flavin cofactors, respectively.
40
CHAPTER 3 MOLECULAR UNDERSTANDING OF EFFICIENT DNA REPAIR MACHINERY OF PHOTOLYASE
41
CHAPTER 3
MOLECULAR UNDERSTANDING OF EFFICIENT DNA REPAIR
MACHINERY OF PHOTOLYASE
3.1 Introduction Life under the sun is endangered by ultraviolet (UV) radiation that causes the formation of genotoxic photoproducts in DNA. The most abundant UV-induced lesion is the cyclobutane pyrimidine dimer (CPD) formed between two adjacent pyrimidine bases, mainly thymine, through a [2+2] photocycloaddition. The formation of this lesion blocks
DNA replication and transcription and is a leading cause of skin cancer. 16,18-20 In prokaryotes, plants and many animals, photolyase, a photoenzyme which contains an active cofactor flavin adenine dinucleotide in fully reduced form (FADHˉ), is mainly responsible for the repair of CPD by catalyzing the cleavage of the cyclobutane ring, using blue light as energy source.10,21
With femtosecond (fs)-resolved spectroscopy, we recently followed the entire repair dynamics from the reactants, to intermediates, and finally to products and thus resolved the complete repair photocycle by photolyase.12,34,52 Briefly, photolyase binds the DNA at a T<>T lesion independent of light. The excited flavin cofactor FADHˉ* injects an
42
electron to the CPD to generate a charge-separated radical pair (FADH+T<>Tˉ) upon excitation. Then the anionic ring (T<>Tˉ) of the dimer is split by a sequential [2+2] cycloreversion and the excess electron returns to the flavin radical FADH to restore the catalytic active form FADHˉ and normal bases (Figure 3.2). However, the central questions of what determines the repair quantum yield and how the repair machinery efficiently functions in photolyase still need to be resolved. The repair of CPD by photolyase achieves a very high efficiency (0.82) compared to those (0.004-0.41) of the synthesized chemical systems, indicating that the amino acids at the active site must play a critical role in the repair efficiency by optimizing the redox potentials of the flavin/CPD species and modulating the electron transfer (ET) rates in the repair photocycle.30-33
Shown in Figure 1 is the X-ray structure of A. nidulans photolyase complexed with the CPD-like lesion after in situ repair.53 At the binding pocket, four charged/polar residues R232, E283, N348, and R350 have hydrophilic interactions with the thymine dimer. On the flavin side, N386 is located near the N(5) position of flavin, which may form a hydrogen bond with the N(5)H group of FADHˉ or FADH.54 M353 is in the middle of flavin and 3’-thymine. The sulfur atom may have interaction with the ring of
3’-thymine. These six residues are conserved in many photolyases. In E. coli photolyase, the corresponding residues are R226, E274, N341, R342, N378 and M345. Moreover, recent mutation studies of S. cerevisiae photolyase showed significant change in the repair quantum yield by mutation of the counterparts of E274, and R342 ——E384A
(0.4, relative to wild-type) and R452A (0.4).55
43
To reveal how these residues modulate the catalytic repair reaction, we prepared six mutant proteins of E. coli photolyase (R226A, E274A, R342A, N378C, N378S, and
M345A) and perform a complete series of fs-resolved transient absorption studies from visible to UV light region. With five typical wavelength detections as well as the careful determination of enzyme/substrate binding constants and repair quantum yields, we were able to resolve the dynamics of forward ET, bond splitting, backward ET without repair, and electron return. Based on the Marcus two-dimensional ET theory,56 we evaluated the driving force, intramolecular and solvent reorganization energies.
3.2 Materials and Methods
3.2.1 Preparation of CPD Photolyase and Mutants. The purification of E. coli photolyase with depletion of the antenna cofactor has been reported elsewhere.41,57 For the mutant studies, we mutated a series of key residue at the active site (R226A, E274A, N341A, R342A, N378C, N378S and M345A). All the mutant plasmids were constructed using QuikChange II XL kit (Stratagene) based on the plasmid of wild-type enzyme. All mutated DNA plasmids were sequenced to ensure correct results. After the standard purification, all mutant proteins were obtained except N341A
(It lacks the FAD cofactor). In femtosecond spectroscopic studies, 100 μM of enzyme (or
50 μM in experiments with probe wavelengths of shorter than 300 nm) was used in a reaction buffer containing 100 mM NaCl, 50 mM Tris-HCl at pH 7.5, 20 mM dithiothreitol, 1 mM EDTA, and 50% (v/v) glycerol.
44
3.2.2 Preparation of CPD Substrate.
We prepared the CPD substrate of oligo(dT)15 as described elsewhere with some
58 modification. Briefly, we dissolve 3 mg oligo(dT)15 (synthesized by Integrated DNA
Technologies) in 1 mL 15% aqueous acetone (v/v). The argon-purged DNA solutions were irradiated over ice with a 302-nm UVB lamp (General Electric) at a 2-cm distance for 50-70 min. CPD formation was monitored by decreases in absorbance at 260 nm. In the final products, there are about 5 CPD in each strand of oligo(dT)15 on average. The concentration of the oligo-substrates used in the femtosecond studies is 3 mM (or 1.5 mM in experiments with probe wavelengths of shorter than 300 nm).
3.2.3 Enzyme Activities. The enzyme activities of the wild-type and mutant photolyases were quantitatively measured. Procedures for determination of dissociation constant and relative quantum yields of the mutants were carried out as described.34,52 For each mutant, a set of mixtures of 1 μM enzyme with different concentrations (111, 333, 500 and 1000 μM) of
CPD-containing oligo(dT)15 substrate were prepared. These samples were put into cuvettes and irradiated at the room temperature using white-light lamp (General Electric) with the same distance of 6 cm. The absorption spectra of different mixtures were recorded at different illumination times. The increased absorbance around 266 nm indicated the formation of thymine base. The absorbance change at 266 nm was plotted vs. illumination time, and the slope is proportional to the binding complex concentrations and the repair efficiency of the enzyme.
45
3.2.4 Femtosecond-resolved Spectroscopy. All the femtosecond-resolved measurements were carried out using transient absorption methods. The laser experimental layout and procedure have been detailed elsewhere.35,44 1-mm quartz cuvettes (Starna) were used as the sample cell in experiments with probe wavelengths of shorter than 300 nm, while 5-mm cuvettes were used for other experiments. Samples were kept stirring during irradiation to avoid heating and photobleaching. All experiments in the femtosecond-resolved measurements were carried out under anaerobic conditions.
3.2.5 Electron Transfer Fitting Model. The forward ET was analyzed by Sumi-Marcus ET model.56 In this model, the free energy surface of reactant and product along the fast q (intramolecular distortion) and slow X (solvent reorganization) coordinates are given by:
1 V r (q, X ) q2 V (X ) (3.1) 2
p 1 2 V (q, X ) (q q ) V (X X ) G (3.2) 2 p p where V (X ) is the free energy as a function of X and assumed to be a harmonic
1 potential X 2 . G is the standard free energy of reaction. The total reorganization 2
1 1 energy is the sum of q 2 (intramolecular distortion) and X 2 i 2 p 0 2 p
(solvent reorganization). At a given X , the activation barrier and the rate k are:
* 2 G (X ) (G 0 i X 20 ) / 4i (3.3)
46
J 2 G* (X ) k(X ) exp( ) (3.4) ikBT 1.35kBT where J is the electron coupling matrix element, is the reduced Planck constant,
kB is the Boltzmann constant, T is the temperature in kelvin, 1.35 is the Pempirical correction of Frank-Condon term by Dutton et al. considering the electron tunneling effect in biological system.59 The population distribution of reactant P(X,t) can be given by reaction-diffusion equation:
P(X ,t) L [k(X ) k ]P(X ,t) (3.5) t lifetime
dV (X ) L D(t) [k T ] (3.6) X B X dX
P(X ,t) 2P(X ,t) dV (X ) D(t)kBT 2 D(t) [P(X ,t) ][k(X ) klifetime ]P(X ,t) (3.7) t X X dX
Where klifetime is the lifetime of excited flavin cofactor FADHˉ* in photolyase along, and the D(t) is the time-dependent diffusion coefficient that can be determined by the protein solvation correlation function C(t) by:60
1 dC(t) D(t) (3.8) C(t) dt
C1(t) 0.4670exp(t) 0.265exp(t / 25) 0.165exp(-t/750) 0.100exp(-t/3000) (3.9)
C2 (t) 0.636exp(-t/1.3) 0.149exp(-t/20) 0.115exp(-t/580) 0.100exp(-t/3000) (3.10)
The normalized solvation correlation function (9) was used for the fitting of N378C and
N378S, while the correlation function (10) was used for wild type and other mutants. For the initial condition, a thermal equilibrium distribution of reactant is assumed:
47
2 P(X,0) exp(X / 2kBT) / 2kBT (3.11)
Then P(X,t) can be obtained from equation (5). Given that, the time-dependent reactant population Q(t) is:
Q(t) P(X,t)dX ,Q(0) 1,Q() 0 (3.12)
The four parameters of ET reaction ( J , G , i and 0 ) can be obtained when the numerical solution of population matches the normalized observed ET transients.
The average reaction time constant is calculated from:
t Q(t)dt (3.13) t0
The extended Sumi-Marcus ET model developed by Barbara was used to analyze the electron return. Considering the high-frequency vibrational modes of product, the activation barrier and the ET rate k at the nth vibrational state are given:
* 2 G (n, X ) (G 0 i X 20 n) / 4i (3.14)
sn J 2 G* (n, X ) k(n, X ) exp(S) exp[ ], S (3.15) n! ikBT 1.35kBT
Here and are the angular frequency and reorganization energy of the high-frequency vibrational mode, respectively. Then the population distribution of reactant P(X,t) can be given by:
P(X ,t) 2P(X ,t) dV (X ) D(t)k T D(t) [P(X ,t) k(n, X )P(X ,t)] (3.16) t B X 2 X dX n
48
3.3 Results and Discussion
3.3.1 Dissociation Constants and Relative Quantum Yields. The binding complex percentage ([ES]/[E]) for each enzyme is plotted against different substrate concentrations (Figure 3.3B), and the dissociation constants were fit by
-6 the equation [ES]/[E]=[S]/([S]+Kd). The derived dissociation constants are 8.3x10 ,
1.2x10-5, 1.4x10-5, 2.0x10-4, 5.0x10-5, 2.4x10-5, and 1.2x10-5M for wild type (WT),
E274A, R226A, R342A, M345A, N378C and N378S, respectively. R342A has the largest dissociation constant, which is consistent with the crystal structure observation that R342 has many H-bonds with the water network at the binding site and direct interaction with the phosphate of DNA backbone. The mutation of R342 will result in the weaker binding.
With the dissociation constants and under the same conditions, the repair quantum yields for each mutant were calculated by comparing the slopes of mutants with that of
WT (Figure 3.3A), and taking account of the binding-complex concentrations. Knowing the repair quantum yield of WT to be 0.82,30 the repair quantum yields for each mutant were obtained (Table 3.1). All the mutants have lower quantum yields compare to WT, suggesting that these residues play a significant role in the repair efficiency.
3.3.2 Reaction Times of Each Elementary Step. Figure 3.4 and 3.5 show the femtosecond-resolved dynamics of CPD repair by the six mutant photolyases with typical detection wavelengths from 800 nm to 266 nm to map out the complete dynamics evolution of the repair. At 800 nm, only excited flavin cofactor FADHˉ* can be detected. In presence of substrate, the transients drastically
49
change and become much faster compared to those transients in absence of substrate, indicating the quenching dynamics by forward ET from FADHˉ* to the bound CPD. The forward dynamics do not follow a single-exponential decay, but a stretched-single-exponential decay, Ae (t / ) ， with the stretched parameters of 0.64 (β) for R342A and 0.71 for other mutants. As pointed out elsewhere,12,34,61 the stretched-exponential dynamic behavior results from the modulation of ET by active-site
1 solvation. Based on ( ) and taking the lifetime contribution into account,62 the timescales of forward ET for each mutant were obtained (Table 3.1). The repair by WT has also been examined (data not shown). It has the similar dynamics as the previous repair study with the substrate of dinucleotide CPD. Compared to WT, both the binding-site mutants (E274A, R226A, R342A) and the N5 position mutants (N378S,
N378S) have the much slower forward ET. Only M345A shows the faster dynamics.
Knowing the forward ET dynamics of FADHˉ*, we can map out the temporal evolution of the flavin intermediate FADH by probing at 620 nm. To detect the thymine-related intermediates and products, we extended the probe wavelength to UV region. Three typical results probed at 300, 270 and 266 nm are shown in Figure 3.4 and
3.5. All these transients can be systematically fit only with the sequential model,34 and the resulting reaction times for all elementary steps are listed in Table 3.1.
After the forward ET, the formed anionic CPD could evolve in two channels: repair by the sequential splitting of C5-C5’ and C6-C6’ bonds and direct back ET without repair.
Similar to previous studies, we did not observe any signal of T<>Tˉ at any probe 50
wavelength, indicating that the splitting of C5-C5’ bond is instantaneous and trace amount of T<>Tˉ is accumulated. Since the back ET from intact CPD has a driving force about -2.0 eV and should occur within hundreds of picoseconds or longer in Marcus inverted region63,64, the back ET without any bond splitting could not compete with the
C5-C5’ bond splitting, leading to the almost unity evolution along the first bond splitting.
The C6-C6’ bond splitting was detected by observing the formation and decay of T-Tˉ intermediate. The N5 position mutants (N378C and N378S) and M345A have the same second-bond splitting time in 90 ps as the wild type, while the binding-site mutants
(E274A, R226A and R342A) have the faster bond splitting (Table 3.1).
After the sequential splitting of both C-C bonds, the excess electron on thymine base will return back to the flavin cofactor FADH to restore the catalytically active state
FADHˉ and complete the photocycle. In UV region, we observed the decay of Tˉ at 300 nm and the formation of the final product T at 270 and 266 nm, reflecting that the electron return dynamics of wild type and mutants differ from 75 ps to 1.4 ns.
3.3.3 Analysis of Forward ET and Electron Return by Sumi-Marcus Model. The electron transfer in protein is a complex process, strongly influenced by the protein environment. The protein environment of reaction center represents an inhomogeneous solvent, which relaxes upon excitation or charge separation over many different time scales.61,65,66 Thus, a complete mode to describe electron transfer should take the active- site solvation into account. Sumi and Marcus developed a model,56 in which the reaction coordinate of ET is divided into two dimensions: one for the
51
intramolecular distortion (q) and the other for solvent reorganization (X). A typical free energy surface and its contour for the forward ET have been shown in our previous study,67 and the ones for the electron return are shown in Figure 3.6B. Upon excitation or charge separation, the reactants relax along both the intramolecular distortion coordinate and the solvent reorganization coordinate. However, the intramolecular distortion is much faster than ET while the solvent reorganization occurs on the time scale of ET or longer.
At different positions along the solvent reorganization coordinate, the activation barriers of ET are different. In this model, the dynamics of ET can be described by the reaction-diffusion equation:
P(X ,t) ˆ L k(X )P(X ,t) (3.17) t where P(X,t) is the population distribution of the reactants along the reaction coordinate X as a function of time t; Lˆ is an operator that describes the diffusion of solvent relaxation along the coordinate and can be determined by the solvation correlation function; k(X ) is the rate constant for electron transfer at the reaction coordinate position :
J 2 (G 2X )2 k(X ) exp i 0 0 (3.18) ikBT 4ikBT where J is the electron coupling matrix element; G is the standard free energy
difference between the reactants and the products; i is the reorganization energy for
the intramolecular distortion and 0 is the reorganization energy for solvent relaxation.
With integrating along , we can get the time-dependent population of the
52
reactantsQ(t) . Since the solvation dynamics have been characterized in our previous
61 ˆ study, L is determined. Thereafter, the four parameters of ET reaction ( J , G , i and
0 ) can be obtained when the numerical solution of population Q(t) matches the normalized observed ET transients.
To get more understanding how the active-site residues modulate the ETs in the repair reaction, we analyzed the forward ET and electron return using this Sumi-Marcus model. Given that the reduction potentials of T<>T/T<>Tˉ and FADH/FADHˉ for wild
63,64,68,69 type are -1.96 V and +0.08 V vs. NHE, respectively, and assuming the S1←S0 transition of FADHˉ at 500 nm (2.48 eV), the free energy for the forward ET of wild type is -0.44 eV. Additionally, the electron coupling between FADHˉ* and CPD has been solved as 0.003 eV in our recent study.52 With these two restrictions, the non-single-exponential decay of forward ET in wild type was well fit by Sumi-Marcus
model (Figure 3.7A), and the other two parameters of i and were obtained (Table
3.2). For the mutants, assuming the electron couplings same as wild type, all the transients of forward ET can be well fit (Figure 3.7A). The fitting parameters are listed in
Table 3.2.
For the electron return, we used the extended Sumi-Marcus model developed by
Barbara and coworkers,70-72 in which the high-frequency vibrational modes of products are considered (Figure 3.6A). The free energy for electron return can be derived by
0 subtracting forward ET G from -2.70 eV, based on the energy of the S1←S0 transition of FADHˉ at 500 nm (2.48 eV) and the reduction potential difference (0.22 eV)
53
between T<>T/T<>Tˉ and T/Tˉ.63,73 The electron coupling J between FADH and Tˉ has been solved as 0.0026 eV in our recent study.52 Figure 3.7B shows the electron return transients of wild type and mutants simulated by the fitting parameters from stretched-exponential model. All the transients can be systematically fit using the extended Sumi-Marcus model. The fitting results of electron return are listed in Table
3.2. Clearly, the driving force ( G ) is much larger than the total reorganization
energy ( i 0 ), thus the electron return is in Marcus inverted region.
The energy parameters involving in forward ET and electron return reactions in all samples are plotted in Figure 3.8A. The driving forces ( G ) and the solvent
reorganization energies ( 0 ) are diverse (see discussion below), while the energies from high-frequency vibrational modes ( ) and the intramolecular distortion reorganization
energies ( i ) are almost in variant except R342A.
R342A has a different (~0.2 eV lower) for forward ET from others, because of its structural specificity. According to the crystal structure (Figure 3.1), the side chain of
R342 directly interacts with the P+1 phosphate on the DNA backbone. Besides, R342 forms the hydrogen bonds with a cluster of water molecules at the binding pocket. This cluster of water builds up a network, which has many interactions with both the side chain of N341 and the 3’-thymine of CPD. The mutation of this residue may strongly disturb the water network, and make the binding pocket much looser. This is consistent with the observation the R342A has more than one order lower binding constant than wild type. With less restriction by the protein environment, the bound CPD can fluctuate
54
more flexibly. Thus, R342 requires less energy for the structure distortion during the charge transfer.
As pointed elsewhere, the active-site solvation plays a critical role in protein recognition and enzymatic reaction.12,74,75 To explore the difference between the ET models with and without the treatment of dynamic solvation, we fit the reaction rates and energies obtained from Sumi-Marcus model by using the classic Dutton’s equation of lg k A3.1(G )2 / , where A represents the ET rate of barrierless reaction.76,77
The free energies of electron return in inverted region are corrected as G ne by the excited vibrational energies of the high-frequency mode of the ground state, in which
ne is the effective quanta of excited vibrational states. The parameter A were fitted as
10.7 for forward ET and 10.5 for electron return, which are lower than those of 11.1 for forward ET and 11.0 for electron return determined by one-dimensional Marcus model in our recent studies,52 indicating that both forward ET and electron return are slowed down
by active-site solvation. The ne value was fitted as 1.9 and the similar quanta of excited vibrational states in backward ET reactions were observed in other model system recently.59
In Sumi-Marcus model, the population distribution of reactant along the solvent reorganization coordinate X can be diffused by the dynamic solvation. The reactant population at time zero is assumed as a thermal equilibrium at X 0 approximately.
When this thermal equilibrium distribution is perturbed, the reorientation of solvent molecules will attempt to restore it. During the ET reactions, the population at the
55
positive side reacts faster due to the lower activation barrier, which is clearly shown in
Figure 3.6B. If the solvation is much faster than the ET reaction, the thermal equilibrium of X is maintained and the dynamics of ET would be single-exponential decay.56 If the solvation is comparable or slower than ET, the distribution to reactant will move to the negative side of X coordinate due to the faster loss of positive-side population. Since the activation barrier of ET is increasing at the negative X coordinate, the ET process is slowed down by the slow active-site solvation.
To examine the dependence of the ET rate on the driving force and the reorganization energy, we plotted the wild-type and mutated photolyases along the coordinates of the corrected driving force and the total reorganization energy (Figure 3.8B). The contours of
ET rates are calculated from above fitting by Dutton’s equation. In the forward ET, the rates are mostly correlated with the driving force. The rates of ET is in a decreased order of M345A > WT > R226A > E274A > N378S > N378C, which is consistent with the order of the driving force. However, both the reorganization energy and driving force play important role in the electron return. Interestingly, R342A is out of the trend and its rates of the forward ET and electron return are much faster than the experimental values.
This inconsistency is resulted from that the Dutton’s equation does not take the solvation into account. Compared to wild type and other mutants, R342A has the much smaller
value of i and larger value of 0 . The larger ratio of 0 i causes the higher activation barrier and leads the slowdown of the ET reaction.56
56
3.3.4 Modulation of Driving Force and Solvent Reorganization by Active-site Residues. As mentioned above, the driving forces ( G ) for the forward ET are diverse among wild type and mutants, suggesting the modulation by active-site residues. At the binding pocket, salt bridges and hydrogen bonds are extensively formed between the
CPD and the amino acid residues: protonated E274 could form two hydrogen bonds with
N3 and O4 of the 5’-thymine; R226 has both electrostatic interaction and hydrogen bond with 5’-thymine; R342 interacts with 3’-thymine not only through salt bridge but also through the water network. These hydrophilic interactions could help stabilize the radical anion of the CPD after electron transfer from FADHˉ*. Thus, the mutations of the binding-site residues would diminish the stabilization and lead to the decrease in the reduction potential of T<>T/T<>Tˉ. Since these mutations do not affect the reduction
potential of flavin cofactor ( ), the free energy FADH / FADH
G ( G 2.48 ev) becomes less negative than that of FET FET FADH// FADH T T T T wild type. On the flavin cofactor side, N378 is opposite to the N5 position of flavin, and its side carbonyl may form a hydrogen bond with the N(5)H group of FADH or FADHˉ.
Destroying the interaction would abolish the ability of photolyase to stabilize the FADH radical.54 After purification, the flavin cofactors in N378C and N378S are in oxidized form FAD, in contrast to the radical FADH in wild type and other mutants. The FAD can be photoreduced to the fully reduced form FADHˉ in presence of external electron donors.
However, no accumulation of radical FADH were observed during the photoreduction, indicating that the formed FADH is very unstable and easy to go further to FADHˉ.
Additionally, FADHˉ in N378C or N378S is more difficult to be reoxidized compared to 57
wild type under aerobic conditions. These observations suggest that the mutation of N378 would increase the reduction potential of FADH/FADHˉ and leading to a less negative
GFET compared to wild type. M345 locates between the flavin and the 3’-thymine, and the sulfur atom on its side chain is in van der waals contact with the thymine ring. The sulfur-aromatic interactions have been extensively studied,78,79 and sulfur-containing amino acids were found to be week electron donors.80,81 The mutation of M345 could make the formed CPD anion radical more stable after forward ET, and result in the increase in reduction potential of T<>T/T<>Tˉ. Therefore, the free energy is more negative and the forward ET becomes more favorable.
The solvent reorganization is also manipulated by the protein environment. At either the CPD binding site or flavin site, electrostatic interactions and hydrogen bonds exist extensively between the polar/charged residues and polar solvent molecules. The mutation of these polar/charged amino acids to the nonpolar or smaller ones will destroy such interactions and make the surrounding solvent molecules more flexible.
Consequently, the solvent molecules reorient with more magnitude during the relaxation, leading to the larger solvent reorganization energies. Typically, the water network is violently disturbed at 3’-thymine side in mutant R342A, and the water molecules could be more flexible to reorganize in the relatively loosen pocket during the ET reactions.
Thus, R342A has a much larger 0 than others.
58
3.3.5 Ring Splitting and Back Electron Transfer. After forward ET from the excited state FADHˉ*, the excess negative charge makes the anionic radical T<>Tˉ really unstable and the C5-C5’ bond splits instantaneously. The formed T-Tˉ evolves along either the C6-C6’ bond splitting or the backward ET without repair. For the N5 position mutants (N378C and N378S), we observe the same second-bond splitting time in 90 ps as the wild type, in agreement with the fact that the mutation only affects the flavin cofactor. At the binding site, the mutation of the charged/ polar residues E274 and R226 at the 5’ side diminishes the stabilization of anionic radical, resulting in the much faster second bond-breaking time of 30 and 50 ps respectively.
Interestingly, R342A and M345A have the similar or the same time scales as the wild type. This observation suggested that the excess electron mainly remains at the 5’side after the C5-C5’ bond breakage, which is consistent with our recent study.52
Besides, we observed the shorter time scales of the backward ET for all six mutants compared to the wild type (2.4 ns). For N378C or N378S, the faster backward ET process is mainly caused by the modulation of redox potential of the flavin cofactor. Since both the forward ET and backward ET are in the Marcus normal region (Figure 3.9B), the slower forward ET results in the faster backward ET. At the binding site, with the strong hydrophilic interactions with T-Tˉ, the residues of E274, R226 and R342 can delocalize the excess negative charge and stabilize the anionic radical. The mutations of these residues will destroy the stabilization and make the backward ET much faster.
59
3.3.6 Maximization of The Quantum Yield. The total repair quantum yield is the product of the efficiencies of two steps: the forward ET bifurcation and the complete splitting bifurcation. As shown in Figure 3.9A, the total quantum yields (grey columns) change in a decreased order of WT > M345A >
N378C > N378S > R226A > R342A > E274A. In the first step, the forward ET competes with the lifetime. Both the binding-site mutants (E274A, R226A and R342A) and N5 position mutants (N378C and N378S) modulate the ET redox potentials, the binding-site mutants for the CPD potential and N378 mutants for the flavin potential, leading to the slower forward ET and thus resulting in the lower efficiencies for the first step. M345A is the only exception that has faster forward ET than the wild type, therefore it has the highest quantum yield (0.89) for the first step. However, M345A has a much faster backward ET, which results in a big loss in the second step quantum yield. Thus, the total quantum yield of M345A is still lower than the wild type. Similar to the wild type, both
N378C and N378S have much slower backward ET compared to the second-bond breakage, leading to the nearly unity of second-step efficiency. For the binding-site mutants, both the second-bond splitting and the backward ET become faster due to the destabilization of anionic radical, but the effect of the faster backward ET dominates and thus causing the decrease in the second-step quantum yield. The total quantum yields of the N378 mutants are mainly lost in the first step while the quantum yield loss of the binding-site mutants result from the combination of both steps.
60
3.4 Conclusions Here we report our systematic study of the ultrafast dynamics of DNA repair by photolyase. With femtosecond-resolved laser spectroscopy, we mapped out the complete evolution of functional dynamics with 6 active-site mutated photolyase in real time. The mutation of the active-site residues results in the significant change in the repair dynamics. We also analyzed the electron transfer processes in the catalytic reaction with
Sumi-Marcus model. The results suggest that photolyase controls the electron transfer and ring-splitting of CPD through the modulation of the redox potential, reorganization energies and stabilization of the anionic intermediates by the interactions form its active-site residues, balancing all the reaction steps and maximizing the enzyme activity.
61
b c c Φ β τlifetime <τFET> ΦFET <τSP> <τBET> ΦSP <τER> WT 0.82 0.71 1300 236 0.85 88 2400 0.96 625 M345A 0.72 0.71 1169 140 0.89 88 368 0.81 200 N378C 0.67 0.71 3374 1181 0.74 88 839 0.91 500 N378S 0.62 0.71 1351 675 0.67 88 1242 0.93 462 R226A 0.53 0.71 1309 480 0.73 50 126 0.72 1412 R342A 0.48 0.64 1600 595 0.73 83 166 0.66 416 E274A 0.38 0.71 1044 615 0.62 31 52 0.62 75
Table 3.1 Results of reaction times, efficiencies of the elementary steps and overall repair quantum yields of wild-type and mutant photolyasesa aAll times are in unit of picosecond. Here β is the stretched parameter and Φ is the overall repair quantum b yield of wild-type and mutant photolyases. <τFET> is the forward ET time calculated from 1 1 [( )( )1 ]1 . τ is the observed timescale of FADH−* decay with substrate probed at 800 nm lifetime c and τlifetime is that in absence of substrate. ΦFET and ΦSP are efficiencies of the forward ET and the ring -1 -1 -1 -1 -1 -1 splitting and calculated by ΦFET = <τFET> /(<τFET> + τlifetime ) and ΦSP = <τSP> /(<τSP> + <τBET> ), respectively. Thus, Φ = ΦFET x ΦSP.
62
b ΔGFET λo, FET λi, FET λo, ER λi, ER ћω WT -0.440 0.226 0.840 0.395 0.840 0.165 M345A -0.510 0.245 0.840 0.455 0.840 0.170 N378C -0.310 0.255 0.860 0.490 0.880 0.168 N378S -0.365 0.250 0.845 0.475 0.850 0.170 R226A -0.383 0.245 0.830 0.400 0.820 0.162 R342A -0.380 0.365 0.650 0.690 0.680 0.174 E274A -0.379 0.245 0.870 0.640 0.880 0.174
Table 3.2 The energies in electron transfer reactionsa aAll energies are in unit of eV. bω is the frequency of the high-frequency vibrational mode of the product, and the quantum number of the high-frequency mode is 3 for wild type and all mutants.
63
Figure 3.1 Enzyme-substrate complex structure of photolyase
(A) X-ray structure of A. nidulans photolyase (light-grey ribbon) containing the catalytic FADHˉ (orange stick) complexed with a repaired CPD lesion. E. coli photolyase has a similar structure. Five critical conserved residues in the active-site are R232, E283, M353, R350 (R226, E274, M345 and R342 in E. coli photolyase) near the substrate, and N386 (N378 in E. coli photolyase) near the cofactor. (B) Diagram of the interactions between the CPD lesion and the active-site residues. Distances are given in angstrom. .
64
Figure 3.2 Schematic representation of the hypothetical dynamic process in DNA photorepair
Shown in the repair scheme are lifetime emission (LT, reaction rate kL) of excited FADHˉ* in the absence of substrates, forward electron transfer (FET, reaction rate kFET) from excited FADHˉ* to thymine dimer upon light excitation, followed by backward electron transfer (BET, reaction rate kBET) without repair, and the repair channel including the sequential splittings of C5-C5’ bonds (SP1, reaction rate kSP1) and C6-C6’ bond (SP2, reaction rate kSP2) in thymine dimer with subsequent electron return (ER, reaction rate kER) from repaired thymine to neutral FADH.
65
Figure 3.3 Determination of dissociation constants and CPD repair quantum yields by mutant photolyase
(A) The relative repair quantum yields of CPD by mutant photolyases were measured by monitoring the formation of thymine bases at 266-nm absorbance with certain visible light irradiation of enzyme-substrate solution. With the knowledge of the dissociation constants for wild-type and mutant photolyases and the repair quantum yields of 0.82 for wild type, the repair quantum yields for each mutant were obtained. (B) The binding percentages of enzyme-substrate complex were measured for each wild type or mutant with different substrate concentrations. The dissociation constants were obtained by fitting the titration curve with the equation [ES]/[E]=[S]/([S]+Kd).
66
Figure 3.4 Femtosecond-resolved transient absorption dynamics of CPD repair by mutant photolyases of R226A, N378S and M345A
The repair dynamics are probed systematically from 800 to 266 nm and the typical transients in UV region with distinct patterns are deconvoluted by total flavin-related species (dashed green), substrate intermediates of Tˉ (dashed dark blue) and T-Tˉ (dashed blue), and thymine products (dashed dark yellow).
67
Figure 3.5 Femtosecond-resolved transient absorption dynamics of CPD repair by mutant photolyases of E274A, R342A and N378C
The repair dynamics are probed systematically from 800 to 266 nm and the typical transients in UV region with distinct patterns are deconvoluted by total flavin-related species (dashed green), substrate intermediates of Tˉ (dashed dark blue) and T-Tˉ (dashed blue), and thymine products (dashed dark yellow).
68
Figure 3.6 Schematic representation of ET coupled with solvation substrates
(A) Free energy curves of both the forward ET from FADHˉ* to CPD and the electron return from Tˉ to FADH along the solvent reorganization coordinate, where the free energy change (ΔGFET and ΔGER+neћω), solvent reorganization energies (λo,FET and λo, ER), and solvation energy (ΔE) are shown. neћω is the energy of the excited vibrational states of high-frequency mode, in which ne is the effective quantum number, ћ is the reduced Planck constant and ω is the angular frequency of high-frequency mode. (B) Free energy surface and its contour which represent the reactants and products of the electron return from Tˉ to FADH. Upon the charge separation the reactants relax along the intramolecular distortion coordinate instantaneously, and relax along the solvent reorganization coordinate on the time scale of ET. The activation barriers of ET at different solvent coordinates are different and therefore the reacted populations are different.
69
Figure 3.7 Analysis of the forward ET and the electron return by Sumi-Marcus model
(A) The ultrafast dynamics of the forward ET probed at 800 nm. All the transients are fitted by the Sumi-Marcus ET model. (B) The ultrafast dynamics of the electron return simulated by the fitting parameter from stretched-exponential model. All the transients are fitted by the extended Sumi-Marcus ET model.
70
Figure 3.8 The energies and rates of ET reactions in wild-type and mutated photolyases
(A) The driving forces and solvent reorganization energies are diverse, while the excited vibrational energies from the high-frequency mode and the intramolecular distortion energies are invariant except R342A. (B) The correlation among the rates of ET, driving force and reorganization energy. In the forward ET, the rates are mostly correlated with the driving forces. The rates of ET is in a decreased order of M345A > WT > R226A > E274A > N378S > N378C, which is consistent with the order of the driving forces. In the electron return, the rates of ET are influenced by both the driving force and the reorganization energies.
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Figure 3.9 Quantum yields, reaction times and free energy diagram of the elementary steps in the repair of CPD
(A) The reaction times of each elementary step observed in various wild-type and mutated photolyases are plotted in the upper panel. The vertical dashed lines represent the two bifurcations of the forward ET (FET) competing with lifetime emission (LT) and the second-bond splitting (SP2) competing the direct backward ET (BET). The total quantum yields (grey columns) are determined by the combination of the efficiencies of two bifurcations. (B) The free energy profile along the reaction coordinate upon excitation. On the neutral surface, the bond-breaking activation barrier (dashed curve) is very high according to the theoretical calculations. Note the different regions, normal and inverted, of three ET processes and the ring reclosure after the futile backward ET.
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CHAPTER 4 ULTRAFAST PHOTOREDUCTION OF FLAVIN COFACTORS IN (6-4) PHOTOLYASE
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CHAPTER 4
ULTRAFAST PHOTOREDUCTION OF FLAVIN COFACTORS
IN (6-4) PHOTOLYASE
4.1 Introduction Photolyase, a photoenzyme which contains the anionic hydroquinone flavin adenine dinucleotide (FADHˉ) as the catalytic cofactor, is responsible for repairing the
UV-damaged DNA (CPD and 6-4PP) in many organisms.10 While keeping the active hydroquinone state in vivo, the flavin cofactor is converted to neutral semiquinone
FADH or fully oxidized quinone FAD during the purification in vitro and loses the enzyme activity. By absorption of light in the visible range, both FADH and FAD undergo a photoreduction in the presence of reducing agents to yield the catalytically active form FADHˉ.10,12,29,82-84
The photoreduction process in photolyases has been studied quite extensively. Upon light excitation, the excited FADH and FAD are readily to be reduced via electron transfer (ET) from the neighboring aromatic residues. In E. coli CPD photolyase, a conserved tryptophan residue W306 was identified as the terminal electron donor in the photoreduction by the site-directed mutagenesis study.84 W306 was found to be near the protein surface, however, more than 14 Å away from the flavin moiety of FAD.25 Such a large distance excludes a direct or superexchange-mediated ET on a picosecond (ps) time
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scale.77 Therefore, the ET was proposed to mainly go through a triple tryptophan chain
(W382-W359-W306) via the hopping mechanism.25 Since these three tryptophans are evolutionary conserved throughout the photolyase/cryptochrome family, including CPD photolyases and (6-4) photolyases,10 as well as cryptochromes,85,86 the detailed dynamics and mechanism of the ET process in photolyase photoreduction might be not only important for the understanding of the photoactivation of photolyase, but also relevant for the examination of the reaction mechanisms of the cryptochrome blue-light photoreceptors.87
To reveal entire dynamics and mechanism of this interesting ET process, we examined the photoreduction of oxidized quinone FAD and neutral semiquinone FADH in Arabidopsis thaliana (6-4) photolyase for the first time with integration of site-mutagenesis and femtosecond-resolved transient absorption spectroscopy. With more than ten-wavelength detection of various photolyae mutants in visible region, we were able to identify that five tryptophan residues as well as the adenine moiety involve in the photoreduction (Figure 4.1) and solve the dynamics of each elementary step. Upon light excitation, the flavin cofactor FAD and FADH are reduced by the adenine moiety and the inner residue (W406) of the conserved tryptophan triad (W406-W383-W329) in less than 1 ps and tens of ps, respectively. Subsequently, the tryptophan triad delivers the electron, via the multi-step hopping mechanism, through a long-distance over 13 Å in hundreds of ps. The W329, which is proximal to aqueous surface of the protein,26 finally stays at radical state for longer than tens of nanoseconds (ns). Based on the empirical
Marcus ET equation, we evaluated how the free energy and reorganization energy
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influence the rate of each ET step. Interestingly, the results show that the tryptophan triad has a reduction potential gradient modulated by the local protein environment, which promotes the electron hopping process with a distinctive directionality.
4.2 Materials and Methods
4.2.1 Preparation of (6-4) Photolyase and Mutants. The sample of Arabidopsis thaliana (6-4) photolyase with an amino-terminal His tag was prepared as described with modification.88 Escherichia coli BL21 (DE3) cell transformed with the vector was grown at 37 ºC in LB medium to OD600 of 1.7. The culture was then cooled down to 25 ºC, and IPTG was added to 0.2 mM. The culture was further incubated for 16-18 hours and then harvested by centrifugation. After harvest, the cell pellet was frozen at -80 ºC. During the protein purification, the cell pellet was thawed and resuspended in a lysis buffer (50 mM Tris-HCl, 300 mM NaCl and 10% (v/v) glycerol, pH=7.4). After the sonication, the cell debris was removed by ultracentrifugation at 20,000 rpm for 1.5 hours. The cell-free extract was loaded onto a
Ni-NTA column (GE) and the fusion protein was eluted with elution buffer (50 mM
Tris-HCl, 300 mM NaCl, 500 mM imidazole and 10% (v/v) glycerol, pH=7.4). Then the eluted sample was applied to a Hi Trap Heparin HP column (5 ml) and eluted with a linear gradient of 0.2-1 M NaCl. The protein was dialyzed against the reaction buffer (50 mM Tris-HCl, 100 mM NaCl, 1 mM EDTA and 50% (v/v) glycerol, pH=7.4) and stored in -80 ºC. To prepare the photolyase with the oxidized FAD cofactor, the sample was incubated in the elusion buffer at 4 ºC for 24-48 hours before loading onto the Hi Trap
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Heparin HP column. For the mutant studies, three tryptophan residues (W406, W383 and
W329) were mutated with phenylalanine. All the mutant plasmids were constructed using
QuikChange II XL kit (Stratagene) based on the plasmid of wild-type enzyme, and were sequenced to ensure correct results.
4.2.2 Femtosecond-resolved Spectroscopy. All the femtosecond-resolved measurements were carried out using transient absorption methods. The laser experimental layout and procedure have been detailed elsewhere.35 The excitation wavelengths were 480 nm for the photolyase with oxidized
FAD and 640 nm for that with semiquinone FADH, respectively. The instrument response time is about 250 fs and the experiments were done at the magic angle (54.7º).
5-mm quartz cuvettes (Starna) were used as the sample cell in experiments and samples were kept stirring during irradiation to avoid heating and photobleaching. All experiments in the femtosecond-resolved measurements were carried out under aerobic conditions without any additional reducing agents.
4.3 Results and Discussion
4.3.1 Direct electron transfer between adenine and flavin moiety in oxidized FAD. To dissect the complicated electron hopping process into individual steps, we prepared three single-position mutants of W406F, W383F and W329F. The phenylalanine is redox-inertial, thus the mutation of tryptophan with phenylalanine will block the ET pathway. Figure 4.2 shows four striking pattern of the transient absorption signals of
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photoreduction of oxidized FAD in W406F, W383F and W329F mutants and wild-type photolyase, with probing ten visible wavelengths from 800 nm to 450 nm.
With the mutation of the nearby tryptophan residue (W406) in the tryptophan triad, we blocked all potential ET channels of flavin from tryptophan or tyrosine within 7 Å. At
800 nm, we detected the signal of excited FAD* following a stretched-single-exponential decay (dashed pink in Figure 4.3A), Ae (t / ) , with the time constant τ=8 ps and the stretched parameter β=0.9. Besides, there is a very minor component of hundreds of picoseconds left. Without aromatic residues in proximity, excited FAD or FMN in inert protein environments, as well as FMN* in solution, have a lifetime of several nanoseconds.89-91 Therefore the radiative lifetime of FAD* in photolyase is expected to be on the similar time scale, but it occurs in a much faster manner. Notably in photolyase,
FAD has an unusual U-shaped structure with the isoalloxazine ring and adenine moiety in close proximity. As recently reported, excited FAD in solution exhibits an intramolecular
ET in 9.0 ps with a similar stack conformation between the flavin and adenine moiety.92
Thus, the observed quenching dynamics of FAD* in 8 ps represents the ultrafast intramolecular ET with formation of a charge separated pair of adenine+ and lumiflavin−
(Ade+ and Lf−). Moreover, the stretched-single-exponential decay behavior reflects the modulation of ET by the binding-site solvation. Base on equation 1,
1 ( ) (4.1) we obtained the average charge separation time scale <τ>=8.4 ps.
To capture the intermediate of cationic adenine radical, the probe wavelength was further tuned to 580 nm. Strikingly, we observed the stimulated emission signal of FAD* 78
recovering in 8 ps (dashed pink in Figure 4.3B) and the Ade+ signal with a rise-decay behavior (dashed dark yellow in Figure 4.3B). The rise indicates the formation of Ade+ on the same time scale of the charge separation. With the same stretched β value of 0.9, we obtained the time constant of 250 ps for the decay of Ade+, which represents the charge recombination between Ade+ and Lf−. It is very important for the finding of this intromolecular ET of FAD in protein, which implies that the adenine moiety can involve the photocycle of photolyase/cryptochrome family by electron hopping between two moieties in stack conformation.
Additionally, we detected another species at the wavelengths shorter than 680 nm
(typically dashed blue in Figure 4.3C). According to the crystal structure (Figure 4.1), there is a tryptophan residue (W301) stacked proximal to the adenine moiety with a short distance of 4.29 Å. This W301 can donate an electron to the cationic adenine radical, to form the cationic tryptophanyl radical (W301+). By fitting with the sequential reversible reaction model:
Lf *Ade W301Lf Ade W301Lf Ade W301
Lf Ade W301 (4.2) we obtained the time constant of 1.5 ns for the hopping from W301 to Ade+, and longer than 10 ns for the back hopping from adenine moiety to W301+. Based on the relative extinction coefficient ratio of W301 radical to Ade+ obtained by fitting and the absorption of Ade+ in solution, the absorption of detected W301 radical has a peak at about 520 nm, which matches the absorption of deprotonated tryptophanyl radical rather than that of cationic tryptophanyl radical (absorption peak at 580 nm) in solution.93,94 79
This absorption characteristic implies that the formed W301+ might go through deprotonation and stay at the W301 state.
4.3.2 Electron flow from the nearby tryptophan. Besides adenine, W406 is the only aromatic residue nearby the flavin ring within 7 Å.
To examine the ET dynamics between W406 and oxidized FAD, we mapped out the dynamics of W383F from 800 nm to 450 nm (Figure 4.2B). At 800 nm, we observed an ultrafast decay of FAD* within less than 1 ps, which is much faster than that of W406F
(Figure 4.2A) but similar to W329F and WT (Figure 4.2C and D). The FAD* is quenched by the electron transfers both from adenine moiety and W406. With the knowledge of quenched dynamics by adenine moiety, we deconvoluted the forward ET from W406 to lumiflavin in 0.45 ps. Such ultrafast electron hopping process is mainly caused by the
73,95 favorable driving force of ET ( ). With 95% possibility to form the W /W Ade / Ade tryptophanyl radical W406+, the tryptophan triad is the major ET channel during the photoreduction.
To reveal the further electron flow after charge separation, we examined the absorption transients of W383F at 580 nm (Figure 4.3 F), which is reported as the absorption peak of the cationic tryptophanyl radical.93 Besides the FAD* dynamics
(dashed pink) and trace signal from Adenine ET channel (dashed dark yellow), we captured the strong signal of W406+ (dashed dark red) with a rise-decay behavior. This decay takes 45 ps and is attributed to the charge recombination between W406+ and Lf−.
It is much slower than its counterpart in flavodoxin, and therefore enhances the efficiency
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of photoreduction.
Interestingly, we detected another intermediate with a rise of 5 ps (dashed blue in
Figure 4.3E and F) which is unable to be fitted by the ET processes described above.
Since the nearby W383 was mutated, the remaining possible candidate involving ET flow is W339 which is 3.67 Å away from W406 (Figure 4.1).26 The long distance of 8.76 Å between W339 residue and lumiflavin ring excludes the possibility of direct ET from
W339 to excited lumiflavin in several picoseconds. Thus, this rise component is attributed to the electron hopping between W406 and W339 residues. Based on the dynamics of FAD* and three radical intermediates described above, and the extinction coefficients of Trp+, Ade+, Lf− and oxidized FAD, we unambiguously determined the forward electron hopping from W339 to W406+ occurs in 40 ps while the back ET in the opposite direction completes in 5 ps. However, due to the lack of further ET channels toward the protein surface, the ETs between the W339 and W406 residues make no contribution to the photoreduction.
4.3.3 Electron hopping among the conserved tryptophan triad. To test the role of middle tryptophan W383, we prepared the mutant W329F which keeps the W406/W383 pair intact but excludes any other potential electron donor/acceptor within 5 Å. Although the quenching dynamics of FAD* is similar to that of W383F at 800 nm (insets in Figure 4.2B and C), the signals from tryptophanyl radicals are very different in the longer time window. At 580 nm, we detected the signal of
W383+ with a rise of tens of ps and a decay of hundreds of ps (Figure 4.4). Since the
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W383 moiety is over 8.7 Å away from the flavin moiety, we would expect the much longer time scale than hundreds of ps for the direct charge recombination between
W383+ and Lf−. Base on the dynamics obtained from W383F, we determined the forward electron hopping from W383 to W406+ occurs in 30 ps while the back electron hopping fromW406 to W383+ takes 460 ps. The much slower back hopping rate implies that W383 is easier to lose an electron than W406. As observed in previous study on photoreduction of FADH and our recent study on photoreduction of FAD in E. coli CPD photolyase, the cationic radical of the middle tryptophan W359+ is deprotonated on nanosecond time scale and finally converts to neutral radical W359.96 However, in our study, we only can detect trace signal of W383 due to the much slower deprotonation
(longer than 30 ns).
With the knowledge of the obtained ET dynamics in mutants, we mapped out the whole process of photoreduction of FAD in wild-type (6-4) photolyase. Unlike the
W329F mutant, the cationic radical W383+ can be further reduced by the final electron donor W329 instead of back electron hopping and deprotonation. The generated W329+, which is on the surface of protein, will gain an electron from the solution and finish the photoreduction process. Compared to W329F, the transient signal of WT at all probed wavelengths shows a long plateau (Figure 4.2D) rather than a decay in hundreds of ps
(Figure 4.2C). This difference is mainly attributed to the formation of long-lived W329+.
Based on the acquired dynamics in mutants, we determined the forward electron hopping from W329 to W383+ takes 350 ps with comprehensive fitting (Figure 4.5). Moreover, the long component signal of W329+ has no obvious decay behavior, indicating that the 82
back electron hopping is longer than our detection time window (4 ns). We estimated a lower limit of 10 ns.
4.3.4 Ultrafast photoreduction of FADH. To make a complete work, we also studied the photoreduction dynamics of FADH.
Following the same strategy as oxidized FAD study, we dissected the complex process into individual steps, and measured the dynamics of each step by using mutant photolyases (Figure 4.6). In W406 mutant, all the direct ET channels of FADH from nearby tryptophan or tyrosine residues are blocked. We observed that the adenine moiety donates an electron to the excited FADH in 150 ps while the charge recombination between Ade+ and FADHˉ takes 60 ps (Figure 4.7A, B and C). Similar to the FAD study, we also detected the bifurcation ET channel from W301 to Ade+ in 500 ps. By mutation of W383, we acquired the forward ET from W406 to excited FADH in 40 ps and the back ET from FADHˉ to W406+ in 4 ps (Figure 4.7D, E and F). Since there is little accumulation of W406+ due to the slow formation and fast decay, we did not detect obvious signal of W339+. In the following, we determined the time constants of 3 ps for the electron hopping from W383 to W406+ and 500 ps for the back electron hopping in mutant W329F (Figure 4.8), and 350 ps for the next electron hopping from W329 to
W383+ and longer than 10 ns for the reverse reaction in wild-type photolyase (Figure
4.9).
In summary, both photoreductions of oxidized FAD and semiquinoid FADH utilize the conserved tryptophan triad as the major electron transfer channel while the adenine 83
moiety also involves in the process. The complete dynamics for each flavin state are described in Figure 5.10A&B, respectively. Due to the influence of flavin redox potentials, all the ET dynamics involved the flavin species are very different in the two photoreduction schemes. In contrast, the two photoreductions share the same electron hopping rates between W329 and W383 because the W383 and W329 are far away from the flavin ring and the redox status of flavin has little impact on them. However, the electron hopping fromW383 to W406+ in photoreduction of FADH is much faster than in that of FAD, indicating that the redox property of the nearby tryptophan W406 is modified by the flavin.
4.3.5 Analysis of the ET networking by Marcus model. The photoreduction of the flavin cofactor in (6-4) photolyase involves in more than
10 individual ET steps, and the conserved tryptophan is identified as the major ET channel. To ensure the high efficiency of the photoreduction, the electron hopping rates from W329 to flavin must exceed those of side reactions. Strikingly, the forward electron hopping from W329 to W383+ and from W383 to W406+ are much faster than the back electron hopping in the reverse direction, which implies a significant variation of redox properties of these chemically-identical tryptophan residues. To get more understanding how the protein modulates the ET in the photoreduction, we analyzed the electron hopping along the tryptophan triad as well as the ETs between the lumiflavin and the adenine moiety using the empirical Marcus ET equation,77,97
2 log kET 13 0.6(r r0 ) 3.1(G ) / (4.3)
-1 where kET is the ET rate in s , r is the edge-to-edge distance in Å, ΔG is the free energy 84
in eV, λ is the reorganization energy in eV. The coefficient of 0.6 reflects the empirical distance-dependence of electron tunneling rate in protein and r0 is the van der Waals distance at 3.0 Å. With the solved X-ray crystal structure of the protein, we obtained the edge-to-edge distance between various electron donors and acceptors and further the
terms related to the electron coupling ( 0.6(r r0 ) ). Thus, the ET rate of each step is mostly determined by the Franck-Condon factor, which consists of two parameters of driving force (-ΔG) and reorganization energy.
First, we analyzed the ETs in the photoreduction of the oxidized FAD. For the electron hopping between the W329 and W383 residues, the driving forces for the forward and reverse reactions have the same magnitude but with different sign
( G G ), while the reorganization energy are forward W 329 /W 329 W 383 /W 383 backward identical in both steps. By applying r=3.41 Å, we obtained the ΔG=-0.12 eV for the forward electron hopping from W329 to W383+ and ΔG=0.12 eV for the back hopping with an identical reorganization energy λ=1.29 eV for both reactions. This indicates that the reduction potential of W329+/W329 is 0.12 V lower than that of W383+/W383.
Following the same strategy, we acquired the reduction potential gradient of 0.10 V from
W406+/W406 to W383+/W383 and the reorganization energy of 0.58 eV for the hopping between W383 and W406. For the bifurcation hopping between W339 and W406, the reduction potential of W339+/W339 is 0.07 V higher than that of W406+/W406 and the reorganization is 0.55 eV. Clearly, there is a reduction potential degression along the tryptophan triad which promotes the electron flow from W329 to W406. According to the crystal structure,26 W329 is at the loop region and exposed to the solution while the
85
W383 and W406 are buried inside the protein. The high dielectric environment and the direct interaction with bulk water could help to stabilize the formed cationic radical
W329+ and therefore lead to the lowest reduction potential of W329+/W329.98,99
Compared to W406, W383 has more hydrophilic residues surrounded and a structured water molecule within 2.9 Å,26 which results in the lower reduction potential. In our recent study on the photoreduction of FAD in E. coli photolyase, we obtained the reduction potential of 1.56 V for W382+/W382. Since the protein environmenst of flavin-binding site in both E. coli CPD photolyase and A. thaliana (6-4) photolyase are highly homologous, it is reasonable to assume the W406+/W406 has the same reduction potential as its counterpart. Then we can get the reduction potentials of 1.46 V for
W383+/W383, 1.34V for W329+/W329, and a highest value of 1.63 V for
W339+/W339. Moreover, the much larger reorganization energy of the hopping process between W329 and W383 is probably due to the significant conformation change of
W329 on the flexible loop rather than W406 and W383 constrained by the tight hydrophobic environment.
We further examined the driving forces and reorganization energies of ET processes involving flavin moiety. Since the electron acceptor is at the excited state, the free energy can be expressed as following,
Gh (4.4) forward Donor// Donor Acceptor Acceptor
G (4.5) backward Acceptor// Acceptor Donor Donor where the h is the S1←S0 transition energy (-2.58 eV at 480 nm). The reduction potential of oxidized FAD was reported as -0.3 V.100 Considering that the backward ET 86
has a larger reorganization energy than the forward ET, we analyzed all the ET rates related to the flavin and obtained the free energies and reorganization energies of all charge separation and recombination processes. The driving forces of charge separation between adenine/W406 and the excited flavin are 0.32 and 0.72 eV while those of corresponding charge recombination are 2.26 and 1.86 eV (Table 4.1). The acquired reduction potential of 1.96 V for Ade+/Ade is exactly matched the reported value in nonpolar solvent.73 The reorganization energies of these processes vary from 0.9 to 1.2 eV (Table 4.1), close to the reported values in E. coli photolyase.34
Following the same procedure, we analyzed the ET processes in the photoreduction of the oxidized FADH. The obtained free energies and reorganization energies are listed in Table 4.2. The tryptophan triad shows a similar pattern of reduction potential degression. Assuming the reduction potential of W329+/W329 has the same value as that in the photoreduction of FAD, we get the reduction potentials of 1.63 V for
W406+/W406, 1.46 V for W383+/W383 and 1.34 V for W329+/W329. Compared to the
FAD study, the larger value of W406+/W406 is probably resulted from the influence of the radical FADH. In addition, the reorganization energies are similar to those in the photoreduction of FAD.
4.4 Conclusions We report our systematic study on the ultrafast photoreduction dynamics of flavin cofactors in Arabidopsis thaliana (6-4) photolyase. With femtosecond-resolved laser spectroscopy and the site-mutagenesis methods, we revealed the dynamics of electron transfer from the adenine and a neighbor tryptophan to the oxidized flavin (FAD) and 87
neutral semiquinone (FADH) and electron delivery along the conserved tryptophan triad.
Upon light excitation, the excited oxidized FAD and neutral semiquinone FADH are reduced by the nearby W406 and adenine moiety in less than 1 ps and 30 ps, repectively.
Subsequently, the conserved tryptophan triad transported an electron, via the multistep hopping mechanism, through a long-distance over 12 Ǻ in a few hundred picoseconds.
The electron hopping process has a distinctive directionality toward the flavin due to a unique reduction potential gradient along the tryptophan triad, which is modulated by the local protein environment to maximize the photoreduction efficiency.
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64PL-FAD τ (ps) ΔG (eV) λ (eV) FAa 8 -0.32 1.03 BAa 250 -2.26 1.19 FETb 0.45 -0.72 0.93 BETb 45 -1.86 1.02 HOP1c 30 -0.10 0.58 BHPO1c 460 0.10 0.58 HOP2d 350 -0.12 1.29 BHOP2d 10000 0.12 1.29 F339e 40 0.07 0.55 B339e 5 -0.07 0.55
Table 4.1 Results of reaction times, free energies and reorganization energies of the individual electron transfer steps in the photoreduction of FAD aFA and BA are the charge separation and recombination between the excited lumiflavin and adenine moiety. bFET and BET are the forward electron transfer from W406 to FAD* and the backward electron transfer in the reverse direction. cHOP1 and BHOP1 are the forward electron hopping from W383 to W406+ and the backward electron hopping in the reverse direction. dHOP2 and BHOP2 are the forward electron hopping from W329 to W383+ and the backward electron hopping in the reverse direction. eF339 and B339 are the forward electron hopping from W339 to W406+ and the backward electron hopping in the reverse direction.
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64PL-FADH τ (ps) ΔG (eV) λ (eV) FAa 150 0.01 0.88 BAa 60 -1.95 1.05 FETb 40 -0.20 1.04 BETb 4 -1.74 1.11 HOP1c 3 -0.18 0.35 BHPO1c 500 0.18 0.35 HOP2d 350 -0.12 1.29 BHOP2d 10000 0.12 1.29
Table 4.2 Results of reaction times, free energies and reorganization energies of the individual electron transfer steps in the photoreduction of FADH aFA and BA are the charge separation and recombination between the excited lumiflavin and adenine moiety. bFET and BET are the forward electron transfer from W406 to FADH* and the backward electron transfer in the reverse direction. cHOP1 and BHOP1 are the forward electron hopping from W383 to W406+ and the backward electron hopping in the reverse direction. dHOP2 and BHOP2 are the forward electron hopping from W329 to W383+ and the backward electron hopping in the reverse direction.
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Figure 4.1 Electron transfer network of FAD in A. thaliana (6-4) photolyase
The flavin moiety (green sticks) of FAD is the major electron acceptor. The adenine moiety and residue W406 (cyan sticks) are nearby and have direct electron transfer with flavin. The tryptophan residues W301, W339 and W383 (marine blue sticks) are the secondary electron donors among which W301 donates an electron to cationic adenine while W339 and W383 have direct electron transfer with W406. The W329 (purple sticks) is the final electron donor along the tryptophan triad. All the electron hopping distances are shown.
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Figure 4.2 Femtosecond-resolved transient absorption dynamics of photoreduction of FAD in mutant and wild-type (6-4) photolyases
The photoreduction dynamics of FAD in various mutant and wild-type (6-4) phtolyases (W406F, W383F, W329F and WT) are probed systematically from 800 to 450 with distinct patterns. The insets show the transient dynamics probed at 800 nm on short time scale.
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Figure 4.3 Femtosecond-resolved dynamics of direct electron transfer to FAD in (6-4) photolyase
Left column shows the normalized transient absorption signals of W406F mutant probed at 800 nm (A), 580 nm (B) and 500 nm (C), respectively, with deconvolution of flavin species (dashed pink), cationic adenine radical (dashed dark yellow) and neutral W301 radical (dashed blue). The inset shows the transient absorption signal of W406F probed at 500 nm with a longer time window. The long plateau indicates the long-lived neutral W301 radical. Right column shows the normalized transient absorption signals of W383F mutant probed at 800 nm (D), 580 nm (E) and 500 nm (F), respectively, with deconvolution of flavin species (dashed pink), cationic adenine radical and neutral W301 radical (dashed dark yellow), cationic W406 radical (dashed dark red) and cationic W339 radical (dashed blue).
93
Figure 4.4 Femtosecond-resolved dynamics of FAD photoreduction involving W383 in (6-4) photolyase
Normalized transient absorption signals of W329F mutant probed at 580 nm (A), 500 nm (B) and 450 nm (C) are deconvoluted by flavin species (dashed pink), cationic W406 and Adenine radical with neutral W301 radical (dashed dark red), cationic W383 radical (dashed cyan), neutral W383 radical (dashed dark orange) and cationic W339 radical (dashed dark green).
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Figure 4.5 Femtosecond-resolved dynamics of FAD photoreduction involving W329 in (6-4) photolyase
Normalized transient absorption signals of wild-type photolyase probed at 580 nm (A), 500 nm (B) and 450 nm (C) are deconvoluted by flavin species (dashed pink), cationic W406 and Adenine radical with neutral W301 radical (dashed dark red), cationic and neutral W383 radical (dashed cyan), cationic W329 radical (dashed dark orange) and cationic W339 radical (dashed dark green).
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Figure 4.6 Femtosecond-resolved transient absorption dynamics of photoreduction of FADH in mutant and wild-type (6-4) photolyases
The photoreduction dynamics of FADH in various mutant and wild-type (6-4) phtolyases (W406F, W383F, W329F and WT) are probed systematically from 800 to 450 with distinct patterns.
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Figure 4.7 Femtosecond-resolved dynamics of direct electron transfer to FADH in (6-4) photolyase
Left column shows the normalized transient absorption signals of W406F mutant probed at 800 nm (A), 580 nm (B) and 500 nm (C), respectively, with deconvolution of flavin species (dashed pink), cationic adenine radical (dashed dark yellow) and neutral W301 radical (dashed blue). Right column shows the normalized transient absorption signals of W383F mutant probed at 800 nm (D), 580 nm (E) and 500 nm (F), respectively, with deconvolution of flavin species (dashed pink), cationic adenine radical and neutral W301 radical (dashed dark yellow), cationic W406 radical (dashed dark red) and cationic W339 radical (dashed blue).
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Figure 4.8 Femtosecond-resolved dynamics of FADH photoreduction involving W383 in (6-4) photolyase
Normalized transient absorption signals of W329F mutant probed at 580 nm (A), 450 nm (B) and 405 nm (C) are deconvoluted by flavin species (dashed pink), cationic Adenine radical and neutral W301 radical (dashed dark red), cationic W339 radical (dashed dark green), cationic W383 radical (dashed cyan) and neutral W383 radical (dashed dark orange).
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Figure 4.9 Femtosecond-resolved dynamics of FADH photoreduction involving W329 in (6-4) photolyase
Normalized transient absorption signals of wild-type photolyase probed at 580 nm (A), 450 nm (B) and 405 nm (C) are deconvoluted by flavin species (dashed pink), cationic Adenine radical and neutral W301 radical (dashed dark red), cationic W406 radical (dashed dark green), cationic and neutral W383 radical (dashed cyan), cationic W329 radical (dashed dark orange).
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Figure 4.10 Kinetic and energetic scheme of the photoreduction of flavin cofactor in A. thaliana (6-4) photolyase
Kinetic and energetic scheme of the tryptophan triad electron transfer chain in the photoreduction of FAD (A) and FADH (B). The energies are not drawn to scales.
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CHAPTER 5 ULTRAFAST DYNAMICS OF FLAVIN COFACTOR IN BLUE-LIGHT RECEPTOR CRYPTOCHROME
101
CHAPTER 5
ULTRAFAST DYNAMICS OF FLAVIN COFACTOR IN BLUE-LIGHT RECEPTOR CRYPTOCHROME
5.1 Introduction Cryptochromes are the flavoproteins that exhibit high sequence and structural similarity to the light-driven DNA-repair enzyme, photolyase.10,41,101 However, cryptochromes lose the ability to repair the DNA damage; instead, they can use the energy from blue light to regulate the growth and development in plants and the circadian clock in plants and animals.10,28,41,101,102 Although the photocycle of DNA repair by photolyases has been completely resolved,12,34,103 the photochemical mechanism of cryptochromes is still not well understood.
A chain of three tryptophan residues near the flavin adenine dinucleotide
(FAD)-binding pocket, which are referred to as the “Trp-triad”, are evolutionarily conserved throughout the photolyase/cryptochrome family.25,44,85 This Trp-triad has been reported as the major electron transfer (ET) channel for the photoreduction of oxidized flavin cofactors in photolyases and cryptochromes in vitro.94,104,105 Recently, some biochemical studies on Arabidopsis thaliana cryptochrome 1 and cryptochrome 2
(AtCRY1 and AtCRY2) proposed that the photoactivation of cryptochromes involves the
Trp-triad-dependent photoreduction.94,106,107 According to this hypothesis, cryptochromes at the resting state contain oxidized FAD; upon excitation, the excited FAD is reduced to
102
semiquinone FADH via the electron transfer (ET) through the Trp-triad; FADH is the functional state that triggers the subsequent conformational changes and the signal transduction of the photoreceptor; FADH can be further reduced to hydroquinone
FADHˉ by green light or reoxidized to FAD by a dark phase under aerobic conditions to complete the photocycle. However, it is well known that photoreduction is not necessary for the DNA repair activity of E. coli photolyase in vivo.84,108 Moreover, recent study on insect type 1 cryptochromes has demonstrated that the Trp-triad mutations that block the
FAD → FAD− photoreduction in vitro do not affect the photoreceptor function of insect type 1 cryptochromes.105,109
To examine this Trp-triad-dependent photoreduction model and determine the functional state of flavin cofactor in AtCRY2, we perform systematic steady-state and ultrafast study on the wild-type AtCRY2 and the mutants of Trp-triad (W397A, W374A and W321A) and the residue opposite to the N5 position of the isoalloxazine ring
(D393A).
5.2 Materials and Methods
5.2.1 Preparation of AtCRY2 and Mutants. The purification of AtCRY2 with His tag has been reported elsewhere.110 For the mutant studies, we replaced each of the three Trp-triad residues (W397, W374 and W321) and the residue opposite to the N5 position of isoalloxazine ring (D393) with alanine. All the mutant plasmids were constructed using QuikChange II XL kit (Stratagene) based on the plasmid of wild-type enzyme. All mutated DNA plasmids were sequenced to ensure correct results. 103
5.2.2 Photoreduction of AtCRY2 and Trp-triad mutants. The purified AtCRY2 and the Trp-triad mutants were illuminated by the 450-nm light in vitro and the absorption spectra were recorded under aerobic conditions in presence of
10 mM β-mercaptoethanol as the external electron donor at 20 ˚C. The light intensity is 2 mW•cm-2.
5.2.3 Femtosecond-resolved Spectroscopy. All the femtosecond-resolved measurements were carried out using up-conversion fluorescence and transient absorption methods. The laser experimental layout and procedure have been detailed elsewhere.35 The excitation wavelengths were 620 nm for the sample with FADH and 400 nm for those with oxidized FAD, FAD− and FADHˉ, respectively. The instrument response time is about 250 fs and the experiments were done at the magic angle (54.7º). 5-mm quartz cuvettes (Starna) were used as the sample cell in experiments and samples were kept stirring during irradiation to avoid heating and photobleaching. The experiments in the femtosecond-resolved measurements were carried out under aerobic conditions for FAD and FADH while under anaerobic conditions for FAD− and FADHˉ.
5.3 Results and Discussion.
5.3.1 Steady-state spectroscopic properties of AtCRY2. In Figure 5.1 we present the local X-ray structure of AtCRY1 and the sequence alignment of flavin binding sites of both AtCRY1 and AtCRY2, with E. coli CPD photolyase and A. thaliana (6-4) photolyase. The alignment shows that the Trp-triad
104
residues are conserved in all these four flavoproteins. The residues opposite to the N5 position of the isoalloxazine ring are aspartic acid (D) in AtCRYs and asparagine (N) in the two photolyases. Interestingly, the corresponding residues are cysteine in insect type 1 cryptochromes, and the oxidized flavin cofactor can only be reduced to FAD− without further protonation and photoreduction to FADH and FADHˉ, indicating that the proton donor must be related to this residue.44 In Figure 5.2 we show typical absorption and emission spectra of wild-type AtCRY2 and the N5 position mutant D393A.
After purification, the flavin cofactors in all wild-type and mutant AtCRY2 are at the oxidized state FAD. The photoreduction activity was examined among the wild-type
AtCRY2 and Trp-triad mutants with 450-nm illumination. Figure 5.3 shows that the wild-type AtCRY2 was photoreduced rapidly in vitro under aerobic conditions while the mutation at any of the Trp-triad residues effectively reduced or abolished the photoreduction activity, which indicated that the Trp-triad is necessary for the in vitro photoreduction of AtCRY2. However, the recent study by our collaborators demonstrated that this Trp-triad-dependent photoreduction is not required for the physiological functions of AtCRY2.110
In contrast to the wild-type AtCRY2 in which the oxidized FAD is first reduced to
FAD−, then to FADH, and finally to FADHˉ, the flavin cofactor in mutant D393A can only be reduced to FAD−, without further going to FADH and FADHˉ as observed in insect type 1 cryptochromes. This D393A mutant is a good candidate to study the dynamics of FAD− in AtCRY2 (see details below).
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5.3.2 Ultrafast photoreduction dynamics of flavin cofactor in AtCRY2. We carried out a systematic investigation on the photoreduction dynamics of FAD in
AtCRY2. Figure 5.4 shows the fluorescence transients gated at 520-nm emission peak for wild type and the three Trp-triad mutants. Since the solvation dynamics mostly appear in the transients gated at the blue side of the emission peak,111,112 the ultrafast decays truly reflect the quenching dynamics of excited FAD with negligible solvation contribution.
The mutation of the nearby tryptophan residue W397 blocks all the potential ET channels of flavin from tryptophan or tyrosine within 5.5 Å. Thus, the observed quenching dynamics of FAD* represent the ultrafast intramolecular ET from adenine moiety to lumiflavin (as discussed in Chapter 4). The fluorescence transient of W397A exhibits double exponential decay of 5 ps (60%) and 34 ps (40%), implying that there might be two population of FAD* with different protein configuration. For wild type and the other two Trp-triad mutants, the fluorescence transients exhibit the faster double exponential decay with nearly the same time scales and amplitudes: 1.2 ps (75-80%) and 11 ps
(20-25%), which represent the quenching of FAD* by the ET from both adenine and
W397 to the excited flavin for the two population of FAD. By deconvolution with the time scale of ET from adenine to lumiflavin, the time scales of ET from W397 to the excited lumiflavin are 1.6 ps for the configuration 1 (~80%) and 16 ps for the configuration 2 (~20%).
To capture the related tryptophan or adenine radical intermediate and resolve the entire dynamics, we examined the excited-state dynamics of FAD with transient absorption methods. Figure 5.5A and B show the absorption transients of FAD* in
W397A probed at 750 and 620 nm. Besides the quenching dynamics of excited 106
lumiflavin in two protein configurations, there is a plateau of transient signal which attributes to the formed intermediate Ade+ radical. The time scales of formation of the
Ade+ radical in two different protein configurations are consistent with the quenching of the corresponding excited lumiflavin, and the charge recombinations between Ade+ and lumiflavin− take longer than 5 ns in both configurations, which implies that there might be further ET from other aromatic residues to Ade+. Figure 5.5C and D show the absorption transients in W374A. At 620 nm, besides the FAD* dynamics and the signal from Adenine ET channel (dashed cyan for configuration1 and dashed dark orange for configuration 2), we captured the strong signal of W397+ with rise-decay behavior
(dashed pink and dashed dark green). The decay takes 25 ps in configuration 1 and is attributed to the charge recombination between W397+ and lumiflavin−. In configuration
2, the charge recombination completes in 115 ps. For the mutant W321A, we detected the strong transient signal of W374+ at 620 and 510 nm (Figure 5.6B and C). Since the
W374 is over 9.5 Å away from the flavin moiety, we can exclude the possibility of direct charge recombination between W374+ and lumiflavin− on nanosecond time scale. Based on the dynamics obtained from W374A, we determined that the forward electron hopping from W374 to W397+ occurs in 22 ps while the back electron hopping from W397 to
W374+ takes 2 ns in configuration 1. In configuration 2, the time scales for the forward and backward electron hopping are 120 ps and 10 ns, respectively. With the knowledge of the obtained dynamics in mutants, we mapped out the whole process of photoreduction of
FAD in wild type. Unlike the mutant W321A, the radical W374+ can be further reduced by the final electron donor W321. The generated radical W321+, which is on the surface 107
of protein, will gain an electron from the solution and finish the photoreduction process.
Compared to W321A, the transient signal of wild type at all probed wavelengths shows a long plateau (Figure 5.7A, B and C) rather than a decay in 2 ns. This difference is mainly attributed to the long-lived W321+. Based on the acquired dynamics in mutants, we determined the forward electron hopping from W321 to W374+ takes 280 ps in configuration 1 and ~2 ns in configuration 2. Moreover, the long plateau signal of
W321+ has no obvious decay behavior, indicating that the back electron hopping takes much longer than our detection window. The time scales of each elementary step of the ultrafast photoreduction of FAD are listed in Table 5.1.
In addition, we also examined the photoreduction of FADH in wild-type AtCRY2
(Figure 5.7D). Upon 620-nm excitation, only FADH can be excited. The absorption transient probed at 800 nm truly reflects the ultrafast quenching dynamics in 13 ps.
Hence, it could be argued that if the photochemistry of excited flavin cofactor is the initial signaling step, both FAD and FADH can not be the physiologically functional state because their excited states are readily quenched on picosecond or tens of picosecond time scale by the neighboring aromatic residues. In contrast, the anionic
FAD− and FADHˉ are the good candidates for the functional states because their excited states can not be quenched by accepting electron from the neighboring aromatic residues and therefore have much longer lifetimes.
5.3.3 Deactivation dynamics of anionic flavin cofactors in AtCRY2. Figure 5.8A shows the fluorescence transient gated at 620 nm for the wild-type
108
AtCRY2. The transient exhibits three exponential decay components: 9.2 ps (30%), 183 ps (27%) and 1547 ps (43%). Without the substrate and the possibility of quenching by electron transfer from the neighboring aromatic amino acids or intrachromolphore electron transfer, the observed multiple exponential decays truly reflect the deactivation dynamics of the excited FADHˉ in AtCRY2. As shown in Figure 5.2B, the peaks of emission from the excited FADHˉ shift to the blue side with the higher excitation energy, indicating that the excited molecules do not completely relax to the lowest excited state as the Kasha’s law but partially deactivate to the ground state during the relaxation. Such multiple deactivation processes mostly occur through the conical intersections often observed in the large organic molecules such as DNA bases.113,114
To confirm such photophysical deactivation processes, we examined the dynamics of the excited FADHˉ in AtCRY2 using transient-absorption detection with a variety of probing wavelengths. Figure 5.8B (left panel) shows the typical transients with several probing wavelengths. All transients can be globally fitted with three exponential decays:
3-9 ps, ~100 ps, and a long component (2-3 ns). These multiple-decay dynamics are consistent with the excited-state deactivation processes observed by the fluorescence up-conversion detection. In our recent studies on the excited-state dynamics of free
FADHˉ in solution as well as FAD− and FADHˉ bound to the insect cryptochromes,44,115 we observed the similar multiple-decay processes. A flexible “butterfly” bending motion of the isoalloxazine ring was proposed to access conical intersections for the multiple deactivation processes.44
In the mutant D393A, the flavin cofactor can be stable at FAD−. The excited-state
109
dynamics of FAD− were recorded by the transient-absorption methods. Figure 5.8B
(right panel) shows three typical absorption transients probed from 750 nm to 510 nm.
The transients exhibit multiple exponential decays of 2-8 ps, 20-90 ps, and ~4 ns, and reflect the deactivation dynamics of FAD− through a similar mechanism of conical intersections as observed for FADHˉ in AtCRY2 and FAD− in insect cryptochromes.44
In contrast, the excited FADHˉ in E. coli photolyase has a single lifetime of 1.3 ns without any fast decay.12 These excited-state dynamics differences probably lie in the fact that the photoreceptor cryptochromes have higher plasticity and the isoalloxazine ring in cryptochromes is not as rigidly held in the active site as in photolyases.53,116,117 With less constraint, the ring undergoes the “butterfly” bending motion in the ground or excited state that leads to multiple deactivation process through conical intersections in which the rapid decay pathways are dominant. As a general rule, chromophores must have a long-lived excited-state to perform photochemical catalysis with a reasonable quantum yield. In line with this notion, the long-lived excited FADHˉ ensures E. coli photolyase to repair the damaged DNA with a quantum yield of near unity.10,12 Although the fraction of the short-lived excited FADHˉ (9 ps and 183 ps) in AtCRY2 are dominant, there are still considerable percentages of excited FADHˉ with long lifetime to perform function. To this point, the excited FADHˉ is more likely to be the functional state rather than the short-lived FAD* (a few ps) or FADH* (tens of ps).
5.4 Conclusions We systematically examined the excited-state dynamics of flavin cofactor in four different redox states (FAD, FAD−, FADH and FADHˉ) in Arabidopsis thaliana 110
cryptochrome 2. With femtosecond resolution, we observed the ultrafast photoreduction of oxidized FAD and neutral semiquinone FADH occurring in a few picoseconds and tens of picoseconds, respectively, through the intraprotein electron transfer along the conserved Trp-triad. Such ultrafast quenched dynamics exclude their potential role as the functional state and therefore overthrow the Trp-triad-dependent photoreduction model.
The anionic hydroquinone FADHˉ and semiquinone FAD− have multiple deactivation dynamics on the time scales from a few picoseconds to a few nanoseconds, which is believed to occur through the conical intersections with a flexible bending motion of the isoalloxazine ring. These results imply that the anionic hydroquinone FADHˉ is more likely to be the functional state rather than the two neutral states.
111
FAb BAb FETc BETc HOP1d BHOP1d HOP2e BHOP2e
f (AtCRY2-FAD)1 5.0 5000 1.6 25 22 2000 280 >10000
g (AtCRY2-FAD)2 34 >10000 16 113 120 5000 2000 >10000
Table 5.1 The time scales of the individual electron transfer steps in the photoreduction of FAD in A. thaliana cryptochrome 2a aAll times are in unit of picosecond. bFA and BA are the charge separation and recombination between the excited lumiflavin and adenine moiety. cFET and BET are the forward electron transfer from W397 to FAD* and the backward electron transfer in the reverse direction. dHOP1 and BHOP1 are the forward electron hopping from W374 to W397+ and the backward electron hopping in the reverse direction. eHOP2 and BHOP2 are the forward electron hopping from W321 to W374+ and the backward electron hopping in the reverse direction. fThe population of FAD in the protein configuration1 in which the excited FAD has the faster quenching. gThe population of FAD in the protein configuration 2 in which the excited FAD has the slower quenching.
112
Figure 5.1 Sequence alignment and local X-ray structure of A. thaliana cryptochrome.
Sequence alignment of two plant cryptochromes and two photolyases. At, Arabidopsis thaliana; Ec, Escherichia coli. Note the conserved tryptophan triad (in red) for photoreduction through electron transfer across all cryptochromes and photolyases. Shown at the bottom is the local X-ray structure around the flavin binding site in A. thaliana cryptochrome 1. The residues in parentheses are the corresponding residues in A. thaliana cryptochrome 2.
113
Figure 5.2 Steady-state spectra of A. thaliana cryptochrome 2
(A) Absorption spectra of oxidized (FAD), anionic semiquinone (FAD−) and hydroquinone (FADHˉ) flavin cofactor in AtCRY2. The stable anionic semiquinone was obtained by mutation of D393 with alanine. (B) Excitation-wavelength dependence of weak FADHˉ emission spectra, indicating deactivation during relaxation. The Raman scattering signals at the blue side of emission peaks were all removed for clarity.
114
Figure 5.3 Photoreduction of A. thaliana cryptochrome 2
Photoreduction of the wild-type AtCRY2 protein and the lack of photoreduction of the three AtCRY2 Trp-triad mutant proteins. The absorption spectra were recorded at indicated times after blue-light illumination (450 ± 15 nm, 2 mW•cm-2) under aerobic conditions at 20 °C in presence of 10 mM β-mercaptoethanol as the external electron donor.
115
Figure 5.4 Femtosecond-resolved oxidized flavin dynamics in A. thaliana cryptochrome 2
Normalized fluorescence transients for the wild-type AtCRY2 and the three Trp-triad mutants, all gated at 520 nm around the emission peak. These signals represent the ultrafast quenching of excited FAD by ET from the neighboring adenine or tryptophans.
116
Figure 5.5 Femtosecond-resolved dynamics of direct electron transfer to FAD in A. thaliana cryptochrome 2
Left column shows the normalized transient absorption signals of W397A mutant probed at 750 nm (A) and 620 nm (B), respectively, with deconvolution of flavin species (dashed cyan for configuration 1 and dashed dark orange for configuration 2), cationic adenine radical (dashed pink for configuration 1 and dashed dark green for configuration 2). Right column shows the normalized transient absorption signals of W374A mutant probed at 750 nm (C) and 620 nm (D), respectively, with deconvolution of flavin species and cationic adenine radical (dashed cyan for configuration 1 and dashed dark orange for configuration 2), cationic W397 radical (dashed pink for configuration 1 and dashed dark green for configuration 2).
117
Figure 5.6 Femtosecond-resolved dynamics of FAD photoreduction involving W374 in A. thaliana cryptochrome 2
Normalized transient absorption signals of W321A mutant probed at 750 nm (A), 620 nm (B) and 510 nm (C) are deconvoluted by flavin species and cationic W397 with Adenine radical (dashed cyan for configuration 1 and dashed dark orange for configuration 2), cationic W374 radical (dashed pink for configuration 1 and dashed dark green for configuration 2).
118
Figure 5.7 Femtosecond-resolved dynamics of photoreduction of FAD and FADH in A. thaliana cryptochrome 2
Normalized transient absorption signals of FAD photoreductio in wild-type AtCRY2 probed at 750 nm (A), 620 nm (B) and 510 nm (C) are deconvoluted by flavin species and cationic W397 and W374 with Adenine radical (dashed cyan for configuration 1 and dashed dark orange for configuration 2), cationic W321 radical (dashed pink for configuration 1 and dashed dark green for configuration 2). Normalized transient absorption signal of FADH photoreduction in wild-type AtCRY2 was probed at 800 nm (D). Note that the excitation wavelengths are 400 nm for the photoreduction of FAD and 620 nm for the photoreduction of FADH. 119
Figure 5.8 Femtosecond-resolved dynamics of anionic flavin cofactors (FADHˉ and FAD−) in A. thaliana cryptochrome 2
(A) Normalized fluorescence transient of FADHˉ for wild-type AtCRY2 gated at 600 nm. (B) Transient-absorption detections of excited FADHˉ for wild-type AtCRY2 (left panel) and excited FAD− for the mutant D393A at various probe wavelengths, showing the similar excited state dynamics probed with fluorescence in panel A.
120
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APPENDIX: PROTOCOLS
I. DNA-Photolyase Protein Purification Protocol II. Photoreduction and Oxidation of Flavin Cofactor and Photodecomposition of MTHF cofactor in Photolyase III. Oligo Cyclobutane Pyrimidine Dimer Preparation Protocol
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I. DNA Photolyase purification Protocol
Transformation Take 100 l of UNC523 cell, place it into an eppendorf tube and keep the tube on ice. Add 1 l of plasmid DNA in the tube which contains 100 l of UNC523 cells. Incubate the whole tube on ice for 30 minutes. Heat-shock the UNC523 cell + plasmid DNA mixture by placing the tube in a heat block at 42 ºC for 45 seconds. Take out the eppendorf tube from heat block and incubate it over ice for 2 minutes. Pipet out the UNC523 cell + plasmid DNA mixture and spread it over a LB-Ampicillin plate. Keep the LB-Ampicillin plate at 37 ºC incubator for 10 minutes. (Keep it face up) Turn the plate upside down (face down) and keep it overnight at 37 ºC.
Preparing the Overnight Culture Take 5 ml autoclaved LB medium and place it in a sterile test tube. Add 10 l of 50 mg/ml Ampicillin in this test tube. (Final Ampicillin 100 g/ ml) Pick single colony from yesterday’s LB-Ampicillin plate and dilute it in the test tube which contains LB-Ampicillin mixture. Keep this test tube for 6~8 hours at 37 ºC incubator with sharking speed at 220 rpm. Add 200 l of 50 mg/ml Ampicillin in the flask which contains 150 ml autoclaved LB medium. Add 1 ml of previous test-tube culture in this flask. Keep this 100 ml culture overnight at 37 ºC incubator with shaking speed at 220 rpm. (This is the overnight culture)
Preparing Large Quantity of Cell Culture and Harvesting Cell Prepare 1 liter LB medium in 2-liter flask and autoclave the whole flask. (Prepare such autoclaved LB medium one day ahead) Add 2 ml of 50 mg/ml Ampicillin in 2-liter sterile flask which contains 1 liter autoclaved LB medium. Take 10 ml of overnight culture which shows O.D. of 1.8~1.9 at 600 nm and dilute it to LB-Ampicillin mixture in the 2-liter flask. Let the culture grow at 37ºC incubator with shaking speed at 220 rpm until 600-nm O.D. value of the culture reaches to 0.6 (0.6~0.8). (It usually takes 2 to 3 hours in the incubator in our lab) After the 600-nm O.D. value of culture reaches to 0.6, cool down the culture in cold room for about 30 min. Add 2 ml of 0.5M IPTG to the each flask. (Final IPTG 1mM) Let it grow at 22 ºC with 220 rpm overnight. Transfer the culture to centrifuge bottles. Centrifuge for 10 minutes at 4 ºC with rotor speed at 5500 rpm. Discard the supernatant. Dissolve the pellet with PBS buffer. (About 10 ml PBS buffer per 1 liter culture) 130
Combine the entire dissolved pellet in one bottle and centrifuge the bottle for 15 minutes at 4 ºC with rotor speed at 6000 rpm. Discard the supernatant. Quick freeze the pellet by liquid nitrogen. Keep the frozen pellet in -80 ºC.
Cell Lysis and Ammonium Sulfate Precipitation Thaw the frozen pellet into ice-water bath at 4 ºC. Add low-salt buffer (10 ml per liter of cell culture) + lysozyme (1.0 mg per ml of low-salt buffer). Mix well and keep mixing on the rocker or stir mixer overnight at 4 ºC. Transfer the lysised cell solution to a metal beak and place the beak over ice in an ice bucket. Sonicate lysised cell with Duty cycle of 50% and pulse output of 4 for 45 seconds. Repeat 8-10 times with 45 seconds interval in between. (Make sure that the dissolved pellet forms nice slurry and no more clogs) Transfer sonicated solution to centrifuge tubes and centrifuge for 45 minutes at 4 ºC with rotor speed at 20000 rpm. (Make sure that the tubes have been cool down before transfer) Transfer the supernatant to another centrifuge tubes and centrifuge for 45 minutes at 4 ºC with rotor speed at 20000 rpm again. Take clean supernatant and combine it in a large beaker. Measure the volume of the supernatant and weight out the ammonium sulfate. (0.43 mg of ammonium sulfate per ml of supernatant) Keep the beaker in ice bucket with ice bath at 4 ºC. Use stir mixer and stir the supernatant while add weighted ammonium sulfate evenly within one hour. After finish adding the ammonium sulfate, stir for extra one hour. Transfer the mixture to centrifuge tubes and centrifuge for 30 minutes at 4 ºC with rotor speed at 14000 rpm. (Make sure that tubes have been cool down before transfer) Discard the supernatant and gently resuspend the pellet with <5 ml of low-salt buffer per tube. (The pellet contains all salt-out proteins) After resuspension, we now get protein mixture. (Simply call it as protein) A) Dialyze the protein against the low-salt buffer for (4+4) hours. Change the buffer after 4 hours during 8 hours dialysis. B) Load the protein (Make sure all pellet is dissolved well and there is no any precipitate) onto Sephadex G25 desalting column and collect all flow-through which shows color. After either process A) or B), the slat concentration in the protein has been lower down. Protein Purification Dilute the protein with an equal volume of low-salt buffer. (If the volume of protein mixture is 60 ml, add extra 60 ml of low-salt buffer into the mixture solution and bring the total volume to 120 ml ) Load the protein onto a blue sepharose column that has been equilibrated with low-salt buffer, adjust the flow rate of 0.30 ml per minute. After loading whole protein sample, wash the blue sepharose column with 2 column volumes of low-salt buffer and keep the flow rate of 0.4-0.5 ml per minute. Collect fractions while washing. Set the fraction collector to 160 drop per tube. Continue such washing process until there is no more yellow color in fraction tubes.
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Elute the protein with high-salt buffer. The fractions with blue color are dominant photolyase protein. Based on the absorption spectrum in the range of 300 ~ 700 nm, combine all fractions which show typical semiquinone flavin absorption bands. If MTHF-depletion photolyase enzyme is wanted, flow the step (6) and (7), otherwise directly go to step (8). Add DTT to the protein sample with final concentration of 10 mM. Equally distribute protein sample to several large test tubes and keep the test tubes titled in an ice bucket with ice underneath. Expose the protein sample to 365 nm light (UVP UVLS-28) in cold room to photodegrade the MTHF cofactor of the photolyase. Rotate the tubes manually to avoid over heating the protein and monitor both absorption and emission spectra during photodegradation. When the emission at 480 nm is completely gone, the photo-degradation process in completed. Combine protein from all tubes and concentrate it in Amicon-tube to final about 4-6 ml. It usually takes 2-4 hours. Load the concentrated protein on to gel-filtration Bio-Gel P100 column that has been equilibrated with low-salt buffer. Once the protein completely gets into the resins, connect the column with low-salt buffer reservoir. Set fraction collector to 160 drops per tube. The fractions with blue color are dominant photolyase protein. Based on the absorption spectrum in the range of 300 ~ 700 nm, combine all fractions of sample which show typical semiquinone flavin absorption bands. Dilute the sample twice with Heparin A buffer, and load the sample onto Heparin column in FPLC (Fast Protein Liquid Chromatography). Elute the column with a gradient of Heparin A and Heparin B buffer with flow rate of 1.0 ml per minute. Set each fraction of 1.5 ml. Combine all fractions which show typical semiquinone flavin absorption bands. Depending on concentration of the protein, concentrate the protein if needed. Dialyze 8-hours against the pre-cooled storage buffer. Transfer photolyase enzyme to eppendorf tubes and quick freeze by liquid nitrogen. Store in -80 ºC.
Buffers
(10X) PBS Buffer (Phosphate Buffered Saline) 1L Na2HPO4 10.9 g NaH2PO4 3.2 g NaCl 80 g KCl 2 g Add DIW to final 1 liter and adjust pH to 7.2 Dilute 1:10 (v/v) with sterile DIW before use and adjust pH if necessary
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Low-salt Buffer Final concentration 1 L 2 L 3 L Tris-HCl pH 7.4 50 mM 6.6 g 13.2 g 19.8 g EDTA 1.0 mM 0.96 g 1.92 g 2.88 g KCl 100 mM 7.43 g 14.9 g 22.29 g DIW 900 ml 1800 ml 2700 ml AUTOCLAVED After sterilization add Glycerol 10 % (v/v) 100 ml 200 ml 300 ml (use autoclaved glycerol) BME 10 mM 0.72 ml 1.43 ml 2.16 ml
High-salt Buffer Final concentration 1 L 2 L 3 L Tris-HCl pH 7.4 50 mM 6.6 g 13.2 g 19.8 g EDTA 1.0 mM 0.96 g 1.92 g 2.88 g KCl 2.0M 149.12 g 298.24 g 447.36 g DIW 900 ml 1800 ml 2700 ml AUTOCLAVED After sterilization add Glycerol 10 % (v/v) 100 ml 200 ml 300 ml (use autoclaved glycerol) BME 10 mM 0.72 ml 1.43 ml 2.16 ml
Heparin A Buffer Final concentration 1 L 2 L 3 L Tris-HCl pH 7.4 50 mM 6.6 g 13.2 g 19.8 g DIW 900 ml 1800 ml 2700 ml AUTOCLAVED After sterilization add Glycerol 10 % (v/v) 100 ml 200 ml 300 ml (use autoclaved glycerol) BME 10 mM 0.72 ml 1.43 ml 2.16 ml
Heparin B Buffer Final concentration 1 L 2 L 3 L Tris-HCl pH 7.4 50 mM 6.6 g 13.2 g 19.8 g
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NaCl 1.0M 58.44 g 116.88 g 175.32 g DIW 900 ml 1800 ml 2700 ml AUTOCLAVED After sterilization add Glycerol 10 % (v/v) 100 ml 200 ml 300 ml (use autoclaved glycerol) BME 10 mM 0.72 ml 1.43 ml 2.16 ml
Storage Buffer Final concentration 1L Tris-HCl pH 7.4 50 mM 6.6 g EDTA 1.0 mM 0.96 g NaCl 100 mM 5.84 g DTT 10 mM 1.54 g Glycerol 50 % (v/v) 500 ml Add DIW to final 1 liter, filter with 0.2-μm membrane
Column Regeneration Sephadex G25 desalting column Low-salt buffer for 2-4 column volumes
Blue-Sepharose (1) High-salt buffer for 2-column volumes (2) 2 M guanidine-HCl for 2-column volumes (3) 1L sterilize DIW (4) Low-salt buffer for 2-column volumes Gel-Filtration (Bio-Gel P100) Low-salt buffer for 2-column volumes
Heparin (1) Heparin B buffer for 5-column volumes (2) Sterile DIW for 5-column volumes (3) Heparin A buffer for 5-column volumes
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II. Photoreduction and Oxidation of Flavin Cofactor and Photodecomposition of MTHF cofactor in Photolyase
Photoreduction of Flavin Cofactor in Photolyase Reduce the semiquinone flavin (FADH) to fully-reduced hydroquinone (FADH) Take 200 l of 300-500 M photolyase enzyme and add final 10 mM DTT. Place protein in a 5 mm quartz purging cell and add a mini stir bar. Place the cap of quartz purging cell and inset purging tubing. Purge the sample with argon in the purge system for about 2-3 hours and maintain slow gas flow rate to avoid drying or diluting the sample. Slowly remove the purge tubing and avoid presence of oxygen. Now the protein sample is in anaerobic condition. Measure the absorption spectrum. The semiquinone flavin shows a broad absorption spectrum with peaks at 484, 580 and 625 nm. Place the quartz cell horizontally with ice underneath. Irradiate the sample with high-intensity table lamp attached a 550-nm long pass cut-off filter. Keep lamp about 2 cm above the quartz cell and rotate the quartz cell manually to avoid over heating the sample. Measure the absorption spectrum every 5 minutes of irradiation and monitor the decrease of semiquinone absorption. Continue the irradiation until all semiquinone converts to fully-reduced flavin which shows a narrower absorption band with single peak at 360 nm and zero absorption above 500 nm. It takes about 10-20 minutes for all process.
Note: Reduction rate is highly dependent on (a) the concentration of reducing agent DTT, (b) distance between light source and sample and (c) presence of oxygen in the quartz cell.
Oxidation of Flavin Cofactor in Photolyase Oxidize the semiquinone flavin (FADH) to oxidized flavin (FAD) Dialyze the photolyase enzyme in oxidation buffer which contains 250 mM imidazole. Monitor the absorption spectrum every 5 hours during the dialysis. Continue dialysis until the absorption of semiquinone is completed gone and oxidized flavin absorption spectrum is clearly shown. The semiquinone flavin shows a broad absorption spectrum with peaks at 484, 580 and 625 nm while the oxidized flavin shows a narrower absorption spectrum with peaks at 475, 454, 432 and 384 nm and zero absorption above 510 nm. (It may takes 1-2 day to completely oxidize flavin cofactor) If precipitation occurs, either quick spin sample or filter sample with 0.2 μm filter. Take the clean supernatant and load it onto Heparin column that has been equilibrated with wash Heparin A buffer. Wash the Heparin column with 5-column volume of Heparin A buffer. Elute the Heparin column with a gradient of Heparin A and Heparin B buffer with flow rate of 1.0 ml per minute. Set each fraction of 1.5 ml. The fractions with yellow color are photolyase protein. Based on the absorption spectrum in range on 300 ~ 700 nm, combine all fractions which show typical oxidized flavin 135
absorption bands. Depending on concentration of the protein, concentrate the protein if needed. Dialyze 8-hours against the 1 liter pre-cooled storage buffer. Transfer photolyase enzyme to eppendorf tubes and quick freeze by liquid nitrogen. Store in -80 ºC.
Oxidation Buffer Final concentration Stock solution 1L Tris-HCl pH 7.4 50 mM 1M 50 ml EDTA 1.0 mM 0.5 M 2 ml NaCl 50 mM 5.0 M 10 ml Imidazole 250 mM 1M 250 ml Glycerol 10 % (v/v) sterile 100 ml Add DIW to final 1 liter
Photodecomposition of MTHF Cofactor in Photolyase Prepare MTHF-depleted photolyase First, photoreduce the flavin cofactor in photolyase as described above. Measure the absorption and emission spectra. The absorption spectrum of MTHF shows peak at 384 nm and absorption peak of fully-reduced flavin is at 360 nm. The emission spectrum of MTHF shows a peak at 474 nm while that of FADHˉ shows a structured fluorescence profile peaking at 515 and 545 nm. Place the quartz cell horizontally with ice underneath. Irradiate the sample on ice with a high intensity 365 nm lamp (UVP UVLS-28). Keep lamp about 2 cm above the quartz cell and rotate the quartz cell manually to avoid over heating the sample. Measure the absorption and emission spectra every 30 minutes and monitor the decease of MTHF absorption and emission peak at 384 nm and 480 nm, respectively, during photodegradation. Continue the irradiation until the emission at 474 nm is completely gone.
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III. Oligo Cyclobutane Pyrimidine Dimer (CPD) Preparation Protocol
Oligo CPD Preparation by Acetone-sensitized Irradiation Dissolve ~ 3 mg oligo(dT)15 (synthesized by Integrated DNA Technologies) in 0.85 ml deionized H2O, and add 0.15 ml pure acetone right before shining with UV light. Place the sample into quartz purging cell, and place the quartz cell on the ice in the cold room. Purge the sample with argon 15 min before irradiation. Maintain a very slow flow gas rate to avoid the evaporation of acetone. Irradiate the sample with a 302-nm UVB lamp (General Electric). Keep the light source about 2 cm above the quartz cell. Frequently rotate the quartz cell over ice and keep the temperature as low as possible. During the irradiation, both temperature of quartz cell and the presence of acetone are critical to produce CPD and to avoid other photoproducts. Check the absorbance after 50-min irradiation (the time is dependent on the light intensity). Take 2 μl of sample out and dry it with speed vacumn to remove the acetone. Dissolve the sample with 200 μl deionized H2O and check the absorbance at 260 nm and 220 nm. Keep irradiation until the ratio of A220/A260 reaches about 10. Take the sample out and transfer it to a 1.7 ml ependorf tube. Dry out the sample in speed vacumn and combine together. Keep it in -20ºC.
Reactivation Assay Prepare a photolyase-CPD complex sample with approximately concentration ratio of 1: 200 (Photolyase: oligo(dT)15-CPD) in storage buffer. The minimum volume of sample is 200 l. It usually takes 2 μM of photolyase to 400 μM of oligo(dT)15-CPD. Prepares it under red light and avoid exposure of <500 nm light. Transfer it to 5 mm quartz cuvette and warp the top of the cuvette with parafilm. Measure UV-Vis absorption spectrum. Define it as time-zero (T=0) spectrum. Incubate the complex sample on ice for 5 minutes. Irradiate the complex sample with 365 nm light. Monitor the absorption spectrum every 5-minutes irradiation. Once the CPD is repaired by photolyase enzyme, the characteristic absorption peak of thymine at 266 nm is shown. Continue irradiation until no more increase in the 266-nm absorption. Substrate the T=0 spectrum out from each T>0 spectrum. Based on the change of 266-nm absorbance before and after irradiation, calculate the total amount of CPDs that have been repaired, and estimate the number of CPDs formed in each strand of oligo(dT)15 on average.
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