Dynamics and mechanism of repair of ultraviolet-induced (6–4) photoproduct by photolyase

Journal name:
Nature
Volume:
466,
Pages:
887–890
Date published:
DOI:
doi:10.1038/nature09192
Received
Accepted
Published online
Corrected online

One of the detrimental effects of ultraviolet radiation on DNA is the formation of the (6–4) photoproduct, 6–4PP, between two adjacent pyrimidine rings1. This lesion interferes with replication and transcription, and may result in mutation and cell death2. In many organisms, a flavoenzyme called photolyase uses blue light energy to repair the 6–4PP (ref. 3). The molecular mechanism of the repair reaction is poorly understood. Here, we use ultrafast spectroscopy to show that the key step in the repair photocycle is a cyclic proton transfer between the enzyme and the substrate. By femtosecond synchronization of the enzymatic dynamics with the repair function, we followed the function evolution and observed direct electron transfer from the excited flavin cofactor to the 6–4PP in 225picoseconds, but surprisingly fast back electron transfer in 50picoseconds without repair. We found that the catalytic proton transfer between a histidine residue in the active site and the 6–4PP, induced by the initial photoinduced electron transfer from the excited flavin cofactor to 6–4PP, occurs in 425picoseconds and leads to 6–4PP repair in tens of nanoseconds. These key dynamics define the repair photocycle and explain the underlying molecular mechanism of the enzyme’s modest efficiency.

At a glance

Figures

  1. Enzyme-substrate complex structure and a possible repair scheme.
    Figure 1: Enzyme–substrate complex structure and a possible repair scheme.

    X-ray structure of Drosophila melanogaster (6–4) photolyase6 (blue) bound to DNA (yellow) containing a (6–4) photoproduct. Arabidopsis thaliana photolyase has a similar structure11 and conserved histidine residue in the active site (His364 in A. thaliana and His365 in D. melanogaster). The 6–4PP is flipped out of DNA and inserted into the active site. The close-up view shows the relative positions of the catalytic cofactor FADH, the conserved His364 (His365) residue and the 6–4PP substrate with a proposed scheme for electron and proton transfers in the repair reaction. Shown in the repair scheme (inset) are forward electron transfer (FET, reaction rate k1) after light excitation, back electron transfer (BET, reaction rate k2) without repair, and the repair channel, including initial proton transfer (PT, reaction rate k3) and late proton and electron return (PR and ER, reaction rate k4) after repair.

  2. Femtosecond-resolved dynamics of flavin species involved in the repair of damaged DNA by (6-4) photolyase enzyme (E6-4PL).
    Figure 2: Femtosecond-resolved dynamics of flavin species involved in the repair of damaged DNA by (6–4) photolyase enzyme (E6–4PL).

    a, Absorption spectra and coefficients of purified protein with FADH· (red), converted active form FADH (blue), damaged 6–4PP (yellow) and FADH* (green) determined by this study. Also determined is a 6–4PP-related reaction intermediate (Inter., purple) around 325nm in Fig. 3. Inset, fluorescence emission of FADH* , with an arrow indicating the gated wavelength. b, Normalized signals detected by both fluorescence (gated around the emission peak of 550nm) and absorption (probed at 800nm) methods without (red) and with (blue) the substrate in the active site show the same lifetime and forward electron transfer decays. The electron transfer dynamics are best represented by a stretched-single-exponential decay. λfl, wavelength of fluorescence; λpr, probing wavelength; a.u., arbitary units. c, Transient absorption signal probed at 640nm with both FADH* (blue) and FADH· detection (green). The total FADH· signal is from the two contributions of the initially formed FADH· signal (dashed purple; k1 formation and k2+k3 decay in Fig. 1) and the branched FADH· signal in the repair channel (dashed light-blue line; k3 formation and k4 decay). Inset, flat signal in tens of picoseconds, reflecting an apparent fast signal rise. ΔA, absorption change. d, Transient absorption signals probed at 640nm of the His364Asn mutant (green line) and wild-type (WT) enzymes in D2O (dark-red line) compared with the wild-type in H2O (light-red line). The corresponding relative steady-state quantum yield measurements are shown in inset.

  3. Femtosecond-resolved transient absorption dynamics of various species involved in the damaged DNA repair.
    Figure 3: Femtosecond-resolved transient absorption dynamics of various species involved in the damaged DNA repair.

    Repair was probed at a, 325nm and b, 315nm for the His364Asn mutant (green line) and wild-type enzymes in D2O (dark-red line) and H2O (light-red line). The signals from the wild type include three contributions of overall flavin species (FADH, FADH* and FADH·, blue line), 6–4PP (yellow line) and a captured intermediate (purple line), shown in insets. The mutant signals decay to zero with a futile electron transfer cycle. The determined absorption coefficients of FADH* and the intermediate are shown in Fig. 2a.

  4. Repair photocycle of (6-4) thymine photoproduct by (6-4) photolyase.
    Figure 4: Repair photocycle of (6–4) thymine photoproduct by (6–4) photolyase.

    The resolved elementary steps include a forward electron transfer in 225ps on excitation (I to II), a back electron transfer in 50ps without any repair (II to I) and a parallel, catalytic proton transfer between the enzyme (His364) and the substrate (II to III), induced by the initial electron transfer, in 425ps. This proton transfer is the determinant step in repair and determines the overall repair quantum yield. The subsequent repair reactions involve a series of atom arrangements with bond breaking and bond making (III to IV), and final proton and electron returns (to His364 residue and flavin cofactor) to convert the 6–4PP to two thymine bases on time scales of longer than ten nanoseconds (IV to V).

Change history

Corrected online 11 August 2010
A correction was made to the x-axis labels of the Fig. 2a inset.

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Affiliations

  1. Departments of Physics, Chemistry and Biochemistry, Programs of Biophysics, Chemical Physics and Biochemistry, 191 West Woodruff Avenue, The Ohio State University, Columbus, Ohio 43210, USA

    • Jiang Li,
    • Zheyun Liu,
    • Chuang Tan,
    • Xunmin Guo,
    • Lijuan Wang &
    • Dongping Zhong
  2. Department of Biochemistry and Biophysics, University of North Carolina School of Medicine, Chapel Hill, North Carolina 27599, USA

    • Aziz Sancar

Contributions

D.Z. designed the research. J.L., Z.L., C.T., X.G. and L.W. performed the research. J.L., Z.L., C.T. and D.Z. analysed the data. J.L., Z.L., A.S. and D.Z. wrote the paper. All authors discussed and edited the manuscript.

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The authors declare no competing financial interests.

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    This file contains Supplementary Kinetic Data Analysis, Supplementary Table 1 and Supplementary Figures 1S-3S with legends.

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