Chapter 9
Cyclobutylthymidine Dimer Repair by DNA Photolyase in Real Time
Advances in Biopolymers Downloaded from pubs.acs.org by YORK UNIV on 12/05/18. For personal use only.
RobertJ.Stanley Department of Chemistry, 201 Beury Hall, Temple University, Philadelphia, PA 19122
DNA photolyase (PL) is a monomeric flavoprotein that binds cyclobutylpyrimidine dimers (CPDs) and repairs them via photoinduced electron transfer from a reduced flavin adenine dinucleotide cofactor (FADH ) to the CPD. In spite of significant effort, the repair mechanism remains poorly understood. We have used femtosecond transient absorption spectroscopy to explore the electron transfer and repair kinetics of A . nidulans DNA photolyase with oligothymidine substrates in real time. Dimeric substrates show a concentration-dependent mixture of kinetics representing bound and unbound substrate. A longer ρentameric substrate shows faster electron transfer than previously observed. Repair of the carbon-carbon double bonds (C=C) in the CPD appears to be complete by 1,500 ps. Target analysis of the kinetics is unable to distinguish between sequential and concerted models for the repair reaction, although the data are suggestive of the former. -
© 2006 American Chemical Society
138 DNA photolyase is a monomeric flavoprotein that binds and repairs cyclobutylpyrimidine dimers (CPDs) in DNA by a unique light-driven electron transfer mechanism. These CPDs are known to cause cell death and mutations and consequently photolyase is found widely distributed in all three kingdoms, including crop plants such as rice. Given the importance of these crops to human welfare, a thorough knowledge of the CPD repair mechanism is warranted. Previous experiments have shown that repair of CPDs by photolyase is extremely fast and efficient. Repair is completed within about 2 nanoseconds after the absorption of blue light by the fully reduced anionic flavin adenine dinucleotide cofactor (FADH). However, in spite of significant efforts, the details of the repair mechanism remain elusive. The structures of E. coli and A. nidulans DNA photolyase have been solved in the absence of substrate. Recently, the photolyase-product complex structure has become available. The CPD appears to be within 4 Â of the FAD at the bottom of the cavity, confirming the hypothesis that the CPD must flip out of the DNA double helix into a cavity in the protein. ' 16
8
14
19
13
14 2,3
P L
r
i
>
d
- - T o r 6
Scheme 1
This static picture provides a framework against which the dynamics of the reaction must be reconciled. The 1 step in the putative repair mechanism (see Scheme 1) involves the formation of a CPD radical anion (TT *") by electron transfer from ^ L ^ " + TT -> PL * + TT * " with rate constant ke , leaving the FAD cofactor in its semiquinone state, PL *. The thymidine dimer radical subsequently undergoes reorganization spontaneously at room temperature to form the monomer base and base anion radical (T-T* ). Bond scission is st
sq
T
sq
-
10
139 thought to compete with recombination of the electron from TT * ~ + PL * - » TT + PL ~ but this rate (k ) must be much slower than the (initial) bond scission rate, since the quantum yield for repair is nearly unity. Bond breaking may involve a sequential mechanism where the C5-C5' bond breaks with rate ki followed by scission of the C6-C6' bond (k ) (or vice versa), or that bond scission is concerted (k ). It is generally accepted that the electron from T - T *~ sq
red
rec
2
con
is transferred back to FADH* to re-reduce the flavin to FADH (k r) although when this back electron transfer step occurs has not been firmly established. Dissociation of the protein-substrate complex is thought to occur at a much slower rate. While this mechanism is reasonable, there is a paucity of experimental data to test its validity. This Proceedings offers experimental evidence to flesh out this mechanism. be
Results and Discussion We probed the temporal evolution of the ultraviolet spectral region with femtosecond resolution following absorption of blue light by photolyase because the repair of the CPD involves breaking the cyclobutane ring with concomitant reformation of the 5-5' and 6-6' C=C bonds of the adjacent thymidines. These bonds absorb strongly at 265 nm (ε = 9,500 M" cm" ). A minimal substrate length of 5 thymidines was used to obtain tight binding between the protein and substrate. The kinetics probed at 265 nm after excitation at 396 nm were measured for FADH", PL f, PL ":d(TT) , and PL ":d(TT) with ~ 850 fs resolution. Transient absorption decays of PL ~ and the PL ~:d(TT) complex (n = 2, 5) are shown in Figure 1. The lifetimes for the exponential modelfitshave been compiled in a previous publication. As a control, the decay of 'FADH" in pH 8.2 phosphate buffer was monitored at X = 265 nm (data not shown). FADH" was generated from F A D by photoreduction in the presence of oxalate as described by Heelis et al J A negative-going transient was obtained which was assigned as a ground state bleach. The amplitude of the 'FADH" decay was linear with pump power above 5.2 μΙ/pulse. The ^ADH" decay can not befittedto a single exponential. A two-component exponential fit gives a X\ ~ 4 ps component and a longer-lived component whose value depends on the length of the acquisition window used. The fast component agrees well with that obtained by Enescu et al. 1
9,11
re(
red
2
red
red
5
red
n
12
probç
0X
4
1
5
140 l
The signal for photolyase without substrate ( PL f) goes negative as well due to the ground state bleach of the FADH" cofactor and then rises slowly and approaches ΔΑ = 0 after about 3,000 ps ( V , Figure 1). Two lifetimes were necessary to fit this decay, with most of the amplitude in the longer τ - 851 ps component, and a small amplitude component with a lifetime of τι ~ 7 ps. It re(
2
0.003 0.002 0.001 Q
0.000
Ο ^ -0.001
% -0.002 -0.003 500
1000
1500
2000 2500 3000
Time (ps) Figure 1: Transient absorption kinetics ofphotolyase: CPD complexes
should be noted that this is not really a proper control for the substrate studies as no electron transfer is expected in the absence of the CPD. Interestingly, a recent femtosecond fluorescence upconversion study of the ^ADH" cofactor in photolyase by Zhong and coworkers returned a single lifetime of 850 ps. The small difference between these measurements has yet to be reconciled. The decay of the PL ":d(TT) complex was obtained as a function of mole ratio of the protein to the substrate. This ratio was varied between 1:5, 1:10, and 1:20 (•,#, and Δ respectively, figure 1). The solid lines represent a three-exponential function fit to the data. The decay of PL ~:d(TT) with a mole ratio of 1:5 shows a very fast component (τ\ ~ 9 ps) that is similar to the 7 ps component of 'PLrecf. The fast components for the d(TT) 1:10 (τ ι ~ 49. ps) and 1:20 (τι - 38 ps) both trend towards shorter times with increasing 18
red
2
1
red
2
2
141 concentration of the substrate. The zero crossing changes from about 1,600 ps->400 ps-> 100 ps for the 1:5, 1:10, and 1:20 mole ratios respectively. The shift in the zero-crossing as a function of substrate suggests that the PL d~:d(TT) transients represent a sum of decays from protein:substrate complexes and free protein. This can be seen in Figure 2, where the transients for the dinucleotide substrate at different mole ratios to the protein are fitted using a linear combination of the PL d~ and the 'PL^cf :d(TT) decays. 1
re
2
!
re
5
Time (ps) Figure 2: Fit (solid lines) of the dinucletoide kinetics to a linear combination of the pentameric protein: substrate and protein-only kinetics. The initial rise time from the excitation pulse is particularly interesting because it contains information regarding the electron transfer lifetime from the photoexcited FADH* to the initial acceptor in the protein:substrate complex. Both Tj and the zero-crossing are fastest for the PL d~ .d(TT) complex with an average mole ratio of 1:4. The initial negative signal recovers promptly with a lifetime of τι = 28 ± 13 ps. We assign this as the electron transfer lifetime, τι = (key)" , which is about four times shorter than previously thought. This in turn requires a re-thinking of the electronic coupling between the 'FADH" and the initial acceptor, suggesting that the initial acceptor may be a nearby molecule (e.g. adenine in FAD, a Trp residue) that the CPD is closer to the flavin than predicted. If the CPD is the initial acceptor then our results support the experimental work of Park et al and Berg and Sancar and the theoretical treatment of Antony et at (all of these studies used the E. coli protein). I
,
re
5
10
1
617,20
14
2
142 To better explore the mechanism for repair, the first 100 ps kinetics for the proteimpentameric substrate is shown in figure 3. The zero crossing occurs around 60 ps for the repair of the pentameric substrate. If no repair took place then the signal should decay to ΔΑ ~ 0. Instead, the absorbance change becomes positive and increases until a maximum is reached around 1,500 ps (see figure
0.001 o.ooo Q -0.001 Ο % -0.002 -0.003 0
20
40
60
Time (ps)
80
100
Figure 3: Initial rise of265 nm transient for the PL:(TT) complex. 5
1). These data suggest that there are two quite different time scales at play and that scission of the cyclobutane bond of the CPD takes place in two steps. The first step occurs directly after electron transfer to the CPD with little or no activation barrier. The second bond scission takes place at least an order of magnitude slower with a much larger activation barrier. This is at least consistent with the observations that an activation barrier exists for bond scission but not for electron transfer. A target analysis based on Scheme 1 was performed in an attempt to account for these observations. The differential equations implicit in Scheme 1 were integrated using reasonable estimates for the 265 nm extinction coefficients for the putative species. The transient absorption data were fitted to this model in a nonlinear least squares sense using ki and k as adjustable parameters. The fits for three scenarios are shown in figure 4. Both a linear time base (top panel) and a logarithmic time base (bottom panel) are shown. Roughly equivalent fits are obtained for two different sequential models (250 ps/250 ps and 60 ps/600 ps), and the (quasi) concerted model (300 ps/6 ps). Clearly more experimental information is needed to distinguish between these different 10
15
2
143
mechanisms mechanism. We are currently engaged in visible light continuum probe studies to obtain this additional information.
Summary In this Proceedings, we have used femtosecond transient absorption kinetics to understand the mechanism of A. nidulans DNA photolyase with oligothymidine substrates in real time. The two color experiments suggest that the repair mechanism may be sequential, although quantitative modeling of the transient absorption data do not lead to a definitive picture of the process. The concentration dependence of the minimal substrate suggests that a longer substrate should henceforth be considered "minimal". The use of a pentameric substrate appears to bind adequately to the protein. Thus, in spite of significant effort, including our own, a detailed understanding of the repair function of DNA photolyase is still in its infancy.
Acknowledgements We gratefully acknowledge research supportfromNSF MCB-0347087.
144
References (1) Antony, J.; Medvedev, D.M.;Stuchebrukhov, A. A. J. Am. Chem. Soc. 2000, 122, 1057-1065. (2) Berg, B. J. V.; Sancar, G. B. J. Biol. Chem. 1998, 273, 20276-20284. (3) Christine, K. S.; MacFarlane IV, A. W.; Yang, K.; Stanley, R. J. Journal of Biological Chemistry 2002, 277, 38339-38344. (4) Enescu, M.; Lindqvist, L.; Soep, B. Photochem. Photobiol. 1998, 68, 150-156. (5) Fisher, G. J.; Johns, H. E. Pyrimidine Photodimers. In Photochemistry and Photobiology of Nucleic Acids; Wang, S. Y., Ed.; Academic Press: New York, 1976; Vol. 1; pp 226-294. (6) Hahn, J.; Michel-Beyerle, M.-E.; Rösch, N. J. Mol. Model. 1998, 4, 7382. (7) Heelis, P. F.; Hartman, R. F.; Rose, S. D. Photochemistry and Photobiology 1993, 57, 1053-1055. (8) Heelis, P. F.; Kim, S. T.; Okamura, T.; Sancar, A. J. Photochem. Photobiol., Β 1993, 17, 219-28. (9) Jorns, M . S.; Wang, B.; Jordan, S. P.; Chanderkar, L. P. Biochemistry 1990, 29, 552-61. (10) Langenbacher, T.; Zhao, X.; Bieser, G.; Heelis, P. F.; Sancar, Α.; Michel-Beyerle, M. E. J. Am. Chem. Soc. 1997, 119, 10532-10536. (11) MacFarlane IV, A. W.; Stanley, R. J. Biochemistry 2001, 40, 1520315214. (12) MacFarlane IV, A. W.; Stanley, R. J. Biochemistry 2003, 42, 85588568. (13) Mees, Α.; Klar, T.; Gnau, P.; Hennecke, U.; Eker, A. P. M.; Carell, T.; Essen, L.-O. Science 2004, 306, 1789-1793. (14) Park, H.-W.; Kim, S.-T.; Sancar, Α.; Deisenhofer, J. Science 1995, 268, 1866-72. (15) Rustandi, R. R.; Jorns, M. S. Biochemistry 1995, 34, 2284-8. (16) Sancar, A. Chemical Reviews 2003, 103, 2203 (17) Sanders, D. B.; Wiest, O. J. Amer. Chem. Soc. 1998, 121, 5127-5134. (18) Saxena,C.;Sancar, Α.; Zhong, D. J. Phys. Chem. Β 2004, 108, 18026. (19) Tamada, T.; Kitadokoro, K.; Higuchi, Y.; Inaka, K.; Yasui, Α.; De Ruiter, P. E.; Eker, A. P. M.; Miki, K. Nature Structural Biology 1997, 4, 887-891. (20) Weber, S.; Moebius, K.; Richter, G.; Kay, C. W. M. J. Am. Chem. Soc. 2001, 123, 3790-3798.
