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Hole Transport Dynamics in Mixed Sequence DNA Can Vary with Salt Concentration: An Experimental and Theoretical Analysis† William B. Davis,* Chad C. Bjorklund,‡ and Paul S. Cho School of Molecular Biosciences, Washington State UniVersity, Pullman, Washington 99164-7520, United States ReceiVed: July 30, 2010; ReVised Manuscript ReceiVed: October 4, 2010
Long-range hole transport (HT) through DNA is a well-established phenomenon with important applications in the fields of material science, biochemistry, and cell biology. However, the role that the surrounding environment plays in modulating DNA HT dynamics is not well understood and will impact the development of DNA HT theories. Here, we report that varying the bulk salt concentration can affect the dynamics of long-range HT in a mixed sequence DNA duplex. Using a previously described AQ-601 construct, we show that HT to distal guanines is maximal at physiological salt concentrations (100-200 mM), and declines as [NaCl] increases or decreases. Using circular dichroism, we observe that the 601 sequence has the unusual property of underwinding with increasing salt, but the connection of these changes in secondary structure to DNA HT dynamics is not clear. Using all-atom molecular dynamics simulations on decamer duplexes from AQ-601, we show that the changes in HT dynamics do not correlate with salt-dependent fluctuations in local DNA structure and/or electrostatic environment. Since the HT results best correlate with known changes in oxygen diffusion rates vs [NaCl], the rate of hole trapping, involving reaction of the guanine radical cation with oxygen species, appears to control the DNA HT dynamics in this duplex. 1. Introduction DNA occupies a unique position at the intersection of biological and materials chemistry because of its powerful combination of tunable electronic and structural properties. One of these electronic properties is the ability of DNA to transport charge through its semiregular, π-stacked nucleobase core. Prior studies have established that long-range DNA hole transport (HT) between guanine (G) nucleobases is efficient over 60 base pairs (200 Å)1,2 with the effects of sequence and the initial oxidizing agent on these reactions extensively studied using a combination of experimental and theoretical approaches.3-7 For systems with well-characterized structures, short transport distances, and one hole acceptor, HT rates generally fall off exponentially with increasing donor-acceptor distance.8,9 HT rates and dynamics in DNA-based donor-bridge-acceptor systems are generally analyzed using the Marcus-Jortner-Levich equation
kET )
[
2π 2 exp(∆G0′ + λ)2 H p DA 4λKBT
]
(1)
where HDA is the electronic coupling between donor and acceptor, ∆G0′ is the Gibbs free energy for the reaction and is related to the difference in redox potentials between the donor and acceptor, and λ is the reorganization energy. The exponential distance dependence of the rates in these systems is accounted for by using the superexchange framework10 to describe the dependence of HDA on the number of base pairs separating donor and acceptor. †
Part of the “Mark A. Ratner Festschrift”. * To whom correspondence should be addressed. Tel: 509-335-4930. Fax: 509-335-4159. E-mail:
[email protected]. ‡ Current Address: MD Anderson Cancer Center, Houston, TX.
In systems with long transport distances between neighboring guanines4 (g4 AT base pairs) and/or multiple guanine sites1,2,11,12 an algebraic dependence of HT rates/efficiency on distance is observed. These experiments require theoretical frameworks beyond superexchange to describe their HT dynamics, and there are several currently competing models in the literature. The first model is thermally induced hopping (TIH) in which resonance between the guanine donor and the adenines (A) in the bridge will lead to movement of a localized hole onto the bridge and result in efficient long-range HT.13 Supporting this model are theoretical studies that have shown that the Boltzmann factor arising from the ∼0.4 eV gap between the oxidation potential of G and A14 can be overcome by fluctuations of DNA structure and/or the surrounding ionic environment.15-17 Two alternatives to TIH are the conformational gating model (CG)18 and ion-gated phonon-assisted hopping (IGPAH) model.19 Both of these models rest on the premise that structural fluctuations lead to transport-competent states in DNA characterized by a hole delocalized over multiple base pairs, and that hole transport occurs between these states and not the individual bases. Where these two models differ is in the origin of the structural fluctuations that control transport. In CG, fluctuations in DNA secondary structure lead to hole transport between preformed, transient 3-5 base pair (bp) DNA structures that maximize electronic coupling. In contrast, IGPAH posits that after a delocalized hole (polaron) is formed in DNA the fluctuations between guanine-counterion bound states control the dynamics of polaron migration. Since all of these theoretical approaches rely on fluctuations in both the local DNA structure and the interaction of DNA with the surrounding polar media, experiments that explore how changes in the environment surrounding the nucleobase stack influence long-range HT are needed for further model refinement. As one example, the effects of backbone neutralization on DNA HT have been explored using multiple approaches
10.1021/jp107191m 2010 American Chemical Society Published on Web 10/20/2010
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including duplexes containing methylphosphonate groups and DNA-peptide nucleic acid (PNA) hybrids. Surprisingly, the loss of negative charge in a small region of a methylphosphonatemodified duplex can lead to a dramatic loss in long-range HT efficiency,19 whereas DNA-PNA hybrids with complete loss of charge along the backbone of the PNA strand are expected to support robust hole transport.20,21 A more challenging class of experiments to interpret has focused on how DNA CT is influenced by interactions with the packaging and regulatory biomolecules found in cells, including bound transcription factors,22 DNA repair enzymes,23 restriction endonucleases,22,24 polyamines,25 and the nucleosome core particle.26-28 Overall, these studies have indicated that HT in DNA is often sensitive to the binding of other biomolecules with differences in HT dynamics typically attributed to global changes in DNA structure, for example, bending, or to electrostatic and polar changes arising from specific molecular contacts.22,24,27 The complexity of the molecular changes occurring in these experiments makes it difficult to tease apart the relative contributions of changes in the local DNA electrostatic environment due to, for example, counterion release, the ordering of water at contact regions, and interactions of charged amino acid side chains with nucleobases, from the changes in DNA structure induced by binding. Furthermore, since these experiments have only interrogated the final distribution of guanine damage arising from competition between hole hopping and hole trapping at the guanines, it is not clear if the environment is perturbing DNA HT rates or the rates of the chemical reactions involving the guanine radical that are important for hole trapping. In our previous experiments on DNA HT in nucleosomes,26 we utilized a duplex consisting of an anthraquinone (AQ) photooxidant covalently coupled to the 601 nucleosome binding sequence (duplex AQ-601). To understand DNA HT dynamics in a DNA-protein complex as sophisticated as the nucleosome, it was necessary to first understand the dynamics of DNA HT in naked AQ-601. The experimental results reported here demonstrate that the ionic strength of the environment surrounding AQ-601 can strongly influence the dynamics of DNA CT in this mixed sequence duplex, even in the absence of any bound proteins. To investigate how the concentration of NaCl affects the global and local structure of naked AQ-601 and the local electrostatic environment surrounding the guanine oxidation sites, we report the results of near-UV circular dichroism (CD) spectroscopy and an analysis of molecular dynamics simulations, similar to that previously carried out on repeat sequence DNA16 on this mixed sequence DNA system. 2. Experimental Procedures A. DNA Charge Transport Experiments. The 162 base pair duplex AQ-601 was prepared by PCR as described previously.26 After purification by gel electrophoresis, the duplex was labeled on its free 5′-terminus using [γ-32P]ATP (Perkin-Elmer) and T4 DNA Kinase. Aliquots containing approximately 100 pmol of AQ-601 were diluted into 100 µL of 10 mM phosphate, pH ) 7.0 buffer containing NaCl at concentrations of either 10, 100, 200, 400, 600, 800, or 1000 mM. The DNA solutions were then dialyzed against 300 mL of buffer with the desired [NaCl] at 4 °C and transferred to 1.5 mL nonsilanized tubes. Samples were irradiated at room temperature for 60 min in the rotating sample carousel of a Luzchem photoreactor with 10 UV-A lamps. Following the addition of glycogen, the samples were precipitated using ethanol. The resulting pellets were resuspended in 10% piperidine and incubated at 90 °C for 30 min. Next, the samples were dried by evaporation under vacuum and washed twice with water. After dissolving the pellets in loading
Davis et al. buffer, samples were loaded and resolved on 6 M urea/8% PAGE gels. Autoradiography of these dried gels was used to quantify the yields of piperidine-sensitive guanine oxidation products. B. UV CD Spectroscopy. A plasmid (pLMG601-23)29 containing 23 repeats of the 147 bp 601 nucleosome binding sequence30 was purified from E. coli DH5R cells using a midi prep kit (Wizard). After digestion of the plasmid with EcoRV, the desired 601-147 duplex was purified from preparative agarose gels. One micromolar samples of 601-147 were made in 10 mM phosphate, pH ) 7.0 buffer (1 mL) with the following NaCl concentrations: 10, 200, 400, 600, 800, and 1000 mM. The samples were dialyzed against 300 mL of the appropriate buffer for 1 h at 4 °C. Each sample was next placed in a 4 × 10 mm quartz cuvette and the CD spectrum recorded using an Aviv CD Spectrophotometer. The final concentration of 601147 in each sample was calculated from A260 using Beer’s Law, and these values were used to transform the raw CD data into ∆ε values.31 C. Molecular Dynamics Simulations. NPT simulations on two decamer sequences from AQ-601 were carried out using NAMD software (ver. 2.7b1)32 on Windows-based computers. CHARMM 27 parameters33,34 were used in all calculations. The two DNA decamers were (1) duplex 601-GG1 with the strand sequences 5′-CAGGGCGGCC-3′ (GG1 and GGG2 underlined) and 5′-GGCCGCCCTG-3′, and (2) duplex 601-GG4 with strands 5′-CCTCGGCACC-3′ (GG4 underlined) and 5′-GGTGCCGAGG-3′. These duplexes were constructed in an ideal B-form geometry using Hyperchem (Hypercube), and the VMD plugin Solvate was used to create a box of explicit waters (TIP3) extending 12.5 Å away from the DNA in all directions. Charge neutralization was carried out by manually replacing one water molecule within 5.0 Å of each phosphate in the backbone with a sodium atom (18 total). For the 100 mM NaCl simulations, three additional sodium atoms and three chloride atoms were randomly placed in the waterbox, while the 400 mM simulations required the addition of 12 sodium and 12 chloride atoms. The four initial structures were subjected to 1000 steps of conjugate gradient minimization three times, the first with all DNA atoms fixed, the second with the DNA backbone fixed, and the final round with the entire system unrestrained. During NPT simulations, periodic boundary conditions were applied and a 1 fs time step was used during propagation. To begin these calculations, each system was slowly heated (4 ps) to 298 K at 1 atm pressure by coupling the system to a Langevin piston. The backbone atoms were kept fixed during heating, and Particle Mesh Ewald electrostatics with a 8.5 Å switching distance and 10 Å cutoff distance were used. Next, the system was equilibrated for 4 ps with the backbone atoms fixed, followed by 400 ps of NPT simulation with no restraints. At this point, all four simulations were deemed equilibrated since system volume, system temperature, and total energy were stable. Ten nanoseconds of NPT simulation were next collected with the structural coordinates recorded every 1 ps. VMD35 was used for analysis of the MD simulation output and visualization of the simulation trajectories. The values of base pair and base step structural parameters were obtained from MD snapshots using the program X3DNA.36 To calculate the total electrostatic potential (Vtot) acting upon the center of mass of selected guanine nucleotides in 601-GG1 and 601-GG4, we modeled the surrounding atoms from the waters, ions, and surrounding DNA as point charges and used the equation
Hole Transport Dynamics in Mixed Sequence DNA
Vtot )
1 4π∈0
q
∑ rii
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(2)
i
where qi is the MM charge on point charge i, and ri is the distance from the point charge i to the center of mass of the guanine of interest. Since electrostatic potential is a linear function, we split the potential into its component parts
Vtot ) VDNA + VI + VW
(3)
with VDNA the potential arising from the surrounding DNA, VI the potential due to the sodium and chloride ions, and VW the potential generated by the water molecules. 3. Results A. DNA Hole Transport Dynamics in AQ-601 Vary With [NaCl]. The selective excitation of AQ in AQ-601 by UV-A irradiation leads to the formation of a guanine radical cation (G•+) that then hops between neighboring guanines until competing reactions with water or oxygen trap the hole.37 Hot piperidine generates strand breaks at the sites of hole trapping, and the damage distribution and site-dependent oxidation yields are evaluated by the autoradiography of DNA sequencing gels (Supporting Information Figure 1). In this study, we have monitored the yields of guanine oxidation in AQ-601 samples irradiated in 10 mM phosphate, pH ) 7.0 buffers with the following NaCl concentrations: 10, 100, 200, 400, 600, 800, and 1000 mM. In AQ-601, we observe oxidative damage at seven guanine sites in the 601 sequence (labeled GG1-G7; Figure 1A) after UV-A irradiation at all salt concentrations. Although not shown, we also confirmed the findings of our previous report26 that there is no guanine damage observed in AQ-601 if either UV-A irradiation is omitted, or the AQ photooxidant is replaced by a 5′-amino linker. A standard approach for visualizing DNA oxidation dynamics is to normalize the damage yields at distal sites by that measured at the site proximal to the hole injection site. Therefore, we divided the guanine oxidation levels at sites GGG2-G7 by the damage yield for GG1 (Gx/GG1; Figure 1B) at each salt concentration studied. The data points in Figure 1B are average values calculated from three independent experiments, and the error in the data is not shown since it was 0.5 for all three guanine oxidation sites at both salt concentrations. Therefore, the total electrostatic potential at these guanine oxidation sites is largely dictated by fluctuations in the surrounding solvent, and not local fluctuations of DNA structure. Furthermore, this trend is independent of bulk salt concentration over the range of values studied. We next calculated the correlation coefficients among the electrostatic potentials arising from the surrounding DNA and ionic media (Table 5). The first observation is that even though fluctuations in VDNA are not correlated to either VW or VI, there is a significant negative correlation between VDNA and VW+I. This partial cancelation of the fluctuations in the electrostatic potentials arising from DNA structure and the ionic media helps explain why Vtot remains small throughout the 10 ns simulations of 601GG1 and 601-GG4. As expected from the qualitative observations in the previous paragraph, there is a high negative correlation between the fluctuations in VW and VI at all three guanine sites regardless of salt concentration. To see if these coupled fluctuations in VW and VI were due to interactions of guanine residues with individual sodium atoms or, conversely, with the entire ionic atmosphere, we calculated the correlation coefficients between the fluctuations in VW and the minimum guanine-sodium atom distance (RNa). As the data in Table 5 shows, there is a strong positive correlation between RNa and VW for all three guanine sites at both salt concentrations. Although not shown, the correlation between RNa and VI is negative and nearly identical in absolute value as that between RNa and VW. The simplest molecular explanation for this observation is when a Na+ comes into close contact with a guanine base, VI increases because of the proximity of the sodium ion and VW decreases due to the altered local water environment. The fact that the correlation coefficient between RNa and VW is not 1.0 (or -1.0 in the case of RNa and VI) indicates that there is a nontrivial contribution from the bulk salt atmosphere to VI. There is, however, no correlation between RNa and VDNA, VW+I, or Vtot (Table 4 and data not shown). The lack of correlation between RNa and Vtot indicates that specific guanine-sodium contacts do not significantly influence the overall electrostatic environment of a guanine nucleobase in DNA, similar to the results reported previously by Kubar and Elstner.16 In summary, the electrostatic potential at any guanine residue in DNA is largely dictated by fluctuations in the surrounding ionic medium, and not local structural fluctuations in DNA. Furthermore, it is best to view the influence of the ionic medium as arising from the collective interactions of a nucleobase with both water molecules and ions. Given the correlation between fluctuations in Vtot and guanine ionization potential,16 the results of this study indicate that the experimental changes in DNA HT dynamics observed in AQ-601 are not expected to arise from salt concentration dependent fluctuations in hole-hopping rates. 4. Discussion The experiments described here demonstrate that simple changes in the surrounding environment can have measurable effects on the dynamics of long-range hole transport in duplex DNA. For AQ-601, varying the concentration of NaCl between 10 mM and 1 M affects both the efficiency of long-range HT and the distribution of damage among the seven resolvable guanine oxidation sites. The largest change is observed between 10 mM and 100 mM salt, since the yields of guanine oxidation at the two AQ-proximal sites (GG1 and GGG2) decrease
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Figure 4. Fluctuations in the base pair parameter rise from 10 ns MD simulations for the oxidation sites GG1, GGG2, and GG4 at (A) 100 mM NaCl and (B) 400 mM NaCl. Note that the y-axis is scaled the same on each figure to highlight the decreased fluctuations in rise at higher salt concentrations.
TABLE 3: Salt Dependence of the Average Values of the Electrostatic Potential (in Volts) at Three Guanine Oxidation Sites Obtained from 10 ns MD Simulationsa site GG1 GGG2 GG4 site GG1 GGG2 GG4 a
[NaCl] (mM)
VDNA
VW
VI
100 400 100 400 100 400
-22.2 ( 0.3 -22.2 ( 0.3 -22.4 ( 0.4 -22.5 ( 0.4 -22.0 ( 0.4 -22.1 ( 0.3
7.2 ( 1.4 7.3 ( 1.8 7.5 ( 1.3 7.3 ( 1.5 8.8 ( 1.3 7.1 ( 1.4
16.2 ( 1.3 16.1 ( 1.8 15.9 ( 1.1 16.4 ( 1.6 14.5 ( 1.2 16.5 ( 1.4
[NaCl] (mM)
VW+I
Vtot
100 400 100 400 100 400
23.4 ( 0.4 23.4 ( 0.4 23.7 ( 0.5 23.7 ( 0.5 23.4 ( 0.4 23.6 ( 0.4
1.2 ( 0.3 1.2 ( 0.3 1.3 ( 0.3 1.2 ( 0.4 1.3 ( 0.3 1.5 ( 0.3
The standard deviations of these parameters are also indicated.
sharply. This type of behavior is unusual in the literature of DNA HT, most likely because (1) most studies are performed
at a single salt concentration, and (2) most transport studies utilize DNA duplexes with nonrandom, repetitive sequences. We note that the maximal yield of long-range DNA HT occurs around physiological salt concentrations (∼150 mM NaCl) in AQ-601. Even though the 601 DNA sequence is not found in any organism, it has phased A/T and G/C tracts shared by a majority of the nucleosome binding sequences found in the genomes of eukaryotes like chicken and yeast.56 Although there is only limited evidence for DNA HT in the eukaryotic nucleus,57 if it does occur then the yields of long-range HT may be optimized at physiological salt concentrations in nucleosome binding sequences in the nucleus. Further, these expectations would help support the recent contention that DNA HT plays a role in allowing DNA repair proteins to identify damaged nucleobases in a coordinated fashion.58 The CD spectral changes observed here indicate that the 601 sequence becomes underwound with increasing NaCl. It is difficult to correlate the secondary structure changes in AQ601 with the hole transport dynamics in this duplex since the secondary structure appears to change smoothly, whereas there
Hole Transport Dynamics in Mixed Sequence DNA
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Figure 5. A comparison of the 100 ps time-scale fluctuations in the electrostatic potential components at site GGG2 from 100 mM and 400 mM NaCl MD simulations. (A) VDNA, (B) VW, (C) VI, (D) VW+I, and (E) Vtot.
are abrupt changes in the HT dynamics at physiological salt concentrations. However, we do want to comment on this unusual behavior since other DNA sequences whose structures have been interrogated by CD, for example, calf thymus DNA, show the opposite behavior (they become overwound) with increasing salt. We speculate that there is a relationship between the underwinding of the 601 sequence and its ability to form stable nucleosome core particles.30 When histone proteins are reconstituted with DNA to form a nucleosome, a salt dependent dialysis procedure is typically utilized. Since one of the hallmarks of nucleosome packaging is DNA underwinding,59 the propensity of the 601 duplex to become underwound at high salt may predispose this construct to form stable nucleosomes. It would be of interest to test this hypothesis by comparing the salt-dependence of the CD spectra of high- and low-affinity nucleosome binding sequences. If the salt-dependence of DNA HT dynamics were measured in these sequences concomitantly, then a correlation between secondary structure changes and DNA HT dynamics could be investigated in genomic sequences. The underwinding of the 601 sequence is not observed in the MD simulations of 601-GG1 and 601-GG4; for instance neither the average values nor the standard deviations of the base step parameters rise and twist change appreciably between
100 and 400 mM NaCl. Most likely, 601 underwinding is a cooperative phenomenon involving larger DNA regions, perhaps even the entire sequence. Evidence for this is provided by recent MD simulations showing that R-satellite DNA, another nucleosome binding sequence, can spontaneously form bent structures with short end-to-end distances.60 Further, the relative stability of these bent conformers increases as the salt concentration increases from 10 mM to 1 M. Spontaneous bending of duplexes shorter than the classical persistence length of DNA (∼150 bp) has also been observed experimentally, although the origin of this behavior is in some dispute.61,62 Taken together, these prior studies indicate that the structural changes monitored in 601147 by CD may arise from spontaneous, cooperative bending of large regions of this duplex with increasing salt. If this is the case, then using MD simulations on short oligos may miss DNA structural dynamics important for understanding HT dynamics in longer transport systems. The only environmental parameter we have changed in these experiments and MD simulations is the concentration of bulk salt. The effect of salt on DNA structure and function has been of intense interest since the earliest biochemical studies of nucleic acids.63 The simplest model describing the interaction between DNA and the surrounding ionic atmosphere is coun-
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Figure 6. Comparison of the 10 ns fluctuations in the electrostatic potentials at site GGG2 in MD simulations at 100 and 400 mM NaCl. (A) VDNA, (B) VW, (C) VI, (D) VW+I, and (E) Vtot.
TABLE 4: Correlation Coefficients of the Fluctuations in Vtot and the Four Components of the Electrostatic Potential at Sites GG1, GGG2, and GG4a Site GG1 GGG2 GG4 Site GG1 GGG2 GG4
[NaCl] (mM)
Vtot/VDNA
Vtot/VI
Vtot/VW
100 400 100 400 100 400
0.22 0.056 0.063 0.22 0.30 0.22
0.051 0.15 0.10 0.014 0.015 0.068
0.16 -0.0011 0.14 0.18 0.16 0.12
TABLE 5: Correlation Coefficients of the Fluctuations in the Electrostatic Potentials Arising from DNA (VDNA), Ions (VI), and Water (VW)a site GG1 GGG2 GG4
[NaCl] (mM)
Vtot/VW+I
Vtot/RNa
site
100 400 100 400 100 400
0.71 0.68 0.56 0.60 0.52 0.61
-0.073 -0.13 -0.086 -0.078 0.0072 -0.033
GG1 GGG2 GG4
[NaCl] (mM)
VDNA/VI
VDNA/VW
VDNA/VW+I
100 400 100 400 100 400
-0.097 0.14 -0.065 -0.24 -0.35 -0.22
-0.061 -0.28 -0.26 0.033 0.15 0.033
-0.53 -0.69 -0.79 -0.64 -0.66 -0.64
[NaCl] (mM)
VW/VI
VW/RNa
100 400 100 400 100 400
-0.96 -0.98 -0.91 -0.95 -0.94 -0.95
0.71 0.73 0.51 0.62 0.74 0.58
a Also shown is the correlation coefficient between Vtot and the minimum sodium-guanine distance (RNa).
a Also shown is correlation between the minimum sodium atom-guanine distance (RNa) and VI and VW.
terion condensation theory.64 In this theory, DNA is treated as a cylinder with a linear charge density along its surface and the counterions “condense” with DNA to lower the effective surface charge density to unity. Experimental results65 have confirmed the basic tenets of this theory. Further, they have shown that
the condensed sodium atmosphere surrounding DNA is not expected to change between 3 mM and 1.3 M NaCl, a range that encompasses the experiments described here. In general, we find that our simulation results are largely consistent with the expectations of counterion condensation theory. For instance,
Hole Transport Dynamics in Mixed Sequence DNA we observe that the average values of VI and VW+I in these simulations are largely independent of [NaCl] (Table 3), indicating that we should not expect large changes in the electrostatic environment surrounding the guanine oxidation sites in AQ-601 in our experiments. The MD simulations also provided a molecular-level picture of DNA-ion interactions. For instance, the fluctuations in VI during the simulations are due to the movements of the sodium ions, and we observe that individual ions can play a significant role in the fluctuations of the electrostatic potential. This conclusion comes from the high correlation coefficient calculated between RNa and VI (and VW) for the guanine oxidation sites in 601-GG1 and 601-GG4. Further analysis of our MD simulations shows that during time steps when VI is large (RNa small) specific molecular contacts are made between the N7/O6 atoms on guanine and a Na+ ion (data not shown). This specific interaction between the major groove face of guanine and alkali metals has been observed in previous MD simulations19,66,67 and in a previous crystal structure of DNA complexed with Tl+ ions.68 In the IGPAH Theory of Landman and Schuster,19 these ion-guanine interactions were proposed to control the motion of the hole polarons in DNA since there would be charge repulsion if a positively charged hole were to come in contact with a neighboring Na+ ion. Given the facts that (1) the fluctuations and average values of VI for sites GG1 and GGG2 are identical at 100 and 400 mM NaCl, and (2) there is no correlation between the fluctuations in Vtot and RNa at any guanine site in 601-GG1 or 601-GG4 (Table 3), we anticipate that changes in these guanine-Na+ interactions are not expected to contribute extensively to the salt-dependent oxidation yields in AQ-601. We also note that our findings are consistent with simple DNA models that predict that the time-averaged major groove occupancy of a monovalent cation is independent of salt concentration over the range of concentrations we utilized.69 Therefore, we find that our theoretical results are consistent with previous studies of DNA-ion dynamics, and that they do not support the fundamental tenants of the IGPAH model of DNA hole hopping. Previous MD simulations15,16 have indicated that water fluctuations appear to contribute more to the electrostatic potential energy experienced by a guanine residue than fluctuations in the distribution of ions. Furthermore, the work of Kubar and Elstner16 has shown that the counterion atmosphere interacts with DNA in a collective, not site-specific, manner. Our simulations completely concur with these previous studies since we also observe that there is no correlation between fluctuations in RNa and Vtot, indicating that interactions between guanines and individuals ions are not responsible for changes in the local electrostatic potential. Therefore, if fluctuations in Vtot do make significant contributions to DNA HT reactions, via their influence on the oxidation potential of guanine residues,16 there should be minimal, if any effect of bulk salt on long-range DNA HT reactions. This expectation is contrary to our experimental studies on AQ-601 since there are significant changes in the hole transport efficiency to the sites furthest from AQ (i.e., GGG3-G7) as salt decreases from 100 to 400 mM. Furthermore, the sharp changes in the site oxidation yields in AQ-601 between 10 and 800 mM NaCl are not explained by the trends in Vtot obtained from these simulations. The other time dependent property of DNA that is expected to influence both long-range HT and site-dependent damage distributions are base step structural fluctuations. There are currently two pictures of how DNA fluctuations can influence DNA HT dynamics. First, the global DNA fluctuations are
J. Phys. Chem. C, Vol. 114, No. 48, 2010 20831 expected to contribute to the electrostatic potential felt by a guanine residue and thus influence that site’s oxidation potential. Even though the fluctuations in VDNA are negatively correlated to the fluctuations in the environment (VW+I but not VW or VI), the correlation coefficient between Vtot and VDNA is near zero. This indicates that there is little if any effect of global DNA structural fluctuations on the electrostatic potential felt by an individual guanine residue. In a more local picture, DNA structural fluctuations are expected to influence HT rates (eq 1) through their influence on the parameters: (1) ∆G0′, via changes in guanine oxidation potential mediated by changes in base step parameters,44 and (2) HDA, through changes in wave function overlap integrals due to fluctuations in base-step parameters. Since the ∆G0′ for hole hopping between guanines in DNA is expected to be nearly zero,70 eq 1 indicates that small changes in free energy can lead to measurable changes in the hole hopping rates. In our simulations of duplexes 601-GG1 and 601GG4, salt concentration does not strongly affect the average values or standard deviations of the base step parameters. Rather, increasing [NaCl] from 100 to 400 mM leads to significant changes in the range of fluctuations displayed by the base steps in DNA. For instance, the damping of fluctuations in the parameter rise at the four GG steps investigated here would lead to a smaller range of ∆G0′ values, based on the results of prior investigations.44 In the case of HDA, theoretical calculations have shown that electronic coupling decreases for all base pair steps with increases in rise, whereas decreases in twist lead to varying effects.41-43 Since the overall coupling between any two guanine oxidation sites in DNA is affected by changes in the coupling between all of the intervening base steps, the connection between changes in the structural fluctuations in AQ-601 is currently uncertain and future theoretical analyses of this system are required. Thus far in our Discussion, we have followed the typical line of reasoning that changes in guanine oxidation yields arise from changes in the hole hopping rates. However, our experimental guanine damage distributions arise from a competition between hole-hopping rates (khop) and the kinetics of hole-trapping reactions (ktrap). The guanine damage distributions observed in any DNA construct arise from differences in the relative magnitudes of khop and ktrap with both a kinetic and a thermodynamic limit identified for simple, repetitive DNA sequences.3 The situation for a mixed sequence like AQ-601 is more complicated since there are multiple types of guanine oxidation sites (e.g., G, GG, GGG) with different ktrap, and the different A-T bridge lengths between resolvable hopping sites lead to a distribution of khop. Chemically, hole trapping arises from reaction of a guanine radical cation with water71,72 and/or oxygen species73 to form lesions like oxazolone (Oz) and 8-hydroxyguanine (8OG). In our experiments, we are largely quantifying the levels of Oz and other highly reduced guanine productssthe contribution from 8OG is negligible since this nucleobase is not reactive with hot piperidine.74 The formation of Oz first requires the deprotonation of the guanine radical cation, then the addition of superoxide, and finally a complex series of rearrangements and fragment loss.72 Any process that slows down the diffusion of superoxide, and thus lowers ktrap, would be expected to increase the efficiency of long-range HT in mixed DNA sequences due to an increased probability of charge hopping over trapping. One of the effects of increasing the concentration of salt in aqueous media is the diffusion coefficient of oxygen species is changed. In the case of molecular oxygen, previous studies75,76 have shown that as [NaCl] increases from 0 to 100 mM, the diffusion coefficient of O2 decreases by
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5-10%. Around 800 mM [NaCl], the diffusion coefficient for O2 has increased back to its value at 0 mM.75 The similarity between the changes in oxygen diffusion coefficients and the DNA HT dynamics observed in AQ-601 is striking. The sharp increase in long-range hole transport to sites GGG3-GG5 between 10 and 100 mM NaCl coincides with a decreased oxygen diffusion coefficient, which would be expected to slow down hole trapping. As salt increases from 200 to 800 mM the increasing oxygen diffusion coefficient could accelerate hole trapping at the proximal guanine sites, and thus decrease the efficiency of long-range HT in AQ-601. Therefore, future experiments investigating the salt concentration dependence of hole transport in periodic DNA sequences in the ktrap > khop and khop > ktrap limits are of interest. 5. Conclusions We have demonstrated that the site-specific yields of guanine oxidation products arising from DNA HT can show sensitivity to changes in the local environment, in this case the bulk concentration of counterions. The increase of long-range (>150 Å) hole transport in AQ-601 at physiological salt concentrations, and the ensuing falloff in transport at higher salt, indicates that charge transport in mixed sequence DNA may be tuned by varying the surrounding medium. Even though CD spectroscopy indicates that the 601 DNA sequence displays the unusual property of underwinding with increasing salt, there is no apparent correlation of the HT dynamics in AQ-601 with these global changes in secondary structure. Ten nanosecond NPT MD simulations carried out on fragments of AQ-601 containing experimentally observed guanine oxidation sites show that bulk salt concentration is expected to have subtle, if any, effect on the average values or fluctuations of the DNA base step parameters responsible for controlling DNA HT rates. Further, the electrostatic potential felt by a guanine residue in these simulations is independent of salt concentration between 100 and 400 mM NaCl, consistent with the expectations of counterion condensation theory. Even though there is a strong correlation between fluctuations in electrostatic potential and molecular contacts between sodium counterions and guanine nucleobases, the observed fluctuations are independent of bulk salt concentration, similar to the results reported in a previous related study.16 Therefore, we propose that salt-dependent changes in hole-trapping rates, related to the diffusion of oxygen species in aqueous media, are controlling the observed distribution of guanine lesions in AQ-601. Acknowledgment. The authors wish to thank Dr. Lisa Gloss and Mrs. Traci Topping for assistance with and access to their Aviv CD spectrometer. Dr. Michael Smerdon (WSU) and Dr. Jonathon Widom (Northwestern) are thanked for discussions about data prior to publication. Funding for this work was provided by a grant from the National Science Foundation (CAREER-0347370). Supporting Information Available: Two tables with the base pair parameters calculated from 10 ns NPT simulations of duplexes 601-GG1 and 601-GG4 at 100 and 400 mM NaCl, and one figure showing an autoradiograph of AQ-601 samples irradiated in buffers containing varying concentrations of NaCl. This material is available free of charge via the Internet at http:// pubs.acs.org. References and Notes (1) Henderson, P. T.; Jones, D.; Hampikian, G.; Kan, Y.; Schuster, G. B. Proc. Natl. Acad. Sci. U.S.A. 1999, 96, 8353–8358.
Davis et al. (2) Nu´n˜ez, M. E.; Hall, D. B.; Barton, J. K. Chem. Biol. 1999, 6, 85– 97. (3) Kanvah, S.; Joseph, J.; Schuster, G. B.; Barnett, R. N.; Cleveland, C. L.; Landman, U. Acc. Chem. Res. 2010, 43, 280–287. (4) Giese, B. Annu. ReV. Biochem. 2002, 71, 51–70. (5) Genereux, J. C.; Barton, J. K. Chem. ReV. 2010, 110, 1642–1662. (6) Long-Range Charge Transfer in DNA; Schuster, G. B., Ed.; Springer: Heidelberg, 2004; Vol. 236-237. (7) Charge Transfer in DNA; Wagenknecht, H.-A., Ed.; Wiley-VCH: Weinheim, 2005. (8) Lewis, F. D.; Letsinger, R. L.; Wasielewski, M. R. Acc. Chem. Res. 2001, 34, 159–170. (9) Davis, W. B.; Hess, S.; Naydenova, I.; Haselsberger, R.; Ogrodnik, A.; Newton, M. D.; Michel-Beyerle, M. E. J. Am. Chem. Soc. 2002, 124, 2422–2423. (10) McConnell, H. M. J. Chem. Phys. 1961, 35, 508–515. (11) Takada, T.; Kawai, K.; Fujitsuka, M.; Majima, T. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 14002–14006. (12) Vura-Weis, J.; Wasielewski, M. R.; Thazhathveetil, A. K.; Lewis, F. D. J. Am. Chem. Soc. 2009, 131, 9722–9727. (13) Bixon, M.; Jortner, J. Chem. Phys. 2006, 326, 252–258. (14) Steenken, S.; Jovanovic, S. V. J. Am. Chem. Soc. 1997, 119, 617– 618. (15) Voityuk, A. A.; Siriwong, K.; Rosch, N. Angew. Chem., Int. Ed. 2004, 43, 624–627. (16) Kubar, T.; Elstner, M. J. Phys. Chem. B 2008, 112, 8788–8798. (17) Kubar, T.; Kleinekathofer, U.; Elstner, M. J. Phys. Chem. B 2009, 113, 13107–13117. (18) O’Neill, M. A.; Barton, J. K. J. Am. Chem. Soc. 2004, 126, 11471– 11483. (19) Barnett, R. N.; Cleveland, C. L.; Joy, A.; Landman, U.; Schuster, G. B. Science 2001, 294, 567–571. (20) Armitage, B.; Ly, D.; Koch, T.; Frydenlund, H.; Orum, H.; Batz, H. G.; Schuster, G. B. Proc. Natl. Acad. Sci. U.S.A. 1997, 94, 12320– 12325. (21) Hatcher, E.; Balaeff, A.; Keinan, S.; Venkatramani, R.; Beratan, D. N. J. Am. Chem. Soc. 2008, 130, 11752–11761. (22) Rajski, S. R.; Barton, J. K. Biochemistry 2001, 40, 5556–5564. (23) Boal, A. K.; Yavin, E.; Barton, J. K. J. Inorg. Biochem. 2007, 101, 1913–1921. (24) Nakatani, K.; Dohno, C.; Ogawa, A.; Saito, I. Chem. Biol. 2002, 9, 361–366. (25) Kanvah, S.; Schuster, G. B. Pure Appl. Chem. 2006, 78, 2297– 2304. (26) Bjorklund, C. C.; Davis, W. B. Biochemistry 2007, 46, 10745– 10755. (27) Bjorklund, C. C.; Davis, W. B. Nucleic Acids Res. 2006, 34, 1836– 1846. (28) Nunez, M. E.; Noyes, K. T.; Barton, J. K. Chem. Biol. 2002, 9, 403–415. (29) Hoch, D. A.; Stratton, J. J.; Gloss, L. M. J. Mol. Biol. 2007, 371, 971–988. (30) Lowary, P. T.; Widom, J. J. Mol. Biol. 1998, 19–42. (31) Bloomfield, V. A.; Crothers, D. M.; Tinoco, I. J. Nucleic Acids: Structures, Properties, and Functions; University Science Books: Sausalito, CA, 2000. (32) Phillips, J. C.; Braun, R.; Wang, W.; Gumbart, J.; Tajkhorshid, E.; Villa, E.; Chipot, C.; Skeel, R. D.; Kale, L.; Schulten, K. J. Comput. Chem. 2005, 26, 1781–1802. (33) MacKerell, A. D., Jr.; Banavali, N. J. Comput. Chem. 2000, 21, 105–120. (34) Foloppe, N.; MacKerell, A. D., Jr. J. Comput. Chem. 2000, 21, 86–104. (35) Humphrey, W.; Dalke, A.; Schulten, K. J. Molec. Graphics 1996, 14, 33–38. (36) Lu, X.-J.; Olson, W. K. Nucleic Acids Res. 2003, 31, 5108–5121. (37) Gasper, S. M.; Schuster, G. B. J. Am. Chem. Soc. 1997, 119, 12762– 12771. (38) Baase, W. A.; Johnson, W. C. J. Nucleic Acids Res. 1979, 6, 797– 814. (39) Johnson, B. B.; Dahl, K. S.; Tinoco, I. J.; Ivanov, V. I.; Zhurkin, V. B. Biochemistry 1981, 20, 73–78. (40) Tierney, M. T.; Grinstaff, M. W. J. Org. Chem. 2000, 65, 5355– 5359. (41) Ivanova, A.; Shushkov, P.; Ro¨sch, N. J. Phys. Chem. A 2008, 112, 7106–7114. (42) Kubar, T.; Woiczikowski, P. B.; Cuniberti, G.; Elstner, M. J. Phys. Chem. B 2008, 112, 7937–7947. (43) Senthilkumar, K.; Grozema, F. C.; Guerra, C. F.; Bickelhaupt, F. M.; Lewis, F. D.; Berlin, Y. A.; Ratner, M. A.; Siebbeles, L. D. J. Am. Chem. Soc. 2005, 127, 14894–14903. (44) Voityuk, A. A.; Davis, W. B. J. Phys. Chem. B 2007, 111, 2976– 2985.
Hole Transport Dynamics in Mixed Sequence DNA (45) Voityuk, A. A. J. Chem. Phys. 2008, 128, 115101. (46) Troisi, A.; Orlandi, G. Chem. Phys. Lett. 2001, 344, 509–518. (47) Grozema, F. C.; Tonzani, S.; Berlin, Y. A.; Schatz, G. C.; Siebbeles, L. D.; Ratner, M. A. J. Am. Chem. Soc. 2008, 130, 5157–5166. (48) Cornell, W. D.; Cieplak, P.; Bayly, C. I.; Gould, I. R.; Merz, K. M. J.; Ferguson, D. M.; Spellmeyer, D. C.; Fox, T.; Caldwell, J. W.; Kollman, P. A. J. Am. Chem. Soc. 1995, 117, 5179–5197. (49) Cheatham, T. E. I. Molecular modeling and atomistic simulation of nucleic acids. In Annual Reports in Computational Chemistry; Spellmeyer, D. C., Ed.; Elsevier: Oxford, 2005; Vol. 1; pp 75-90. (50) Peticolas, W. L.; Wang, Y.; Thomas, G. A. Proc. Natl. Acad. Sci. U.S.A. 1988, 85, 2579. (51) Ponomarev, S. Y.; Thayer, K. M.; Beveridge, D. L. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 14771–14775. (52) Voityuk, A. A. J. Chem. Phys. 2008, 128, 045104. (53) Voityuk, A. A. J. Phys. Chem. B 2009, 113, 14365–14368. (54) Bonvin, A. M.; Sunnerhagen, M.; Otting, G.; van Gunsteren, W. F. J. Mol. Biol. 1998, 282, 859–873. (55) Pal, S. K.; Zhao, L.; Xia, T.; Zewail, A. H. Proc. Natl. Acad. Sci U.S.A. 2003, 100, 13746–13751. (56) Segal, E.; Fondufe-Mittendorf, Y.; Chen, L.; Thåstro¨m, A.; Field, Y.; Moore, I. K.; Wang, J.-P. Z.; Widom, J. Nature 2006, 442, 772–778. (57) Nunez, M. E.; Holmquist, G. P.; Barton, J. K. Biochemistry 2001, 40, 12465–12471. (58) Boal, A. K.; Genereux, J. C.; Sontz, P. A.; Gralnick, J. A.; Newman, D. K.; Barton, J. K. Proc. Natl. Acad. Sci. U.S.A. 2009, 106, 15237–15242. (59) Luger, K.; Mader, A. W.; Richmond, R. K.; Sargent, D. F.; Richmond, T. J. Nature 1997, 389, 251–260. (60) Ruscio, J. Z.; Onufriev, A. Biophys. J. 2006, 91, 4121–4132. (61) Du, Q.; Smith, C.; Shiffeldrim, N.; Vologodskaia, M.; Vologodskii, A. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 5397–5402.
J. Phys. Chem. C, Vol. 114, No. 48, 2010 20833 (62) Cloutier, T.; Widom, J. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 3645–3650. (63) Jayaram, B.; Beveridge, D. L. Annu. ReV. Biophys. Biomol. Struct. 1996, 25, 367–394. (64) Manning, G. S. Acc. Chem. Res. 1979, 12, 443–449. (65) Bleam, M. L.; Anderson, C. F.; Recore, M. T. J. Proc. Natl. Acad. Sci. U.S.A. 1980, 77, 3085–3089. (66) Feig, M.; Pettitt, B. M. Biophys. J. 1999, 77, 1769–1781. (67) Korolev, N.; Lyubartsev, A. P.; Laaksonen, A.; Nordenskio¨ld, L. Biophys. J. 2002, 82, 2860–2875. (68) Howerton, S. B.; Sines, C. C.; VanDerveer, D.; Williams, L. D. Biochemistry 2001, 40, 10023–10031. (69) Allahyarov, E.; Lo¨wen, H.; Gompper, G. Phys. ReV. E 2003, 68, 061903. (70) Lewis, F. D.; Wasielewski, M. R. Dynamics and Equilibrium for Single Step Hole Transport Processes in Duplex DNA. In Long-Range Charge Transfer in DNA I; Schuster, G. B., Ed.; Springer Berlin: Heidelberg, 2004. (71) Reynisson, J.; Steenken, S. Phys. Chem. Chem. Phys. 2002, 4, 527– 532. (72) Cadet, J.; Douki, T.; Ravanat, J.-L. Acc. Chem. Res. 2008, 41, 1075– 1083. (73) Yun, B. H.; Lee, Y. A.; Kim, S. K.; Kuzmin, V.; Kolbanovskiy, A.; Dedon, P. C.; Geacintov, N. E.; Shafirovich, V. J. Am. Chem. Soc. 2007, 129. (74) Cullis, P. M.; Malone, M. E.; Merson-Davies, L. A. J. Am. Chem. Soc. 1996, 118, 2775–2781. (75) van Stroe, A. J.; Janssen, L. J. J. Anal. Chim. Acta 1993, 279, 213– 219. (76) Ju, L.-K.; Ho, C. S. Biotechnol. Bioeng. 1985, 27, 1495–1499.
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