Thermodynamic Stability of DNA Duplexes Comprising the Simplest T

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Thermodynamic Stability of DNA Duplexes Comprising the Simplest T # dU Substitutions Carolyn E Carr, Irine Khutsishvili, Barry Gold, and Luis A. Marky Biochemistry, Just Accepted Manuscript • DOI: 10.1021/acs.biochem.8b00676 • Publication Date (Web): 05 Sep 2018 Downloaded from http://pubs.acs.org on September 8, 2018

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Biochemistry

Thermodynamic Stability of DNA Duplexes Comprising the Simplest T → dU Substitutions

Carolyn E. Carr, Irine Khutsishvili†, Barry Gold‡ and Luis A. Marky* Department of Pharmaceutical Sciences, University of Nebraska Medical Center, 986025 Nebraska Medical Center, Omaha, NE 68198-6025

*

To whom correspondence should be addressed. Tel.: (402) 559-4628. Fax: (402) 559-9543.

E-mail: [email protected] †

Current Address: Institute of Physics, Javakhishvili State University, Tbilisi, Goergia.

‡

Current Address: Department of Pharmaceutical Sciences, University of Pittsburgh, Pittsburgh,

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Abstract The family of enzymes called uracil-DNA glycosylases (UDGs) recognize and bind uracil, sequestering it within the binding site pocket and catalyzing the cleavage of the base from the deoxyribose, leaving an abasic site. This recognition and binding is passive and relies on innate dynamic motions of DNA wherein base-pairs undergo thermally induced breakage and conformational fluctuations. Once the uracil breaks from its base-pair, it can be recognized and bound by the enzyme, which then alters its conformation for sequestration and catalysis. Our results suggest that the thymine to uracil substitution, which differs only by a single methyl group, causes a destabilization of the duplex thermodynamics, which would lead to an increase in the population of the extra-helical state and increase the probability of uracil being recognized and excised from DNA by UDG. This destabilization is dependent on the identity of the nearestneighbor base-pair stacks; a G•C nearest-neighbor leads to less of a thermal and enthalpic destabilization than having two A•T neighbors. In addition, uracil substitution yields a nearest neighbor increase in the counterion uptake of the duplexes but decreases the immobilization of structural water for all substituted duplexes regardless of neighbor identity or number of substitutions.

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Introduction Deoxyuracil (dU) is incorporated into DNA through one of two ways, either through cytosine deamination, which occurs ~102 times per cell per day, or through incorporation of dUTP instead of dTTP by DNA polymerases, which occurs once every 2000 to 3000 dNTP additions.1, 2 dU incorporation occurs because polymerases cannot distinguish between the structure of dU and T, so dU is occasionally incorporated instead of T. Because T is synthesized from dU, dU is present in the cell at all times.3 Cytosine deamination is mutagenic, as the C•G base-pair eventually becomes a T•A base-pair through successive rounds of replication (C•G → U•G → U•A → T•A).4 dU incorporation by DNA polymerases opposite an A is not mutagenic, as DNA polymerases cannot discriminate between uracil and thymine, and so the polymerase continues to read the sequence as normal.5 While direct dU incorporation is not mutagenic it may have effects due to changes in DNA-DNA and DNA-protein interactions and it occurs much more frequently than cytosine deamination. Uracil excision from DNA is performed by Uracil-DNA glycosylases (UDGs), a group of DNA repair enzymes.6, 7 UDGs are part of an overall base excision repair pathway and remove uracil and other unusual bases from DNA. UDGs use water as a nucleophile and cleave the Nglycosidic bond between the base and the deoxyribose, generating an abasic site that is then recognized for further repair. The UDG superfamily has seven subfamilies, all with varying specificity. UDGs function by first searching for uracil within DNA, generally thought to occur via a 3dimensional hopping and sliding motion along the DNA.6,

7

Once the enzyme encounters a

uracil, its conformation is altered as it introduces a sidechain that replaces the extra-helical uracil and sequesters it deep within the active site pocket that allows nucleophilic attack by water. The


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result is DNA with an abasic site and a liberated uracil. UDGs passively recognize uracil, i.e., they do not actively open base-pairs to search for uracil. Instead, UDGs rely on the thermally induced dynamic motions of DNA, which include spontaneous base-pair dissociation and twisting of nucleotides out of the helical conformation.8, 9 Kinetic studies have shown that once UDGs bind uracil, they increase the lifetime of these open states, but do not increase the rate at which the open state occurs. Studies have shown that dU•A base-pairs open seven times faster than T•A base-pairs.1 UDGs are the most heavily studied family of all DNA glycosylases and their mechanism mirrors that of all DNA glycosylases, regardless of the DNA damage they recognize.10-13

Figure 1. Proposed structures of the duplexes. Uracil substitutions in bold. Because the dynamics of DNA opening, and thus the probability of UDG encountering a dU, is affected by its thermodynamic properties, we chose to study a series of DNA duplexes with dU substitutions (Fig. 1) and investigated changes in its structural and thermodynamic properties as a result of the substitutions. We found that the substitutions cause a decrease in the thermal


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Biochemistry

and enthalpic stability and a partial compensation by a favorable increase in the entropy term, which would increase the population of extra-helical dU. We also found that the identity of the nearest-neighbors dictated the magnitude of this effect. In addition, incorporation of uridines increased the counterion uptake based on the identity of the nearest neighbors and decreased the immobilization of structural water for all duplexes. Materials and Methods Materials. All oligonucleotides were synthesized by the Eppley Institute Molecular Biology Core Facility at University of Nebraska Medical Center, reverse-phase HPLC purified, desalted on a G-10 Sephadex column, and lyophilized to dryness prior to use in experiments. The concentration of the oligomer solutions was determined at 260 nm and 80 °C using the following molar extinction coefficients in mM-1cm-1 of strands:

d(CCGGAATTCGCC), 112.2;

d(GGCGTTAACCGG), 119.5; d(CCGGAATUCGCC), 107.8; d(GGCGUTAACCGG), 114.2; d(CCGGAAUTCGCC), 110.2; d(GGCGTUAACCGG), 117.0; d(CCGGAAUUCGCC), 108.7; d(GGCGUUAACCGG), 115.9. The extinction coefficients were calculated by extrapolation of the tabulated values of the dimer and monomer bases from 25 °C to high temperatures, using procedures reported earlier.14 Buffer solutions consisted of 10 mM sodium phosphate (NaPi) at pH 7.0 and adjusted to different sodium and ethylene glycol concentrations with NaCl and ethylene glycol. All chemicals used in this study were reagent grade and used without further purification. Temperature-dependent UV spectroscopy. Absorbance versus temperature profiles (UV melting curves) were measured at 260 nm and the temperature was scanned at a heating rate of ~0.6 °C/min. Shape analysis of the melting curves yielded transition temperatures, TM.15 UV melting curves were carried out as a function of strand concentration to determine transition


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molecularities; the TM of an intramolecular complexes is independent of strand concentration, while higher molecularities display concentration dependent TMs. Circular dichroism (CD) spectroscopy. Simple inspection of the CD spectra allowed analysis of the conformation of each oligonucleotide at temperatures at which the structure of each oligonucleotide is 100% helical. The CD spectra was obtained using an Aviv Model 202SF spectrometer (Lakewood, NJ) equipped with a peltier temperature control system. Spectra were obtained at 5 °C from 320 to 220 nm in 1 nm increments and the reported spectra correspond to the average of at least two wavelength scans. Differential scanning calorimetry (DSC). Heat capacity functions of the helix-coil transition of duplexes were measured with a VP-DSC from Malvern MicroCal (Northampton, MA). The thermograms were obtained with a temperature ramp of 0.6 °C/min at a concentration of 200 µM in total strands. The thermograms were buffer subtracted, normalized by strand concentration and deconvoluted using a non-two-state zero ∆Cp model with the MicroCal software. Analysis of these thermograms yielded model-independent thermodynamic profiles (∆Hcal, ∆Scal and ∆G°cal). These parameters are measured with the following relationships: ∆Hcal=∫∆Cp(T)dT and ∆Scal=∫∆Cp(T)/TdT, where ∆Cp is the heat capacity of the oligonucleotide solution during the unfolding process. The free energy, ∆G°cal, is calculated at 20 °C using the Gibbs relationship ∆G°(T)=∆Hcal−T∆Scal. Additional DSC thermograms were obtained at several salt concentrations in order to indirectly obtain the associated heat capacity contributions (∆Cp) from the slopes of the lines of ∆Hcal vs. TM plots. Differential binding of counterions and water molecules. The helical and coil states of an oligonucleotide are associated with a different number of bound ions and water molecules. Therefore, the helix→coil transition is accompanied by a differential release (or uptake) of


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Biochemistry

counterions and water molecules.16 The following equations were used to measure the thermodynamic uptake of counterions, ∆nNa+, and water molecules, ∆nW, upon folding of each duplex17, 18: +

2

∆nNa+ = 1.11[∆Hcal/RTM ](∂TM/∂ln[Na ])

(1)

2

(2)

∆nW

= [∆Hcal/RTM ](∂TM/∂ln[aW])

where 1.11 is the correction factor that corresponds to conversion of concentrations into ionic activities. The first term in brackets of Eqs. (1)-(2), [∆Hcal/RTM2], is a constant determined directly from DSC experiments while the second term in parenthesis is determined experimentally from UV melting curves by measuring the TM dependencies on the concentration of counterions or water. In the determination of ∆nNa+, UV melting curves were measured with a [NaCl] ranging from 0.01 M to 0.2 M at pH 7.0. UV melting curves as a function of ethylene glycol concentration (0.5 M to 2.5 M) were carried out to determine ∆nW as this co-solute does not interact specifically with DNA.19 The osmolality of the latter solutions was obtained with a UIC vapor pressure osmometer Model 830, which was calibrated with standardized solutions of NaCl. These osmolalities were then converted into water activities (aw) using the following equation:20 ln aw=−Osm/Mw, where Osm is the solution osmolality and Mw is the molality of pure water equal to 55.5 mol/kg of H2O. Results and Discussion Discussion of Duplexes. We used a non-self-complementary dodecamer duplex with a central core of four A•T base pairs. For the purposes of discussion, the duplexes are arranged in the tables with the T → dU substitutions on the 5’-strand first, followed by the duplexes with substitutions on the 3’-strand and then substitutions on both strands. In all figures the data is


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displayed as a set, with the T → dU substitutions on the 5’-strand displayed with the control duplexes in a single graph to highlight differences. All reactions are bimolecular, indicating duplex formation. The UV melting curves were obtained over a total strand concentration of 4 – 40 µM in 10 mM NaPi buffer at pH 7.0 to confirm the transition molecularity of each duplex. The corresponding TM-dependencies are shown in Figure S1. All duplexes show a TM-dependence on strand concentration, indicating that there is a bimolecular association reaction occurring under these solution conditions. The TMdependence confirms that all strands are forming duplexes regardless of sequence differences and the monophasic transitions indicate that no other species are present. All duplexes are in the B-conformation. The CD spectrum of each duplex was recorded in 10 mM NaPi buffer at pH 7.0 and is shown in Figure S2. All duplexes display typical B-form DNA spectra, which consists of a negative peak at 250 nm and a positive peak at 280 nm which are approximately equivalent in magnitude. All spectra are nearly identical to the control (shown in black in all graphs). The peak at 280 nm is related to sugar puckering and is nearly identical in all duplexes. The peak at 250 nm corresponds to the extent of base-pair stacking; while several of the T → dU substitution duplexes appear to have a reduction in the strength of base-pair stacking, overall it can be said that the substitutions do not significantly alter the strength of the base-pair stacks. Calorimetric unfolding of duplexes. DSC curves in 10 mM NaPi at pH 7.0 for all duplexes are shown in Figure 2 and in 10 mM NaPi, 0.2 M NaCl at pH 7.0 in Figure S3 and the resulting thermodynamic parameters are shown in Table S1. As can be seen in Table S1, standard folding thermodynamic profiles consist of a favorable free energy term resulting from the compensation of a favorable enthalpy and an unfavorable entropy. The favorable enthalpy is


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due to the formation of base-pairs and base-pair stacks and the unfavorable entropy is due to the ordering of strands and the uptake of ions and water.

Figure 2. DSC thermograms of duplexes in 10 mM NaPi buffer at pH 7.0. (Left) AATT/TTAA (black squares), AATU/TTAA (red squares), AAUT/TTAA (blue squares), AAUU/TTAA (green squares) (Middle) AATT/TTAA (black squares), AATT/UTAA (red circles), AATT/TUAA (blue circles), AATT/UUAA (green circles) (Right) AATT/TTAA (black squares), AATU/UTAA (red triangles), AAUT/TUAA (blue triangles), AAUU/UUAA (green triangles).

The differential thermodynamic profiles at two salt concentrations, where the thermodynamic parameters of the substituted duplexes are subtracted from the unsubstituted control duplex, are shown in Table 1. The unfolding of all duplexes was highly reproducible, and all unfolded as a single transition. To analyze the effects of the T → dU substitutions, only Table 1 will be considered during this discussion. There are several obvious trends that can be seen from Table 1 at low salt. The first trend is that the nearest neighbors of the substitution have the most critical effect on the thermodynamics. The first duplex, AATU/TTAA is the least affected by the substitutions, yielding a ∆TM of -2.8 °C and a ∆∆Hcal of -4.3 kcal/mol. The next duplex, AAUT/TTAA, has a greater reduction in its TM and enthalpy, indicating the substitution causes a more significant effect. The major difference between these two duplexes is that in AATU/TTAA the dU is flanked by G•C and A•T base-pairs, while in AAUT/TTAA it is flanked by two A•T base-pairs. It is a well-known fact that a G•A/C•T base-pairs stack leads to an increase in TM as compared to A•T/T•A or A•A/T•T base-pair stacks, and it is clear from these results that the added strength mitigates the 9 ACS Paragon Plus Environment

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effect of a T → dU substitution. This trend can be seen regardless of whether the substitution occurs on the 5’-strand, 3’-strand (AATT/UTAA, AATT/TUAA and AATT/UUAA), or both strands (last three duplexes of Table 1). Table 1. Differential Unfolding Thermodynamic Profiles for Dodecamers with T → dU Substitutions Relative to the Unsubstituted Duplex. Duplex AATU/ TTAA AAUT/ TTAA AAUU/ TTAA AATT/ UTAA AATT/ TUAA AATT/ UUAA AATU/ UTAA AAUT/ TUAA AAUU/ UUAA

[Na+] (mM) 16 216 16 216 16 216 16 216 16 216 16 216 16 216 16 216 16 216

∆TM (°C) -2.8 -0.1 -5.2 -0.7 -5.8 -1.3 -2.9 -0.2 -6.0 -0.4 -6.1 -1.9 -4.0 -1.4 -6.0 -2.4 -6.9 -3.0

∆∆Hcal (kcal/mol) -4.3 -16.7 -6.4 -16.4 -8.9 -16.6 -6.0 -16.7 -11.8 -16.7 -12.1 -15.8 -5.0 -16.8 -9.8 -16.8 -10.0 -17.4

∆(T∆Scal) (kcal/mol) -3.1 -14.3 -4.4 -13.9 -6.6 -14.0 -4.6 -14.3 -9.2 -14.3 -9.4 -13.2 -3.5 -14.1 -7.3 -13.9 -7.3 -14.3

∆∆G°20 (kcal/mol) -1.1 -2.4 -1.9 -2.5 -2.2 -2.6 -1.3 -2.4 -2.5 -2.4 -2.6 -2.6 -1.4 -2.7 -2.4 -2.9 -2.6 -3.1

∆∆nNa+ (mol Na+/mol)

∆∆nW (mol H2O/mol)

0.1

-20

0.8

-17

0.8

-18

0.1

-21

0.6

-19

0.6

-20

0.2

-19

0.7

-26

0.7

-21

All parameters are measured in 10 mM NaPi buffer at pH 7.0. Experimental uncertainties are as follow: ∆TM (± 0.7 °C), ∆∆Go (± 10%), ∆∆Hcal (± 7%), T∆S (± 7%), ∆∆nNa+ (± 17%), ∆∆nW (± 17%), ∆CP (± 20%). The other observation that can be seen in Table 1 is that having two adjacent substitutions does not produce a cumulative or cooperative effect on the thermodynamics. While having two substitutions is slightly worse, for the most part the difference between AAUT/TTAA and AAUU/TTAA is statistically insignificant and this difference decreases for the next two sets (having the substitutions on the 3’ or both strands). This suggests that, as mentioned above, the nearest neighbors of the substitution is critical, and the thermodynamics of the duplex are determined by the most detrimental substitution. This conclusion can also be seen when comparing the different sets against each other; set one, with substitutions on the 5’ strand, is the


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least affected, while set two and three (substitutions on the 3’ or both strands) are nearly identical, indicating that not only are the 5’- and 3’-strands inherently different, but the thermodynamics of the duplex is dictated by mutations on the 3’ strand, which cause the most detrimental effect. At higher salt concentrations, the trends in all thermodynamic parameters disappear, with the exception of TM which becomes progressively worse as more dU are substituted in. However, the change in TM is small, with a maximum of 3 °C. Thus, at higher salt thermodynamic differences are minute and would not explain the higher rate of base-pair opening in dA•dU base-pairs versus in dA•dT base-pairs. However, while differences between substitution sets are small, the difference in enthalpy and entropy between all substituted duplexes and the control is large, with an average ∆∆Hcal of 16.7 kcal/mol. This is much larger than a single base-pair stack, and is extremely unlikely to be caused by complete lack of base-pairing between the dU and A, as U•A base-pairs are canonical in RNA and should have two hydrogen bonds even in DNA. We attribute this to a general weakening of the dU•dA base-pair as well as the surrounding base-pair stacks, which would facilitate more rapid movement of the uridine outside of the helix, increasing the probability of being located by UDGs. However, all duplexes in the study have at least one A/T base-pair surrounding the substituted base-pair. Previous research on T→dU substitutions flanked by two C/G base-pairs21 yielded no significant difference in the thermodynamic profiles, indicating that the nearest-neighbor thermodynamics contributes to the mechanism of dU recognition and removal. The ∆∆G° of low and high salt are very similar (2.0 and 2.6 kcal/mol, respectively); at higher salt the duplexes display a larger ∆∆Hcal which is compensated by an equally large ∆∆Scal, leading to a similar free energy. Thus, at physiological salt concentrations there is no increase in folding free energy that is usually associated with 11 ACS Paragon Plus Environment

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increased salt concentrations. Heat capacity effects. The heat capacity of an oligonucleotide can provide qualitative information on the folded state of the molecule and is useful when comparing small sequence changes to understand how that change affects the structure. Oligonucleotide unfolding typically has a positive heat capacity, indicating that more hydrophobic residues are exposed to solvent upon unfolding than hydrophilic residues.22-25 The larger the positive heat capacity, the more hydrophobic surface area is sequestered in the folded state. Because both the sign and magnitude of the heat capacity is important, the ∆CP values in Table 1 will not be converted into ∆∆CP. Heat capacities were determined from the slope of a ∆Hcal vs. TM plot at two different salt concentrations. It can be immediately seen that the control duplex has the largest positive heat capacity, indicating that it is the most well folded molecule, i.e., it is sequestering the most hydrophobic surface area in its folded state. This indicates that the substitutions are having an impact on the folded state of the molecule. Interestingly, the substitutions near the G•C basepairs are consistently the most affected in terms of heat capacity (two are negative and one is positive but small in magnitude) despite being the least affected in terms of thermodynamics and counterion binding. Heat Capacity Modification of the Calorimetric Unfolding of Duplexes. We corrected the thermodynamic profiles using the associated heat capacity values shown in the last column of Table S1. The corrected values are displayed in Table S2 and the differential thermodynamic profiles obtained from the corrected values are listed in Table 2. One major difference of the heat capacity corrected values from the original is that at low salt the values of each individual set (substitution on the 5’, 3’ or both strands) are not statistically significantly different. In addition, each set as a whole is not statistically different


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from the others. Thus, at low salt with thermodynamic values corrected for heat capacity effects, there is no relevant difference in regard to where the substitution is placed and how many there are. The opposite is true at physiological salt concentration; when the thermodynamic values are corrected for heat capacity, there is differences between the sets. The first set, with substitutions on the 5’ strand, display only minor differences in their thermodynamics, but there is a trend Table 2. ∆CP Modified Differential Unfolding Thermodynamic Profiles for Dodecamers with T → dU Substitutions Relative to the Unsubstituted Duplex. Duplex AATU/ TTAA AAUT/ TTAA AAUU/ TTAA AATT/ UTAA AATT/ TUAA AATT/ UUAA AATU/ UTAA AAUT/ TUAA AAUU/ UUAA

[Na+] (mM) 16 216 16 216 16 216 16 216 16 216 16 216 16 216 16 216 16 216

∆TM (°C) -2.8 -0.1 -5.2 -0.7 -5.8 -1.3 -2.9 -0.2 -6.0 -0.4 -6.1 -1.9 -4.0 -1.4 -6.0 -2.4 -6.9 -3.0

∆∆Hcal (kcal/mol) -38.3 -58.8 -35.3 -55.4 -33.7 -49.2 -33.0 -52.9 -32.4 -42.2 -29.8 -37.3 -37.6 -61.3 -33.2 -47.4 -34.5 -49.4

∆(T∆Scal) (kcal/mol) -31.9 -51.8 -28.8 -45.2 -27.3 -50.0 -27.4 -43.2 -26.1 -34.4 -23.8 -30.0 -31.0 -49.9 -26.8 -38.2 -27.1 -39.7

∆∆G°20 (kcal/mol) -6.4 -11.5 -6.5 -10.2 -6.4 -9.2 -5.6 -9.7 -6.3 -7.8 -6.0 -7.3 -6.6 -11.4 -6.4 -9.2 -6.8 -9.7

All parameters are measured in 10 mM NaPi buffer at pH 7.0. Experimental uncertainties are as follow: ∆TM (± 0.7 °C), ∆∆Go (± 10%), ∆∆Hcal (± 7%), T∆S (± 7%). showing decreasing effects of the substitution, i.e. when next to a G•C base-pair there is a greater detriment to the thermodynamics than when there is an A•T base-pair on both sides, or when the two uridine substitutions are sequential. This is more striking in the next two sets (substitutions on the 3’ strand and both strands) wherein there is a large difference in enthalpy between the first substitution in the set, which is between an AT and GC base-pair, and the next substitution which is between two AT base-pairs. This indicates that when corrected for heat capacity effects, being 13 ACS Paragon Plus Environment

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next to a GC base-pair is actually detrimental to the free energy, indicating that this would cause the uridine to be extra-helical with a higher frequency, increasing the probability that it will be found by a UDG. However, our thermodynamic results in total indicate that, regardless of salt concentration or heat capacity effects, if a dU is incorrectly incorporated in place of any thymine at any position, the probability of it being in an extra-helical conformation is increased substantially due to the lower energy of a base-pair stack incorporating dU than dT. The differential binding of ions and water. The differential binding of counterions was calculated using Eq. 1 and the differential binding of structural water was calculated using Eq. 2. We obtained the slope of the TM-dependencies on salt or osmolyte concentration from UV melting curves as a function of [Na+] (Fig. S4) or ethylene glycol (Fig. S5) concentration. The average ∆Hcal/RTM2 term is obtained from DSC melts at several salt or ethylene glycol concentrations. The folding of each molecule is accompanied by an uptake of ions due to a shift in the equilibrium toward the conformation with a higher charge density parameter,26 as well as an overall uptake of water molecules due to the shift towards the conformation with a higher hydration state.18, 27 The ∆nNa+ are presented in Table S1, and the ∆∆nNa+ values are shown in Table 1. The trends seen for the thermodynamic profiles are also seen for the ∆nNa+ values except instead of reducing the counterion binding, the T → dU substitutions increase counterion binding. Replacing a T with a dU near a G•C base-pair yields no significant change in counterion uptake. However, substitution in between two A•T/T•A base pairs yields a significant increase in the ∆nNa+ value and two consecutive substitutions are not cumulative but adopt the ∆nNa+ value of the substitution with the greatest effect. However, the reason for an increase in counterion uptake is unclear, as a dU differs from a T by one methyl group, which should not positively affect


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Biochemistry

counterion uptake. In addition, the reduction in thermodynamic stability suggests that the double helical structure is less compact and thus would have a lower charge density parameter, not a higher one. Howevver, previous studies have shown that oligonucleotides with greater instability have a higher counterion uptake. Our hypothesis is that a higher number of ions bind, increasing the favorable enthalpy at the expense of unfavorable entropy, leading to an overall higher free energy of folding. The ∆nW values are shown in Table S1 and the ∆∆nW values are listed in Table 1. Interestingly, there is no obvious trend regarding position or number of T → dU substitutions. Instead, all substituted molecules display the same marked decrease in immobilization of structural water compared to the control duplex. This is surprising; while lack of a methyl group would yield less surface area and be expected to bind less structural water, it should not decrease water immobilization by 33%. In addition, because these duplexes do not have significant homogenous sequence tracts, any ordering of the water (such as seen in continuous A-tracts) would be miniscule and would not be broken by addition of a single dU. It might be expected that this is related to counterion uptake, as an increase in counterions would displace structural water in favor of electrostricted water. However, if this were true then the single substitutions at the ends (near the G•C base-pairs), which have almost no increase in counterion uptake, would have unaffected water immobilization, which is not the case. Thus, T → dU substitutions increase counterion uptake based on the identity of the substitution neighbors, and decrease water immobilization, regardless of location. Conclusions Uracil-DNA glycosylases rely on the dynamic conformational sampling of individual DNA base-pairs to find dU for excision. The base-pairs unstack, which allows them to rotate into an


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extra-helical state; the rate of this unstacking and rotation is influenced by the thermodynamic parameters (∆H, ∆S, ∆G) of the duplex. The more favorable enthalpy and less unfavorable entropy, the less often the base-pair stacks will sample other conformations, which reduces the frequency of binding to DNA repair enzymes. Base-pair mismatches will of course have a lower enthalpy than canonical Watson-Crick base-pair stacks but a U•A base-pair is canonical in RNA and has identical hydrogen bonding interactions in DNA. Uracil and thymine differ only by the addition of one methyl group in T. However, our studies have shown that not only do T to dU substitutions decrease the thermal and enthalpic stability of duplexes, which would decrease the penalty for dU being extra-helical and increasing the probability of being bound by a UDG, but the identity of the nearest-neighbors is a factor in base-pair stability. Being flanked by at least one G•C base-pair provides stability to the substitution; previous studies show that being flanked by two, which is statistically very unlikely, may decrease the effect of the uracil even more, decreasing the likelihood of being available for binding by a UDG. Thus, this work provides a thermodynamic basis for the increased rate of opening for a dU•A base-pair, and a reason why UDGs so frequently find dU bases.

Conflict of Interest The authors declare no conflict of interest. Acknowledgements This work was supported by Grant MCB-1122029 from the National Science Foundation. Supporting Material Five figures and two table are available at http://


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Biochemistry

References (1) Friedman, J. I., and Stivers, J. T. (2010) Detection of damaged DNA bases by DNA glycosylase enzymes, Biochemistry 49, 4957-4967. (2) Tye, B.-K., Chien, J., Lehman, I. R., Duncan, B. K., and Warner, H. R. (1978) Uracil incorporation: A source of pulse-labeled DNA fragments in the replication of the Escherichia coli chromosome, Proc Natl Acad Sci U S A 75, 233-237. (3) Reidy, J. A. (1988) Role of deoxyurifine incorporation and DNA repair in the expression of human chromosomal fragil sites, Mutat Res 200, 215-220. (4) Schormann, N., Zhukovskaya, N., Bedwell, G., Nuth, M., Gillilan, R., Prevelige, P. E., Ricciardi, R. P., Banerjee, S., and Chattopadhyay, D. (2016) Poxvirus uracil-DNA glycosylase-An unusual member of the family I uracil-DNA glycosylases, Protein Sci 25, 2113-2131. (5) Visnes, T., Doseth, B., Pettersen, H. S., Hagen, L., Sousa, M. M., Akbari, M., Otterlei, M., Kavli, B., Slupphaug, G., and Krokan, H. E. (2009) Uracil in DNA and its processing by different DNA glycosylases, Philos Trans R Soc Lond B Biol Sci 364, 563-568. (6) Schormann, N., Ricciardi, R., and Chattopadhyay, D. (2014) Uracil-DNA glycosylasesstructural and functional perspectives on an essential family of DNA repair enzymes, Protein Sci 23, 1667-1685. (7) Zharkov, D. O., Mechetin, G. V., and Nevinsky, G. A. (2010) Uracil-DNA glycosylase: Structural, thermodynamic and kinetic aspects of lesion search and recognition, Mutat Res 685, 11-20. (8) Parker, J. B., Bianchet, M. A., Krosky, D. J., Friedman, J. I., Amzel, L. M., and Stivers, J. T. (2007) Enzymatic capture of an extrahelical thymine in the search for uracil in DNA, Nature 449, 433-437. (9) Cao, C., Jiang, Y. L., Stivers, J. T., and Song, F. (2004) Dynamic opening of DNA during the enzymatic search for a damaged base, Nat Struct Mol Biol 11, 1230-1236. (10) Grasso, S., and Tell, G. (2014) Base excision repair in Archaea: back to the future in DNA repair, DNA Repair (Amst) 21, 148-157. (11) Wallace, S. S. (2014) Base excision repair: a critical player in many games, DNA Repair (Amst) 19, 14-26. (12) Prakash, A., and Doublie, S. (2015) Base Excision Repair in the Mitochondria, J Cell Biochem 116, 1490-1499. (13) Schermerhorn, K. M., and Delaney, S. (2014) A chemical and kinetic perspective on base excision repair of DNA, Acc Chem Res 47, 1238-1246. (14) Cantor, C. R., and Schimmel, P. R. (1980) Biophysical Chemistry, Part 2: Techniques for the study of biological structure and function, 1st ed., W. H. Freeman and Company, New York. (15) Marky, L. A., and Breslauer, K. J. (1987) Calculating thermodynamic data for transitions of any molecularity from equilibrium melting curves, Biopolymers 26, 1601-1620. (16) Zieba, K., Chu, T. M., Kupke, D. W., and Marky, L. A. (1991) Differential Hydration of dAdT Base Pairing and dA and dT Bulges in Deoxyoligonucleotides, Biochemistry 30, 8018-8026. (17) Kaushik, M., Suehl, N., and Marky, L. A. (2007) Calorimetric unfolding of the bimolecular and i-motif complexes of the human telomere complementary strand, d(C3TA2)4, Biophys Chem 126, 154-164. 17 ACS Paragon Plus Environment

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Biochemistry


Thermodynamic Stability of DNA Duplexes Comprising the Simplest T

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