Fluorescence of εA-containing DNA in single-strand, duplex, and in complex with AAG. Fluorescence emission spectra of 400 nM DNA were collected with excitation at 320 nm, as described in the Materials and Methods. (A) Both single-strand (○) and duplex –TEC– (●) are compared to single-strand (□) and duplex (ν) –AEA– DNA. The εA in both sequences is more fluorescent in the single-strand and becomes quenched in the duplex. (B) Fluorescence emission spectra were collected in the presence of saturating concentration of AAG and compared to the spectra in the absence of AAG. The –TEC– duplex DNA exhibited very similar fluorescence in the absence (●) or presence (○) of AAG. In contrast, the –AEA– duplex DNA, which was strongly fluorescent in the absence of protein (ν), was strongly quenched in the presence of AAG (□). The spectra were taken within 1 minute of AAG addition (1 µM), during which time negligible excision can occur.

Kinetics of DNA binding and nucleotide flipping, determined by stopped-flow fluorescence at equimolar concentrations of AAG and DNA. AAG was rapidly mixed with an equimolar solution of –TEC– (A) or –AEA– (B) DNA to yield final concentrations after mixing of 0.15 (bottom) to 1.5 µM (top). Each trace is the average of 3 independent binding reactions, and the lines indicate the best fits of equation 3. For the –TEC– substrate the raw data is plotted, indicating the increased signal observed at higher concentrations of DNA, but for the –AEA– substrate the larger fluorescence change required that the gain be adjusted between different concentrations of DNA and therefore the individual traces are arbitrarily offset.

Stopped-flow fluorescence measurements of the binding of εA-DNA to AAG with excess protein. Representative data is shown from experiments in which a fixed concentration of DNA was mixed with increasing concentrations of AAG. Samples were excited at 320 nm and emission was collected with a 360 nm long-pass filter (See Materials and Methods for details). Several individual reactions were monitored at each concentration of AAG, and the traces shown are the averages of all reactions. Since the fluorescence changes were superimposed at the different concentrations of AAG, the traces are offset for clarity. (A) The binding of 0.5 µM –TEC– duplex DNA by 0.5 (bottom), 1, 2, 4, and 8 µM AAG (top). (B) The binding of 0.15 µM –AEA– duplex DNA by 0.5 (bottom), 1, and 2 µM (top) AAG. The lines indicate the best fits of equation 2 to each binding reaction.

We measured the rate of dissociation of the abasic DNA product to compare it to the rate of εA product release. The steady state reaction of AAG with –TEC- substrate gave a k_cat value that is essentially identical to the single turnover rate constant under these reaction conditions (data not shown), suggesting that the very slow rate of hydrolysis of the N-glycosidic bond is rate limiting for multiple turnover excision of εA at high ionic strength. Since AAG shows much faster rates of excision of Hx from inosine lesions, we measured the k_cat value for multiple turnover on an inosine-containing substrate. It has previously been shown that AAG exhibits burst kinetics for the excision of Hx and that the steady state rate constant, k_cat, reflects the rate of dissociation of the abasic DNA product (34). We determined that the rate constant for the dissociation of abasic DNA is 0.24 min⁻¹ under these reaction conditions, ~6-fold faster than the rate constant for N-glycosidic bond cleavage (Scheme 4; see Supporting Information). This is only slightly faster than the observed rate constant for dissociation of the εA substrate (k_{off, obs} = 0.12 min⁻¹), suggesting that the abasic DNA product is also likely to be stably bound in an extrahelical conformation.

Minimal mechanism for AAG-catalyzed nucleotide flipping and base excision. AAG (crescent) binds nonspecifically to DNA and rapidly slides in search of DNA damage. Once a lesion (filled rectangle) is encountered in an initial damage recognition complex it can be flipped out into the active site to form the specific recognition complex. In this specific complex the DNA is bent, the damaged nucleotide is flipped out by 180°, and Y162 (red circle) is intercalated into the DNA to take the place of the extrahelical lesion (15, 16). From this specific complex, AAG catalyzes hydrolysis of the N-glycosidic bond (k_chem). The lesion base is subsequently released (k_{εA release}). The abasic product is an inhibitor of AAG, so presumably the sugar can be stably bound in a flipped out conformation analogous to the abasic pyrrolidine conformation observed by x-ray crystallography (15). Dissociation of AAG from the abasic product requires rotation of the abasic sugar back into the duplex (k_unflip). From the less tightly bound complex, AAG can translocate along the DNA in search of additional sites of damage and can ultimately dissociate (k_off).

It is possible to relate these observed rate constants to the microscopic rate constants in Scheme 3, with a bimolecular binding step followed by a unimolecular conformational change that occurs prior to the chemical step. The initial binding step appears to be reversible (see below), and therefore the observed rate constant is given by the sum of the association and dissociation steps in Scheme 3 (k_1,obs = [AAG]k₁ + k₋₁). It should be noted that this equation applies when [E]≫[S]. Under our conditions the enzyme was in only modest excess (2–7-fold). Nevertheless, the resulting values of k₁ are in reasonably good agreement with the values measured by mixing equimolar solutions of enzyme and substrate (see below). We suggest that the unimolecular step corresponds to nucleotide flipping. Since this is an approach to equilibrium, the observed rate constant is equal to the sum of the rate constants for the forward and reverse flipping steps (k_2,obs = k₂ + k₋₂). Substrate dissociation experiments, measured by pulse-chase suggest that the value of k₂ is much greater than k₋₂, indicating that k_2,obs is approximately equal to k₂, the microscopic rate constant for flipping.

The rate constants for binding and nucleotide flipping by AAG in two different sequence contexts. The observed rate constants for –TEC– DNA (●, solid line) and – AEA– DNA (ν, dashed line) were taken from the fits of equation 3 to the data for equimolar binding reactions (Figure 6). (A) The observed bimolecular rate constant for the initial binding of AAG to DNA is very similar for both substrates (~2 × 10⁸ M⁻¹s⁻¹; Table 1). According to the simple two-step binding mechanism depicted in Scheme 3, this observed rate constant is simply k₁, the microscopic rate constant for binding. (B) The observed rate constant for the second step is independent of concentration, and yields an average value of 4.3 s⁻¹ for the –AEA– sequence context compared to a value of 2.6 s⁻¹ for the –TEC– sequence context. According to Scheme 3, this observed rate constant is the sum of the forward and reverse rate constants for flipping (k_{2, obs} = k₂ + k₋₂). Additional experiments described below indicate that the equilibrium constant for flipping is highly favorable (k₂ ≫ k₋₂), and therefore the observed rate constant for flipping is simply the forward rate constant for flipping (see text for additional discussion).

Determination of the forward and reverse rate constants for the initial binding of AAG to εA-DNA. The observed rate constant for DNA binding (k_{1, obs}) is from the first phase of the binding reactions shown in Figure 4. The average and standard deviations are shown for 2–4 determinations. The dissociation rate constant for the initial AAG•DNA complex can be determined from the y-intercept of this plot (k_{1, obs} = k⁻¹ + k₁[AAG]; Scheme 3). For –AEA– DNA (ν), the best linear fit (solid line) gives an intercept of 48 s⁻¹ (k₋₁) and a slope of 1.5×10⁸ M⁻¹s⁻¹ (k₁). This second order rate constant is identical to the value determined from equimolar binding experiments (Table 1 and Figure 7). For –TEC– DNA (●), only two concentrations of protein gave reliable data due to the weak fluorescence and more rapid binding. A linear fit (dotted line) of the two data points that were obtained gives an intercept of 22 s⁻¹ and a slope of 2.4 × 10⁸ M⁻¹s⁻¹. By forcing the slope of the line to be 2.1× 10⁸ M⁻¹s⁻¹ (dashed line), the value that was independently determined from equimolar binding reactions (Table 1 and Figure 7), an intercept of 43 s⁻¹ was obtained. We take the average (33 ± 15 s⁻¹) as a rough estimate for the k₋₁ value for this substrate.

The human enzyme, alkyladenine DNA glycosylase (AAG), is responsible for repairing a diverse set of alkylated and oxidized purines, including purines alkylated at the N7 and N3 positions, and the deaminated purines hypoxanthine (Hx) and oxanine [(2, 3) and references therein]. Chemical structures of damaged and undamaged adenine bases are shown in Scheme 1. AAG also recognizes the bulkier alkylation adduct, 1,N⁶-ethenoadenosine (εA) that can be formed from lipid peroxidation products and exposure to exogenous agents such as urethane and vinyl chloride (4, 5). This lesion is highly mutagenic in mammalian cells and leads to misincorporation of all possible nucleotides if it is replicated prior to repair (6, 7). AAG is the only known human glycosylase that catalyzes the excision of εA, but an alternative direct reversal pathway has recently been discovered in which εA can be directly oxidized to restore the adenine base by AlkB family members (8). This backup pathway may account for the observation that mice lacking AAG remain viable (9, 10). Nevertheless, mice lacking AAG have decreased rates of repair of εA and are unable to tolerate the burden of increased levels of DNA alkylation (11, 12).

The 25mer oligonucleotides or the constituent phosphoramidites were obtained from commercial sources and contained a central lesion that was placed opposite of a T. The sequences are given in Scheme 2. Oligonucleotides containing only normal deoxynucleotides or deoxyinosine were synthesized and deprotected with standard phosphoramidite chemistry, whereas those containing a single εA lesion were synthesized and deprotected using Ultra-Mild chemistry according to the supplier’s recommendations. The εA-containing oligonucleotides were either synthesized by the W.M. Keck Facility at Yale University or the phosphoramidite was obtained from Glen Research and incorporated using an ABI 394 DNA synthesizer. The oligonucleotides were desalted using Sephadex G-25 and purified using denaturing polyacrylamide gel electrophoresis as previously described (13). Oligonucleotides for gel-based assays were labeled on the lesion containing strands with a 5’-fluorescein (6-fam) label. The concentrations of the single stranded oligonucleotides were determined from the absorbance at 260 nm, using the calculated extinction coefficients for all oligonucleotides except those containing εA. For εA containing oligonucleotides, the extinction coefficient was calculated for the same sequence with an A in place of the εA and corrected by subtracting 9400 M⁻¹cm⁻¹ to account for the weaker absorbance of εA as compared to A (31). The lesion-containing oligonucleotides were annealed with a 1.1-fold excess of the complement by heating to 90°C and cooling slowly to 4°C.

Pulse-chase experiment to measure the macroscopic dissociation rates for εA-containing DNA substrate. (A) The experimental design is indicated. Fluorescein (fam)-labeled DNA (100 nM –TEC–) was mixed with excess AAG (200 nM) for 20 seconds (incubation time, t₁) then 20 µM unlabeled –TEC– DNA was added as a chase. The reactions were quenched at the indicated time points (t₂), and the fraction of abasic DNA product determined by alkaline hydrolysis and gel electrophoresis (See Materials and Methods for additional details). (B) Single turnover excision of εA from –TEC– DNA by AAG in the absence (●) or presence (ν) of chase under the standard reaction conditions. In the absence of chase, the line indicates the best fit to a single exponential (Equation 1). In the presence of chase, the line indicates the best fit of an exponential burst followed by a slow steady state reaction (Equation 4). This gives a macroscopic dissociation rate constant of 0.12 ± 0.01 min⁻¹. (C) Comparison of pulse chase experiments performed at concentrations of sodium chloride. The data from experiments at (●) 70 mM, (ν) 120 mM and (σ) 320 mM ionic strength were fit by equation 4 (lines). Analysis of the observed rate constant and the endpoint gave essentially identical macroscopic rate constants and the average and standard deviation of duplicate reactions determined by both methods yielded macroscopic dissociation rate constants of 0.08 ± 0.005, 0.12 ± 0.01, and 0.35 ± 0.05 min⁻¹, respectively for 70, 120, and 320 mM ionic strength.

Comparison of εA release and N-glycosidic bond cleavage from single turnover assays. Fluorescence emission spectra monitoring the increase in fluorescence that is observed upon single turnover excision of εA from (A) –TEC– and (B) –AEA– DNA catalyzed by AAG. The spectra for samples containing 400 nM duplex DNA were collected prior to the addition of protein (○), and within 1 minute of the addition of 1.2 µM AAG (●). Additional spectra were collected after 3 (σ), 6 (∆), 12 (ν), 30 (□), 60 (υ), 90 (◊), and 120 (τ) minutes. Note that the high fluorescence of the –AEA– substrate is strongly quenched upon binding to AAG. Representative experiments are shown, but duplicate reactions gave essentially identical results, and the rate of reaction was independent of the concentration of AAG. (C) The gel-based glycosylase assay relies on selective hydroxide-catalyzed cleavage of abasic sites and the different mobility of intact substrate and nicked product to monitor formation of the abasic DNA product (See Materials and Methods for details). A representative denaturing polyacrylamide gel shows the separation of the fluorescein-labeled substrate (25-mer) and product (12-mer) and a time course for the formation of the abasic DNA product. The first lane shows a no enzyme control. The small amount of nicked DNA (~4%) results from nonenzymatic ring-opening and depurination during synthesis, purification, and the hydroxide heating step. The next 5 lanes indicate samples taken from a reaction containing 100 nM –TEC– DNA and 200 nM AAG that were quenched 2–120 minutes after the initiation of the reaction by the addition of AAG. (D) Quantification of the results from εA release and abasic site formation indicates that both assays are limited by a common step, presumably N-glycosidic bond cleavage. The fraction product was calculated from the εA fluorescence assay for –TEC– (●) and –AEA– (○) as well as the gel based excision assay for –TEC– (υ), as described in the Materials and Methods. The error bars indicate the standard deviation from the mean for 2–5 independent determinations. The reaction progress curves for –TEC– monitored by εA fluorescence and abasic site formation are fit by a single exponential (Equation 1; k_max = 0.038 min⁻¹). The reaction of –AEA– is slightly slower (k_max = 0.031 min⁻¹).