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Published online on January 2, 2008, 10.1073/pnas.0710363105
PNAS | January 8, 2008 | vol. 105 | no. 1 | 70-75
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BIOLOGICAL SCIENCES / BIOCHEMISTRY
A continuous hyperchromicity assay to characterize the kinetics and thermodynamics of DNA lesion recognition and base excision

Conceição A. S. A. Minetti, David P. Remeta, and Kenneth J. Breslauer*

Department of Chemistry and Chemical Biology, Rutgers, The State University of New Jersey, 610 Taylor Road, Piscataway, NJ 08854

Communicated by I. M. Gelfand, Rutgers, The State University of New Jersey, Piscataway, NJ, November 2, 2007 (received for review October 3, 2007)


   Abstract
Top
 Abstract
Results
 Discussion
 Materials and Methods
 ACKNOWLEDGMENTS.
 References
 
We report a continuous hyperchromicity assay (CHA) for monitoring and characterizing enzyme activities associated with DNA processing. We use this assay to determine kinetic and thermodynamic parameters for a repair enzyme that targets nucleic acid substrates containing a specific base lesion. This optically based kinetics assay exploits the free-energy differences between a lesion-containing DNA duplex substrate and the enzyme-catalyzed, lesion-excised product, which contains at least one hydrolyzed phosphodiester bond. We apply the assay to the bifunctional formamidopyrimidine glycosylase (Fpg) repair enzyme (E) that recognizes an 8-oxodG lesion within a 13-mer duplex substrate (S). Base excision/elimination yields a gapped duplex product (P) that dissociates to produce the diagnostic hyperchromicity signal. Analysis of the kinetic data at 25°C yields a Km of 46.6 nM for the E·S interaction, and a kcat of 1.65 min–1 for conversion of the ES complex into P. The temperature dependence reveals a free energy ({Delta}Gb) of –10.0 kcal·mol–1 for the binding step (E + S {leftrightarrow} ES) that is enthalpy-driven ({Delta}Hb = –16.4 kcal·mol–1). The activation barrier ({Delta}G{ddagger}) of 19.6 kcal·mol–1 for the chemical step (ES {leftrightarrow} P) also is enthalpic in nature ({Delta}H{ddagger} = 19.2 kcal·mol–1). Formation of the transition state complex from the reactants (E + S {leftrightarrow} ES{ddagger}), a pathway that reflects Fpg catalytic specificity (kcat/Km) toward excision of the 8-oxodG lesion, exhibits an overall activation free energy ({Delta}GT{ddagger}) of 9.6 kcal·mol–1. These parameters characterize the driving forces that dictate Fpg enzyme efficiency and specificity and elucidate the energy landscape for lesion recognition and repair.

8-oxoguanosine | activation energy | energetics | glycosylase/lyase

The number of DNA lesions implicated in genetic diseases and degenerative cell disorders has increased over the past decades (1, 2). Such findings have stimulated efforts to characterize specific enzyme families that function in the recognition and repair of DNA damage (3–7). The majority of existing methods used to determine kinetic parameters for base/nucleotide excision repair (BER/NER) enzymes employ labeling and quenching techniques that measure either product formation or substrate disappearance (3, 8–11). Although of great value, many of these assays require chemical modifications and/or sampling of the enzyme-catalyzed reaction at discrete time intervals. These features may potentially compromise successful monitoring of intrinsic system properties. To address this challenge, we have developed a continuous hyperchromicity assay (CHA) that exploits the intrinsic free-energy differences of a damaged DNA duplex substrate and the resultant processed product. The method requires minimal sample manipulation and provides a real-time, continuous optical signal to quantify product formation. Because the presence of a damaged base or lesion often exerts profound effects on DNA duplex energetics and stability (12), the solution conditions of the assay are optimized to ensure that the duplex substrate harboring a defect is sufficiently stable, whereas the excised product containing a nick or gap readily dissociates.

In this study, we use the CHA method to characterize the kinetics and thermodynamics of Fpg-catalyzed 8-oxodG base excision/elimination, which yields a gapped duplex by means of concerted glycosylase/lyase activities. Published studies on the kinetics of 8-oxodG removal by Fpg exhibit significant differences in the magnitudes of the kinetic parameters reported. These disparities generally are attributed to variability in the experimental conditions used as well as to the use of discontinuous methods for monitoring reaction progress (13). By contrast, the temperature-dependent CHA method described herein is not so encumbered and furnishes kinetic (kcat, Km) and energetic binding ({Delta}Hb, {Delta}Sb, and {Delta}Gb) parameters associated with enzyme–substrate interactions as well as activation parameters ({Delta}H{ddagger}, {Delta}S{ddagger}, and {Delta}G{ddagger}) for enzyme catalysis. Characterization of these kinetic and thermodynamic properties is needed to map the energy landscapes for Fpg-mediated base excision repair and define the energetic origins of substrate specificities for this and other glycosylases.


   Results
Top
 Abstract
 Results
Discussion
 Materials and Methods
 ACKNOWLEDGMENTS.
 References
 
Experimental Design and Method Validation. The CHA method exploits the differential physicochemical properties of deoxyoligonucleotides in which lesion-containing duplex substrates are processed in a site-specific manner to produce a DNA strand break and concomitant duplex dissociation. In this assay, an n-mer-duplex containing a central damaged base yields a nicked/gapped duplex with a stability that is significantly lower than the original substrate and that readily dissociates into an n-mer complementary single strand plus two [(n-1)/2]-mer single strands. In the specific example reported here, Fpg-catalyzed excision/elimination of an 8-oxodG base centered within a 13-mer oligonucleotide duplex results in an unstable gapped product that dissociates into three single strands (Scheme 1). We have validated the assay by using model compounds that simulate incremental product/substrate ratios, and we have confirmed that the CHA method selectively and quantitatively tracks the hyperchromicity range reflective of product formation [Materials and Methods and supporting information (SI) Fig. 6]. The resultant hyperchromicity at 260 nm, therefore, serves as an accurate reporter of the enzyme reaction and is proportional to product formation as monitored by duplex dissociation. A typical baseline-corrected progress curve is depicted in Fig. 1. The linear portion of the curve provides the initial velocity of the reaction (dashed line in Fig. 1), which constitutes the basis for our steady-state analysis of the kinetic and thermodynamic parameters. The assay solution conditions have been optimized to ensure maximal enzyme activity (SI Fig. 7) and DNA substrate stability over the selected concentration (10–250 nM) and temperature (15–25°C) ranges. Because determination of the enzyme turnover number (kcat) requires precise characterization of the active enzyme fraction, we confirmed that Fpg is essentially 100% active at the concentrations used in this study (SI Fig. 8).


Figure 1
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  		Scheme 1. Representation of the model system used in the present study to determine the energetic and kinetic parameters of Fpg-glycosylase/lyase activity. Fpg-mediated catalysis of a 13-mer oligonucleotide duplex harboring an 8-oxodG lesion in the central position undergoes dissociation into a single-strand 13-mer and two 6 mers upon 8-oxodG base excision and β/{delta}-elimination.


Figure 1
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  		Fig. 1. Progress reaction of Fpg-mediated 8-oxodG base excision/elimination monitored by the time-dependent hyperchromicity at 260 nm. The initial velocity is calculated from the slope of the linear phase of the progress curve as depicted by the dashed line. (Inset) Raw data of the catalytic reaction (red), enzyme control in the absence of substrate (blue), and baseline-corrected progress curve (magenta).

Conceptual Framework: Transition State Theory and Catalytic Specificity. Much of quantitative modern enzymology is based on Eyring's transition state theory (14), which describes absolute reaction rates in terms of rate-limiting formation of an activated enzyme–substrate complex (E·S{ddagger}), the latter representing the highest energy level in which chemical bonds are formed or broken (15). Considering the usefulness of transition-state analysis for elucidating enzyme mechanisms, it is somewhat surprising that relatively few studies have focused on the recognition and repair of damaged DNA by glycosylases. A refreshing exception is a recent review that examines the transition state mechanisms associated with N-glycoside hydrolysis/transfer (16), with particular emphasis on uracil glycosylase, a BER enzyme for which electrostatic stabilization of the transition state has been demonstrated (17).

Classical views on thermodynamics of catalysis have been discussed in detail (18). Transition-state theory relates the activation energy of the enzyme-catalyzed conversion of substrates (S) to products (P). Expressed in thermodynamic terms as illustrated in Scheme 2, the free energy difference between the enzyme and substrate (E + S) relative to the enzyme–substrate (Michaelis) complex in the ground state (E·S) is defined as {Delta}Gb. The activation free energy ({Delta}G{ddagger}) represents the energy required to promote the enzyme–substrate complex from its ground state (E·S) to the transition state (E·S{ddagger}). The free-energy difference between the free substrate and enzyme (E + S) relative to the activated transition-state complex (E·S{ddagger}) is defined as {Delta}GT{ddagger}. This net activation free energy, {Delta}GT{ddagger}, for the E + S to E·S{ddagger} transformation therefore corresponds to the sum of the positive free energy barrier, {Delta}G{ddagger}, for the E·S to E·S{ddagger} activation and the negative favorable binding free energy, {Delta}Gb, for the E + S to the E·S (i.e., Michaelis) "ground state" complex. In the sections that follow, we present the kinetic and thermodynamic parameters derived from our measurements and describe how these can be used to map the energy landscape of Fpg-mediated 8-oxodG base excision.


Figure 2
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  		Scheme 2. Relationship between the kinetic and thermodynamic parameters associated with enzyme–substrate interactions and catalysis.

Kinetic Parameters for Fpg-Mediated 8-oxodG Base Excision/Elimination. The present study reports kinetic parameters for Fpg glycosylase base excision/gap formation under conditions that satisfy the Michaelis–Menten steady-state approximation. The relevant reaction may be summarized as follows:

Formula

In this scheme, interaction of the substrate with the enzyme (E + S {leftrightarrow} ES) is fast relative to the chemical reaction (ES {leftrightarrow} E + P) (19). Accordingly, k–1 >> kcat and the Michaelis constant (Km) corresponding to the ratio (k–1 + kcat)/k1 reflects the true dissociation constant (Km = Kd = k–1/k1) for the E·S complex. The chemical reaction therefore represents the limiting step in the pathway and corresponds to the first-order catalytic rate constant kcat. Under the conditions of the assay, the coupled duplex dissociation event that occurs upon product formation (KD->SS) is rapid and therefore does not affect the overall kinetic parameters for catalysis. The initial velocity (dashed line in Fig. 1) derived from least-squares analysis of the linear segment of the progress curve is recorded as a function of substrate concentrations to obtain Michaelis–Menten kinetic parameters. The data are fit to a rectangular hyperbola (Fig. 2) to derive both Vmax and Km for the Fpg-catalyzed reaction (20, 21) and recast in the form of a Hanes plot (Fig. 2 Inset) that enhances graphical visualization (i.e., data points spaced equidistantly). Table 1 presents a summary of kinetic parameters for Fpg-catalyzed 8-oxodG base excision/elimination.


Figure 2
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  		Fig. 2. Steady-state Michaelis–Menten analysis of Fpg-mediated 8-oxodG base excision/elimination. Typical kinetic experiments performed at 25.0°C in which the initial velocities are recorded as a function of DNA substrate concentration. The data are fit to a rectangular hyperbola to derive the kinetic parameters Vmax and Km. (Inset) The initial velocities are recast in the form of a linearized Hanes plot.


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  		Table 1. Kinetic parameters for Fpg-mediated 8-oxodG base excision/elimination

 
Thermodynamic Parameters for Fpg-Mediated 8-oxodG Base Excision/Elimination. The activation free energy ({Delta}G{ddagger}) associated with kcat is estimated by using the relationship: {Delta}G{ddagger} = RT (ln kB/h – ln kcat/T), in which kB and h are the respective Boltzmann and Planck constants, R is the universal gas constant, T is the absolute temperature in Kelvin, and kcat is expressed in sec–1. Substituting the relevant value of 0.0276 sec–1 (1.654 min–1) for kcat, an activation free energy of 19.6 kcal·mol–1 is determined for Fpg catalysis of the cognate 8-oxodG substrate at 25.0°C. Steady-state analysis of Fpg catalysis reveals that Km represents an equilibrium microscopic dissociation constant derived from the ratio k–1/k1. Accordingly, the "binding" free energy associated with Km (i.e., {Delta}Gb) may be estimated by using the relationship: {Delta}Gb = –RT ln (1/Km). Based on the Km value of 46.6 nM measured at 25.0°C for Fpg catalysis of the cognate 8-oxodG substrate (Table 1), we calculate an overall enzyme–substrate binding free energy ({Delta}Gb) of –10.0 kcal·mol–1 for this reaction.

Enthalpic and Entropic Contributions to Fpg-Mediated Catalysis. In principle, activation ({Delta}G{ddagger}) and binding ({Delta}Gb) free energies may be derived from single temperature measurements under a standard set of conditions. The CHA method offers the advantage of facilitating kinetic measurements at multiple temperatures, thereby allowing us to resolve the relative contributions of enthalpic and entropic forces to both the kinetic and thermodynamic parameters associated with enzyme catalysis. Typical progress curves for Fpg catalysis of a 100 nM 8-oxodG substrate acquired as a function of temperature are presented in Fig. 3. These data reflect the temperature dependence of the initial rates. By monitoring the concentration dependence of the initial rates as illustrated in Fig. 2, one obtains the Michaelis constant (Km), the turnover number (kcat), and the specificity constant (kcat/Km) at each temperature. The temperature dependences of the kinetic parameters listed in Table 1 allow one to characterize the thermodynamic parameters associated with catalysis and elucidate the underlying enthalpic and entropic driving forces associated with Fpg enzyme activity.


Figure 3
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  		Fig. 3. Temperature dependence of the Fpg-catalyzed reaction rates. Typical progress reaction curves recorded for the 8-oxodG cognate substrate under continuous stirring in a quartz cuvette thermostatted at the desired temperature. Absorbance changes are monitored at 260 nm immediately after Fpg addition as described in Materials and Methods.

Temperature Dependence of kcat: Activation Energy, Activation Enthalpy, and Activation Entropy. The temperature dependence of kcat is cast in the form of an Eyring plot in which ln kcat/T is presented as a function of the reciprocal of the temperature (1/T) in Fig. 4. Analysis of the temperature dependence of kcat permits determination of the activation enthalpy ({Delta}H{ddagger}) and activation entropy ({Delta}S{ddagger}) according to the relation: ln kcat/T = ln kB/h + ({Delta}S{ddagger}/R) – ({Delta}H{ddagger}/RT), in which the slope of the plot is –{Delta}H{ddagger}/R, and the intercept at 1/T = 0 is {Delta}S{ddagger}/R + 23.76. Analysis of the resultant data reveals an activation enthalpy ({Delta}H{ddagger}) of 19.2 kcal·mol–1 and a corresponding entropic contribution (T{Delta}S{ddagger}) of –0.4 kcal·mol–1. Thus, the activation barrier to catalysis is overwhelmingly enthalpic in nature with a negligible entropic contribution. The activation energy (Ea) associated with Fpg catalysis, as derived from the relation Ea = {Delta}H{ddagger} + RT, is therefore 19.8 ± 0.7 kcal·mol–1.


Figure 4
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  		Fig. 4. Eyring plot depicting the temperature dependence of kcat to derive the activation enthalpy ({Delta}H{ddagger}) and activation entropy ({Delta}S{ddagger}).

Temperature Dependence of Km: Driving Forces of Fpg–Substrate Interactions During Catalysis. We have resolved the enthalpic and entropic contributions of Fpg binding to its cognate substrate from analysis of the temperature dependence of the enzyme–substrate association constant, which corresponds to the reciprocal of Km: dln(1/Km)/d(1/T) = –{Delta}H/R. The resultant slope of the plot in Fig. 5 yields a binding enthalpy ({Delta}Hb) of –16.4 kcal·mol–1 at 25.0°C. Substituting values for the relevant thermodynamic parameters into the relation {Delta}Gb = {Delta}Hb – T{Delta}Sb, we calculate a negative entropic contribution of –6.4 kcal·mol–1. Thus, association of Fpg with its cognate substrate is enthalpically driven, with unfavorable entropic contributions. Table 2 summarizes the thermodynamic parameters and driving forces associated with Fpg-catalyzed 8-oxodG base excision/elimination derived by using data from our continuous hyperchromicity assay.


Figure 5
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  		Fig. 5. Van't Hoff analysis of the temperature dependence of Km to derive the binding enthalpy ({Delta}Hb) and entropy ({Delta}Sb).


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  		Table 2. Thermodynamic parameters for Fpg-mediated 8-oxodG base excision/elimination

 

   Discussion
Top
 Abstract
 Results
 Discussion
Materials and Methods
 ACKNOWLEDGMENTS.
 References
 
Advantages of the CHA Method and Comparisons with Published Data. Conventional methods used to characterize enzymes that process nucleic acids in a site-specific manner routinely incorporate labeling steps followed by monitoring product formation by reaction quenching at discrete time intervals. Discrimination of the reaction product(s) generally is based on physical separation of the species by using analytical techniques such as gel electrophoresis. Continuous methods for monitoring site-specific enzymatic processing of DNA duplexes are less frequently used and often require substrate prelabeling steps or coupled detection systems. To address these challenges, we have developed an assay that enables continuous, direct, in-solution measurement of the site-specific cleavage of a DNA substrate by means of an intrinsic optical signal that requires no labeling. In the CHA method, we selectively and continuously monitor product formation by means of the hyperchromicity at 260 nm, thereby avoiding complications inherent in dissecting contributions arising from the coupled effects of binding, catalysis, and duplex dissociation as well as potential perturbations caused by substrate labeling. SI Table 3 summarizes some of the advantageous characteristics of the CHA method for assessing the kinetics and energetics of BER and related enzymes that specifically process phosphodiester bonds.

Previous studies employing conventional assays have provided kinetic parameters for Fpg-catalyzed 8-oxodG base excision/elimination, thereby furnishing insight into the origins of Fpg catalytic specificity (3, 22, 23). However, the data reported differ substantially because of variability in the experimental conditions used (13). To determine the origin and reconcile the range of reported results, we have standardized the experimental solution conditions for the CHA method, particularly with respect to temperature and ionic strength. We also have selected a phosphate buffer system that has a low temperature dependence. Screening of ionic strength reveals that optimal Fpg activity is achieved at concentrations not exceeding 100 mM NaCl (SI Fig. 6), a finding consistent with previous reports (24). Over the temperature range of 15.0–25.0°C selected for our continuous assay, Km and kcat (and by extension kcat/Km) exhibit significant temperature-dependent behaviors. Values range from {approx}18–47 nM, 0.5–1.6 min–1, and 0.028–0.036 nM–1min–1 for Km, kcat, and kcat/Km, respectively. Comparison of our CHA-determined kinetic parameters with those derived from gel-based assays reveals the variables of temperature, sequence context, and solution conditions to be critical determinants of Fpg catalytic activity, thereby contributing to some of the variability in the published data.

Energetics of Fpg-Mediated 8-oxodG Recognition and Excision. Despite progress in the elucidation of the biochemical, structural, and dynamic features of several members of the BER/NER enzyme family, there is a dearth of studies that characterize the energetic driving forces for enzyme recognition and catalysis (25, 26). Such data are needed for mapping the energetic landscape of glycosylase catalysis. The CHA method allows acquisition of temperature-dependent kinetic parameters under solution conditions in which both the enzyme and DNA duplex substrate preserve their overall native conformations. We therefore have restricted our studies to a temperature range in which there are no noticeable thermal effects arising from either the enzyme (i.e., partial inactivation/denaturation) or the substrate (i.e., dissociation) that might compromise the targeted energetic characterizations.

Activation Barrier to Fpg-Mediated Catalysis Is Enthalpic in Origin. The temperature dependence of the rate constants is of fundamental importance to characterize the underlying driving forces associated with energy barriers for Fpg-catalyzed base excision. The latter facilitates partitioning of the enthalpic and entropic contributions to the free energy of activation ({Delta}G{ddagger}), a resolution afforded by the CHA method. The data reveal that the activation free energy of 19.6 kcal·mol–1 is almost exclusively enthalpic in origin ({Delta}H{ddagger} = 19.2 kcal·mol–1), with no significant departure from linear behavior observed when the kcat values are analyzed via Eyring plots over the temperature range of 15.0–25.0°C (Fig. 4). This finding is consistent with the active site of Fpg being preorganized for effective substrate binding, with minimal conformational changes involved. With the exception of the active-site loop region (residues 221–234), studies reveal that the structural features of Fpg are, in fact, not significantly altered during catalysis, supporting the proposition that Fpg functions as a typical enzyme by exploiting the intrinsic reactivity of the substrate to promote catalysis (16). It is worth noting that the Eyring plot acquires a "convex" shape when the measurements are extended to temperatures >35°C (data not shown). These findings are consistent with UV melting experiments that reveal a population of dissociated substrate species at temperatures >35°C because of the reduced effective duplex concentration and low ionic strength used in the assay. To alleviate this constraint, longer DNA sequences may be used to shift the functional window of the measurements to higher temperatures. This adaptation of the assay is of importance for evaluating the energetics of base excision and/or phosphodiester bond cleavage by hyperthermophilic enzymes, which exhibit optimal temperature ranges higher than those of their mesophilic counterparts.

Energetics of Enzyme–Substrate Interactions in the Presence and Absence of Turnover. Despite the similarities in Fpg binding affinities observed for the cognate 8-oxodG substrate ({Delta}Gb = –10 kcal·mol–1) and the nonhydrolyzable abasic duplex ({Delta}Gcal = –9.7 kcal·mol–1) as noted in Table 2, their relative driving forces differ significantly. Binding to the cognate 8-oxodG lesion is enthalpic in nature ({Delta}Hb = –16.4 kcal·mol–1), whereas Fpg association with the nonhydrolyzable abasic analog is entropy driven ({Delta}Hcal = 0.8 kcal·mol–1) at 25.0°C (27). Fpg affinity for the synthetic nonhydrolyzable abasic intermediate state analog in the absence of turnover is typical of protein–minor groove interactions dominated by binding-induced desolvation of apolar surface areas (27–29). The abasic site analog conceivably undergoes "facile" accommodation into the enzyme pocket because the physical absence of the base provides a larger surface area and consequently a greater hydrophobic effect, which is consistent with the observed entropic contribution to binding and the large {Delta}Cp. By contrast, reactive groups in the cognate 8-oxodG substrate (and presumably its transition state) may establish optimal contacts with the enzyme active site during catalysis. Therefore, it is plausible that a more extensive hydrogen-bonding and/or electrostatic-interaction network within the transition state accounts for both the large enthalpic stabilization ({Delta}Hb = –16.4 kcal·mol–1) and significant negative entropic contribution (T{Delta}Sb = –6.4 kcal·mol–1) observed for cognate substrate binding during catalysis.

Thermodynamic Forces Governing Fpg Catalytic Specificity. Theoretical and experimental approaches stress the importance of evaluating kcat/Km as it pertains to free enzyme properties and as a measure of enzyme efficiency and catalytic specificity (18, 20). Accurate and reliable kcat/Km values represent a useful probe of the chemical nature of the enzyme–substrate interactions that lead to productive binding and/or transition state stabilization (20). Inspection of the temperature dependence of kcat/Km listed in Table 1 reveals that this ratio increases moderately as function of temperature. Such a trend indicates that the thermally induced reduction in affinity (increase of Km) is more than compensated by an increase in turnover number. As illustrated in Scheme 2, the thermodynamic parameters associated with kcat/Km may be evaluated in terms of transition state theory such that {Delta}G{ddagger} is the activation free energy relative to kcat, {Delta}Gb corresponds to the binding affinity defined as 1/Km, and {Delta}GT{ddagger} represents the activation free energy relative to kcat/Km. Invoking the relation {Delta}GT{ddagger} = {Delta}G{ddagger} + {Delta}Gb, we calculate an overall energy barrier ({Delta}GT{ddagger}) of 9.6 kcal·mol–1 associated with Fpg catalyzed 8-oxodG removal at 25.0°C. The latter results from a large activation free energy ({Delta}G{ddagger}) of 19.6 kcal·mol–1 that is partially counteracted by a favorable binding free energy ({Delta}Gb) of –10.0 kcal·mol–1. Conceivably, a significant portion of this binding energy is used in catalysis, because evidence suggests that Fpg prefers the transition-state relative to the ground-state forms of the substrate. This proposition is consistent with our finding that substrate analogs bind poorly to Fpg (27), whereas transition state mimics and abasic/intermediate analogs exhibit high binding affinity (27, 30, 31), effectively inhibiting Fpg catalytic activity (SI Fig. 9). Collectively, these observations support our proposal (27) that the primary affinity of the enzyme for the duplex substrate is dictated by interactions with a state that more resembles the product of base excision than the initial cognate substrate. Such a model implicitly assumes a coupling of the recognition/binding and base excision steps.

The free energies measured in the present study and their corresponding enthalpic components are cast in the form of an energy diagram in Scheme 3 that depicts the reaction coordinates of Fpg recognition and excision of an 8-oxodG lesion. Implicit in this scheme is the reality that favorable enzyme and substrate interactions ({Delta}Gb) contribute to transition state stabilization by lowering the enzyme-catalyzed energy barrier ({Delta}G{ddagger}) relative to the uncatalyzed reaction, thereby resulting in enzyme-induced rate enhancement. Analogous to the activation free energy, the overall activation enthalpy ({Delta}HT{ddagger}) for the apparent second order rate constant (kcat/Km) can be represented by the relation {Delta}HT{ddagger} = {Delta}H{ddagger} + {Delta}Hb (Scheme 3). The temperature dependence of the Michaelis constant yields a van't Hoff enthalpy of –16.4 kcal·mol–1 for {Delta}Hb, and the temperature dependence of the catalytic constant yields an activation enthalpy {Delta}H{ddagger} of + 19.2 kcal·mol–1. Consequently, we calculate a {Delta}HT{ddagger} of + 2.8 kcal·mol–1 and a corresponding activation entropy term (T{Delta}ST{ddagger}) of –6.8 kcal·mol–1. The modest enthalpy barrier is consistent with the lower temperature dependence of Fpg catalytic specificity. It is tempting to hypothesize that the relative substrate preferences of Fpg are largely governed by entropic factors, whereas specific group interactions required for transition state stabilization may not assume a critical role in Fpg catalytic specificity toward various substrates. Accordingly, substrates exhibiting a higher degree of "flexibility" would be preferred over those exhibiting greater "rigidity," particularly in the transition state. Solvent screening effects and/or the hydration properties of various substrates could also contribute to such differential preferences.


Figure 3
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  		Scheme 3. Energy diagram illustrating the overall reaction coordinate for Fpg-DNA binding and catalysis. The relevant thermodynamic quantities are the activation free energy ({Delta}G{ddagger}) and activation enthalpy ({Delta}H{ddagger}), the free energy ({Delta}Gb) and enthalpy ({Delta}Hb) associated with the Fpg–substrate interaction, and the resultant activation free energy ({Delta}G{ddagger}T) and enthalpy ({Delta}H{ddagger}T) associated with kcat/Km.

Concluding Remarks. We have developed a continuous hyperchromicity assay (CHA) to map the energy landscape of a BER enzyme. The CHA method is general and can be used to characterize a myriad of enzyme systems that target nucleic acid substrates in a site-specific manner. The CHA approach is adaptable to virtually any site-specific enzyme assay in which a phosphodiester bond is hydrolyzed, including but not limited to restriction enzymes, endonucleases, repair enzymes such as those that recognize and process lesions (i.e., mismatches, adducts), chemical modifications of bases and nucleotides (bases, sugars, or phosphate backbone), and DNA folding anomalies/motifs (e.g., bulges). Because of the well known impact of a lesion or adduct on duplex energetics as well as the modulatory effects of sequence context and solution conditions, the method provides a window of applicability that is sufficiently broad to be useful in the characterization of base (BER) and nucleotide (NER) excision repair enzymes. The assay in its current form yields both kinetic and thermodynamic parameters for an enzyme reaction, thereby providing a rapid and precise methodology for expanding our knowledge of the forces that dictate and control enzymatic catalysis. The data reported herein provide a well resolved mapping of the energetic landscape associated with Fpg recognition and catalysis. Such profiles are required to define the origins of enzyme specificity, information that is essential for rational modulation of enzymatic activities.


   Materials and Methods
Top
 Abstract
 Results
 Discussion
 Materials and Methods
ACKNOWLEDGMENTS.
 References
 
Enzyme and Substrate Preparation. Wild-type Fpg was overexpressed and purified from B834 (DE3) Escherichia coli harboring a PET13a-fpg plasmid as described (32). Concentrated Fpg preparations were stored at –20°C in a buffer system comprising 50 mM Hepes-KOH (pH 7.6), 250 mM NaCl, 1 mM EDTA, and 1 mM DTT diluted 1:1 (vol:vol) with glycerol (Buffer A). Fpg protein concentrations were determined by UV absorption spectroscopy using a calculated molar extinction coefficient of 39,410 M–1 cm–1 at 280 nm. The oligonucleotides used in this study were synthesized using standard phosphoramidite chemistry. The 8-oxodG lesion was incorporated as the central base (X) within the sequence d(GCGTACXCATGCG). The modified deoxyoligonucleotide was synthesized and purified as described (33), and the extinction coefficients determined by complete nuclease digestion, followed by spectroscopic analysis of the products.

Kinetics Method. Kinetics measurements were performed on either an Aviv Model 14 (Aviv Biomedical) or a Varian Model 4000 (Varian) UV/Vis spectrophotometer, both of which are equipped with a peltier-controlled thermostatted sample compartment for accurate temperature regulation. Substrate deoxyoligonucleotide standards ranging in concentration from 10 nM to 250 nM were prepared in a prefiltered and degassed buffer system composed of 10 mM sodium phosphate and 100 mM NaCl adjusted to pH 7.0 (Buffer B). An appropriate volume (2.50 ml) of each duplex substrate solution was dispensed into a 10 x 10-mm quartz cuvette and equilibrated at the desired temperature (±0.05°C) under constant stirring for 10 min. The kinetic measurement was initiated by pipetting an aliquot of enzyme stock solution into the equilibrated duplex substrate solution, followed by continuous monitoring of the absorbance increase at 260 nm. Progress reaction curves were collected over the time interval of 0–600 sec to monitor enzymatic processing of the substrate. Product formation was estimated by using the relative extinctions of substrate and product, the latter evaluated quantitatively based on model compounds (refer to SI Fig. 6). In this control experiment, product formation was mimicked by decreasing the fraction of fully associated 13-mer duplex and proportionally increasing the amounts of a stoichiometric ratio of product (i.e., a mixture of 13-mer single strand, 5'- and 3' 6-mer complementary strands, and dGTP) while maintaining a constant deoxyoligonucleotide concentration of 1.0 µM (SI Fig. 6). The resultant data confirm the validity of employing the hyperchromicity at 260 nm to quantitatively determine product formation.


   ACKNOWLEDGMENTS.
Top
 Abstract
 Results
 Discussion
 Materials and Methods
 ACKNOWLEDGMENTS.
References
 
We thank Konstantin Kropachev for expression and purification of the Fpg protein preparations used in this study. This work was supported by National Institutes of Health Grants GM-23509, GM-34469, and CA-47795 (to K.J.B.).


   Footnotes
 
*To whom correspondence should be addressed. E-mail: kjbdna{at}rci.rutgers.edu

Freely available online through the PNAS open access option.

Author contributions: C.A.S.A.M., D.P.R., and K.J.B. designed research; C.A.S.A.M. and D.P.R. performed research; C.A.S.A.M. and D.P.R. analyzed data; and C.A.S.A.M., D.P.R., and K.J.B. wrote the paper.

The authors declare no conflict of interest.

This article contains supporting information online at www.pnas.org/cgi/content/full/0710363105/DC1.

© 2008 by The National Academy of Sciences of the USA


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 Results
 Discussion
 Materials and Methods
 ACKNOWLEDGMENTS.
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