Probing DNA Polymerase Fidelity Mechanisms Using Time-Resolved Fluorescence Anisotropy

METHODS 25, 62–77 (2001) doi:10.1006/meth.2001.1216, available online at http://www.idealibrary.com on

Probing DNA Polymerase Fidelity Mechanisms Using Time-Resolved Fluorescence Anisotropy Michael F. Bailey, Elizabeth H. Z. Thompson, and David P. Millar1 Department of Molecular Biology, The Scripps Research Institute, 10550 North Torrey Pines Road, La Jolla, California 92037

Prior to undergoing postsynthetic 3⬘–5⬘ editing (proofreading), a defective DNA primer terminus must be transferred from the 5⬘–3⬘ polymerase active site to a remote 3⬘–5⬘ exonuclease site. To elucidate the mechanisms by which this occurs, we have used time-resolved fluorescence spectroscopy to study the interaction of dansyl-labeled DNA primer/templates with the Klenow fragment of Escherichia coli DNA polymerase I. The dansyl probe is positioned such that when the DNA substrate occupies the polymerase active site, the probe is solvent-exposed and possesses a short average fluorescence lifetime (4.7 ns) and extensive angular diffusion (42.5⬚). Conversely, when the DNA substrate occupies the exonuclease active site, the probe becomes buried within the protein, resulting in an increase in the average lifetime (14.1 ns) and a decrease in the degree of angular diffusion (14.4⬚). If both polymerase and exonuclease binding modes are populated (lower limit ⬃5%), their markedly different fluorescence properties cause the anisotropy to decay with a characteristic “dip and rise” shape. Nonlinear least-squares analysis of these data recovers the ground-state mole fractions of exposed (xe) and buried (xb) probes, which are equivalent to the equilibrium proportions of the DNA substrate bound at the polymerase and exonuclease sites, respectively. The distribution between the polymerase and exonuclease binding modes is given by the equilibrium partitioning constant Kpe (equal to xb/xe). The important determinants of the proofreading process can therefore be identified by changes made to either the protein or DNA that perturb the partitioning equilibrium and hence alter the magnitude of Kpe. 䉷 2001 Academic Press Key Words: DNA polymerases, DNA/protein interactions, fluorescence anisotrophy, time-resolved fluorescence, polymerase fidelity, DNA labeling.

The application of fluorescence spectroscopy to the study of nucleic acid–protein interactions is accompa1 To whom correspondence should be addressed. Fax: (858) 7849067. E-mail: [email protected].

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nied by many advantages. Among these are the sensitivity, time scale, and versatility of the fluorescence phenomena that may be studied. Sensitivity allows for detection of events at low concentrations. For example, with the advent of single-molecule techniques, an encounter between a single enzyme and substrate may be observed. Even bulk methods, however, can allow detection of nanomolar concentrations of appropriately labeled material, enabling access to binding phenomena at and below the Kd values characteristic of many DNA–protein interactions. The time scale of fluorescent events is similar to that of many interesting dynamic processes, including charge transfer and quenching, therefore enabling study of these phenomena. Additionally, since fluorescence studies are performed in solution, the effects of a wide range of experimental conditions may be explored. When applied to a particular system and studied in depth, a wealth of information is available from fluorescence spectroscopy. In this article, we apply time-resolved fluorescence anisotropy to study the complex inner workings of a model DNA polymerase. DNA polymerases perform the faithful replication of an organism’s genetic material, a task that requires incorporation of a specific base across from its complement. Cooperation between two separate activities, one that extends the nascent DNA primer strand and one that removes misincorporated bases, is necessary to ensure accurate replication. While a wide variety of DNA polymerases have evolved to carry out faithful replication, polymerases from various organisms appear to share structural motifs and many highly conserved residues (1, 2). Given the sequence and structural similarities, it is likely that insights into structure–function relationships gleaned from a model polymerase would be widely applicable (3). 1046-2023/01 $35.00 Copyright 䉷 2001 by Academic Press All rights of reproduction in any form reserved.

FLUORESCENCE ANALYSIS OF DNA POLYMERASE FIDELITY

The model polymerase used in these studies is the Klenow fragment of Escherichia coli DNA polymerase I. Klenow fragment is relatively small and simple, requiring no accessory proteins to synthesize DNA with high fidelity. Its single 68-kDa polypeptide chain contains both a 5⬘–3⬘ polymerase active site and a 3⬘–5⬘ ˚ apart in space (4). exonuclease site, located about 30 A While these features alone indicate a tractable model system, a wealth of structural and biochemical information about the Klenow fragment is also available, providing an extensive framework that may prove useful when interpreting findings. The schematic in Fig. 1 illustrates the interaction between a DNA substrate and the two binding sites of

63

the polymerase. In a mixture of polymerase and substrate, there exists an equilibrium distribution of the substrate between the two binding sites. Using a system where each binding mode is characterized by distinct fluorescence behaviors, the distribution of the substrate can be quantitatively assessed. By examining the substrate distribution in mutant polymerases in which specific amino acid side chains have been altered, the polymerase–DNA interactions that stabilize each mode of binding can be dissected in detail. A series of studies based on this approach have assessed the energetic contribution individual amino acid side chains make to binding the DNA substrate. Other studies address the effect of particular features of the DNA substrate on the partitioning equilibrium. The theory of fluorescence anisotropy and the instrumentation necessary to make these measurements are discussed in some depth in this article. In addition, an overview of the experimental design and a summary of the findings on the features of the polymerase and DNA that dictate a given mode of interaction are given.

RATIONALE OF EXPERIMENTAL SYSTEM

FIG. 1. Diagram depicting the interaction of primer/template DNA with the Klenow fragment of E. coli DNA polymerase I (KF). The primer is represented by the shorter, lighter-shaded strand and the template by the longer, darker-shaded strand. The dansyl probe, attached to the primer strand via a 5-(aminopropyl)-2⬘-deoxyuridine residue seven nucleotides from the 3⬘ end, is represented by the filled sphere. The regions designated “P” and “E” refer to the 5⬘–3⬘ polymerase and the 3⬘–5⬘ exonuclease domains of KF, respectively. Formation of the replicative complex, described by the equilibrium constant Kp, entails the binding of the primer’s 3⬘ terminus to the polymerase site as a duplex. In this case the probe is solvent-exposed and capable of extensive motion about its point of attachment. Formation of the editing complex, described by the equilibrium constant Ke, entails the binding of the primer’s 3⬘ terminus to the exonuclease site as a frayed end. In this case the probe is buried within the protein, which significantly reduces its degree of local rotation. Interconversion of the two complexes (by an as yet undetermined mechanism) is described by the equilibrium partitioning constant Kpe.

To differentiate between the polymerase and exonuclease modes of DNA binding, our experimental system requires the use of an extrinsic fluorophore. In principle, this could be attached to either the protein or the DNA. The ␧-amino group of lysine residues and the ␤-sulfhydryl group of cysteine residues provide convenient sites for the fluorescent labeling of proteins. However, the relatively large number of these residues in most proteins makes it difficult to control the location and degree of labeling, meaning that preparations are invariably heterogeneous and difficult to purify. A range of modified nucleoside phosphoramidites bearing reactive amines or thiols can be incorporated into synthetic oligonucleotides to provide a unique site for fluorescent labeling, either within the chain or at the termini (5). For several reasons, deoxyuridine residues bearing nucleophilic functions at position 5 of the pyrimidine ring have become common vehicles for the internal labeling of oligonucleotides. First, rather than introducing a novel functional group into one of the other DNA bases, derivatization of position 5 of the uracil ring builds on a previously existing functionality, namely, the methyl group of thymine. Second, since position 5 substituents of the pyrimidine ring are not involved in hydrogen bonding, subsequent fluorescent

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BAILEY, THOMPSON, AND MILLAR

modification will not directly affect helix stability. Although there are several commercially available 5-aminodeoxyuridine phosphoramidites (5), we use a 5(aminopropyl)-2⬘-deoxyuridine phosphoramidite synthesized in-house (6). This has a three carbon spacer separating the amino group from the pyrimidine ring, which is significantly shorter than those of the commercial derivatives. We feel that the closer the probe is tethered to the DNA, the less likely it is to sterically hinder binding to the Klenow fragment. The sequences of the oligonucleotide primer/templates described in these studies are shown in Fig. 2A. For time-resolved anisotropy data to reflect the presence of heterogeneous fluorophore populations and thereby report the abundance of the polymerase and exonuclease binding modes, it is mandatory that each population possess unique fluorescence and rotational properties. This requires judicious choice of both the probe and its location within the DNA. We have chosen the 5-dimethylamino-1-naphthalenesulfonyl (dansyl)

probe for two reasons. First, it is comparatively small and therefore less likely than other, bulkier fluorophores to perturb the intrinsic polymerase s exonuclease partitioning equilibrium. Second, it is extremely sensitive to its local environment, having a short average fluorescence lifetime in aqueous media and a significantly longer average fluorescence lifetime in nonpolar media. The dansyl probe is attached to the 5-(aminopropyl)-2⬘-deoxyuridine residue, located seven nucleotides from the 3⬘ end of the primer strand (Fig. 2). When positioned thus, the probe inhabits vastly different environments depending on where the primer’s 3⬘ terminus is located within the enzyme (Fig. 1). When the terminus is bound to the polymerase site as a duplex, the probe is exposed to the aqueous solvent and capable of extensive local motion. However, when the terminus is bound to the exonuclease site as a frayed end, the concomitant translocation of the DNA draws the probe into a nonpolar environment within the protein, resulting in a significant restriction in its local

FIG. 2. Details of the oligonucleotide primer/templates used to study partitioning of DNA between the polymerase and exonuclease sites of Klenow fragment. (A) Nucleotide sequences and nomenclature of primer 17-mers and template 27-mers. The primer strand, designated 17*-mer, contains a 5-dimethylamino-1-naphthalenesulfonyl (dansyl) probe covalently attached to a 5-(aminopropyl)-2⬘-deoxyuridine residue, denoted by the letter X. The template strands, designated 27-mer, 27G-mer, 27GG-mer, 27ATG-mer, and 27CCTT-mer are designed so that the duplexes resulting from their hybridization with 17*-mer will contain either zero, one, two, three, or four terminal mismatches, respectively. The duplexes are designated 17*/27-mer, 17*/27G-mer, 17*/27GG-mer, 17*/27ATG-mer, and 17*/27CCTT-mer, respectively. Mismatched base pairs, if present, are underlined. (B) Chemical structure of the dansyl-labeled 5-(aminopropyl)-2⬘-deoxyuridine residue. The 5-(aminopropyl)2⬘-deoxyuridine and dansyl moieties are delineated for clarity.

FLUORESCENCE ANALYSIS OF DNA POLYMERASE FIDELITY

rotation (Fig. 1). This combination of factors is particularly conducive to producing the “dip and rise” anisotropy decays that are needed to report the equilibrium fractions of substrate bound at each active site. All Klenow fragment derivatives used in these studies contain the D424A (aspartic acid-to-alanine) mutation. This essentially abolishes the 3⬘–5⬘ exonuclease activity, but does not compromise substrate binding, enabling investigation of active site switching without the added complication of substrate degradation (7). This is a particularly important concern because exonucleolytic shortening of the primer strand could give rise to a subset of primer/templates that have the dansyl probes buried within the protein when bound at the polymerase site. This would report a larger proportion of exonuclease-bound substrate than is actually present and would result in an overestimate of the partitioning constant Kpe. Although the D424A derivative is a mutant form of the Klenow fragment, it is referred to throughout the text as the wild type for the purpose of comparison with derivatives carrying further mutations.

PREPARATION OF OLIGONUCLEOTIDE PRIMERS/TEMPLATES Oligonucleotides are synthesized using standard ␤ cyanoethyl phosphoramidite chemistry. The 5-(aminopropyl)-2⬘-deoxyuridine phosphoramidite is incorporated into the primer strand and derivatized with dansyl chloride as described (8). Oligonucleotides are purified from 20% denaturing polyacrylamide gels and transferred into the experimental buffer (50 mM Tris–HCl, pH 7.5, 3 mM MgCl2) by gel filtration. Concentrations are determined spectrophotometrically at 260 nm, using molar extinction coefficients calculated by the nearestneighbor method (9). No correction is made for the small contribution the dansyl probe makes to the 260-nm absorption. Primer/template duplexes are prepared by mixing equal volumes of 6.4 ␮M primer and 6.8 ␮M template solutions, heating at 80⬚C for 10 min, then allowing to cool slowly to room temperature (2–3 h).

PREPARATION OF KLENOW FRAGMENT The host strain CJ376 and the plasmid pVD2 encoding the D424A mutant of the Klenow fragment of E. coli DNA polymerase I were kindly provided by Dr. C. Joyce (Yale University, New Heaven, CT). Site-directed mutagenesis of pVD2 was carried out using a commercially available kit (Stratagene, La Jollo, CA). The entire coding region of each clone is sequenced to confirm

65

that the desired mutation(s) is present and that extraneous mutations have not arisen as a result of polymerase chain reaction (PCR) amplification. Klenow fragment is overexpressed and purified as described (10), except that the protein is finally transferred into a buffer of 10 mM Tris–HCl, pH 7.5, 1 mM EDTA. To avoid loss of binding activity caused by repeated freeze– thaw cycling, the protein is concentrated to a stock of 100–150 ␮M, aliquotted into 50-␮l portions, flash frozen in liquid N2, and stored at ⫺80⬚C. Protein concentrations are determined spectrophotometrically, using ␧280 ⫽ 5.88 ⫻ 104 M⫺1 cm⫺1, calculated from the predicted amino acid sequence by the shareware program SEDNTERP.

FORMATION OF KLENOW FRAGMENT– PRIMER/TEMPLATE COMPLEXES The fractions of buried and exposed probes recovered from the analysis of anisotropy decays exhibiting a “dip and rise” are especially sensitive to the local rotational properties of the dansyl probe. Since the local rotational properties are the same for dansyl probes attached to free DNA and to DNA bound at the polymerase site of Klenow fragment (11), the presence of free DNA in the sample could weight the data artificially in favor of the polymerase binding mode and lead to an underestimate of the partitioning constant. To eliminate this possibility, the amount of protein required to saturate the DNA is determined empirically by a steady-state fluorescence anisotropy titration. Titrations are performed at 20⬚C with an SLM Aminco 8100 spectrofluorometer using the L-format method. A 150-␮l solution of 3.2 ␮M dansyl-labeled primer/template is pipetted into a quartz fluorescence microcuvette and titrated with small (1–2 ␮l) aliquots of freshly thawed Klenow fragment. Following each addition, the cuvette is gently shaken and returned to the sample compartment for 5 min to equilibrate. The intensity of the vertically polarized component of the fluorescence emission (IVV) is measured at 545 nm (8-nm spectral bandpass) for 2 s using vertically polarized excitation of 345 nm (8-nm spectral bandpass). The emission polarizer is then rotated 90⬚ and the intensity of the horizontal component of the fluorescence emission (IVH) measured as above. After the emission polarizer is returned to the vertical position, the excitation polarizer is rotated 90⬚ to the horizontal position. The intensity of the vertically and horizontally polarized components of the emission, in this case denoted IHV and IHH, respectively, are measured sequentially as above. Each of the four polarized intensity measurements is corrected for background

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(obtained with a reagent blank) and the fluorescence anisotropy calculated according to Eq. [8]. Each anisotropy measurement represents the average of 10 individual determinations and has a typical standard deviation of 0.001 unit. Binding is monitored by the increase in fluorescence anisotropy accompanying addition of Klenow fragment and is considered complete when this parameter no longer increases with added protein (Fig. 6A, inset). The high (micromolar) concentrations of primer/template used in these experiments are required to measure the relatively weak emission of the dansyl fluorophore (quantum yield Q ⫽ 0.1–0.3) with an acceptable signal-to-noise ratio. Since the primer/ template concentration is approximately three orders of magnitude above the Kd describing the interaction (7 nM) (8), the binding is essentially stoichiometric (i.e., equivalence point ⬇ endpoint). This, in conjunction with the ability to prepare highly concentrated protein stocks, means that the endpoint can be attained with minimal (ⱕ 5%) dilution of the components of the original solution.

The fluorescence anisotropy, r, is defined as the ratio of the intensity of the polarized component of the emission to the total intensity of the emission. Division of Eq. [1] by Eq. [2] gives Ip I 㥋 ⫺ I⬜ ⫽r⫽ . IT I㥋 ⫹ 2I⬜

[3]

As anisotropy measurements are conventionally carried out using vertically polarized excitation, I㥋 and I⬜ represent the vertical (V) and horizontal (H) components of the emission, respectively. It is convenient to refer to I㥋 as IVV and I ⬜ as IVH, as this notation indicates both the order and orientation of the excitation and emission polarizers. Substitution of IVV for I㥋 and IVH for I⬜ in Eq. [3] yields r⫽

IVV ⫺ IVH . IVV ⫹ 2IVH

[4]

In most cases the emission wavelength chosen to measure the polarized emission intensities is selected with a monochromator. The holographic gratings in these devices generally do not transmit the vertically and horizontally polarized components of the emission with

DESCRIPTION OF METHOD 1. Fluorescence Anisotropy When a fluorophore is excited with polarized light, oriented for example along the z axis as shown in Fig. 3, some fraction of its emission is polarized in the same direction. The emission radiated along the z axis is parallel to the direction of the polarized excitation. Its intensity (I㥋) can be measured through a vertically oriented polarizer, positioned on the x axis. As implied above, I㥋 is composed of both polarized and unpolarized components. The unpolarized component of the emission can be isolated by rotating the polarizer 90⬚ and measuring the intensity of the emission radiated along the y axis (I⬜), which is perpendicular to the direction of the polarized excitation. The intensity of the polarized component of the emission, Ip, is given by I p ⫽ I 㥋 ⫺ I ⬜.

[1]

When excitation is polarized along the z axis, the excited-state distribution is symmetric around this axis (12). Thus, the intensity of the emission radiated along the y axis is equal to that radiated along the x axis. Using the equality Ix ⫽ Iy ⫽ I⬜, the total emission intensity IT , given by Ix ⫹ Iy ⫹ Iz , can be expressed as IT ⫽ I㥋 ⫹ 2I⬜.

[2]

FIG. 3. Setup for fluorescence anisotropy measurements using the L-format (single-channel) method. After passing through a polarizer (P1) and monochromator (M1), vertically polarized light (oriented parallel to the z axis) of the desired wavelength travels along the y axis and excites the sample contained in the cuvette. The emission oriented parallel to the direction of polarized excitation (I㥋) is composed of a polarized component (shaded double-headed arrow) and an unpolarized component (thick solid double-headed arrow). The emission oriented perpendicular to the direction of polarized excitation (I⬜) is completely depolarized. Passage of I㥋 through a vertically aligned polarizer (P2) and monochromator (M2) yields vertically polarized emission of the desired wavelength (IVV), which is recorded by a detector positioned along the x axis. After sufficient acquisition time, polarizer P2 is rotated 90⬚ to the horizontal position and IHV is measured by collecting I⬜ as above.

FLUORESCENCE ANALYSIS OF DNA POLYMERASE FIDELITY

equal efficiencies, which, if not corrected, will result in inaccurate data. This problem may be overcome by placing a polarization scrambler (depolarizer) between the emission polarizer and emission monochromator. A more common method, however, is to experimentally determine the difference in transmission efficiencies and incorporate the correction factor into Eq. [4]. For the more common L-format (single channel) method used to measure fluorescence anisotropy, the correction factor (termed the grating correction factor or G factor) is acquired by measuring the vertically and horizontally polarized emission intensities obtained with horizontally polarized excitation (i.e., IHV and IHH). When the excitation is horizontally polarized, the excited-state distribution of the fluorophore is rotated to lie along the x axis as shown in Fig. 3 (12). In this situation both polarizer orientations are perpendicular to the direction of polarized excitation, hence both the vertical and horizontal emission components are equal to I⬜. Any difference in the measured intensities will therefore be due to the instrument and not the polarization of the sample. For horizontally polarized excitation, the observed polarized emission intensities are IHV ⫽ SV I⬜

[5]

IHH ⫽ SH I⬜,

[6]

and

where SV and SH are the sensitivities with which the vertically and horizontally polarized emission components are detected, respectively. Division of Eq. [5] by Eq. [6] yields the G factor S V I⬜ SV IHV ⫽ ⫽ ⫽ G. IHH SH I⬜ SH

[7]

For example, a G factor of 1.2 indicates that the emission monochromator passes vertically polarized light at the desired wavelength with a 1.2-fold greater efficiency than it passes horizontally polarized light. Incorporation of the G factor into Eq. [4] (12) yields the more familiar formulation used to calculate fluorescence anisotropies: r⫽

IVV ⫺ GIVH . IVV ⫹ 2GIVH

[8]

Equation [8] indicates that the anisotropy has a value of 1 when IVH ⫽ 0 and a value of zero when IVV ⫽ IVH. In practice, a value of unity is observed only with

67

scattered light. The anisotropy of a solution of fluorophores is considerably less than this due to the intrinsic effect of photoselection and the extrinsic effects of transition dipole displacement and rotational diffusion. In dilute vitrified solution, depolarizing factors such as nonradiative energy transfer and rotational diffusion (discussed later) are eliminated. Under these conditions the theoretical maximum anisotropy of a solution of fluorophores can be expressed as r⫽

3具cos2␪E典 ⫺ 1 , 2

[9]

where ␪E is the angle the emission dipole of the fluorophore makes to the z axis (Fig. 3). For a solution of randomly oriented fluorophores, the probability of excitation is proportional to cos2␪A, where ␪A is the angle the absorption dipole of the fluorophore makes to the z axis (Fig. 3). Thus, fluorophores having their absorption dipoles aligned parallel to the z axis have the greatest probability of being excited, an effect known as photoselection. Given random orientation, the maximum value of 具cos2␪A典 is 3/5. Assuming that the absorption and emission dipoles within the fluorophore are colinear (i.e., ␪A ⫽ ␪E), the maximum anisotropy for a solution of fluorophores is 2/5 (0.4). In reality, the absorption and emission dipoles are not colinear, but rather displaced by an angle ␣A,E within the plane of the fluorophore. The intrinsic anisotropy of a dilute vitrified solution of fluorophores is given by the product of the loss of anisotropy resulting from photoselection and from the angular displacement of the absorption and emission dipoles, r ⫽ r0 ⫽

3具cos2␪E典 ⫺ 1 3 cos2␣A,E ⫺ 1 2 2

冢

冣冢

冣

⫽ 0.4

冢

3 cos2␣A,E ⫺ 1 , [10] 2

冣

where r0 is the limiting anisotropy, that is, the anisotropy observed when photoselection and dipolar displacement are the only factors responsible for depolarization. In most cases, fluorophores are studied in dilute nonviscous solutions and undergo significant rotational diffusion on the fluorescence time scale. Molecular motion causes further angular displacement of the emission dipole relative to the z axis, yielding a further loss of anisotropy. The effect of rotational diffusion on the fluorescence anisotropy of a spherical molecule is described by the Perrin equation.

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BAILEY, THOMPSON, AND MILLAR

r⫽

r0 , 1 ⫹ (␶ /␾)

[11]

where ␶ is the fluorescence lifetime and ␾ is the rotational correlation time. Both of these terms, which typically have units of nanoseconds, are defined in greater detail in the proceeding section. Although more complicated expressions are required to account for the anisotropy of nonspherical molecules, it is useful to examine the limiting behavior predicted by the Perrin equation. When ␾ À ␶, r ⬇ r0. However, when ␶ À ␾, r ⬇ 0. The rotational correlation time is given by

␾⫽

␩V , RT

the vertical, meaning that the total intensity will be measured irrespective of the polarization of the sample. Second, the fluorescence anisotropy decay is obtained using the same L-format geometry shown in Fig. 3 and described earlier, except that the vertically and horizontally polarized components of the emission are measured in a time-resolved fashion. The information obtained from each type of time-resolved decay measurement is described below. 2.1. Decay of Isotropic Fluorescence The time-dependent isotropic fluorescence intensity I(t), can generally be represented as a sum of exponential decays according to

[12]

where ␩ is the viscosity of the solvent (cP), V is the volume of the rotating unit (liters mol⫺1), R is the gas constant (J mol⫺1 K⫺1), and T is the absolute temperature (K). Provided ␶ and ␾ are reasonably well matched, the anisotropy will be a function of the molecular weight of the fluorescent species. This dependence has made fluorescence anisotropy an attractive parameter for monitoring interactions between biological macromolecules in solution. In general, the anisotropy of the fluorescent species will increase on formation of a higher molecular weight complex, and decrease upon its dissociation. In summary, the anisotropy will be sensitive to any factor causing rotation of the emission dipole of the fluorophore relative to the z axis (Fig. 3) during the lifetime of the excited state.

2. Time-Resolved Fluorescence To appreciate the methods we use to investigate DNA polymerase fidelity mechanisms, it is necessary to review some of the principles behind the acquisition, interpretation, and analysis of time-resolved fluorescence data. Time-resolved fluorescence data can be acquired either in the time domain, using time-correlated singlephoton counting, or in the frequency domain, using phase demodulation. In the former method, which is practiced in our laboratory, the direct time decay of emission is recorded following a brief excitation pulse. Two types of fluorescence decay can be measured using time-resolved techniques. First, the isotropic fluorescence decay contains information on the fluorescence lifetime(s) of a sample and is recorded with the emission polarizer oriented at 54.7⬚ to the vertically polarized excitation. When the emission polarizer is oriented thus (magic angle conditions), the intensity of the horizontal component of the emission will always be twice that of

I(t) ⫽

N

兺 ␣j exp(⫺t/␶ ) , j⫽1 j

[13]

where N is the number of decay components and ␣j and ␶j are the fractional amplitude and lifetime associated with the jth decay component, respectively. Further, N 兺j⫽1 ␣j ⫽ 1. The simulated isotropic intensity decay shown in Fig. 4A is for a single component, indicative of a homogeneous fluorophore population. In this case, ␣ is equal to unity and ␶, defined as the time for the intensity to decay to 1/e of its initial value, is 10 ns. In many cases, however, fluorescence decays are more complex, even for a homogeneous probe population, due to emission from multiple excited states or solvent relaxation. The simulated isotropic intensity decay in Fig. 4B is for two components, for which ␣1 ⫽ 0.6, ␶1 ⫽ 1 ns, ␣2 ⫽ 0.4, and ␶2 ⫽ 10 ns. When more than one decay component exists, it is convenient to refer to the average fluorescence lifetime ␶, which is given by

␶ ⫽ 兺j ␣j␶j2 /兺j ␣j␶j .

[14]

2.2. Decay of Fluorescence Anisotropy The time-dependent fluorescence anisotropy, r(t), is calculated from the polarized components of the intensity decay, r(t) ⫽

IVV(t) ⫺ GIVH(t) , IVV(t) ⫹ 2GIVH(t)

[15]

where IVV(t) and IVH(t) are the time-dependent decays of IVV and IVH defined in Eq. [4]. For a homogeneous population of fluorophores, the time-dependent fluorescence anisotropy can also be represented as a sum of exponential decays according to r(t) ⫽

M

兺 ␤k exp(⫺t/␾ ) , k⫽1 k

[16]

FLUORESCENCE ANALYSIS OF DNA POLYMERASE FIDELITY

where M is the number of decay components and ␤k and ␾k are the anisotropy amplitude and rotational correlation time associated with the kth decay component, respectively. The total limiting anisotropy is r0, where r0 ⫽ 兺M k⫽1 ␤k. The simulated anisotropy decay shown in Fig. 4C is for a single component. This is a unique case, predicted only for a molecule with spherical geometry. In this case, ␤ (equivalent to r0) is 0.25 and ␾ (defined as the time for the anisotropy to decay to 1/e of its initial value) is 20 ns. The simulated anisotropy decay shown in Fig. 4D is for two components and is more typically observed. The faster decay component, characterized in this case by ␤1 ⫽ 0.1 and ␾1 ⫽ 1.2 ns, is usually interpreted as reorientation of an intrinsic fluorophore within a locally flexible domain (such as a tryptophan residue in a protein) or rotation of an extrinsic fluorophore about its point of attachment to the macromolecule (such as a dansyl probe attached to DNA). The slower decay component (␤1 ⫽ 0.15 and ␾1 ⫽ 20 ns) is due to rotational diffusion of the macromolecule itself. A quantitative description of the angular range of the restricted internal rotation of the probe can be calculated by using a model of isotropic diffusion within a cone (13). The semiangle of the cone is given by

␪ ⫽ cos⫺1[(1/2)(1 ⫹ 8S)1/2 ⫺ 1],

[17]

where S ⫽ (␤2 /r0)1/2 is the generalized order parameter (14). For a heterogeneous population of fluorophores, the anisotropy decay can be expressed as r(t) ⫽ fa(t)ra(t) ⫹ fb(t)rb(t)

[18]

where fa(t) are ra(t) are the fraction of the total fluorescence and the anisotropy of a population of probes inhabiting environment a at time t, respectively. Accordingly, fb(t) and rb(t) are the corresponding quantities for the probes inhabiting environment b (15). The function fa(t) is given by Na

兺 ␣ja exp(⫺t/␶

ja)

xa

j⫽1

fa(t) ⫽

Na

xa

兺 ␣ja exp

j⫽1

(⫺t/␶ja)

Nb

⫹ xb

兺 ␣jb exp

,

[19]

(⫺t/␶jb)

j⫽1

where x is the ground-state mole fraction of the fluorophore. The a and b subscripts on the x, ␣j , ␶j , and N parameters refer to the fluorophores in environments a and b, respectively. The ␣ja , ␣jb , ␶ja , ␶jb , Na , and Nb

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parameters are as defined in Eq. [13]. An analogous expression applies for fb(t). Furthermore, xa ⫹ xb ⫽ 1. The anisotropy decay functions ra(t) and rb(t) are each given by expressions of the form shown in Eq. [16] and are given an additional subscript to denote each fluorophore population. The curves in Fig. 4E and 4F essentially recapitulate a subset of our experimental data and serve to show how the distribution of a fluorescent probe between two environments can affect the anisotropy decay. Data were simulated with 90% of the fluorophores in environment a and the remaining 10% in environment b. A full listing of the parameters used to generate the data is provided in the caption to Fig. 4. The salient features differentiating the two populations are that the probes in environment a are characterized by short-lived fluorescence and an extensive range of local rotation (␶a ⫽ 4.7 ns, ␪a ⫽ 42.5⬚), whereas the probes in environment b are characterized by a considerably longer-lived fluorescence, but a more restricted range of local rotation (␶b ⫽ 14.1 ns, ␪b ⫽ 14.4⬚). It must be stressed that although the range of local rotation for the probes in each environment is different, the rate of motion (both local and global) is the same (i.e., ␾1a ⫽ ␾1b , ␾2a ⫽ ␾2b). Figure 4E shows the fraction of fluorescence contributed by each fluorophore population as a function of time after excitation. Due to their significant excess, the probes in environment a make the greatest contribution to the fluorescence at early times in the decay (Fig. 4A). As time progresses, however, this more rapidly decaying population contributes less to the observed fluorescence, its share eventually being overshadowed by the more slowly decaying probes in environment b. This differential contribution to the fluorescence by the two populations over the course of the decay, combined with their distinct local mobilities, gives rise to a “dip and rise” shape in the anisotropy decay (Fig. 4B). At early times, the anisotropy decays rapidly, reflecting reorientation of the probes in environment a. At later times, when the fluorescence from this population has decayed away, the anisotropy rises to approach the value characteristic of the restricted probes in environment b, before slowly decaying to zero. The hypothetical fluorophores inhabiting environments a and b are analogous to the exposed (e) and buried (b) dansyl probes in our experimental system. Consequently, the subscripts a and b attached to the decay parameters defined above are replaced by the subscripts e and b when referring to these parameters in the context of our system.

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3. Data Acquisition For a more detailed description of pulsed laser sources and time-correlated single-photon counting than is provided below, the reader is referred to monographs by Svelto (16) and O’Connor and Phillips (17).

A schematic depiction of the time-correlated single-photon counting instrument used in our laboratory is shown in Fig. 5. Fully bound DNA samples from the steady-state anisotropy titrations are placed in a temperature-controlled housing thermostated at 20⬚C and

FIG. 4. Simulated isotropic and anisotropic fluorescence decays. (A) The simulated isotropic fluorescence decay is generated according to Eq. [13] for N ⫽ 1, ␣ ⫽ 1, and ␶ ⫽ 10 ns. (B) As for (A), except that N ⫽ 2, ␣1 ⫽ 0.6, ␶1 ⫽ 1 ns, ␣2 ⫽ 0.4, and ␶2 ⫽ 10 ns. (C) The simulated fluorescence anisotropy decay is generated according to Eq. [16] for M ⫽ 1, ␤ ⫽ 0.25, and ␾ ⫽ 20 ns. (D) As for (C), except that M ⫽ 2, ␤1 ⫽ 0.1, ␾1 ⫽ 1.2 ns, ␤2 ⫽ 0.15, and ␾2 ⫽ 20 ns. (E, F) Phenomena associated with fluorescence decays arising from a heterogeneous population of fluorescent probes. The two probe populations are assumed to inhabit either environment a or environment b. (E) The continuous curve, describing the fraction of the total fluorescence contributed by the probes in environment a as a function of time, was generated according to Eq. [19] for xa ⫽ 0.9, Na ⫽ 3, ␣1a ⫽ 0.3, ␶1a ⫽ 1.01 ns, ␣2a ⫽ 0.674, ␶2a ⫽ 4.13 ns, ␣3a ⫽ 0.026, ␶3a ⫽ 11.9 ns, xb ⫽ (1 ⫺ xa) ⫽ 0.1, Nb ⫽ 2, ␣1b ⫽ 0.7, ␶1b ⫽ 3.06 ns, ␣2b ⫽ 0.3, and ␶2b ⫽ 18.4 ns. The broken line, describing the fraction of the total fluorescence contributed by the probes in environment b as a function of time, is generated by an expression analogous to Eq. [19] using the parameters above. (F) The simulated fluorescence anisotropy decay is generated according to Eq. [18] using ␤1a ⫽ 0.132, ␾1a ⫽ 1.2 ns, ␤2a ⫽ 0.071, ␾2a ⫽ 57 ns, ␤1b ⫽ 0.039, ␾1b ⫽ 1.2 ns, ␤2b ⫽ 0.208, ␾2b ⫽ 57 ns. The mole fractions of each population and their associated lifetime parameters are as described in (E).

FLUORESCENCE ANALYSIS OF DNA POLYMERASE FIDELITY

repetitively excited using the frequency-doubled output of a synchronously mode-locked and cavity-dumped dye laser (Coherent 702). The frequency-doubled light is tunable from 285 to 340 nm, depending on the dye solution used, and the repetition rate is adjustable from 150 kHz to 3.8 MHz. The pulse duration is typically ⬃5 ps. For measurements of dansyl-labeled DNA, an excitation wavelength of 318 nm and a repetition frequency of 1.9 MHz are typically used. In a recent modification of the system shown, the dye laser has been replaced by a mode-locked titanium–sapphire laser (Coherent Mira), which provides shorter pulses (⬍2 ps) and a wider tuning range. When used with a suitable frequency doubling or tripling system, the titanium– sapphire laser produces light pulses that are continuously tunable in the wavelength range 240 to 525 nm, which allows for excitation of a wide variety of fluorophores. The pulse repetition frequency, determined by an external pulse selector, is similar to that of the cavity-dumped dye laser. The fluorescence from the sample is collected at 90⬚ to the excitation beam, collimated by a lens, passed through a polarizer that is mounted on a motor-driven rotary stage, and then focused on the entrance slit of a 10-cm-focal-length single-grating monochromator (JY H-10). The polarizer direction can be set to vertical, horizontal, or magic angle orientations, under computer control. A polarization scrambler placed between the emission polarizer and the monochromator is used to eliminate the polarization bias of the grating, obviating the need to determine a G factor. For measurements of dansyl emission, the monochromator is set to 535 nm with a spectral bandpass of 8 nm. A microchannel plate photomultiplier (MCP, Hamamatsu R2809U-01) placed at the exit slit is used to detect the fluorescence. The single-photon pulses at the output of the MCP are amplified by 20 dB in a wide-bandwidth amplifier (Mini Circuits Z1042-J) and used to trigger a constant fraction discriminator (CFD, Tennelec 455). The CFD output is sent to the START input of a time-to-amplitude converter (TAC, Tennelec 934). An electrical pulse is derived from a photodiode viewing a portion of the excitation beam, processed by a second CFD, and then delayed by 20–50 ns and sent to the STOP input of the TAC. For each detected fluorescence photon, the TAC generates a pulse whose amplitude is proportional to the time elapsed between START and STOP events. These are sorted and stored in the memory of a multichannel analyzer (MCA, Oxford PCA3) in order of increasing delay time. For the arrangement shown in Fig. 5, the resulting decay profile is reversed in the MCA memory, with low channel numbers corresponding to the tail of the decay and high channel numbers corresponding to

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the beginning of the decay. This “inverted” configuration is more efficient than the “normal” configuration in which the laser pulse is used to start the TAC, and the decay profile can be readily reversed once the memory contents of the MCA are transferred to a computer. The isotropic emission decay of samples, used in the determination of fluorescence lifetimes, is measured with the emission polarizer at the magic angle and is accumulated until at least 40,000 photon counts have been collected in the peak channel. For measurements of time-resolved fluorescence anisotropy, two separate decays are recorded in different segments of the MCA memory, one with the emission polarizer parallel to the excitation polarization and the other with a perpendicular orientation. The polarizer is switched between these two directions every 15 s and the two decays are accumulated until at least 40,000 photon counts are collected in the peak of the vertical polarization decay. An advantage of the time-correlated single-photon counting technique is that excellent time resolution can be achieved. Because the measurement is based on the time delay between excitation and detection events, the time resolution is determined by the transit time spread in the photomultiplier, which is typically shorter than the rise time. With a microchannel plate detector, this time spread is on the order of 50 ps (full width at halfmaximum) or less. It is important to note that because the excitation pulse is not infinitely narrow and photon detection is not instantaneous, the measured intensity decay is a convolution of the ideal intensity decay with the instrument response function (the response of the instrument to a zero lifetime sample such as the excitation pulse). As a result of this convolution, the measured data represent the sum of many intensity decays, starting with different amplitudes and at different times across the excitation pulse. Clearly, this complicates determination of the decay amplitude(s) and fluorescence lifetime(s). To obtain the ideal intensity decay (i.e., that which would be observed with instantaneous excitation and detection), the measured decay is mathematically deconvolved from the instrument response function. In our system, however, convolution effects can be safely ignored because the response of the detection system (50 ps) is essentially instantaneous in comparison to the fluorescence lifetimes and rotational correlation times characteristic of the dansyl-labeled primer/templates bound to Klenow fragment. 4. Data Analysis Owing to the large number of parameters required to fit a single dip and rise anisotropy decay (ⱖ 15), multiple data sets are globally fit to a form of Eq. [18] in which the exposed population possesses three isotropic

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decay components, the buried population possesses two isotropic decay components, and both populations possess two anisotropy decay components. Global fitting imposes strict limits on the range of values the fitting parameters can assume and therefore greatly improves the precision with which they are recovered. The global data set is obtained by constructing an n ⫻ m matrix of anisotropy decays, where n is the number of Klenow fragment mutants and m is the number of dansyl-labeled primer/templates (containing from zero to four mismatches, Fig. 2A). This produces a data set containing from 0 to 100% buried probes and ensures that the contributions from both probe populations are equally weighted in the analysis. The analysis is also simplified by optimizing only the parameters that the fit is most sensitive to and constraining the values of the other parameters to those determined in separate time-resolved experiments. Nonlinear least-squares analysis of the data (18) is carried out by optimizing the linked anisotropy amplitudes (␤1e, ␤2e, ␤1b and ␤2b) across all data sets, while simultaneously optimizing the fraction of buried probes (xb) for each data set (the

fraction of exposed probes, xe, is computed using the relationship xe ⫽ 1 ⫺ xb). The lifetime parameters of the exposed and buried probes are held constant during the fit and are obtained in separate isotropic decay experiments. The values for the exposed probes may be extracted from the isotropic decay of unbound primer/ template, while those of the buried probes may be determined from the isotropic decay of a complex of Klenow fragment with a quadruply mismatched primer/template, which binds exclusively in the exonuclease mode (Figs. 2A, and 6). The values of the lifetime parameters are as follows: ␶1e ⫽ 1.01 ns, ␣1e ⫽ 0.30, ␶2e ⫽ 4.13 ns, ␣2e ⫽ 0.67, ␶3e ⫽ 11.9 ns, ␣3e ⫽ 0.03, ␶1b ⫽ 3.06 ns, ␣1b ⫽ 0.70, ␶2b ⫽ 18.4 ns, ␣2b ⫽ 0.30. The rotational correlation times are also held constant in this analysis. Their values were determined in a previous study and are as follows: ␪1e ⫽ ␪1b ⫽ 1.2 ns and ␪2e ⫽ ␪2b ⫽ 57 ns (19). In all cases, the anisotropy data are assigned appropriate weighting factors according to the values of IVV (t) and IVH(t) at each time point (17, pp. 270–272). The quality of the fit is judged by the global reduced

FIG. 5. Picosecond laser and time-correlated single-photon counting system used for fluorescence decay measurements. Excitation pulses of a few picoseconds’ duration are derived from a synchronously mode-locked and cavity-dumped dye laser (pumped by an actively modelocked argon ion laser). FD, frequency-doubler for excitation in the ultraviolet region (285–340 nm); MCP, microchannel plate photomultiplier; PD, photodiode; CFD, constant fraction discriminator; TAC, time-to-amplitude converter; MCA, multichannel analyzer.

FLUORESCENCE ANALYSIS OF DNA POLYMERASE FIDELITY

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␹ 2 value and the local reduced ␹ 2 value associated with each data set. Implicit in this analysis are the assumptions that (i) the buried and exposed probes possess the same molar extinction coefficients and radiative rate constants, and (ii) since the time scale of the fluorescence decay (nanoseconds) is significantly faster than that of population interconversion (milliseconds) (20), the ground-state fractions of buried and exposed probes are frozen during the experiment and therefore represent the equilibrium fractions of substrate occupying the polymerase and exonuclease sites, respectively. Given the fractions of buried and exposed probes, the equilibrium constant describing partitioning of the DNA substrate between the polymerase and exonuclease sites (Kpe) can be computed according to Kpe

xe s xb

[20]

and Kpe ⫽

xb . xe

[21]

As xb approaches unity, the value of Kpe becomes poorly defined. Consequently, any measured fraction ⱖ 0.95 is reported as a Kpe of ⱖ 19. FIG. 6. Partitioning of matched and mismatched DNA substrates between the polymerase and exonuclease sites of Klenow fragment. The inset to (A) shows steady-state anisotropy titrations of 17*/27mer (䢇), 17*/27G-mer (䡵), 17*/27GG-mer (䉱), 17*/27ATG-mer (䉲), and 17*/27CCTT-mer (⽧) with D424A Klenow fragment. The sequences of these primer/template duplexes are shown in Fig. 2A. (A) Anisotropy decays, from bottom to top, are for the complexes of D424A Klenow fragment with 17*/27-mer, 17*/27G-mer, 17*/27GG-mer, 17*/ 27ATG-mer, and 17*/27CCTT-mer, respectively. The number under each decay curve indicates the number of mismatches in the DNA substrate. The decay curves represent the averages of three separate experiments; however, the individual decays of the triplicate set are used in the global fit. The smooth curves represent the best global fit of the data to a form of Eq. [18] in which the exposed probe population possesses three isotropic decay components, the buried probe population possesses two isotropic decay components, and both probe populations possess two anisotropy decay components. The anisotropy amplitudes are optimized for best fit across all data sets, while the ground-state mole fraction of buried probes (xb) is optimized for best fit for each data set. The anisotropy amplitudes recovered from the global analysis are: ␤1e ⫽ 0.132, ␤2e ⫽ 0.071, ␤1b ⫽ 0.039, and ␤2b ⫽ 0.208. The average xb values are, from bottom to top, 0.138 ⫾ 0.003, 0.279 ⫾ 0.007, 0.520 ⫾ 0.021, 0.832 ⫾ 0.015, and 0.990 ⫾ 0.012. The other lifetime and anisotropy parameters required to fit the data have been determined in previous studies and are held constant in this analysis: ␶1e ⫽ 1.01 ns, ␣1e ⫽ 0.3, ␶2e ⫽ 4.13 ns, ␣2e ⫽ 0.674, ␶3e ⫽ 11.9 ns, ␣3e ⫽ 0.026, ␶1b ⫽ 3.06 ns, ␣1b ⫽ 0.7, ␶2b ⫽ 18.4 ns, ␣2b ⫽ 0.3, ␾1e ⫽ ␾1b ⫽ 1.2 ns, ␾2e ⫽ ␾2b ⫽ 57 ns. The global reduced ␹ 2 value associated with the fit was 1.1 and the local reduced ␹ 2 values associated with each fit were ⱕ 1.3. (B) Equilibrium partitioning constants are presented as a function of the number of mismatches present in the DNA substrate. The partitioning constant

RESULTS AND CONCLUSIONS Application of the previously detailed methodologies and instrumentation has allowed for the elucidation of a number of factors that affect the partitioning of a DNA substrate between the polymerase and exonuclease sites of the Klenow fragment polymerase. These include features of both of the active sites and of the DNA itself. Figure 6A shows a set of time-resolved anisotropy decays obtained for a set of dansyl-labeled primer/template duplexes bound to Klenow fragment (21). The decays are shown with fit lines that describe the distribution of the substrate between the two sites, and the inset details the steady-state titrations of the duplex necessary to ensure complete binding of the DNA. As

Kpe is obtained using Eq. [21]. The partial sequences shown highlight the positions and identities of mismatches; the full sequence of each duplex substrate is shown in Fig. 2A. Due to the significant increase in exonuclease site partitioning for duplexes containing three and four mismatches, the data for these complexes are shown on a separate set of axes.

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noted previously, the degree of polymerase site partitioning might be overestimated if the system contained a significant fraction of unbound duplex. The experimental data (from bottom decay to top) correspond to complexes of Klenow fragment with DNA containing 0, 1, 2, 3, and 4 consecutive mismatches. The corresponding primer/template sequences are shown in Fig. 2A. The substantially mismatched substrates are not of direct physiological relevance, as the polymerase would be extremely unlikely to undergo two or more misincorporation events in a row without an excision event. However, their use allows for a more accurate description of the fluorescence properties of the buried probes. The equilibrium partitioning constants (Kpe) derived from the analysis of the curves in Fig. 6A are presented in bar-graph form in Fig. 6B. As may be expected, the Kpe values increase with increasing number of mismatches in the primer/template, indicating progressively greater occupancy of the exonuclease site. In all likelihood, the reason for this behavior is twofold. Certainly, the destabilization of the primer terminus due to mismatches should yield a substrate that is increasingly suited for binding as a single strand at the exonuclease site, rather than as a duplex at the polymerase site. Furthermore, other factors that would be expected to lead to destabilization of the terminus, such as increased temperature, result in higher exonuclease site partitioning for a given substrate (unpublished data). However, it has been widely speculated that the polymerase employs mechanisms of its own to detect the correct base paired structure of the DNA substrate, rather than relying entirely on the increased melting capacity of a mispaired terminus. In fact, several factors that might be expected to have little if any effect on the stability of the duplex terminus, including internally positioned bulges and mismatches, and certain perfectly base paired sequences exhibiting unusual helix geometry, show a marked increase in exonuclease site binding (19, 22, 23). As the DNA substrate binds to the exonuclease site as a frayed end, any identification of mismatches must occur in the polymerase site, and therefore rely on interactions between matched or mismatched base pairs and side chains in the polymerase site. In principle, both favorable interactions with matched base pairs and unfavorable interactions with mismatched base pairs could contribute to the ability of the polymerase to discriminate between a correctly base paired duplex and one in which a misincorporation event has occurred. To probe the functional role of specific residues in mismatch recognition and polymerase fidelity, a series of polymerase site mutations were constructed. Residues for mutation were chosen on the basis of the crystal

structure of a homologous polymerase from bacteriophage T7 (24). All residues selected are contained within the polymerase domain and are well removed from the exonuclease domain. Partitioning data for selected polymerase site mutants are presented in Fig. 7 (25). Two mutations were identified that appear to disrupt the polymerase’s ability to identify matched base pairs, as shown in Fig. 7A. Relative to the wild type, the H734A and N845A polymerases exhibit an increased Kpe for a correctly base paired duplex, reflecting the loss of favorable interactions at the polymerase site. Since the same mutations have no effect on partitioning when the duplex contains a single terminal mismatch, the major role of the H734 and N845 side chains appears to be in stabilizing binding of correctly base paired DNA at the polymerase site, rather than mismatch recognition. Two other mutant polymerases, R835L and R841A, appear to favor polymerase site binding, particularly for a mismatched duplex, as seen in Fig. 7B. Since removal of the side chains of R835 and R841

FIG. 7. Effect of polymerase site mutations on the partitioning of matched and singly mismatched DNA substrates between the polymerase and exonuclease sites of Klenow fragment. All mutations are abbreviated using the following convention: the residue number is preceded by the one-letter code for the wild-type amino acid and followed by the code for the mutant amino acid. (A, B) Kpe values describing the partitioning of matched DNA substrates (dark-shaded bars) and DNA substrates containing a single mismatch (light-shaded bars). The data are presented as the averages of three separate measurements and were derived as described in Fig. 6.

FLUORESCENCE ANALYSIS OF DNA POLYMERASE FIDELITY

actually stabilizes binding of the mismatched DNA to the polymerase site, the interaction between these side chains and the mismatched base pair at the primer terminus must be energetically unfavorable in the wildtype polymerase. Thus, these particular side chains appear to have a direct role in the recognition of mismatches at the primer terminus. Taken together, these findings indicate that fidelity is enhanced through a combination of changes in polymerase site binding, including favoring of matched substrates and discrimination against mismatched substrates. Accordingly, a properly base paired duplex is more likely to remain bound in the polymerase site, where it can undergo subsequent rounds of nucleotide incorporation. A mismatched substrate, the result of a misincorporation event, is more likely to translocate to the exonuclease site where the misincorporated base can be removed. The same approach may be applied to interactions between the single-stranded primer terminus and the exonuclease site of the enzyme. Figures 8A and B show the residues selected for mutation and their geometry in the exonuclease active site. Figure 8C shows the change in free energy of partitioning associated with the mutation of the side chain (defined as ⌬⌬G0 ⫽ ⫺RT ln[Kpe (mutant)/Kpe (wild type)]) and therefore quantifies the contribution of that side chain to substrate binding at the exonuclease site (26). A positive ⌬⌬G0 value indicates that the mutant is unable to bind the DNA substrate at the exonuclease site as avidly as the wild type. Conversely, a negative ⌬⌬G0 value demonstrates that the mutant possesses a greater ability to stabilize binding of the DNA substrate at the exonuclease site than does the wild type. In all but one case, the mutation of a side chain results in decreased exonuclease site binding, and therefore a positive ⌬⌬G0 for a DNA substrate containing a single terminal mismatch. Presumably, this indicates that most side chains that contact the DNA in the exonuclease binding mode are involved primarily in stabilizing the binding of the single-stranded primer terminus. The side chains of L361 and F473 appear to be particularly important in this respect, as their removal results in the greatest destabilization of the exonuclease complex. This is in agreement with structural data that show these residues making intimate contacts with the penultimate and terminal bases of the primer strand (Fig. 8A). Interestingly, mutation of Y497 actually results in a more stable exonuclease complex. The interaction between the Y497 side chain and the DNA substrate must therefore be energetically unfavorable. This may reflect

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a role for the tyrosine side chain in straining the singlestranded substrate toward the transition state of the exonuclease reaction. Having identified side chains that interact with the single-stranded substrate at the exonuclease site, it is desirable to probe the subtleties of these interactions. For example, in the related polymerase of bacteriophage RB69, the L361 and F473 homologs are present, but in a reversed orientation. It is of interest, therefore, to determine whether the presence of the two side chains is sufficient to produce wild-type level exonuclease site partitioning or whether the relative position is also important. A set of mutants was constructed to address this question, including single-position substitutions (L361F and F473L) and a double mutant to mimic the RB69 configuration (L361F/F473L). Figure 8D shows the free energy change relative to the wild type for these mutants interacting with a substrate containing a single terminal mismatch (21). The far right-hand column (designated L361F ⫹ F473L) indicates the theoretical free energy change for the RB69 configuration if the effects of the two single mutations were additive. As the double mutant shows a significant defect in binding DNA at the exonuclease site, it would appear that the position, rather than presence of these two side chains, is the prime determinant in stabilizing exonuclease site binding in Klenow fragment. However, it is also noteworthy that the effects of the two mutations are not additive, suggesting that the reversed configuration compensates somewhat for the loss of the wildtype contacts, perhaps by promoting favorable interactions between the substrate and other side chains in the exonuclease site.

APPLICATION OF TIME-RESOLVED ANISOTROPY TO OTHER SYSTEMS From the examples presented in this article, it is evident that time-resolved fluorescence anisotropy is an excellent technique for dissecting the proofreading mechanism of DNA polymerases such as Klenow fragment. The method could also be extended to other members of the DNA polymerase I family that possess similarly separated polymerase and proofreading domains, such as those of Thermus aquaticus and bacteriophage T7 (24, 27). It is important, however, to note that the time-resolved anisotropy technique is applicable to a variety of other systems as well. This includes multifunctional enzymes that act on nucleic acid substrates at different active sites. Examples of these include the

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FIG. 8. Effect of exonuclease site mutations on the partitioning of a singly mismatched DNA substrate between the polymerase and exonuclease sites of Klenow fragment. (A) Single-stranded oligonucleotide bound at the exonuclease active site, surrounded by potentially important side chains. The two metals ions involved in the 3⬘–5⬘ exonuclease reaction are represented by spheres A and B. Replacement of Asp424 with alanine (yielding D424A) results in loss of metal ion B, which is essential for catalytic activity (7). (B) Different view of the active site, highlighting interactions of the oligonucleotide with other side chains. The view is rotated relative to that of (A), as can be seen from the positions of the Leu-361 and Phe-473 side chains. The His-660 side chain has two conformations in the presence of bound DNA: the major conformation interacts with the third base from the 3⬘ end, and the minor conformation forms a hydrogen bond with the nonesterified oxygen of the third phosphodiester bond from the 3⬘ end. Both of these conformations are shown. (C, D) Energetic contribution made by selected exonuclease site residues to DNA binding. The data are presented as averages of three separate measurements and were derived as described in Fig. 6. Kpe values were converted to ⌬⌬G0 values according to the relationship ⌬⌬G0 ⫽ ⫺RT ln[Kpe (mutant)/Kpe (wild type)]. The far right column in (D), titled L361F ⫹ F473L, indicates the ⌬⌬G0 value expected if the effects of the individual L361F and F473L mutations on the partitioning equilibrium were additive.

FLUORESCENCE ANALYSIS OF DNA POLYMERASE FIDELITY

reverse transcriptase of HIV-1, which catalyzes RNAdependent DNA polymerization at one active site, and can degrade the RNA substrate in a separate structural domain, and DNA polymerase III of E. coli, in which the polymerase and proofreading activities are located within different subunits of the multimeric enzyme. In addition to nucleic acid enzymes, there are a variety of proteins that can bind to DNA with more than one footprint. Many transcriptional regulatory proteins, for example, can exist in different multimeric forms, each of which can bind to the DNA. Examples include transcriptional repressors, such as TrpR and TyrR (28, 29), as well as activators such as NtrC (30). In this case, the various multimeric forms of the protein make contact with different regions of the DNA, potentially giving rise to different environments for a fluorescent probe attached to the DNA. In principle, with judicious choice of the labeling position, it would be possible to design a system with two distinct probe environments for any DNA-binding protein that exhibits multiple forms of binding. An appropriate DNA labeling position could be rationally chosen provided structural information is available for the DNA–protein complex in each mode of binding. In other cases, the labeling position can be established empirically. Preliminary time-resolved anisotropy experiments can be carried out using a panel of DNA molecules labeled at different positions to determine which location gives rise to heterogeneous fluorescence behavior. With an appropriately labeled DNA substrate, the method described herein could be used to obtain quantitative information on the equilibrium distribution of the different binding modes under various solution conditions.

ACKNOWLEDGMENTS The research described in this article was supported by a grant from the National Institute of General Medical Sciences (GM 44060). M.B. is the recipient of C. J. Martin Fellowship 145843 from the Australian National Health and Medical Research Council.

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