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J. Biol. Chem., Vol. 282, Issue 39, 28884-28892, September 28, 2007

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A Pre-equilibrium before Nucleotide Binding Limits Fingers Subdomain Closure by Klentaq1^*

From the Institute of Structural Molecular Biology, Birkbeck College and University College London, Malet Street, London, WC1E 7HX, United Kingdom

Received for publication, June 12, 2007 , and in revised form, July 18, 2007.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Numerous studies have been undertaken to establish the mechanism of dNTP binding and template-directed incorporation by DNA polymerases. It has been established by kinetic experiments that a rate-limiting step, crucial for dNTP selection, occurs before chemical bond formation. Crystallographic studies indicated that this step may be due to a large open-to-closed conformational transition affecting the fingers subdomain. In previous studies, we established a fluorescence resonance energy transfer system to monitor the open-to-closed transition in the fingers subdomain of Klentaq1. By comparing the rates of the fingers subdomain closure with that of the rate-limiting step for Klentaq1, we showed that fingers subdomain motion was significantly faster than the rate-limiting step. We have now used this system to characterize DNA binding as well as to complete a more extensive characterization of incorporation of all four dNTPs. The data indicate that DNA binding occurs by a two-step association and that dissociation of the DNA is significantly slower in the case of the closed ternary complex. The data for nucleotide incorporation indicate a step occurring before dNTP binding, which differs for all four nucleotides. As the only difference between the (E·p/t) complexes is the templating base, it would suggest an important role for the templating base in initial ground state selection.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
DNA polymerases constitute the central component of the DNA replication machinery. The basic role of a DNA polymerase is to catalyze the addition of a dNMP to a primer strand based on its complementarity to a template base provided by a template strand. Various DNA polymerases have been discovered, specialized for different roles, and despite sharing a common catalytic mechanism for the nucleotidyl transfer reaction, vary considerably in their accuracy during template directed DNA synthesis. Most polymerases studied, irrelevant of their specificity, share a rate-limiting conformational change, which occurs before chemistry and is utilized during selection of a correct nucleotide.

A basic model for DNA polymerases was proposed based on early kinetic work on T4 DNA polymerase (1), T7 DNA polymerase (2, 3), HIV-1² reverse transcriptase (HIV-1 RT) (4) and the Klenow fragment of DNA polymerase I from Escherichia coli (Klenow) (5-8). In this model, the first step is binding of the primer/template (p/t) substrate to the unliganded polymerase (E). For polymerases in which spectroscopic techniques have been applied to study p/t binding, a more complex interaction between the enzyme and the substrate has been observed, displaying either multiple binding modes or a sequential binding mode for HIV-1 RT and polymerase (pol ), respectively (9-12). After p/t binding, there follows an initial loose binding of the dNTP substrate. This initial ground state binding of the dNTP is used by some polymerases such as HIV-1 RT and T7 DNA polymerase, which are both replicative enzymes, to discriminate against non-complementary dNTPs by a factor of 250- and 390-fold, respectively (2, 4). For repair polymerases, the differences between correct and incorrect nucleotide binding are much smaller. Klenow only selects against incorrect nucleotide incorporation on average by a factor of 3.4, yeast polymerase by a factor of 4, and polymerase by a factor of 20 (7, 8, 13-15). Binding of the correct nucleotide leads to a conformational change, which converts a loose E·p/t·dNTP ternary complex into a tight activated E'·p/t·dNTP complex capable of undergoing chemistry. The rate of this conformational transition was named k_pol. In general, this step is the point at which the strongest discrimination occurs. It is also the slowest, rate-limiting step in the nucleotide incorporation cycle. After chemistry occurs, there then follows the release of the pyrophosphate product and either translocation or dissociation of the DNA product.

Structural studies of DNA polymerase as exemplified by Klentaq1, a family A repair enzyme, have solved many intermediates in the reaction pathway including the apo form of the polymerase, the E·p/t and E·dNTP binary complexes, as well as the ternary complex, E'·p/t·dNTP, which was formed using a "non-reactive" 3' primer terminus with the next correct nucleotide bound to form a "trapped closed" ternary complex (16-23). The structural states reveal a limited conformational change occurring in the thumb subdomain upon p/t binding and a large conformational change occurring in the fingers subdomain upon binding of the correct nucleotide to the E·p/t complex. The change in conformation of the fingers subdomain results in the formation of the binding site for the nascent base pair. The original idea was that the open-to-closed transition seen structurally corresponds to the kinetically defined conformational change required for selection.

However, a large body of evidence now indicates that the kinetically defined conformational change does not correspond to the motion of the fingers subdomain for most polymerases. For pol , a mutation of Asp-276, a residue that makes contact with the incoming nucleotide only in the closed conformation, results in an increase in free energy of dNTP ground state binding (24), suggesting that the rate-limiting conformational change is not the open-to-closed structural transition but instead is triggered in the closed polymerase conformation (24). Similar conclusions were drawn from inspection of the rate-limiting kinetics of wild-type pol and a pol variant mutated at Tyr-265, a residue that does not make any contact with the DNA or the dNTP (25, 26). Fluorescence studies on pol also indicated that motions in the fingers subdomain appear to be fast, but the extent to which it is not rate-limiting was not addressed (27). The conclusions that the rate-limiting conformational change actually occurs in the closed conformation would appear to also apply to the repair enzymes, such as Klentaq1 and Klenow. It was shown using the Klenow fragment and 2-aminopurine as a reporter that a 2-aminopurine at the templating position (the first unpaired template base) undergoes a sizable and very rapid decrease in fluorescence associated with dNTP binding (28). If the motion of the templating base is coupled to the open-to-closed transition affecting the fingers subdomain, then the open-to-closed conformational transition affecting the fingers subdomain must also be at least as fast. In contrast, for T7 DNA polymerase, recent fluorescence studies support a rate-limiting conformational change affecting the fingers subdomain (29). However, despite the positioning of a fluorophore in the fingers subdomain, it is unclear whether this probe is monitoring the overall movement of the fingers subdomain or more subtle changes occurring in this subdomain. Finally, we developed a FRET system on Klentaq1 in which the motion of the fingers subdomain was directly monitored upon nucleotide binding and incorporation; this study indicates that the fingers subdomain motion is fast and not rate-limiting (23).

We have now used this FRET system to characterize DNA binding as well as complete a more extensive characterization of binding and incorporation of all four dNTPs. The data indicate that DNA binding occurs by a two-step association and that dissociation of the DNA is significantly slower in the case of the closed ternary complex, E'·p/t·dNTP. The data for nucleotide incorporation indicate a step occurring before nucleotide binding and fingers subdomain closure. The dependence of this step on nucleotide concentration differs for each dNTPs. As the only difference between the E·p/t complexes is the templating base, it would suggest an important role for the templating base in initial dNTP ground state binding.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Purification and Labeling of Klentaq1 and KT^V649C—Klentaq1 was purified according to Korolev et al. (16). The mutation Val-649 to Cys (KT^V649C) was introduced using the Stratagene QuikChange kit, and the sequence was confirmed by DNA sequencing. Purification of the mutant protein was performed as for wild-type Klentaq1. The concentration of protein was calculated using a molar extinction coefficient of 69,622 cm^-1 M^-1 at 280 nm. Cysteine 649 was labeled with the acceptor fluorophore Alexa Fluor 594-C₅ maleimide (Molecular Probes, Leiden, Netherlands) according to Ref. 23 to form Alexa Fluor 594-labeled Klentaq1 (KT^V649C(A594)). Labeling efficiencies were typically 90% or greater.

Nucleotides—100 mM solutions of dNTP and dideoxynucleoside triphosphates (ddNTPs) were purchased from GE Healthcare (Amersham, UK).

Oligonucleotides—The Alexa Fluor 488-labeled DNA primer strand was purchased from IBA (Göttingen, Germany). The donor Alexa Fluor 488 probe was coupled using an N-hydroxysuccinimidyl-activated Alexa Fluor 488 dye from Molecular Probes reacted on a commercially available 5-C6-Amino-2'-deoxythymidine. The primer used, p^A488-6, was a 40-mer (sequence 5'-CAGCGCCACTGGGTCAGTCCGAGCCGTCGCAGCCTACCGT-3') in which the donor-labeled base is indicated in bold. The template oligonucleotides used in this study have the following sequences (bold indicates the base-pairing region). The templates are called t_A, t_C, t_G, t_T, with the letters in capital corresponding to the next dNMP to be incorporated. The primers are as follows: t_A, 5'-TGGTTAATCTCTGCCTACGGTAGGCTGCGACGGCTCGGACTGACCCAGTGGCGCTG-3'; t_C, 5'-TGGTTAATCTCTCTAGACGGTAGGCTGCGACGGCTCGGACTGACCCAGTGGCGCTG-3'; t_G, 5'-TGGTTAATCTCTTAGCACGGTAGGCTGCGACGGCTCGGACTGACCCAGTGGCGCTG-3'; t_T, 5'-TGGTTAATCTCTGCTAACGGTAGGCTGCGACGGCTCGGACTGACCCAGTGGCGCTG-3'. The hybridization of primer and template was performed according to Rothwell et al. (23). Stock solutions of 100 µM were made in 50 mM Tris, pH 7, and 20 mM NaCl.

Steady State Titration Measurements—Steady state titration experiments to determine the affinity of the E·p/t interaction were recorded using a Fluoromax-3 (Jobin Yvon Horiba) at 20 °C. A solution of p^A488.6/t_G at a concentration of 100 nM in 20 mM Tris-HCl (pH 7.5), 50 mM NaCl, 2 mM MgCl₂ was made, and the fluorescent signal of the donor was monitored over time until a stable signal was observed. Following the addition of an increasing amount of KT^V649C(A594) (see Fig. 1A), the sample was mixed and was allowed to equilibrate until a stable signal was observed. Alexa Fluor 488 was excited at 496 nm, and the fluorescence emission was recorded at 520 nm with slits set at 4 and 1 nm for excitation and emission, respectively. Data were normalized to 1 using Origin 7.0 (OriginLab Corp.) and the signal of the donor-labeled p^A488.6/t_G as a reference. The FRET system was used here because it yielded better data quality than when the unlabeled enzyme was used. It was also used to remain consistent with the pre-steady state measurements (see below).

Pre-steady State Kinetic Measurements—All stopped flow experiments were performed at 20 °C in 20 mM Tris-HCl (pH 7.5), 50 mM NaCl, 2 mM MgCl₂. Data were recorded using a Hi-Tech scientific DX-2 stopped flow system with excitation of Alexa Fluor 488 at 493 nm. Donor and acceptor fluorescence were separated using the filters XF3084 (Glen Spectra; bandpass range 510-570 nm) for Alexa Fluor 488 and XF3028 (Glen Spectra; bandpass range 615-650 nm) for Alexa Fluor 594. Slits were set at 10 nm for both excitation and emission. The voltage on both photo multipliers varied depending on the conditions and were optimized for signal-to-noise. Each trace shown represents the average of at least three "shots."

To monitor formation of the E·p/t complex, a solution of 20 nM p^A488.6/t_G was mixed rapidly with a varying concentration of KT^V649C(A594) (100-1200 nM), and the signal produced from the decrease in the donor fluorescence of the labeled DNA was recorded. Although a signal decrease is also observed by mixing labeled DNA with the unlabeled wild-type enzyme, larger concentrations of the p^A488.6/t_G are needed, typically 100 nM, to produce a recordable signal. Thus, the labeled enzyme was used for all experiments.

For p/t dissociation experiments from the binary E·p/t or ternary E'·p/t·dNTP complex, the p/t was terminated as described previously (23) to produce a complex of 1 µM KT^649C(A594):500 nM p^A488.6/t_G_TE (where t_G_TE indicates a t_G primer terminated by ddGTP with the next nucleotide to be incorporated being dCTP). For a DNA dissociation experiment from the binary E·p/t complex, the 1 µM KT^649C(A594):500 nM p^A488.6/t_G_TE complex was loaded into one sample chamber with the second sample chamber containing 0.2 mg/ml heparin or 20 µM unlabeled p/t (final concentration). For p/t dissociation experiments from the ternary E'·p/t·dNTP complex, the E'·p/t·dNTP was first produced from the KT^649C(A594):p^A488.6/t_G_TE complex by adding dCTP to a final concentration of 250 µM. The resulting closed ternary complex was mixed rapidly with 0.2 mg/ml heparin to monitor dissociation kinetics. For nucleotide incorporation experiments, preincubated KT^V649C(A594) (1 µM final concentration) and p^A488.6/t_X (500 nM final concentration), where X is the next nucleotide to be incorporated, were loaded into one sample chamber with the second sample chamber containing varying concentrations of the corresponding dNTP to be incorporated.

In all stopped flow experiments except for the DNA dissociation experiments, a "blank" run was performed in which reaction buffer was mixed rapidly with either the labeled p^A488.6/t_G alone for DNA binding experiments or the KT^V649C(A594)·p^A488.6/t_X (where X is the next nucleotide to be incorporated) complex for dNTP incorporation experiments. This was subtracted from each individual data set using Origin 7.0. Data were fitted using Grafit 5.0 (Erithacus Software Ltd.) between 0.02 and 1.5 s according to the models described under "Results."

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
We previously designed a FRET system to monitor the fingers subdomain motion of Klentaq1 upon nucleotide incorporation (23). The system consists of a primer/template (p/t) labeled six bases away from the 3' terminus of the primer with the donor fluorophore Alexa Fluor 488 called p^A488.6/t_X (where X is the next nucleotide to be incorporated). In combination with Klentaq1 labeled with Alexa Fluor 594 at an introduced cysteine at position 649 in the fingers subdomain, called KT^V649C(A594), a measurable change in FRET occurs. This is due to the fingers subdomain closure on nucleotide binding, which results in the acceptor fluorophore in the fingers subdomain moving closer to the donor fluorophore on the p/t DNA. We have now extended the use of this system to monitor the association of the p/t to Klentaq1 and perform a more complete characterization, involving all four dNTPs, of the fingers subdomain motion upon nucleotide incorporation.

Primer/Template DNA Binding to Klentaq1—To investigate the affinity and binding of KT^V649C(A594) to p/t, both steady state and pre-steady state association experiments were performed. For the steady state titration experiments, a fixed concentration of donor labeled p^A488.6/t_G of 100 nM was used, and increasing amounts of KT^V649C(A594) were added (Fig. 1A). The data were fitted with the equation below where F, F_max, and F_min are the recorded, maximum, and minimum fluorescence signals, respectively. E₀ and S₀ are the initial concentrations of KT^V649C(A594) and p^A488.6/t_G, respectively, and K_D is the dissociation constant.

(Eq.1)

Fitting of the data gave a K_D value of 86.5 ± 8.8 nM, which is slightly lower than the published value of 160 nM (31).

We next investigated DNA binding using stopped flow to determine the pre-steady state binding parameters. Fig. 1B shows raw data obtained from the association kinetics of pA488.6/t_G to KT^V649C(A594). The donor labeled p/t was kept at a fixed concentration (20 nM end concentration) and mixed rapidly with increasing amounts of KT^V649C(A594), and the change in donor fluorescence was recorded. From the data, two exponential phases are apparent. The first exponential phase is protein concentration-dependent. The rate constants (k_obs) for this phase obtained at various protein concentrations were plotted against protein concentration (Fig. 1C), and the data were fitted using the linear equation,

(Eq.2)

where k₁ and k_-1 represent the forward and reverse rate constants for the first phase binding step. The rate constants k₁ and k_-1 are derived from the slope and the intercept with the y axis of the plot of k_obs versus enzyme concentration, respectively (Fig. 1C). For this step of p/t binding, values for k₁ of 5.3 x 10⁸ ± 0.2 x 10⁸ M^-1 s^-1 and k_-1 of 16.5 ± 7.5 s^-1 were determined. The k_-1 value is inaccurate due to uncertainty about the intercept at the y axis. Nevertheless, K_D determined from k_-1/k₁ was 31 nM, in the same order of magnitude as that obtained from steady state measurements of 86.5 nM. The second phase is not well resolved over the concentration range measured but appears to show no dependence on KT^V649C(A594) concentration and has an average value of 16 s^-1. This step may represent an additional conformational change occurring after p/t binding or may represent a partitioning into a second binding mode.

Association experiments were also performed with the labeled p/t and wild-type Klentaq1 (data not shown). As indicated under "Experimental Procedures," measurements had to be carried out at higher concentrations of 100 nM p^A488.6/t_G but provided very similar results (not shown). This indicates that the two phases observed are not due directly to FRET, rather the acceptor fluorophore on the labeled KT^V649C(A594) is amplifying the signal obtained from the reduction in the donor fluorescence on the labeled p/t.

View larger version (14K): [in this window] [in a new window]   FIGURE 1. Binding of the acceptor labeled KTV649C(A594) to donor labeled pA488.6/t_G. A, steady state titration of the acceptor labeled KTV649C(A594) against 100 nM pA488.6/t_G. Excitation was at 493 nM, and emission was recorded at 520 nm. The data were fitted as described under "Results" and gave a value of 86.5 ± 8.8 nM. B, pre-steady state kinetic measurement of p/t binding. A typical stopped flow result is shown in which a 20 nM sample of pA488.6/t_G was mixed rapidly with 500 nM KTV649C(A594)). Excitation was at 493 nm, and the donor fluorescence was collected using band pass filter between 510 and 570 nm. The inset shows the first 50 ms of the binding. The data were fitted with a single exponential (dot), single exponential with slope (dash), and a double exponential (solid) with the best fit being a double exponential. C, dependence of the pseudo-first order rate constant on KTV649C(A594) concentration. A constant concentration of 20 nM pA488.6/t_G was mixed with increasing amount of KTV649C(A594). k1 and k-1 were determined by the slope of the linear fit and the intercept of the line with the y axis and yielded values of 5.3 x 108 ± 0.2 x 108 M-1 s-1 and 16.5 ± 7.5 s-1, respectively.

View larger version (16K): [in this window] [in a new window]   FIGURE 2. Kinetics of dissociation of the donor-labeled pA488.6/t_GTE from the acceptor-labeled KTV649C(A594). A complex of 1 µM KT649C(A594):500 nM pA488.6/t_GTE was mixed rapidly with 0.2 mg/ml heparin. The terminated substrate was used to provide a direct comparison between the dissociation of the binary (E·p/t) and the ternary (E'·p/t·dCTP) complexes, although the non-terminated p/t gives essentially the same results. Excitation was at 493 nm. The donor fluorescence (green) was collected using a band pass filter between 510 and 570 nm, and the acceptor fluorescence (red) was collected using a band pass filter between 615 and 650 nm. Two time scales were recorded and superimposed on the graph. The data were fitted with a double exponential (dash) for both donor and acceptor and gave values of 38.70 ± 0.33 and 37.70 ± 0.35 s-1 for the first phase and 3.09 ± 0.01 and 3.42 ± 0.01 s-1 for the second phase for the donor and acceptor, respectively. The relative amplitude of the first phase as compared with the total amplitude is 52% for both donor and acceptor.

View larger version (25K): [in this window] [in a new window]   FIGURE 3. Kinetics of dissociation of pA488.6/t_GTE from the ternary complex. A complex of 1 µM KTV649C(A594):500 nM pA488.6/t_GTE·250 µM dCTP was mixed rapidly with 0.2 mg/ml heparin. Excitation was at 493 nm, the donor fluorescence (green) was collected using a band pass filter between 510 and 570 nm, and the acceptor fluorescence (red) was collected using a band pass filter between 615 and 650 nm. The data were fitted with a double exponential (dot) and a triple exponential (dash) for both donor and acceptor with the triple exponential model providing the best fit. Fitting of the data gave values of 69.40 ± 0.33 and 42.40 ± 0.35 s-1 for the first phase, 0.72 ± 0.01 and 1.01 ± 0.01 s-1 for the second phase, and 0.13 ± 0.01 and 0.26 ± 0.01 s-1 for the third phase for the donor and acceptor, respectively. The relative amplitude of the phases is 22% for phase 1 for both donor and acceptor, 62% for donor and 52% for acceptor for phase 2, and 16 and 26% for donor and acceptor, respectively, for phase 3. The inset shows a comparison between dissociation of the binary complex (lighter shades of green and red for donor and acceptor, respectively) and the ternary complex (green and red for donor and acceptor, respectively).

Nucleotide Dependence of Fingers Subdomain Closure—Fig. 4A shows typical data obtained for single dCTP incorporation into the KT^V649C(A594)·p^A488.6/t_C complex for three different dCTP concentrations. Only the first 1.5 s of the trace are shown, which show the increase in FRET (decrease in donor fluorescence and increase in acceptor fluorescence) due to fingers subdomain closure. The second component of the signal (not shown) consists of an increase in donor fluorescence and, although a corresponding decrease in acceptor fluorescence is observed, is not due to FRET (see Ref. 23 for details). The data shown in Fig. 4A were fitted with a single exponential and slope, and the change in FRET occurring during fingers subdomain closure in response to differing concentrations of dNTPs was determined. The rate constants derived from the single exponential fit (k_obs) were then plotted as a function of dNTP concentration (Fig. 4B). In Fig. 4B, each point is the average of at least three measurements with the error bars being generated from the standard error on averaging for each concentration. For dATP, dTTP, and dCTP, due to a low amplitude of the FRET increase phase at low concentrations, it was only possible to reliably record down to 50 µM.

What can be observed immediately is that the rate of fingers subdomain closure decreases as a function of increasing dNTP concentrations. Such a trend in the data has been previously interpreted as an indication that the rate of a binding reaction is limited by a pre-equilibrium step occurring beforehand where the first step (the pre-equilibrium step) is slow as compared with the second step (the binding step) (30). Thus, we propose here that the rate observed for fingers subdomain closure is limited by a step occurring beforehand. For such a case, the model proposed by Fersht (30) and adapted here is the following.

Here, we omitted showing the (E·p/t)^*·dNTP species because it immediately transitions to the [(E·p/t)^*·dNTP]_closed form. Derivation by Fersht (30) leads to the following.

(Eq.3)

This model envisages two E·p/t complexes, E·p/t or (E·p/t)^*, in equilibrium, only one of which is competent for dNTP binding.

It is interesting that for the different nucleotides, the rates of k₃ and k_-3 vary. For both dATP and dTTP, the forward and reverse rates are similar, having values of 1.49 and 1.58 s^-1 for k₃ and 8.68 and 7.72 s^-1 for k_-3, respectively. However, for dCTP incorporation, the forward and reverse rates are more similar, k₃ being 4.07 s^-1 and k_-3 being 7.40 s^-1, whereas for dGTP incorporation, the forward and reverse rates are almost identical (k₃ = 2.21 s^-1 and k_-3 = 1.56 s^-1). The rates would indicate that in the case of dATP and dTTP, the enzyme is predominantly in the E·p/t state; for dCTP, less so, and dGTP would be predominantly in the (E·p/t)^* state. Although the K_D4 (see Equation 3) values are inaccurate due to the lack of information at lower concentrations for dATP, dCTP, and dTTP, there seem to be almost equivalent K_D values for the purine-based nucleotides dATP and dGTP of 173 and 175 µM, respectively, and for the pyrimidine-based nucleotides dCTP and dTTP of 236 and 215 µM, respectively. These values are higher than previously determined by quench flow measurements and probably do not represent the K_D for ground state binding recorded by this technique (31).

View larger version (18K): [in this window] [in a new window]   FIGURE 4. Dependence of fingers subdomain closure on dNTP concentration. A, raw data obtained for single nucleotide incorporation by Klentaq1. A complex of 1 µM KTV649C(A594):500 nM pA488.6/t_C was mixed rapidly with increasing amounts of dCTP. Excitation was at 493 nm, the donor fluorescence was collected using a band pass filter between 510 and 570 nm, and the acceptor fluorescence was collected using a band pass filter between 615 and 650 nm. Three concentrations of nucleotides are shown. Red is 1000 µM, green is 500 µM, and blue is 250 µM. The rich colors show the donor signal, and the faded colors show the acceptor signal. The data were fitted between 0.02 and 1.5 s with a single exponential with slope. B, secondary data obtained from the rate obtained from the FRET-increase phase for dATP (), dCTP (), dGTP (), dTTP (). The data were fitted using Equation 3 and gave values of k3 and k-3 of 1.49 ± 0.28 and 8.68 ± 0.34 s-1 and KD4 of 172.9 ± 31.6 µM; 4.07 ± 0.37 and 7.40 ± 0.44 s-1 and KD4 of 235.7 ± 66.9 µM; 2.21 ± 0.13 and 1.56 ± 0.12 s-1 and KD4 of 175.2 ± 60.1 µM; and 1.58 ± 0.38 and 7.72 ± 0.35 s-1 and KD4 of 215.9 ± 51.1 µM for dATP, dCTP, dGTP, and dTTP, respectively.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Perhaps a better explanation for the observation of two E·p/t complexes observed from the DNA association and dissociation experiments may be found in the following remarkable feature of the E·p/t complex structure: the stacking of the first paired template base onto the aromatic ring of Tyr-671 (Fig. 5A). This interaction provides a means to register the base-paired portion of the DNA relative to the active site residues. However, upon closure of the fingers subdomain, Tyr-671 escapes this stacking arrangement and, as shown in the ternary E'·p/t·ddNTP complex structure, the templating base of the template strand (i.e. the first unpaired base of the template) swings in its place, positioning itself to form a Watson-Crick base pair with the correct incoming nucleotide (Fig. 5B). If one assumes that the fingers domain in the E·p/t complex (i.e. in the absence of an incoming nucleotide) constantly oscillates between an open and closed form, then the DNA might be released from the stacking arrangement with Tyr-671, and thus, might be free to move within the confines of the p/t-binding site of the enzyme. Indeed, molecular dynamics simulations suggest that the DNA-binding site may form an electrostatic tube within the quasi-cylinder that the p/t-binding site forms and within which no preferential interaction with any particular residue occur leaving the DNA free to slide within the site. Thus, the E·p/t₍₁₎ and E·p/t₍₂₎ states may correspond to two binding modes within the p/t-binding site, these two modes only differing in relation to Tyr-671 and the conformational transition affecting the fingers subdomain. Two p/t-binding modes have also been observed in HIV 1-RT, and a similar interpretation was suggested (9, 12, 32). For HIV 1-RT, the two E·p/t species differ by onlya5Å shift in the position of the nucleic acid. These two states were described as "educt" and "product" states in the polymerization reaction cycle whereby the educt state is a complex in which the nucleic acid is positioned to allow nucleotide incorporation and the product state corresponds to a state formed immediately after nucleotide incorporation but before RT translates to the next nucleotide.

View larger version (48K): [in this window] [in a new window]   FIGURE 5. Crystal structures of the E·p/t (A), E'·p/t·ddNTP ternary closed (B), and E·p/t·ddNTP ternary open (C) complexes. In A, B, and C, the panel on the left is a schematic diagram of the stereo view panel of the structure shown at right. In the stereo view panels at the right, the fingers, palm, and thumb subdomains are shown in C- trace representation, color-coded in green, magenta, and dark blue, respectively. The O-helix is shown in red. The p/t DNA is shown in stick representation, color-coded in yellow and cyan for the template and primer strands, respectively. The first templating base is shown in dark blue. The incoming nucleotide and the magnesium ions bound to it are shown in ball-and-stick representation, color-coded in orange. Tyr-671 is shown in ball-and-stick representation with carbon, oxygen, and nitrogen color-coded in light gray, red, and blue, respectively. In the panels at left, the color-coding is the same as in the corresponding panels at the right. The first paired template base is indicated, as are the various sites mentioned under "Results." In A, a representation of a nucleotide is shown in tuned orange color; it has been added bound to the O-helix, as is observed in the E·dNTP structures of Li et al. (17). Protein Data Bank entry codes for the E·p/t, E'·p/t·ddNTP closed, and E·p/t·ddNTP open complexes are 4KTQ, 2KTQ, and 3KTQ, respectively.

View larger version (15K): [in this window] [in a new window]   FIGURE 6. Models for templating base and dNTP concentration dependence of fingers subdomain closure. In both panels, definition and color-coding of the various players are as in Fig. 5. In each case, the equilibrium is with the pre-insertion state, and thus, only the state of the templating base corresponding to its dNTP interacting state is shown. A, binding of the dNTP to the templating base in its insertion state. B, binding of the dNTP to the templating base in its intermediate pre-insertion state.

Although the pre-insertion versus insertion states equilibrium of the templating base may account for the observed base-specific behavior, the observed dependence on nucleotide concentration can only be explained if one of these states is involved in nucleotide ground state binding. This implies that the templating base might participate in nucleotide binding before the fingers subdomain has closed. One could envisage the incoming nucleotide binding directly in the active site as is shown in Fig. 5C and the templating base in its insertion state interacting with the incoming nucleotide through Watson-Crick pairing as shown in Fig. 6A. In the configuration shown in Fig. 5C, which was observed crystallographically by Li et al. (17), the incoming nucleotide is in the active site, i.e. bound to the catalytic residues Asp-610 and Asp-785 with only one magnesium ion involved; however, the fingers subdomain is not clamped onto the nucleotide as it is in the open conformation and Tyr-671 locates opposite the base of the incoming nucleotide. To account for the dNTP concentration dependence for fingers subdomain closure described here, the Tyr-671 would need to move in and out of stacking arrangement with the first paired template base in the open state of the fingers subdomain to allow the templating base to move in and out of the insertion state site as shown in Fig. 6A.

A second model is suggested by Temiakov et al. (33), who have described a state of T7 RNA polymerase, where the templating base would locate between the pre-insertion and insertion state sites (Fig. 6B). In that third state, termed "intermediate pre-insertion state," the templating base would be close enough to the incoming nucleotide bound to the nucleotide-binding site (the O-helix in the case of Klentaq1) in the open fingers subdomain to interact with the incoming nucleotide through Watson-Crick base pairing. Thus, the pre-equilibrium, which we observed here, as it must be templating base- and dNTP concentration-dependent, would consist of an equilibrium between a pre-insertion state and an intermediate pre-insertion state, the nature of which would differ depending on the identity of the templating base.

Whether the pre-equilibrium revealed here reflects the pre-insertion versus insertion states equilibrium or the pre-insertion versus intermediate pre-insertion states equilibrium of the templating base and Tyr-671, both are sufficient to explain the observed nucleotide concentration dependence of fingers subdomain closure. For both models, there would be little difference in the ground state binding affinity of each of the dNTPs, rather closing of the fingers subdomain would amplify what little differences there are between Watson-Crick and non Watson-Crick base pairs and provide a closed reaction site in which nucleotide incorporation could occur.

Replicative DNA polymerases, such as T7 DNA polymerase, appear to be capable of strong discrimination in the ground state. It would be interesting to perform similar experiments on T7 DNA polymerase to determine how this greater level of discrimination is seen.

	FOOTNOTES

 
^* This work was funded by Grant 067879 from the Wellcome Trust. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement"in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

This article was selected as a paper of the week.

¹ To whom correspondence should be addressed. Tel.: 44-207-631-6833; Fax: 44-207-631-6803; E-mail: g.waksman{at}mail.cryst.bbk.ac.uk or g.waksman{at}ucl.ac.uk.

² The abbreviations used are: HIV, human immunodeficiency virus; FRET, fluorescence resonance energy transfer; RT, reverse transcriptase; p/t, primer/template; pol, polymerase; ddNTP, dideoxynucleoside triphosphate.

	ACKNOWLEDGMENTS

 
We thank Dr. Timothy Lohman (Washington University School of Medicine, St. Louis, MO) and Dr. Thomas Durek (Institute for Molecular Bioscience, University of Quensland, Brisbane, Australia).

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