Real-time measurement of pyrophosphate release kinetics.

There are over 1000 enzymes, including at least one from each of the six major enzyme commission categories, that generate pyrophosphate (PPi) as a product of catalysis (1). Prominent among these are DNA and RNA polymerases in addition to enzymes that utilize ATP and release PPi as one of the products of the reaction. In our ongoing efforts to characterize the kinetics and fidelity of DNA replication, it was necessary to have a sensitive technique for observing pyrophosphate dissociation from the active site of the polymerase following a single nucleotide incorporation reaction. A method was designed for reactions performed under pre-steady-state conditions using a fluorescence stopped-flow apparatus. The coupled assay described here is an adaptation of the methods developed by Webb for sensing the release of phosphate (Pi) using a site-specifically 7-diethylamino-3-((((2-maleimidyl)ethyl)amino)carbonyl)coumarin (MDCC) labeled phosphate binding protein (PBP) (2). The labeled protein exhibits an 8-fold increase in fluorescence upon the binding of phosphate that is both rapid (> 3 × 108 M−1s−1) and tight (Kd ~ 100 nM) (3;4) and has been used to measure the rate of phosphate release from a number of enzymes (5–10). To increase sensitivity, the assay employs a “Pi mop”, consisting of 7-methylguanosine (7-MEG) and purine nucleoside phosphorylase (PNPase) to remove free contaminating Pi from solution by its conversion to ribose 1-phosphate. Under most circumstances the Pi mop does not interfere with the real-time measurement because the rate of Pi consumption is slow in comparison to the rate at which it binds to the PBP, but it does limit the utility of the method for measuring rates less than 0.01 s−1. Here we report a detailed procedure for adapting this sensitive Pi assay for use in measuring PPi release kinetics. The method for the preparation of the MDCC-labeled PBP used during this study was described previously (11). Shown in Fig. 1 are data from a series of experiments performed in a KinTek SF-2004 (Austin, TX) stopped-flow apparatus documenting the utility of the method. Our first experiment (Fig. 1A) was aimed at comparing the time dependence of the fluorescence change associated with directly mixing the PBP with Pi or PPi. In these experiments 1.5 μM PBP was pre-incubated with 100 μM 7-MEG and 0.02 units/mL PNPase for 15 minutes at room temperature. This solution was then rapidly mixed at 20°C with 10 μM Pi or PPi. The sample was excited using a wavelength of 425 nm and observation of the emission was achieved with a 450 nm long pass filter. All concentrations listed refer to the final values after mixing (1:1 dilution) has occurred. The buffer system used for all experiments consisted of 50 mM Tris-base pH 7.5 at room temperature, 100 mM NaCl and 5 mM MgCl2. Fig. 1 Kinetics of the PPi sensor using the PBP The data show that PPi does not induce a significant change in the PBP fluorescence (Fig. 1A), compared to the large signal seen after mixing with excess phosphate. The time dependence of the signal after mixing with Pi was biphasic and was therefore fitted using a double exponential equation to obtain the observed amplitudes (A1 and A2) and rates (k1 and k2) of the two phases: Fluorescence = A1· e−k1·t + A2 · e−k2·t + C. Greater than 90% of the binding reaction occurred at a rate of 327 ± 2 s−1, while the remaining change occurred at 62 ± 2 s−1. The slow phase may represent a small fraction (less than 10%) of less active PBP, but because most experiments are conducted with PBP in excess over Pi, the slower component does not interfere. In contrast, when PPi was mixed with the PBP there was only a small change in fluorescence of about 2% relative to that seen with Pi that could be attributable to small percentage (0.3%) of contaminating Pi in the PPi. To measure PPi quantitatively, PPi must be hydrolyzed to Pi in order to obtain a measurable signal. An experiment was performed in which various concentrations of yeast inorganic pyrophosphatase (PPase) (Sigma-Aldrich St. Louis, MO) were included in the solution containing the PBP. The concentration of the PPase was determined by using the Coomassie Plus assay available from Pierce Biotechnology (Rockford, IL) and assuming a molecular weight of 32,000 Da. In addition, the “Pi mop” components were included in both syringes to eliminate trace Pi contamination and allowed to incubate for 15 minutes at room temperature prior to mixing in the stopped-flow. Two representative traces are shown in Fig. 1B., for the lowest and highest concentrations of PPase inspected (75 nM and 2.9 μM, respectively). The data were biphasic and therefore fitted using a double exponential equation to obtain the observed rates. The concentration dependence of the two phases of the reaction are plotted as a function of PPase concentration in Fig. 1C. The fast phase increased linearly with the concentration of PPase while the slower phase approached a constant value. The kinetics of the coupled PPi sensor reaction can be understood by considering the simplified pathway shown in Scheme I. Scheme I The faster phase of the reaction, under the conditions of the experiments (Figs. 1B and 1C), is defined by the rate of the PPase reaction, while the slower phase is defined by the rate of the Pi binding to the PBP. In general, experiments will be designed so that the concentrations of PPase and the PBP are in excess over the concentration of PPi; therefore, the rate of each step is governed by the product of the concentration of the PPase or the PBP and the corresponding second order rate constant (Scheme I). That is, to a first approximation the rate at which PPi is converted to Pi is defined by the product, k2[PPase], and the rate at which Pi binds to the PBP is defined by the product, k3[PBP]. The slope from the concentration dependence of the fast phase (Fig 1C) defines k2 = 70 μM−1s−1, the apparent second order rate constant governing the conversion of PPi to Pi by PPase and equal to kcat/Km. The binding of Pi to the PBP is a two-step process with the formation of a rapid equilibrium collision complex (Kd = 4.9 μM) followed by isomerization to the highly fluorescent state at a rate of 317 s−1 (4). Pi+PBP⇄4.9μMPi•PBP⇄4.5s−1317s−1Pi•PBP∗ This two-step reaction defines an apparent second order rate constant of 65 μM−1s−1 for the binding of Pi to the PBP. Thus, at a concentration of 1 μM PBP, one expects a rate of slightly less than 65 s−1, which is consistent with the observed rate of 61 ± 5 s−1 (Fig 1C). These data demonstrate that PPase and the PBP can be coupled to monitor PPi at a rate sufficiently fast to be useful for a real-time assay. The overall magnitude of the fluorescence change observed when the PBP is in excess of Pi should also allow for an accurate determination of the concentration of PPi liberated during a reaction. An experiment was performed in which only the concentration of PPi was varied to evaluate whether the amplitude of the signal was a linear function of concentration of PPi. For this experiment, various substoichiometric concentrations of PPi were mixed with 1.5 μM PBP in the presence of 190 nM of PPase. The total change in fluorescence (ΔFobs) varied with the concentration of PPi as shown in Fig. 1D. The data were slightly non-linear and were fitted using following hyperbolic function to generate the standard curve: ΔFobs=ΔFmax·[PPi]K1/2+[PPi]. Fitting the data using this equation afforded a more accurate quantitative description of the trend so that the amount of PPi released during the course of an enzymatic reaction can be reliably determined. However, the reaction was sufficiently linear at concentrations below 400 nM, that one can interpret the fluorescence time course as a linear function of PPi concentration. Experiments shown here and computer modeling show that at concentration of 1 μM PPase and 1 μM PBP, the assay can accurately measure rates of PPi release up to 25 s−1. Under these circumstances, one expects the time dependence to follow three exponentials with rates of 25, 70 and 65 s−1 according to Scheme I. The two faster reactions blend together to form the “lag” phase with a rate of approximately 35 s−1, computed by the approximation: knet = 1/(1/k2 [PPase] + 1/k3 [PBP]). Due to the high signal:noise ratio provided by the fluorescently labeled PBP, the rate of PPi release at 25 s−1 can be readily resolved from the lag phase at 35 s−1. We have successfully measured the rate of release of PPi from the active site of the human mitochondrial DNA polymerase γ following a single nucleotide incorporation reaction that occurred at a rate of approximately 20 s−1 (12). Scheme I serves as a practical guide for understanding the kinetics of the couple assay, but it overlooks the rate-limiting hydrolysis of PPi and release of Pi by PPase and the isomerization of the PBP after Pi binding. The maximum rate of PPi release that can be measured with the PPase/PBP system is limited by the maximum rate of turnover of PPase (kcat = 270 s−1) (13) and by the rate of isomerization of the PBP to the enhanced fluorescence state after Pi binding (317 s−1) (4). We performed more comprehensive modeling by computer simulation including the rates of release of Pi from PPase and the two-step Pi binding to the PBP. Computer simulation was performed using KinTek Global Kinetic Explorer software (KinTek Corp, Austin, Tx). This analysis predicted that an assay with 10 μM PPase and 10 μM PBP would be able to resolve rates of PPi release of at least 100 s−1.