According to McClure (6), when two auxiliary enzymes are used in a coupled assay, the lag time necessary to reach steady-state conditions approaches a minimum when the first-order rate constant (Vmax/Km) of the secondary auxiliary enzyme is about 4-fold the corresponding constant of the primary auxiliary enzyme. This means, taking into account the constants reported here (Vmax/Km equal to 6.4 × 10−4 and 4.2 × 10−3 for XOD and PNPase at 25 mU/mL, respectively), that a 25-fold excess of XOD over PNPase should be used to minimize the lag time. Alternatively, to save auxiliary enzymes, McClure proposed the use of amounts giving equal first-order rate constants (6). In our case, this translates into using XOD 6–7 times in excess over PNPase. Similar considerations arise when the theory of Storer and Cornish-Bowden (7) is applied. Indeed, an excess of about 10-fold of XOD over PNPase should be used to attain maximal velocities for both enzyme-catalyzed reactions in approximately equal times. Therefore, we decided to test PPase activity in the presence of 50 and 500 mU/mL of PNPase and XOD, respectively. As Figure 1C shows, a linear dependence of pyrophosphatase activity as a function of PPase concentration was observed in the range 2–20 mU/mL of PPase. According to this observation, we propose that, to detect a pyrophosphate-liberating enzyme (such as a DNA polymerase), 2–10, 50, and 500 mU/mL of PPase, PNPase, and XOD can be conveniently used, respectively.
As a model system to test our enzyme coupled assay, we selected the Klenow fragment of E. coli DNA polymerase I. According to the reaction velocities observed with different concentrations of coupling enzymes (see above) we decided to use 5, 50, and 500 mU of PPase, PNPase, and XOD, respectively. Furthermore, in order to perform a set of appropriate controls, poly-dA:oligo-dT was chosen as substrate. As a first test, we incubated Klenow enzyme with 2.5 μM oligo-dT, in the absence of both poly-dA and dTTP. No appreciable activity was detected under these conditions (Figure 2A). Moreover, when 2.5 μM poly-dA was added to the reaction mixture (in the presence of 2.5 μM oligo-dT), no significant activity was detected in the absence of dTTP (Figure 2B). On the contrary, when 2.5 μM poly-dA:oligo-dT and 100 μM dTTP were present, a linear increase in absorbance at 293 nm was readily observed (Figure 2C). Under the same conditions, the polymerase activity detected in the presence of 1.25 μM polydA:oligo-dT was slightly lower when compared with the activity observed with 2.5 μM DNA (Figure 2C), indicating that the kinetics is zero-order. To further inspect the kinetics of DNA elongation catalyzed by Klenow, we assayed polymerase activity as a function of poly-dA:oligo-dT concentration (at 100 μM dTTP). Under these conditions, Km and Vmax were calculated equal to 12.4 nM and 6.1 nM/s, respectively (Supplementary Figure S3). Our estimation of Km is about 2-fold higher than previously published values (KD, 5–6 nM), referring to synthetic oligonucleotides (19-21). It is however worthy to note that we used a substrate different from previously used DNAs (19-21). Unfortunately, we were unable to determine the concentration of Klenow enzyme used in our assays. The protein concentration of the dialyzed stock enzyme solution was indeed below the sensitivity limit of the micro-Bradford method (1 μg/mL). Taking into account the volume of stock enzyme solution used per assay, we estimate that the final concentration of Klenow polymerase was below 2.4 nM in each assay. This, in turn, means that the magnitude of kcat is ≥ 2.5 s−1.
To compare our enzyme coupled assay with a reference method, we used the conventional Malachite green procedure to detect phosphates. In particular, the activity of the Klenow enzyme and of a partially-purified E. coli DNA Pol-III (13) were tested using both the coupled enzyme assay and the Malachite green method. Moreover, we wanted to evaluate if the two methods, when applied to Klenow enzyme, yielded activity values compatible with those indicated by the manufacturer (New England Biolabs). To this aim, the polymerase activity was assayed under the conditions indicated by New England Biolabs, i.e., at 37°C, using activated calf thymus DNA as substrate, and incubating the reaction mixture for 30 min. When 0.1 units of Klenow were used, we detected the release of 0.54 ± 0.07 nmol of pyrophosphate in 30 min with the Malachite green assay. When the enzyme coupled assay was performed under the same conditions, 0.1 units of Klenow enzyme produced 0.50 ± 0.01 nmol of pyrophosphate in 30 min. This value indicates that the coupled assay reliably estimates DNA polymerase activity. In addition, the values of activity obtained by the conventional Malachite green method and by the coupled assay are in reasonable agreement with the activity indicated by the manufacturer, i.e., 10 nmol of pyrophosphate released by 1 unit of Klenow in 30 min at 37°C. It should be noted that our enzyme coupled assay can be performed using electron acceptors other than oxygen, e.g., 2-(4-iodophenyl)-3-(4-nitrophenyl)-5-phenyl-2H-tetrazolium chloride (INT). In this case the assay would rely on the detection of INT-formazan, which can be conveniently monitored at 458 (22) or 460 nm (15). Therefore, we determined the activity of Klenow enzyme using our coupled assay in the presence of activated calf thymus DNA as substrate and 0.5 mM INT as electron acceptor. Under these conditions, 0.1 units of enzyme yielded 0.29 ± 0.01 nmol of pyrophosphate in 30 min (Supplementary Figure S4). This value is significantly lower than those obtained with the Malachite Green method and with our coupled assay performed in the presence of oxygen as the terminal electron acceptor (see above). We propose that this discrepancy is linked to the ε of INT-formazan (we used 12.5 × 103 M−1cm−1), which has been difficult to determine (15).