Enzymatic Reaction: A Common Strategy for Initiating an Enzymatic Reaction

This article throws light upon a common strategy for initiating an enzymatic reaction and learn about how it also controls the running of enzymatic reaction.

Initiating, Mixing, and Stopping Reactions:

In a typical enzyme assay, all but one of the components of the reaction mixture are added to the reaction vessel, and the reaction is started at time zero by adding the missing component, which can either be the enzyme or the substrate.

The choice of the initiating component will depend on the details of the assay format and the stability of the enzyme sample to the conditions of the assay.

In either case, the other components should be mixed well and equilibrated in terms of pH, tempera­ture, and ionic strength.

The reaction should then be initiated by addition of a small volume of a concentrated stock solution of the missing component. A small volume of the initiating compo­nent is used to ensure that its addition does not significantly perturb the conditions (temperature, pH, etc.) of the overall reaction volume. Unless the reaction mixture and initiating solutions are well matched in terms of buffer content, pH, temperature, and other factors, the initiating solution should not be more than about 5-10% of the total volume of the reaction mixture.

Samples should be mixed rapidly after addition of the initiating solution, but vigorous shaking or vortex mixing is denaturing to enzymes and should be avoided. Mixing must, however, be complete; otherwise there will be artifactual deviations from linear initial velocities as mixing contin­ues during the measurements.

One way to rapidly achieve gentle but complete mixing is to add the initiating solution to the side of the reaction vessel as a “hanging drop” above the remainder of the reaction mixture, as illustrated in Fig. 7.4. With small volumes (say, < 50 µL), the surface tension will hold the drop in place above the reaction mixture. At time zero the reaction is initiated by gently inverting the closed vessel two or three times to mix the so­lutions.

Fig. 7.4 illustrates this technique for a re­action taking place in a micro-centrifuge tube. It is also convenient to place the initiating solution in the tube cap, which then can be closed, per­mitting the solutions to be mixed by inversion as illustrated in Fig. 7.4. For optical spectroscopic assays, the reaction can be initiated directly in the spectroscopic cuvette by the same technique, using a piece of Parafilm and one’s thumb to seal the top of the cuvette during the inversions.

Regardless of how the reacting and initiating solutions are mixed, the mixing must be achieved in a short period of time relative to the time interval between measurements of the reaction’s progress. With a little practice one can use the inversion method just described to achieve this mixing in 10 seconds or less.

This is usually fast enough for assays in which measurements are to be made in intervals of 1 minute or longer time. A number of parameters, such as temperature and enzyme concentration, can be adjusted to ensure that the reaction velocity is slow enough to allow mixing of the solutions and making of measurements on a convenient time scale.

In some rare cases, the enzymatic velocity is so rapid that it cannot be measured conveniently in this way. Then one must resort to specialized rapid mixing and detection methods, such as stopped-flow techniques (Roughton and Chance, 1963; Kyte, 1995); these methods are also used to measure pre-steady state enzyme kinetics. For assays in which samples are removed from the reaction vessel at specific times for measure­ment, one can start the timer at the point of mixing and make measurements at known time intervals after the initiation point.

In many spectroscopic assays, however, one measures changes in absorption or fluorescence with time. For most modern spectrometers, the detection is initiated by pressing a button on an instrument panel or depressing a key on a computer keyboard. Thus to start an assay one must mix the solutions, place the cuvette, or optical cell, in the holder of the spectrometer, and start the detection by pressing the appropriate button.

The delay between mixing and actually starting a measurement can be as much as 20 seconds. Thus the time point recorded by the spectrometer as zero will not be the true zero point (i.e., mixing point) of the reaction. Again, with practice one can minimize this delay time, and in most cases the assay can be set up to render this error insignificant.

As we shall see, there should always be two control measurements: one in which all the reaction components except the enzyme are present, and a separate one in which everything but the substrate is present. (In these controls, buffer is added to make up for the volumes that would have been contributed by the enzyme or substrate solutions.) With these two control measurements one can calculate what the absorption or fluorescence should be for the reaction mixture at the true time zero.

If the first spectrometer reading (i.e., the time point recorded as time zero by the spectrometer) is significantly different from this calculated value, it is necessary to correct the time points recorded by the spectrometer for the time delay between the start of mixing and the initiation of the detec­tion device. A laboratory timer or stopwatch can be used to determine the time gap.

Many non-spectroscopic assays require measurement times that are long in comparison to the rate of the enzymatic reaction being monitored. Suppose, for example, that we wish to measure the amount of product formed every 5 minutes over the course of a 30-minute reaction and assay for product by an HPLC method.

The HPLC measurement itself might take 20-30 minutes to com­plete. If the enzymatic reaction is continuing during the measurement time, the amount of product produced during specific time intervals cannot be determined accurately. In such cases it is necessary to quench or stop the reaction at a specific time, to prevent further enzymatic production of product or utilization of substrate.

Methods for stopping enzymatic reactions usually involve denaturation of the enzyme by some means, or rapid freezing of the reaction solution. Examples of quenching methods include immer­sion in a dry ice—ethanol slurry to rapidly freeze the solution, and denaturation of the enzyme by addition of strong acid or base, addition of electrophoretic sample buffer, or immersion in a boiling water bath.

In addition to these methods, reagents can be added that interfere in a specific way with a particular enzyme. For example, the activity of many metallo-enzymes can be quenched by adding an excess of a metal chelating agent, such as ethylenediaminetetraacetic acid (EDTA). Three points must be considered in choosing a quenching method for an enzymatic reaction.

First, the technique used to quench the reaction must not interfere with the subsequent detection of product or substrate. Second, it must be established experimentally that the quenching technique chosen does indeed completely stop the reaction. Finally, the volume change that occurs upon addition of the quenching reagent to the reaction mixture must be accounted for. Similarly, meas­urement of product or substrate concentration must be corrected to compensate for the dilution effects of quencher addition.

The Importance of Running Controls:

Regardless of the detection method used to follow an enzymatic reaction, it is always critical to perform control measurements in which enzyme and substrate are separately left out of the reac­tion mixture. These control experiments permit the analyst to correct the experimental data for any time-dependent changes in signal that might occur independent of the action of the enzyme under study, and to correct for any static signal due to components in the reaction mixture.

To illustrate these points, let us follow a hypothetical enzymatic reaction by tracking light absorption decrease at some wavelength, as substrate is converted to product. Let us say that there is some low rate of spontaneous product formation in the absence of the enzyme, and that the enzyme itself imparts a small, but measurable absorption at the analytical wavelength.

We might set up an experiment in which all the reaction components are placed in a cuvette, and the reaction is initiated by the addition of a small volume of enzyme stock solution. For this illustration, let us say that the reaction mixture is prepared by addition of the volumes of stock solutions listed in Table 7.2. The strategy for preparing the reaction mixtures in Table 7.2 is typical of what one might use in a real experimental situation.

Fig. 7.5 A illustrates the time courses we might see for the hypothetical solutions from Table 7.2. For our experimental run, the true absorption readings are displaced by about 0.1 units, as a result of the absorption of the enzyme itself (“No substrate” trace in Fig. 7.5A). To correct for this, we subtract this constant value from all our experimental data points.

If now we were to determine the slope of our corrected experimental trace, however, we would be overestimating the velocity of our reaction because such a slope would reflect both the catalytic conversion of substrate to prod­uct and the spontaneous absorption change seen in our “No enzyme” control trace. To correct for this, we subtract these control data points from the experimental points at each measurement time to yield the difference plot in Fig. 7.5B.

Measuring the slope of this difference plot yields the true reaction velocity.

As illustrated in Fig. 7.5A, the correction for the spontaneous absorption change may appear at first glance to be trivial. However, the velocity measured for the uncorrected data differs from the corrected velocity by more than 10% in this example. In some cases the background signal change is even more substantial. Hence, the types of control measurement discussed here are essential for obtaining meaningful velocity measurements for the catalyzed reaction under study.