Living cells and organisms could not function without enzyme controlled reactions. The more we understand about how enzymes function and the reactions they control, the better we can use the machinery of nature to benefit human endeavours.
How do you measure the rate of enzyme controlled reactions?
Enzymes operate throughout biological organisms, both intracellularly and extracellularly. You will be aware that enzymes are biological catalysts, meaning they increase the rate of chemical reactions without undergoing any permanent change. Enzymes are made from long chains of amino acids, folded precisely into a three dimensional shape (or tertiary structure) with an active site that allows it to operate as a catalyst. Any changes to this three dimensional structure can change the shape of the active site and cause the enzyme to become denatured. This structure is represented in the lock and key and induced-fit models of enzyme action, with the induced-fit model including the changes that can occur in enzyme shape to allow catalysis.
Given the range of enzyme controlled reactions, there is no single best method for measuring reaction rates as the products of reactions vary greatly. For example, catalase is a common intracellular enzyme that speeds the decomposition of hydrogen peroxide (a byproduct of metabolism) into water and oxygen. In this reaction the produced oxygen gas can be collected and used as a way of measuring the reaction rate. Alternatively, the extracellular enzyme tripsin breaks down casein in milk, changing its colour from white to clear. The reaction rate can therefore be measured with a colorimeter, which will indicate the absorbance of light through the product. The spectrophotometer shown below is similar to a colorimeter, although it measures the transmission, rather than the absorbtion of light.
As the dependent variable (the variable being tested) is the rate of reaction, we need to ensure that the measurements that we are taking are plotted against time. The independent variable (the variable we are manipulating, for example, enzyme concentration) could be represented by plotting multiple lines on the same graph.
What kinds of enzymes do researchers investigate?
How do you avoid errors?
Errors can happen in even the best experiments, but attention to detail and good experimental design can help to minimise both random and systematic errors.
Systematic errors arise from either imperfections in the equipment being used, or by improper technique in the laboratory. An example of a systematic error would be if you were using a cuvette that was stained or scratched, so less light pass would through your sample and all readings using that cuvette would be affected. Similarly it is vital to properly clean and dry cuvettes, fill them using a pipette, handle them only using gloves, and if possible, store them in a cuvette rack.
Random errors are most likely to occur because of the limitations of the equipment that you are using. For example, if your balance is only accurate to a value of 0.1 grams but you need to measure out 250 milligrams of a substance. However, selecting the correct tools for the correct job can help minimise random errors. For example, an adjustable pipette will be much better at measuring out a few millilitres of a solution when performing a serial dilution than using a 50 mL beaker. If random errors are unavoidable due to equipment limitations, then the best way to minimise them is to repeat the experiment as many times as possible to average out the error.
In the Laboratory Confessions podcast researchers talk about their laboratory experiences in the context of A Level practical assessments. In this episode we look at the use of appropriate apparatus to record quantitative measurements and the use of qualitative reagents to identify biological molecules.
What can our measurements tell us?
We can plot our results to help us easily identify the factors that can change enzyme activity. There is is a clear link here between the practical and theoretical elements of biology as the impact of concentration (of enzyme and substrate), inhibition, temperature and pH all have characteristic effects on the rate of reaction plot.
By plotting the amount of product against time, you should create a curve that looks a little bit like the one pictured. This plot is useful as it allows you to calculate the initial rate of reaction. The initial rate of reaction is the gradient of the straight line portion of the plot, shown by the dotted red line. The initial rate of reaction is when concentrations of enzyme and substrate are known, so this allows fair comparison if you then change initial concentrations of enzymes or substrate.
Once you have multiple reaction rates at different substrate or enzyme concentrations, it is then possible to take this one step further and plot reaction rate against substrate concentration, enzyme concentration, temperature or pH. Plotting reaction rate against substrate concentration typically gives a curve that is similar in shape to the product/time plot. It is, however, a different curve and can tell you different things. Most importantly the Maximal Velocity (Vmax), which is when the enzyme is saturated with substrate and the rate of reaction is highest, and the Michaelis-Mensten constant (Km), which is a measure of the enzyme's efficiency. Note that it is possible in some reactions for the reaction rate to drop once Vmax has been reached, as excess substrate can act as an inhibitor. A plot of reaction rate against enzyme concentration will usually result in a straight line, as typically the volumes of enzyme used are much lower than the volume of substrate; in other words it is similar to the straight line portion of the reaction rate/substrate plot. Eventually this plot will level off in a similar way to the reaction rate/substrate plot, although this is unlikely to be observed in classroom experiments! Reaction rate/pH plots should produce a classic bell curve, with the optimum pH at the peak of the curve, and reaction rate/temperature plots should show an increasing rate of reaction with temperature until an optimum is reached (often between 45 and 55 degrees Celsius), after that the reaction rate drops off quickly as the enzymes become denatured.
Different enzyme inhibitors will also change reaction rate/substrate curves in different ways. A competitive inhibitor (for example, cyanide) competes with the substrate for the active site of the enzyme, reducing the rate of reaction at lower substrate concentrations. Given a high enough concentration of substrate the inhibitor can be overcome, so the same Vmax as the reaction without inhibition can be reached although the Km will be changed. Noncompetitive inhibitors (such as penicillin) do not use the active site of the enzyme, perhaps binding in another place and changing the conformational shape (an allosteric inhibitor). Increasing substrate concentration should still increase the reaction rate, but because enzymes can be inhibited regardless of how saturated their active sites are, both the Vmax and the Km will be changed.
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