Verify the results using Beer's law:. You will use known amounts of the reaction product p-nitrophenol in citrate buffer to obtain a standard curve. As p-nitrophenol is a skin irritant, you should wear gloves throughout these assays. Calculate and record the concentration of the substock. Also calculate the amount of p-nitrophenol in each vial and complete the 2nd column in Table 3.
Pipette the volumes given in Table 3 , and measure and record the absorbance at nm in the spectrophotometer. Use a pair of plastic cuvettes, and start with the most dilute sample. Now you have to determine enzyme activities in all the original aliquots and for the fractions obtained from gel filtration. Using appropriate pipetman, pipette each sample and buffer in to clean test tubes see Table 4. Add the substrate and start timing. Vortex and leave at room temperature for 30 min.
Measure absorbance at nm. The interactions of proteins in a sample with the stationary phase result in separation based on polarity, and is sensitive enough to uniquely identify proteins that differ by only a few amino acids. HPLC also allows for specific identification and accurate quantitation of protein impurities.
Mass spectrometry is a powerful analytical technique used for monitoring protein purity which can detect post-translational modifications with accuracy and precision. One drawback of the denaturing that occurs in the mass spectrometer is that this technique cannot be used to determine whether the proteins in a sample are intact.
HIC separates proteins by the distinctions in their surface hydrophobicity achieved by utilizing a reversible interaction between the proteins and the hydrophobic surface of a HIC medium. HIC also has the benefit of avoiding the use of organic solvents, which can often denature protein samples. The interaction of hydrophobic proteins and a HIC medium is substantially impacted by specific salts in the running buffer.
A high salt concentration strengthens the interaction while dropping the salt concentration causes separation and elution based on hydrophobicity [4]. If the methods discussed above have displayed protein purity, its dispersion within the sample must be checked.
Dynamic light scattering uses polarized laser light to measure the level of diffraction in a sample with proteins; this is the scattering that occurs as an effect of the hydrodynamic radius of the particles in solution as the sample travels through the instrument.
Fluidics testing is a simple option to measure protein purity based on size and concentration. MDS uses microfluidic chips to run the protein sample through a channel with an auxiliary fluid in a steady state laminar flow without mixing. To move from one stream to another the proteins must diffuse and the rate at which this occurs is proportional to their size, measured as the hydrodynamic radius R h.
The ratio of diffused and undiffused species is used to calculate this value. In microplate absorbance assays, the path length is equal to the depth of the liquid, and it is possible to miniaturise without loss of signal by reducing the diameter of the wells and maintaining a constant depth of liquid. For example, 96 well plates can easily accommodate ul of sample, but with a 50ul assay volume it would be better to use a well plate to increase the path length over that seen with 50ul of sample in a well plate.
Most assays are carried out for a fixed period time end point assays and the reaction is halted by the addition of a stop reagent e. However, in continuous assays the appearance of product less commonly the consumption of substrate is recorded continuously e.
The same basic rules apply; a plot of signal versus time for a fixed amount of enzyme should be linear and the rate should double if the amount of enzyme is doubled. From a practical point of view one key consideration is the amount of product that must be generated in order to give a measurable assay signal.
Another consideration is the Km for the substrate. While adding more substrate generally means that you will see higher activity values, the relationship is not linear and the cost of the substrate may need to be considered.
At this point it is probably useful to introduce the classic Michaelis-Menten equation:. The derivation of this equation and its underlying assumptions may be found in any text book on enzyme kinetics. Although it is normal to measure rather than to calculate the rate of enzyme reactions, it is useful to understand the underlying principles for the purpose of assay design. The Michaelis-Menten equation is useful in that it helps you to select a suitable concentration of substrate if the Km is known.
Clearly this is higher, but not 10 times higher. At very high concentrations of substrate the Km in equation 1 numerically becomes insignificant and the measured rate then equals Vmax. Many assays are run with substrate concentration at or around the Km value but if the Km is very high it may not be possible to use such a high concentration of substrate e.
In some situations it may be even advisable to use a relatively low concentration of substrate. For example, in pharmaceutical drug discovery, the use of a very high substrate concentrations would make it more difficult to identify competitive enzyme inhibitors competitive inhibitors bind to the same site as the substrate. A balance of the various factors has to be struck so that the assay has a measurable signal, can be operated in the linear range and can meet any other assay objectives e.
A standard curve is always required if you wish to calculate enzyme activity. It is not essential if you are only interested in relative activity values. The standard curve is constructed by measuring the assay signal with standard solutions of the reaction product over a suitable range of concentrations.
Ideally you should run a standard curve for every experiment, but if the standard curve is highly reproducible it may be acceptable to run it periodically. A typical standard curve is shown in Figure 2 below. The phosphate is detected by means of a dye binding reagent which changes colour in the presence of phosphate. However, the specifics of this test are not important here; whatever the nature of the product or the method of detection that is used, a standard curve showing the dependence of signal on the amount of product in the assay must be constructed.
Most graphical software packages will automatically calculate values of x from measured values of y. The amount of activity can then be calculated see later. There is no single correct answer here, and the possibility of using either approach can often cause confusion when activity values are calculated.
However, laboratory reagents are usually prepared at known concentrations , and it is often easier to plot these values on the x axis.
However, when you calculate the activity or specific activity of your enzyme you must remember to convert the concentration on the x-axis into the number of nmoles of product formed, which means you must take account of the assay volume. The reason is easy to understand: 50ul and ul volumes of a standard solution will be at the same concentration, but the larger volume will contain twice the amount of product. Generally, it is best to pick one approach i.
Proper controls are vitally important for quantitative work, as is the correct use of the control data in calculations.
Controls tell you indirectly how much of the assay signal is due to the action of the enzyme and how much arises for other reasons. Other assay components may also give rise to a small signal depending on the nature of the components and the type of assay. The purpose of controls is to allow you to remove by subtraction any elements of the total signal that are not related to the action of the enzyme.
If the assay is well-designed and the assay reagents are of good quality the treatment of controls is usually quite simple. If we return to the colorimetric assay for detecting inorganic phosphate we can highlight some potential problems that are readily detected with proper controls. The phosphate detection reagent gives a low reading of about 0.
For any enzyme assay, a key control A is to leave out the enzyme and replace with buffer. This will give you the background signal for all the other assay components as a group, including the substrate which may contain a small amount of product.
This control does not tell you the background signal for the enzyme, but clearly you cannot add the enzyme to the substrate as a control! In most assays the enzyme gives no signal because it is diluted down significantly before use in assay. This can easily be checked, as in B above. The data for sample A above suggest that the mixture is contaminated with inorganic phosphate. The individual components can then be checked to identify the source of the problem.
This situation is quite common for assays of ATPases and other phosphate- generating enzymes, as the substrates are often unstable and are partially hydrolysed to give some inorganic phosphate. It can be awkward to correct raw data if several components give false signals; elimination of the problem at source is generally far better than any mathematical treatment.
Remember the assay plate plus detection reagent give a background signal of around 0. The same small signal must also be hidden within the value for the enzyme control. Thus in the correction applied above we have subtracted this hidden blank twice.
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