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Phosphate detection

Methods for measuring inorganic phosphate using dyes (especially Malachite Green) are well known, however common problems associated with these reagents include precipitation of the dye and inorganic phosphate complexes, and acidic reagents which lead to non-enzymatic hydrolysis of many phosphorylated substrates.

The Innova Biosciences range of phosphate detection products contains PiColorLock phosphate detection reagent, which changes colour in the presence of inorganic phosphate (Pi) and can therefore measure any enzyme that generates Pi.

PiColorlock has special additives that enhance the stability of dye complexes and ensure low backgrounds with acid-labile substrates, overcoming these common problems.

The reagent itself, along with its Accelerator and Stabiliser are available as a product, for you to utilise within your assay, however we also supply this reagent within our ATPase and GTPase assay kits. These contain all the necessary products for you to measure enzyme activity with the added benefits of the PiColorLock reagent. These also come with utra high quality ATP and GTP, which ensures the lowest possible assay background. 

We also have our PiBind Resin, which has very high affinity for phosphate, meaning it can be used to efficiently clean-up phosphate containing samples, to eliminate contamination of samples during phosphate detection experiments.

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  • Are there any other ways of eluting binding proteins from the ATP Agarose than ATP?

    Answer: Yes. While ATP is the obvious choice, ligands that are structurally related to ATP may be used (e.g. NADH, AMP, adenosine) to elute a specific subset of the ATP-binding proteins. Drugs that are known to bind to ATP-binding proteins might also be used. If preservation of biological activity is not required, aliquots of resin may be boiled with SDS sample buffer prior to gel electrophoresis.

  • Why am I getting high backgrounds when there is no free Pi in my enzyme or substrate?

    Answer: Please check that you added all the correct buffers are indicated in the protocol. The PiColorLcok is designed to reduce non-enzymatic hydrolysis in the assay, which cause high backgrounds, however it is essential that all buffers be added at the correct time.

  • What is the difference between the standard PiColorLock kit, and the ATPase/GTPase assay kit?

    Answer: As a basic summary, the PiColorLock kits contain the PiColorLock reagent itself, along with the necessary side products, and a phosphate standard to produce a standard curve.The ATPase/GTPase kits contain the same components, as well as extra buffers to support the enzyme, the Purified ATP/GTP as appropriate, and the suitable 96 well plates necessary to carry out the assay. Please see the following table for details on which components are contained in which kit.

  • What is the best buffer to use?

    Answer: There is no ‘right’ answer here. In the absence of any information on the binding requirements of the protein(s) of interest a good starting point is a buffer containing 20mM Hepes, 100-500mM NaCl (or KCl), 20mM MgCl2 or 1mM DTT, pH 7.5. Alternatively, try experimenting with several buffer conditions using small amounts of resin in 1.5ml tubes. After the wash step, elute with ATP or other competing ligand and analyse the eluted proteins by SDS-PAGE. The purification method can then be scaled up using the preferred buffer conditions.

  • What controls should I include in my assay?

    Answer: We would recommend including at least a negative control in your assay, to ensure any results you see are definitely caused by enzymatic-hydrolysis. You could simply omit the enzyme and replace with enzyme diluent, however, a slightly more rigorous approach would be to include the enzyme in the control, but change the order of addition so that the Gold mix is added to the substrate before the enzyme. This control should be set up when the other assay wells are being stopped, so that all wells receive the Gold mix at the same time. The advantage of this approach is that by subtracting a single control value, you actually correct for all free Pi, whatever its source. Also, this operation subtracts the blank value too, meaning the resulting value can be used to determine Pi from the blank subtracted standard curve.

  • I have a high background but cannot seem to isolate the source of the problem.

    Answer: Detergents used in glass washers may contain high concentrations of phosphate and this may carry over into solutions prepared in beakers and measuring cylinders. If most of your components appear to be contaminated with Pi, try switching to a phosphate-free detergent or segregate assay glassware from the normal laboratory wash.

  • How much enzyme should I use in my assay?

    Answer: Our free guide ‘Enzyme units explained’ should provide some help here. In a 200µl reaction you should aim to add sufficient enzyme to generate 1-8nmol of Pi (5-40µM). For any new enzyme it will be necessary to determine the extent of Pi production with serial dilutions of the enzyme. Plot the amount of Pi released versus amount of enzyme and select a dilution of enzyme that is in the linear range.

  • How much substrate should I use?

    Answer: As a general rule, the amount of substrate hydrolysed to Pi should not exceed 10-20% in an assay; otherwise the rate of Pi released with time may not be linear. However, the linear range for any given set of conditions can only be determined by experimentation and you will need to set up a time course. To get a large assay window with only a modest % conversion of substrate, the initial concentration of the phosphorylated substrate in a Gold assay will usually need to be 50-250µM. If Pi production is between 10µM and 40µM the assay signal will normally be between 0.5 and 2.0 absorbance units (see Fig 2).

  • At what temperature should assays be carried out?

    Answer: Enzyme assays are usually carried out in the range 20-37°C, however the preferred temperature will be determined to some extent by the lab equipment that is available. To compare data obtained on different days you need to make sure you standardise the assay in regards to assay temperature. As far as the Gold detection reagent is concerned the temperature of the initial enzyme assay is not important.

  • I have 5% DMSO in my assay. Can I use PiColorLock?

    Answer: Yes, the reagent is designed for drug screening work and other situations that require DMSO, so this will not cause any problems.

  • Do I need to subtract blanks from the standard curve?

    Answer: Not necessarily, but we would recommend it. The blank is the value obtained in the absence of Pi, and should be ~0.1. So, if the absorbance for a sample is read at 1.0, the signal due to Pi released by the GTPase is actually 0.9. Remember, if you subtract blanks from your standard curve, you need to also subtract blanks from your assay data.

  • I have a high background in my assay and I definitely do not have free phosphate in my sample

    Answer: This is almost always due to inadequate mixing of the Stabiliser component of the kit. It is really important that the stabiliser is added 2 minutes after the PiColorLock, and is mixed into solution using a pipette set to at least 50% of the assay volume.

  • I have phosphate in my enzyme. What can I do?

    Answer: This will of course need to be remnoved prior to using the PiColorLock reagent, but this can easily be achieved by using our accesory product PiBind, which quickly and efficiently binds to free phosphate in the sample. Alternatively, you can dialyse or desalt the enzyme into a phosphate-free buffer.

  • I know my sample has activity but all my wells are yellow. Why is this?

    Answer: The most likely explanation is that the Stabiliser has been added with, or immediately following, the Gold mix. Make sure that the Stabiliser is added two minutes after the Gold mix to produce the required green colour.

  • Should I subtract blanks from my assay samples and standard curve?

    Answer: This comes down to personal preference but the main thing to consider is that for any single absorbance reading there are actually two or more components to that reading. This applies to all absorbance assays, and is not specific to this particular assay. An appreciation of the different components is required in order to determine the best way of handling the controls and blanks, and whether to subtract blanks from the assay wells and standard curve before carrying out calculations. For example, if we assume that the substrate is contaminated with free Pi, the single measured absorbance (Y1) for the assay wells is the sum of three separate components (i) the blank value due to the Gold reagent alone, which is ~0.1 (ii) the signal due to contaminating Pi (iii) the signal due to Pi released from the substrate during the assay. The control wells (in this case wells with substrate but without enzyme) give a single absorbance reading (Y2) that is made up of two components, the blank value and the signal due to contaminating Pi in the substrate. Thus subtraction of Y2 from Y1 subtracts the component due to contaminating Pi and also the blank component. The resulting value can therefore be used to calculate the amount of Pi formed using a blank-subtracted standard curve. Whilst in the above example it was not necessary to subtract the measured blank value directly from the assay data (since subtraction of the control Y2 from Y1 achieved the same result) it is generally safer to subtract the blank value (i.e. water plus 0.25 volumes of the reagent mix and 0.1 volume of stabiliser) from the standards, assay wells and any control wells before calculations on the data are performed. In this way, regardless of how many controls need to be subtracted from the assay data you cannot inadvertently subtract the ‘hidden’ blank value more than once. #Tip: you can do a single control that includes all assay components by using a different order of reagent addition. Add all components except the enzyme (but do not add water instead of enzyme) to triplicate wells, followed by the reagent mix (ignore the fact that the enzyme is missing and add the usual 0.25 volumes of reagent mix). Next, add the enzyme. (There are few, if any, enzymes that are active in the acidic medium). Five minutes later add the stabiliser and read the plates normally. This approach allows you to combine the enzyme and substrate in a single control well.