Hey guys
I'm working with PNP-acetate and PNP-butyrate, two model substrates for esterase/lipase reactions which yield a product which absorbs at 405 nm on hydrolysis. All of the assays I've come across suggest using phosphate buffer in the reaction mixture and advice I've obtained on here suggests dissolving PNP-substrates in DMSO.
However, when I run a few tests with the substrate in various moralities of phosphate buffer, from 0.2 M to 0.05 M I see hydrolysis of the substrate and a appreciable rate correlating positively with the molarity of the buffer. I know I can subtract this rate from my kinetics rate, but it's very large and I'd prefer to find some way to identify what's making it unstable and/or increase the stability.
I'm using sodium phosphate made from a combination of disodium phosphate and monosodium phosphate. I've tested both components individually and I see no hydrolysis when using the monosodium phosphate (~pH 4.7_ on its own and a rate when using the disodium phosphate (~pH 7. ). When I mix both components together to make the phosphate buffer and test for hydrolysis, I see a rate at pH 7, 8 and 9. As before an increase in the molarity of the buffer results in an increased rate of hydrolysis. I'm doing several biophysical assays with this protein in phosphate buffer and would love to stick with that if possible.
Things I've tried:
- Making new buffer
- trying different pH
attempting to subtract the rate of PNP in buffer from that of the enzyme reaction, but rate far too large.
Any help you guys can give would be much appreciated.
ancora imparo
Harry
A common mistake in measuring enzyme activity using these substrates is to perform the assays at neutral or acidic pH without considering that only part of the chromophoric product is ionized.
These substrates are readily hydrolyzed in contact with water, so it is normal to see spontaneous hydrolysis. However, lower autohydrolysis is observed at acid pH due to the ionization state of p-nitrophenolate. Which is colorless at pH below its pka value.
I recommend you J Lipid Res. 2015 May;56(5):1057-67. doi: 10.1194/jlr.D052837. Epub 2015 Mar 7.
It sounds like pH is a factor in the stability, with greater stability at lower pH. Use the lowest pH that is compatible with activity of the enzymes. Also, since you found that stability was worse when the buffer was more concentrated ("molarity," not "morality," by the way), keep the buffer concentration low, such as 25-50 mM. It might also be worth testing out some buffers other than phosphate, such as HEPES and MOPS.
A common mistake in measuring enzyme activity using these substrates is to perform the assays at neutral or acidic pH without considering that only part of the chromophoric product is ionized.
These substrates are readily hydrolyzed in contact with water, so it is normal to see spontaneous hydrolysis. However, lower autohydrolysis is observed at acid pH due to the ionization state of p-nitrophenolate. Which is colorless at pH below its pka value.
I recommend you J Lipid Res. 2015 May;56(5):1057-67. doi: 10.1194/jlr.D052837. Epub 2015 Mar 7.
Did you try isopropyl alcohol to dissolve the substrates? It worked well for me to dissolve paranitrophenol palmitate.
Acids and bases act as catalyst for some substrates, including pNP-esters. Their stability is therefore pH-dependent. The solution is to have reagent blanks in all assays, which include everything but the enzyme under test. To make it quite clear: if you work at several pH-values, you need a separate blank for each! Otherwise, you can come to very wrong conclusions!
In addition, make sure that the solution after stopping the reaction is alkaline enough to convert pNP into the ionic form, as only that, but not the non-ionised form, is coloured. So you need enough base in the stopping solution to have an alkaline pH regardless on how acidic the reaction mix was.
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