Hello all,
I work with a tRNA synthetase and I am setting up a classical tRNA aminoacylation reaction with radioactive amino acid. After letting the reaction to run for 1h 30', I precipitated my tRNA with the addition of 50ul 10% TCA, and then I transfer the reaction mixture to a glass-fiber filter for filtration. However, I don't see any trace of enzyme activity. I have already worked with this tRNA synthetase with a different biochemical assay that used to work, so the buffer, pH and salt concentrations are already optimised. Anyone that can give a hand on this?
Are you sure the tRNA is intact? Was the final TCA concentration high enough to precipitate tRNA? Did you allow enough time for the tRNA to precipitate before filtration? How was the enzyme stored? Can you re-test it in the original assay to see if it still works in that assay?
will an N-terminal His tag keep the synthetase enzymattically active? I run the aminoacylation reaction for an hour and then I ethanol precipitate it and resuspend the pellet in acidic buffer. Still I don't get the enzymatic activity. Where could be the problem?
What pH is the acidic buffer? tRNA will precipitate in strongly acidic conditions.
An N-terminal His tag probably won't affect the aaRS activity, but it is a possibility. If it is a cleavable tag, see what happens if you cut it off.
Do you have positive controls to show (1) that the aaRS is catalytically active and (2) that the tRNA is in good condition and is working as a substrate for the aaRS?
The pH of my acidic buffer is 5.5 which I use and is recommended.
As a positive control I only run the substrate i.e. tRNA on the gel to check its purity.
Another test you could do is to see if the aaRS possesses activity in the aminoadenylation reaction (no tRNA, just ATP and amino acid).
Combine a high concentration of the enzyme, such as 1 µM (control: no enzyme), with its cognate amino acid (1 mM, control: no amino acid), ATP (100 µM), and inorganic pyrophosphatase enzyme (0.1 U/ml). Let the reaction go for at least an hour. Inorganic pyrophosphatase converts the pyrophosphate product of the aminoadenylation reaction into 2 phosphates. Then measure the phosphate by the color change of a Malachite Green/molybdate reagent. Both inorganic pyrophosphatase and Malachite Green/molybdate phosphate detection reagents are commercially available. Note: do not use a phosphate buffer for this reaction.
Some tRNA synthetases will only do one catalytic turnover of the aminoadenylation reaction with cognate amino acid. Others will go through several turnovers in an hour. In some cases, you get more product if you use a noncognate amino acid because of pretransfer editing or because the aminoadenylate product is unstable.
The no-amino-acid control is important because the enzyme prep could be contaminated with other ATP hydrolyzing enzymes.
Instead of using the detection reagents, can absorbance based quantification of the assay be done?
There is no change in the absorbance of ATP when it is converted to AA-AMP. I don't think the reaction can be monitored just by absorbance. You could follow it by some form of chromatography to separate ATP from AA-AMP. Thin-layer chromatography should work. If you use a plate with a fluorescent indicator, you would be able to see the dark ATP and AA-AMP bands under UV illumination.
I have attached the article where they propose the assay using spectrophotometry, as far as I understand. I have highlighted the text. The group has used malachite green to stop the reaction and after incubation over a period of time at RT, they have measured the synthetase activity using spectrophotometer, at 620 nm. Please let me know if what I have understood is correct.
The Malachite Green reagent is what I suggested in a previous answer to this thread 10 months ago. I have used this method myself. It works.