Hello!
I was wondering if anyone had a detailed protocol for measuring β-hexosaminidase activity in isolated lysosome fractions.
I will have 0.5ml fractions of lysosome isolate (after magnetic isolation).
Here is what I know so far:
- samples: lysosome aliquots with and without Triton X-100 (1% final triton concentration) + whole cell lysates + cell culture supernatant + controls without sample
- reaction buffer: 100 mM Na Citrat, 0.2% BSA, 1mM 4-MU-β-N-acetylglucosaminide
- incubation: 10 min 37°C
- stop solution: glycine/NaOH pH 10.4
- fluorimeter: excitation 360nm, emission: 450nm
There's quite a bit on the internet about this. Here's one reference:
Article Measuring Mast Cell Mediator Release
Just looking at your conditions, for the most part you look on target. Unless you're adding a pretty small amount of sample to the reaction buffer, you're going to be diluting the 4MU substrate significantly. Since the Km is ~ 1 mM, which is the concentration in your buffer, you're going to be suboptimal in terms of rate and reproducibility. I would try to increase the substrate concentration 10X (if possible). I would also increase my reaction time- the above paper runs theirs for 90 minutes. You could take one sample and do a time course to determine optimal reaction time.
Helena Milionis Do 25 uL of lysosomal eluate + 50 uL of substrate, incubate 30 min at least, then stop with 200 uL stop solution. This usually works for magnetically-purified lysosomes, but you can prolong the incubation time for sure. In some cases (exocytosis assays) I've increased it up to 24 hours.
I was feeling a bit pessimistic about my isolation so I incubated 25ul of lysosome eluate (50ug protein/ml) with 50ul of substrate (as Vojtěch Dostál kindly suggested) for 20 hours which blew up the signal completely out of range for the instrument!! Even after a 1:100 dilution of the measured sample the signal was still too strong. I will definitely do a dilution and time course experiment as Nick Chacos suggested!
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