I try to read enzymatic activity in tissue homogenates using fluorescence emission in a microplate reader. I hypothesize that a specific treatment can reduce amount of enzyme in tissue and/or inhibit the enzyme itself. Both hypotheses would be interesting to monitor in one read-out.
For characterizing the response curve of my assay, I made a serial dilution of tissue homogenates. Interestingly, I noticed that the enzymatic activity read-out is proportional to the amount of protein I put in my test. This is logical, if I put more material and if I have enough substrate, I got more signal.
I also would like to determine the amount of total protein to normalize activity of the enzyme to the quantity of protein load in the test. Therefore, I used several dyes which all provided me a non-linear response between dilution of the sample and fluorescence read-out (a log response, like a saturation effect). Seemingly these dye do not affect the read-out of enzyme activity.
As we need a certain amount of protein to detect the fluorescence, but this amount saturate the read-out of protein loading I thought about 2 solutions:
1) Do a serial dilution of my sample to determine the protein concentration in my sample on the plate and evaluate the enzyme activity (more accurate because know exactly the amount of protein per test, but presence of an extra compound in the test).
2) First measure protein concentration in a first test, use the result to load the exact same amount of protein per well (cheaper because less replicate to load, but no control on the quality of the initial protein loading).
What do you usually use?
1. way: After measuring your enzymatic activity (or on parallel plate) - assuming you have the same total volume in each sample - fill in for the Bradford method, along with some standard curve measurements. You should expect some dilution effect taking place.
2) Measure the A280 of your plate; you can use BSA or a protein of size similar to yours protein of interest. as standard curve in identical total volume (this assay will give you various A280/ug prot values, depending on the protein size used in the standard curve).
Alternatively, A205/31 as good approximation for protein conc. 1-100ug/ml.
3) see http://lifeserv.bgu.ac.il/wb/zarivach/media/Documentation/Lab%20Faqs-Roche-Diagnostic%20bench%20mate.pdf page 99 and later
Hi Nicolas, fluorescence enzyme assays are fine if you are monitoring appearance of fluorophores with time.
Protein fluorescence is mostly qualitative, not quantitative.
Fluorescent enzyme assays will give you enzyme activity/time. If you have all of the parameters equal for your fluorescence assays then you can compare the results internally.
However, you need a quantitative assay for your protein mixture. Fluorescence of a protein mixture using dyes is not quantitative.
You will have to use a different method to quantify your proteins. Marcin suggested A280 measurements. I suggest you use the BCA kit from Pierce.
Interesting,
Doing the protein measurement after reading activity should work. I have to take care if the reagent needed for the enzyme activity are not cross-reacting with the colorimetric essay.
Are absorption methods also working in complex mixture of protein like protein lysate?
And dye to stain protein, it is a bit a shortcut I have to admit. The idea is more to use hoechst to quantify the amount of DNA following this idea: http://www.ncbi.nlm.nih.gov/pubmed/1706565. I agree that this method does not provide information directly on the amount of protein but allow to quantify amount of DNA in tissue homogenate.
Hi Nicolas, you should not do protein assays and enzyme assays in the same wells.
That is a bad idea. Protein assays like BCA are denaturing and they might react with your substrate.
It would be nice to have a fluorescent protein assay using a dye that reacts equally with all proteins in a complex mixture to give a quantitative value.
Alas, unlike DNA, which has a defined structure that can reliably bind interchelators like Hoechst, protein mixtures contain structures that might or might not bind fluorescent dyes equally.
Thank you Steingrimur for your input. You are right, good lab practice should allow to make from the same alliquot of protein a reproducible protein-concentration measurement and an enzymatic-activity measurement.
It would have been nice to make routine to have in one well both read out to exclude any technical problem.
Therefore I do not give up and I am still looking for ideas :)
Hi Nicolas, I admire your tenacity.
I am old.
Based on my experience, I can state that some things are impossible.
But I truly hope that things that I thought were impossible will soon become routine procedures.
That is very nice from you, thank you!
All the best in your research
Why don't you measure the A280 or A205:31 before adding your substrate? I assume such multiplate readout will not take longer than 30sec, after which you can multi-pipett (I assume again) the same amounts of substrate per well for activity testing.
Alternatively, you could pipet out two plates in parallel: one for the enzymatic assay, one (in buffer/water) strictly for protein measurements.
Hello Nicolas,
From what I understand you have tissue homogenates coming from different treatments and you want to understand how the treatments can impact enzyme activity (if this isn't right, disregard the rest of this response).
I am not sure what organism you are using (human, rodent, other), but for the sake of the response I will assume human.
Enzyme activity (enzyme kinetics) assay require certain restrictions. If you are running human samples, much of the information on different probe substrates have already been determined. Specifically protein concentration, buffer specifications and probe substrate kinetics (Vmax, Km).
Typically when we perform enzyme kinetic assays we perform them using the “linear conditions.” This means a concentration of protein, an incubation time and a probe substrate concentration (Km or below).
From what you described, it sounds like you are putting a particular volume of your tissue homogenates into the reaction and measuring the change over time. If this is the case, there is no way to report your results so that you can compare the specific enzyme activity. Since you want to quantify the activity of a particular enzyme (protein), you need to standardize your reactions to have the same amount of protein. Then, when you report your rates of reaction (usually in moles of substrate converted by time by amount of protein e.g. moles/min/mg protein), the difference between the samples will be the difference in the activity of the enzyme under the same condition (normalized).
To do this, the common practice is to quantify your protein samples after extraction (tissue homogenization). There are many assays for this: Lowry, Bradford, BCA. Once you know the protein content of your samples you can then do dilutions to get the protein concentration you want for your assay. [Note: Two ways to do this. Quantify, then standardize stock to concentration desired prior to storage, or store and then dilute prior to assay]. You may also want to consider the sample you use. It sounded like you are using whole tissue homogenates. Depending on what you want to study, you can further separate out the cellular compartments (S9 fraction, cytosol, microsomes). This can allow you to get more sensitive results and is a lot cleaner to work with.
Now, depending on the enzyme assay, it is always a good idea to determine the linear conditions of the assay in your hands/lab. You started to do this by “For characterizing the response curve of my assay, I made a serial dilution of tissue homogenates. Interestingly, I noticed that the enzymatic activity read-out is proportional to the amount of protein I put in my test. This is logical, if I put more material and if I have enough substrate, I got more signal.” When you change the concentration of the protein you will get slower/faster activity. You need to ensure you are in the linear range as there will be cutoffs to concentrations where turnover of the substrate and under conditions that you cannot distinguish difference from.
As Steingrimur stated in his post, you do not want to do multiple reactions in the same well. If you don’t know how the chemicals react with each other, you may get incorrect results. The solution is as you put it at the end of your post, #2. “2) First measure protein concentration in a first test, use the result to load the exact same amount of protein per well (cheaper because less replicate to load, but no control on the quality of the initial protein loading).” This is the standard in the field of enzyme kinetics and good scientific practice.
I hope this helps. If you need any clarification on something particular let me know. Best of luck with your reseach!
Thank you Luc for the detailed answer, one more point for the 2 step analysis!
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