We are using zymography to look at proteases in skeletal muscle samples. Unfortunately, while the positive controls are working, the samples with muscle extract are not. There are a couple of issues that I could use some help with.
We are homogenizing the tissue in 50 mM Tric-HCl pH 7.5/ 0.25% TX-100, spinning the samples, quantifying protein and then freezing the samples until we are ready to run the gels (both casein and gelatin gels). The positive controls for the gels work, but we are not seeing any signal from our tissue samples. Some of the questions/issues that we could help with include:
1) We are only solubilizing a small percentage of the muscle protein with this extraction protocol and don't see typical protein profiles on Comassie stained gels (egs. MHC, actin, etc). Is there an extraction buffer that will do a better job of solubilizing the protein?
2) We are not including any protease inhibitors in the extraction buffer because it might inhibit enzyme activity on the zymograms. However, these muscles have a lot of proteases and maybe they are degrading some of the enzymes we want to measure. Any recommendations?
3) How much protein do you typically load in a well?
Any help you could offer would be greatly appreciated.
Thanks.
One guess might be that removal of SDS after electrophoresis was incomplete and therefore renaturation of the MMPs in the sample was sub-optimal (although a strong signal from the positive control could still be visible).
Also I am always including protease inhibitors (10 μg/mL aprotinin, 2 μg/mL leupeptin, and 4 mM benzamidine) and never experienced problems - you should get rid of them during electrophoresis anyway I think.
Hope that helps, Christian
Have you read this? Might be useful.
http://books.google.com/books?id=511Ua8BOU6kC&pg=PA267&lpg=PA267&dq=muscle+protease+zymogram&source=bl&ots=emvZxbOdcu&sig=phF9KwfFmx7Ujcqg_NMpmf86s0w&hl=en&sa=X&ei=lPJPVJLHKom5ogSQtYKYAw&ved=0CCQQ6AEwAg#v=onepage&q=muscle%20protease%20zymogram&f=false
How are you homogenizing the muscle? Because I think that it is a hard tissue to deal with, so maybe you need to mince it first (using a polytron) and use a potter-elvehjem (I don't know if it will work, maybe just using a polytron is enough). Also, remind to do the entire procedure at 4°C, otherwise you will have protein degradation and this might be one of your problems. Finally, which kind of proteaseas are you looking for? Because if you are targeting MMPs, I suggest to put in the extraction buffer also 5 mM CaCl2 and at least 1 uM (micromolar) of ZnSO4.
Thanks for these helpful suggestions. I keep the tissue frozen on dry ice until I homogenize it with a polytron, which does a good job of shearing the tissue. Because of the low salt, I am not solubilizing most of the contractile apparatus, which may be fine if the proteases are still being liberated. I didn't have aprotinin in hand (it's on order) but did include leupeptin and benzamidine in the most recent test, which did allow me to get active enzyme on a gelatin zymogram, so I think your suggestions are putting me on the right track. Once I have the aprotinin I will repeat the extractions and see if that does the trick. Many thanks for all the help.
Hi,
I think the key point is to renaturation of the MMPs after electrophoresis. usually this is done using a non-ionic detergent such as Triton-X100 to replace the SDS in the gel. The renaturation buffer i have used previously is of the following composition (200mM NaCl, 5mM CaCl2, 5uM ZnCl2, 2.5% Triton X 100, 0.02% NaN3 and 50mM Tris-HCL, pH7.5). Hope this helps.
All the best!
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