Please try to obtain the aorta samples fresh from almost a living animal, put them in small aluminum foil pockets  the size of half envelopes and snap freeze them by dipping immediately in liquid nitrogen.  After that, immediately put the foil pockets with the aorta samples inside, into a cooler containing dry ice. After you have put all your samples in dry ice, then take all your samples and freeze them in a  -80 freezer. The protein concentration of the sample should stay the same. I hope I have helped.

Thanks for your answer Oladapo. So, do you mean that samples should be freezed in nitrogen liquid before putting them at -80ºC? When we obtain the sample, we directly put it a -80ºC until we use them... Do samples get damaged treating them that way?

we kept the aorta tissues at -80ºC until we use them.  Then, homogenized in lysis buffer [20 mMTris-HCl (pH 7.5), 2 mM EGTA, 5 mM EDTA, 10 mM dithiothreitol, 0.5 mg/ml aprotinin, 0.001 mg/ml pepstatin,0.001 mg/ml leupeptin, and 0.5 mM PMSF] on ice, thencentrifuged at 14,000 rpm at 4°C for 30 min to remove the insoluble pellet.

maybe, you should  check over the determination protocol of protein concentration, pippetting and standarts, this is the key step for WB. 

Just a word of caution re.  Fatma's suggestion: the lysis buffer used will basically remove half your proteins because of its low salt concentration.  Make sure you include enough ionic strength (at least 0.15 M of NaCl or equivalent) and you should probably include some detergent.  If you are interested in nuclear proteins, raising the salt to 0.4 M or so may be needed.  Consider sonication for hard to extract tissues.  Good luck!

Dear Teresa,

Are you still interested in advice?  I can help you a lot but you should first in a couple of my old papers on the subject.  There is a lot of information about your problem.

Apolinary Sobieszek

 Thanks to all for your answers. Yes, I am still trying to figure out where the problem is, so any advice will be welcome :)

I have written this for somebody else. Perhaps you can find useful.

As you can see have been working with the muscle tissues?  In this case addition of Mg ATP (NaATP plus Mg2Cl added at equal molar concentration; do not buy Mg ATP it is very expensive and is exactly the same) is absolutely necessary for muscle relaxation and you have to remove CaCl2 by addition of 2 mM of EGTA.  Actually in smooth you have to remove both Ca++ and Mg++ with EDTA.  More important is what kind of proteins you are trying to extract and what do you need them for?  Do you see them on SDS PAGE of your extracts? 

For a simple extraction I would suggest to use a solution at pH about 10 buffered with 10 mM EDTA. You can be sure to extract everything by adding to this buffer 8 M urea  but this will denaturate your proteins. They may renaturate if rather small.

Fragmentation of smooth muscle tissue is critical!  See my paper Sobieszek & Bremel, 1975).

Perhaps you should also read my paper on Urea-glycerol PAGE and see how the samples were prepared there.  We have routinely applied the samples from this system on SDS PAGE after simple addition of SDS (final more than ~3%).  You have to remember that the samples should be at neutral pH.  This can be judged by blue color of the coomassie blue that is normally added to the samples.  If yellow add 1-3 µl of 2-5 N NaOH.

Thank you very much Apolinary,

I have taken a look to both of your papers but still do not know what information to select in order to fix my problem.

The proteins I had problems with is b-actin, the one I use as a loading control. The thing is that after doing protein extraction and reading my samples concentration by Bradford method, I do obtain very weird results. I mean, I charge a volume of the sample that makes the exact quantity of protein I want to charge, f.e:5ug (according to its concentration). and everytime I perform an SDS-PAGE with that exact volume of those same samples, I obtained different width of the band. It seems as if the protein concentration of the sample it is not the same every time I thaw-freeze them.

This is the method it is used in our laboratory for protein extraction:

With the help of a mortar and liquid nitrogen we make powder from the freezed tissue sample.

After that we grind up the powder of the tissue with a lysis buffer, I have tried using two different buffer lysis:

1) 20mM Tris + 0'1% triton + 1%EDTA 10mM + CIP (anti-proteases from Roche)

Actually, I have made Bradford more than once with the same sample extracted with this buffer, and as I obtained different results, I did centrifuge them for almost an hour at 15000 rpm at 4ºC. After that, I removed the supernatant, leaving the pellet and the lipid layer in the old eppendorf, but still don't obtain same bands in all the samples (when it is supposed that I am charging the same quantity of the proteins in all the samples).

2) Nicholson buffer lysis (x uL of the buffer for 1mg of powdered tissue)

 This buffer obtains 1'2M urea and 4%SDS. In this case, I do not quantify the proteins

Dear Teresa,

 I would like bring to you attention to the fact that the Bradford method is your first problem and you have realized this yourself.  This method is of high sensitivity but specific for particular proteins in contrast to the Biuret method which has low sensitivity but it is not specific; i.e. staining is the same for different proteins.  The disadvantage of the latter is that it is destructive and it requires relative high proteins concentrations (amounts !).  Therefore, in case you have no problem with the amount of the protein; i.e.: you can scarify 1-2 mg of proteins (I would need to think if this is correct amount) then you should use the Biuret method. 

 You can “improve” the Bradford method by “careful” measurements and some precautions during the measurements.  It would take me a lot of time to explain therefore I would prefer to do this online via Skype (my Skype ID is kelop45).  You can sent me SMS when you want to talk so I will go on line (remember the time difference). 

 I concluded that in the end you need the lyses and protein concentrations for blotting and antibody staining.  In this case I would advise you to change the conditions of the lyses drastically.  After all, the proteins are fully denaturated during the SDS PAGE and they will not denaturate more by e.g. increasing the concentration of urea to 6 or 8 M.  Any way the later is removed during the SDS PAGE and there is no difference in the end conditions (in case they would be renaturted during the blotting). 

 I haven’t been doing a lot of antibody staining myself but during one year of experimentation in this area I realized that there are a lot of illogical steps in the established procedures.  I might be wrong; therefore you should take my suggestions with some skepticism.  

 Apolinary Sobieszek

Dear Apolinary,

I completely agree with you, there are a lot of illogical steps and it is more irreproducible than reproducible...anyway...

I will have a look to the Biuret Method. However, I don't know if it is about the Bradford method or about the way I homogenized the samples. I can not understand why they do act differently anytime I thaw them. So the didwhen using the other buffer (and the samples homogenized in this buffer are not quantifyed by Bradford).

I would really appreciate to have the chance to talk to you, let me know which the time difference is and we will see how to arrange a meeting.

I will add you on skype,

Best regards

Thanks to Fatma and Sébastien. I will consider your advice of trying to use a higher salt concentration. Although I am not interested in nuclear proteins, it's a point to take into account for future experiments. I appreciate your help,

By the way, Sebastien, what do you mean by the sonication thing? Do you sonicate your samples once you have performed the protein extraction?

Best regards.

Sometimes, the aorta tissue is really tough to break down into complete homogenization. I personally used to cut them physically into really small pieces under frozen (maybe on dry ice), I used to use cryostat but that requires some experience. Someone use other physical methods but the same concept. Then soak the small pieces of tissue in lysis buffer you choose (maybe stronger like others stated) in 4 degree for a while, which you can test the amount of time you like, maybe 20 minutes. Then I did not use sonication but used physical Bullet Blender homogenizer with metal beads at a relatively high strength plus a number of repeats, all under 4 degree (in a fridge). Eventually I can get better results than regular methods. After centrifuge there is a relatively very small size of left over. Don't forget all kinds of protease inhibitors in the buffer of course. I guess that the most I can think of.

I have used this description for Biuret methods given by WHO and find it useful. This was many years ago and I forgotten why. 

http://apps.searo.who.int/PDS_DOCS/B0218.pdf

Thank you Shuai,

I think I can try by soaking the tissue pieces for a half an hour or so, in order to improve their homogenization; since the left over I obtain after centrifugation it's quite considerable. The buffer I use contains a cocktail of inhibitors from roche, TRIS, TRITON and EDTA, shall I add anything else? Any suggestion regarding this step?

You are welcomed Teresa.

I think your inhibitor cocktail should be good.

The smaller the pieces you can cut, the stronger method you choose (but no too much, of course), the better for aorta, in my opinion.

If you have a considerable size of left over now, that could be quite possible the reason of the your problem, as you do not know if you really got all/most of the protein from the very beginning. I would see if you can try several and find a stronger homogenization method that fits you. Good luck. God bless.

I am also facing problem in getting proteins from aortic tissue. I am homogenizing tissue in ripa buffer.