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1. Concentrating protein lysates (Angela Alexander)
2. antibody crossreactivity problem (Michael Sullivan)
3. Re: antibody crossreactivity problem (peter)
4. Re: antibody crossreactivity problem (Tom Anderson)
5. Re: Concentrating protein lysates (AK)
6. Re: antibody crossreactivity problem (DK)
7. Re: Concentrating protein lysates (DK)
8. S-gal (joel_boumje_boumje from yahoo.com)
9. Re: antibody crossreactivity problem (Pow Joshi)
Date: Mon, 2 Jul 2007 04:27:59 -0700 (PDT)
From: Angela Alexander
Subject: Concentrating protein lysates
To: methods from magpie.bio.indiana.edu
Message-ID: <290513.32287.qm from web54109.mail.re2.yahoo.com>
Content-Type: text/plain; charset=iso-8859-1
I'm working with a new cell line that's giving me very dilute lysates (even with a vastly reduced volume of lysis buffer), such that I can barely load 15-20 micrograms of protein on a western gel. I'm doing a lot of signaling analysis and sometimes need more protein than that to get solid blots for some phospho-antibodies (we usually load 30-40). So I was wondering if you guys had any suggestions on the best method for concentrating my lysates without affecting my data ie losing robustness of phosphorylation.
Speedvac'ing might be reasonable, as we have one in my lab right behind my bench, but I'm concerned the length of time the lysate will not be kept cold that will be required might not be such a great idea. Is dialysis the only way to go, or is there a quicker, easier method?....since I will be generating many long time courses/dose responses with these cells once I figure out the best method/something that works.
University of Texas MD Anderson Cancer Center
easiest way to concentrate your proteins is Amicon/Pall filters. You can get centrifugal filters for various volumes with MWCO 3-100 kDa. We use these to remove proteins from our HPLC samples, but we also use them to concentrate proteins by removing buffer. The centrifugal filters come loaded with azide. To remove azide, we spin 20 minutes with 20-40 uL ddH2O and discard. Then, load the sample on top of the filter and spin at 5 min intervals, 4C, 14K x g. Spin until the volume remaining on top of the filter is the volume you want.
These are expensive so we reuse ours in the lab. After we're done with the filter, we run through with Tris or Kpi buffer, 20% glycerol buffer, (usually 2-3x volume we just used). Then we'll filter again with 20% EtOH, or a very high glycerol buffer 50-80% and leave the membrane wet. Store at 4C until use again. The difference in solutions is based on what they are used for. for instance, I use some for filtering HPLC samples- on a column that is organic intolerant, these I use high glycerol buffer for. In other applications, proteins etc, I use EtOH. we've found these very useful, faster and easier than dialysis.
Grad. Res. Asst.
University of Texas- Austin
Institute for Cellular and Molecular Biology
Department of Medicinal Chemistry
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