Mckenzie at naxos.unice.fr
Mon Jul 24 06:06:19 EST 1995
In article <kalch-2307951827550001 at hayci.generes.ca> Mikek, kalch at ulam.generes.ca writes:
First of all, In my opinion you Should be getting two major bands in the presence
of your monoclonal. An IgG molecule has a molecular mass of about 160,000 Da.
This is made up with two heavy chains (55,000 Da each) and two lighter chains
(25,000 Da each). Try a laemmli running buffer with no DTT or beta-mercapto to
break up the disulphide bridges holding the IgG together and you should find that
your IgG molecules stay together and you have a single band (90% of the protein)
The second question you asked is a bit more tricky.... I presume that you have a
reasonable amount of salt, tris, edta etc in your lysis buffer. If you are hoping that
a protein co-ip's with your protein of interest in your lysis buffer cocktail,
(which I suppose you incubate together for several hours) then why would you want
to change this buffer for the washes? I'd wash with exactly the same buffer and hope
to maintain exactly the same interactions. The problem then becomes, what do you
define as your lysis buffer; do you want to have a high NaCl buffer for example which
may be closer to the conditions outside the cell, or a lower salt buffer (say 10mM NaCl)
which is closer to the intracellular salt concentration. ( but in which the antibodies
may not function as well). It depends on your protein of interest and what you are
I would add just one word about the protein-A-seph washes. Don't vortex the beads
like crazy and pellet them at 20,000 x g. It isn't neccesary and you will break the beads.
When washing beads and wanting to be more or less stringent, then the number of washes is what counts and not the volume.
Hope you co-Ip something of interest!
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