Advice needed for Western blotting

Hiya,

I am just starting up a project investigating two novel proteins in
testis function. My background is in molecular genetics and transcription
profiling, so protein work is all pretty new to me.

We have peptide antibodies prepared (or in the works) for both proteins -
the first step is obviously to characterise the antibodies. We're using
peptide antibodies rather than antibodies to recombinant protein because
there are many related genes, and we want to avoid cross-reactivity.

We have ORFs cloned for each of the genes and their close relatives. The
aim is to use in vitro transcription/translation to generate pure
proteins in order to check the antibody specificity. Once we know
that the antibodies do actually detect our proteins of interest (and not
the relatives), we can move on to other more interesting questions!

So the first plan is Western blots using whole testis extracts to see
whether we have a band of the expected size in testis. We'll then do
further Western blots of testis extract alongside the various in vitro
translated proteins in order to check AB specificity. At this point I'm
looking for advice / sanity checking on how to extract my proteins and
what sort of gels to use for the Western blots.

On looking around in the literature, I'm finding myself a little
bewildered by buffer compositions - many papers refer to "standard RIPA
buffer" or "standard Laemmli sample buffer", and yet there seem to be as
many different recipes as there are laboratories. Secondly, it looks like
the most common buffers simply won't work for my proteins!

Details below for each of the proteins of interest:

*************************************************

Protein 1: H2AL1
----------------

This one is a novel histone. I've looked at the QIAGEN QProteome Nuclear
kit, but I'm loath to commit to using that as it doesn't tell you what any
of the buffers are. I can't even tell if it's an acid extraction or a
salt extraction! Also, it's very expensive for the amount of tissue you
can process with it.

So I'd prefer to do my own histone extraction if possible. I believe that
means I want to prepare purified nuclei and then do an acid extraction.
Previously I've done nuclear preps when preparing high molecular weight
gDNA - will the same buffers be appropriate for protein work? Protocol is
as follows:

* Homogenise tissue at low speed in lysis buffer [0.05M TrisCl pH 7.5, 1mM
EDTA, 5mM MgCl2, 50mM NaCl, 5% glycerol, 1% Triton X-100, 1% B-ME], using
100 mg tissue / ml buffer.

* Centrifuge to spin down nuclei

* Wash in lysis buffer and spin down again


Once I've got the purified nuclei, the histone prep looks admirably
straightforward. According to Abcam, you just resuspend the nuclear
pellet in 0.2M HCl and leave it overnight at 4 degrees C. Centrifuge to
spin down debris and keep the supernatant. However, other protocols seem
to use different acid concentrations, or H2SO4 instead of HCl, and may
even include B-ME in the histone extraction buffer.

One protocol I Googled called for you to neutralise the preparation with
KOH afterwards - is that really called for? Wouldn't it just drop the
proteins straight back out of solution again?

Once I've got my histone prep, what buffer should I use for running them
on an SDS-PAGE gel? Standard Laemmli sample buffer? If so, *which*
standard Laemmli buffer? Will a normal SDS-PAGE gel work OK, or will
there be pH issues due to the sample being in pretty much neat acid?
Generally the lab uses Novex gels in a XCell mini-blot module. For a
histone (size ~17 kDa), it'll be quite a high percentage gel.


*************************************************


Protein 1: mgclh
------------------

Localisation of this protein is unknown, but a closely related gene
(Gmcl1) is known to be a nuclear lamina protein. Gmcl1 is RIPA-insoluble,
so it's likely mgclh will also be RIPA-insoluble. Protein size is
expected to be around 55 kDa.

The plan here is to do a standard protein extraction by homogenising
testis tissue in RIPA + protease inhibitors. Retain the supernatant (to
get the RIPA-soluble fraction), and then dissolve the pellet in something
(to get the RIPA-insoluble fraction). Question is what to dissolve the
pellet in.

The original literature on Gmcl1 said they resuspended the RIPA-insoluble
pellet in "SDS-polyacrylamide gel electrophoresis sample buffer", but
doesn't give the composition. I would assume this is 1x Laemmli sample
buffer. Does this sound plausible? If so, what recipe for Laemmli buffer
would you recommend? I've found half a dozen different recipes, and the
only common factor between them is ~60-100mM Tris pH 6.8 and 2% SDS.
There are variants with and without DTT, with and without B-ME, with
and without loading dye, using sucrose or glycerol for increased density!

I would imagine I'll need to add B-ME to reduce the proteins before
running on the gel, but should I add the B-ME at the point I resuspend the
pellet, or only when I'm about to run an aliquot on a gel? Will I need to
add protease inhibitors, or will the Laemmli buffer itself take care of
that?

*************************************************

Thank you for your time and patience - sorry for asking so many questions,
most of which will undoubtedly have obvious answers. Probably several
mutually incompatible obvious answers, mind you, but that's biology for
you. My theory is that it's better to ask silly questions and look daft
than to forge ahead without asking and cock something up!

Thanks,
Peter Ellis

Am 17.06.2008, 07:06 Uhr, schrieb Peter Ellis <[Only registered users see links. ].uk>:


That is fine, just remember that the epitope may be hidden in native
proteins, thus the AB may not work e.g. in immunoprecipitation. In Western
you deal with denatured proteins, so that is not a problem.


In addition to testing against individual proteins you should test against
a crude tissue lysate. You should see only a single band. Even more
telling would be a blot from a 2D-gel (SDS-PAGE after IEF).


The most common method is SDS-PAGE according to Laemmli (Laemmli: Nature
227 (1970) 680 and Laemmli & Favre, J. Mol. Biol. 80 (1973) 575-599).
Separation is by molecular mass. For more specific detection, this is
preceeded by isoelectric focussing, a method that separates by isoelectric
point. See J. Klose and U. Kobalz: Electrophoresis 16 (1995) 1034-1059


It's not as bad as that. The Laemmli buffer is a fairly good bet.


Histones have a high positive charge, to get a reasonable estimate of
their molecular mass from PAGE, use the positive detergent CTAB rather
than the negatively charged SDS (E. Buxbaum: Anal. Biochem. 314 {2003}
70-76)


I'd probably use KCl rather than NaCl, as the cell interior is high in K
and low in Na. I'd also reduce [Mg] to 1 mM or so to discourage
metal-dependent proteases. The Triton I'd leave out at this step and
reduce bME to 1 mM (or replace it with DTT, which has a more appropriate
redox potential). You'll need a protease inhibitor coctail (EDTA, PMSF,
Pepstatin, Leupeptin, e-aminocaproic acid).


The acid is quite irrelevant, the idea is simply to increase the
solubility of the highly charged histons and to pellet as many of the
other proteins as possible. The bME protects cysteine -SH groups from
oxidation by air. However, this extraction procedure is too selective for
your purposes, you will not know whether there are cross-reacting proteins
in the pellet.

No. But you need a pH near neutral and as low an ion concentration as
possible for the electrophoresis step. Therefore solubilisation in sample
buffer rather than HCl is better. That also gives you an idea about the
cross-reactions of your antibody.


I find gradient gels (5-15 or 20%) most convenient


Again the use of CTAB electrophoresis may help, as CTAB can solubilise
proteins very efficiently.


The buffer is actually double concentrated, once you mix it with an equal
volume of sample you get the working strenght. DTT and bME reduce
disulphide bonds and thus lead to more complete unfolding of the proteins.
DTT (Clelands reagent) smells less and has a better standard redox
potential for this purpose. However, it is more expensive than bME. Wether
you use glycerol or sucrose to increase density really doesn't matter, you
just need either for proper gel loading, its physics, not chemistry. The
dye is important not only to aid in loading, but even more as front
marker, without it you can't calculate Rf-values which you need for
molecular mass determination.


Reducing agent is part of the 2x concentrate, usually the sample/buffer
mixture is heated to get more efficient reduction. Many people do 2 min in
a boiling water bath and for your purposes that is probably fine. With
large proteins I found that 10 min at 60 degrees works better. Very few
proteases work in hot SDS under reducing conditions, but there have been
rare cases reported where that was a problem. The protease inhibitors
should of course be present during the homogenisation of the cells and the
centrifugation steps. Many (like PMSF) actually chemically inactivate
proteases, solving the problem permanently anyway.