Native PAGE stacker pH

Hi all,

Can anyone tell me why the Ornstein and Davis Native PAGE recipe for a
stacking gel is at pH 6.8 when the separating gel and buffers are
usually around 8.5 and up? It seems counter-intuitive to drop the pH in
the stacker... or is this something to do with gel integrity? I have
looked through the original Ornstein papers but can't quite work out the
reasoning. Anyone know the logic behind it?

Cheers,

--
Bean

Remove "yourfinger" before replying

In article <44e2be1f$[Only registered users see links. ].au>, [Only registered users see links. ] wrote:

Isn't it amazing that the basic theory of electrophoresis is no
longer taught and in 99.9% cases people using electrophores
have vague idea why what they are doing works?

For a stacker to work, you want the voltage to drop primarily on it.
E.g, you want the field intensity, E (V/cm), to be high in the stacker
and low in the separating gel. The rate of EF is proportional to E.
That means protein will run fast through stacker and slow down
once reaching the end of it ==> concentrating action.

All of it requires low conductivity in the stacker. Cl- is a fast ion,
glycine is a trailing ion. pI of glycine is somewhere close to neutral.
Which means that close to 50% of glycine molecules in the stacker
will be neutral and won't conduct current. There you have it: low
concentration of slowly moving ions = low conductivity = stacking
action.

DK

Can anyone tell me why the Ornstein and Davis Native PAGE recipe for a

For a stacker to work, you want the voltage to drop primarily on it.

...yes, DK is right.... those are the pH conditions that lead to
stacking of proteins, and that's true even for SDS PAGE.... chloride
act as the "leading ion" and glycine as "trailing" ions....except here
due to the SDS binding (1.4g of SDS/1g protein), the proteins acquire
a net negative charge, and their migration therefore depends on the
sieving action of the gel, and therefore the mobility is proportional
to the size...

it's true DK, "biochemistry" seems no longer in fashion....(
pow


On 8/16/06, DK <dk@no.email.thankstospam.net> wrote:

Bean Long wrote:


That's strange, I thought they explained it pretty well. The stacking
effect of discontinous (DISK) electrophoresis (note that the
Laemli-system belongs here as well) is required to concentrate the
protein on the stacking gel, and in the process to reduce the width of
the bands from several mm in the loading zone (depending on sample
volume) to less than a mm at the transition from stacking to separating
gel. Only this way you can combine large sample volumes with high
resolution.

To achieve this, the proteins move between a fast moving (leading) ion
alpha and a slow moving (trailing) ion beta, with a common counter-ion
gamma. Then Kohlrauschs regulating function applies resulting in
concentration. You can make this effect visible by using prestained
molecular weight markers (e.g. Rainbow Markers), it is very intuitive to
watch the sorting and stacking life. In the stack all protein move with
the same velocity, but at different positions in the stack. Note that it
is possible to separate proteins in the stacking gel only
(isotachyphoresis), separation distance however are small.

Once the stack arrives at the stacking/separating gel boundary the
change in pH increases the ionisation (and hence mobility) of the
trailing ion, which passes the protein stack. The proteins from then on
move in a constant electrical field, their speed now depends on their
size and charge.

You can use different buffer systems for DISK-electrophoresis depending
on the stability of your sample or for optimising the separation between
certain proteins. See Jovin, Ann. N.Y. Acad. Sci. 209 (1973) 477-96 for
how to calculate such systems and [Only registered users see links. ] for the
calculator.

Many thanks all. It's not that basic biochemistry is not taught
properly any more... it's just that I forgot ! :-)

After I sent the message it immediately dawned on me that there would be
consequences for the trailing and leading ions at this pH. It all
suddenly came back!

Isn't science wonderful!!

Cheers,
--
Bean

Remove "yourfinger" before replying

On 8/17/06, Bean Long <ben.long@yourfinger.anu.edu.au> wrote:


...... science is almost magical )) it's why, i suppose, all of us
folks spend the better part of our life holed up in the Lab )) ....
and thanks for your question as well. I, too, learnt some new words
"isotachyphoresis", and recollected some old forgotten terms that were
already vague in the mind, like Kohlrauschs function.... thanks to
Engelbert.


pow

On Wed, 16 Aug 2006 13:39:29 GMT, [Only registered users see links. ] (DK)
wrote:


I must have got confused somewhere - isn't the local voltage supposed
to be high in the stacking part? You need the protein to move fast
here but it can't move faster than the Cl- ion, so it slowly stacks?

A separate question not quite related to the thread on native PAGE to
those who have done native PAGE often - how successful are you at
doing native PAGE? I have met more people who failed to get native
PAGE to work or got the wrong result than those who are successful, so
I'm wondering if the technique is completely sound or if the people
doing it are just incompetent.

In article <[Only registered users see links. ]>, ChenHA
<[Only registered users see links. ]> wrote:

Just a minor terminology confusion. "Drop" above is a verb, not noun.
Voltage drops *on* stacker (The analogy is river with a waterfall).
Most of the "drop" occurs where resistance is highest, e.g. stacker.
So it is the electric field *intensity* (which is a driving force of
ectrophoretic movement), that is high in the stacker. If under "local
voltage" you meant difference in electric potentials between stacker
ends, then you are absolutely correct - it will be larger than the
difference of potentials between, say, ends of separating gel or
power supply end the start of stacker.


IMHO, it's pretty straightforward but heavily depends on your protein.
In many cases, a particular protein is unstable/aggregating
in the buffer conditions chosen (true particularly for many loading
buffers). For most acidic proteins, it's really a piece of cake - just use
you regular SDS-PAGE system and omit SDS from everywhere.
It goes without saying that the gel has to be run in the cold room, with
ice cold buffers and either be cooled with the recirculating bath (as Hoefer
allows) or be completely submersed in large volume of the lower buffer
(as possible with Bio-Rad's mini). Also run at lower voltage (I use 150V
for Bio-Rad) and runs take much longer, particularly if you want
separation between specific proteins.

No matter what, native gels always look a lot uglier than SDS-PAGE.
I find natives indispensable in refolding studies - you can clearly see the
amout of oligomers, smearing, etc and estimate refolding efficiency.
Being able to do it for 26 sample at a time surely beats running
26 HPLC gel-filtrations... (There are some pretty neat things that
can be observed. For example, in one case I clearly that glycerol
and Triton X-100 stabilized some intermediate fold that ran right
in between a properly folded monomer and some weird dimer).

DK

DK wrote:

OK, thanks for the clarification. I thought you meant the voltage is
lower when you said "drops".



I think for some reasons, most of the people I talked to worked on
basic proteins, perhaps that explain why they have difficulties. So
the question will then be, is native PAGE basically not useful for
basic proteins?

It is probably strange that since I work with proteins, I haven't
actually done any native PAGE before. But I am interested to know how
reliable that technique is for basic proteins. One of the problem is
that in papers people don't report things that don't worked - for
example, some people ran a native PAGE that gives the result that
contradicts all the other biophysical analyses (smaller monomer rather
than the larger oligomer expected, or vice versa). The results don't
get reported of course, but given that I have found more people who
can't get the right result (it appears to be random process whether
the protein bands appear in the right place), can I trust the result
of those who use native PAGE to indicate oligomeric states for basic
proteins?

ChenHA wrote:
Just switch the voltage.