My thoughts:
1. Consider what you are trying to preserve, and "when" in the
protocol you need to fix it. Some things like actin stress fibers are
so cross-linked that a minute of 0.5 % Triton doesn't do much damage
before fixation, while a lot of other cytoplasmic, membrane, and some
cytoskeletal material disappear. Point is that the use of detergent
is very sample dependent and requires testing. And as I will say
again below, detergents and antibodies need never be in the same
solution.
2. The more "effective" the detergent at scrubbing, the more loss of
material. Many procedures use 0.2% Tween 20, which is not so nasty as
Triton. Again, what is the target. If I want the best preservation of
microtubules, not only must the sample be pre-fixed first at 37 deg.
C, no detergent can be used at all. After prefixing, I then move on
to the membrane permeable paraformaldehyde, not requiring detergent
for penetration and fixation. Any subsequent detergent treatment,
such as 0.2% Triton, is to ensure that membrane "holes" are large
enough to let antibodies get into the cells, but I use it only
briefly in the blocking step and not subsequently. Antibodies never
see the detergent.
3. I am not sure about Tween, but most common detergents have the
effect of making pore like structures in membranes. Low
concentrations of Triton produce a modestly specific sodium ion
permeability. The average size, permeability behavior, etc will
depend on the effective membrane concentration achieved, and the
amount of lipid and detergent soluble material that can be extracted.
This will affect the size of the domains available permitting
penetration of antibodies. Thus, there are a number of parameters to
consider for each target to be labelled, including probe properties
such as IgG, or Fab or Fab2 fragments. Smaller things get in more
easily and more quickly and are less sticky. The use of solvents such
as DMSO and ethanol have a strong disordering effects on lipid
membranes and, in fact, it is this property that creates space for
probe molecules to get in cells depending on the protocol. Old
procedures for microtubules used cold acetone or methanol to
permeabilize and fix. They also can effectively kill certain epitopes
making them pretty useless.
In Ian's example, unless I read it wrong, he was describing solvent
treatment on sections. He did not describe, the fixation itself which
I think important to the success of his approach and it seems to be
in the context of TEM more than confocal, which usually doesn't
include 50-80 um sections but 0.5 um sections at best. Ian, perhaps
you could clarify? In any event, detergents work by disrupting
hydrophobic structures, which means that they can kill antibody
probes as well as epitopes of interest. So in that I agree, fully.
Keep detergents out of the antibodies. Besides, once detergent has
been used on the sample, the permeabilizing is done. Wash it away,
the holes will stay.
4. Yes, freeze thaw cycles are bad, but it will really depend on the
antibody as to how tolerant it is and as to what the other proteins
it is stored with, if any. The issue is denaturation and loss of
specificity. Inclusion of other passive proteins such as BSA to 10
mgs/ml can help. Our labs practice has been to upon arrival of
antibody from Jackson, Calbio, etc., to aliquot 2 ul into 0.5-1.5 ml
plastic vials and store at -80 deg C. Use a vial for a few days after
diluting with incubation media, which usually involves Pierce
SuperBlock or equivalent and store between uses at 4 deg C. After a
few days, throw it out. One freeze, one thaw. Why even make it an
issue except for antibodies that don't like being put at -80 deg.
There are some. The glycerol approach seems to work for them stored
at -20 deg. C.
>Hi All - Ian's last reply was very interesting, as I've been wondering about
>detergent alternatives to permeabilisation for antibodies some time. Any
>opinions among people about comparative merits of Triton, Tween, saponin, on
>morphology if you have to go the detergent route. Also what alternatives are
>there - freeze-thaw? for thick specimens (say 50 -80 microns
>vibratome cut brain
>sections). Ian, could you give a reference or detailed protocol for the DMSO
>method? How uniform is the labelling through the thickness of the section?
>regards,
>
>Martyn Evans
>SmithKline Beecham
>Harlow UK
--
Mario M. Moronne, Ph.D.
NanoMed Technologies
ph (510) 528-2400
FAX (510) 528-8076
Berkeley, CA
94706
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