Following is a summary of the responses I received about DIG in situ
hybridization. Thank everyone who share me with very helpful information.
With regards to non-radioactive in situ protocol. I have used the one
essentially described in Coen et al. (1990) Cell 63; 1311-1322. Some
modifications can be found in Bradley et al. (1993) Cell 72; 85-95 and in
Fobert et al. (1994) EMBO J 13; 616-624. I have not tried it on Arabidopsis
roots, but it works on Antirrhinum roots and well as various confier
tissues. Unfortunately my copy of the protocol is heavily marked up and
will not photocopy well. I keep meaning to re-type it, but...There is a
relatively new book out (sorry, I don't remember the name, but could find
out if you wish) on in situ hybridization. David Jackson has a chapter in
it which may be of interest to you for details, etc...
Putting the sections through an ethanol series will weaken the signal, in
some cases considerably. However, background hybridization can be a serious
problem if you do not dehydrate. With very few exceptions, I always wash in
ethanol. One could always argue that if the signal washes away, it wasn't
"real" anyway! The only dyes that I have used for staining background
tissues (as described in the references listed above) are fluorescent ones.
The advantage of these is that you do not alter the coloration of the
sections, which may obscure signal in some cases. If you decide that the
tissue doesn't look good with the fluorescent dye, you shut off the U.V. on
I hope that this is of some help to you. I think that you will find that
the radioactive methods are more sensitive than the DIG, but their
resolution may not be as great.
Good luck with your experiments.
Pierre Fobert, Research Officer
>Dear Yi-Fang Tsay, I have a collection of methods for doing dig - in situs
>which I will send you (I have a paper in Feb. 94 "Development" (Jackson et
>al.)- let me know if you don't have access to that and I will send you a
The NBT product is only slightly soluble in ethanol. It's fine to
dehydrate the slides through an ethanol series, just dont spend more than
15-30 seconds in each step. The product is not soluble in clearing agents
(e.g. xylenes, histoclear), so you can let the slides sit in them for a
while before mounting.
Often there is faint brown background over the tissue, so counterstaining
is not necessary; especially if you can use some kind of interference
optics like nomarski. Otherwise you should use a pale pink/red
counterstain , I've used safranin succesfully, but at a much lower
concentration than normal. Obviously you shouldnt use tol. blue, since the
reaction product is also blue,
best wishes, David Jackson.
One thing I would like to point out is how
difficult it is to get decent root sections. We found the roots got a
life of their own while trying to mount them in wax prior to sectioning.
We used Toluene Blue as a background stain, and this showed all
important parts of our sections very clearly. We also found that DAPI
staining, to observe the nucleic acids, gave a good indication of how
well preserved the sections were. The stronger the staining the better
the sections tended to be for probing.
If you want the whole protocol write back and I would be glad to send it
University of Sussex
Brighton, BN1 9QG
I'm actually doing the hybridization on my first attempt at whole mount
in situs with DIG labelled probes right now. I don't have any results yet,
so I can't recommend a protocol, although I do have several detailed protocols.
I'll tell you what I know or have heard from others. My source is from a
lab which is very successful getting whole-mount DIG in situs to work in
mouse embryos. They say that after doing whole mounts they go through a normal
fixation/dehydration schedule for paraffin embedding and then section the
embryos. I assume they use EtOH dehydrations for this, and I know they use
NBT and X-Phos as their color reagents. As for background staining, I learned
from a very senior plant anatomist at UC Davis (Dr. Ernest Gifford) that
the best background stain to use for tissue that will be paraffin embedded
(i.e. taken through EtOH, xylene or tertiary butyl alcohol dehydration) is
saffranine O (a red dye). I don't know about neutral red. I know that TBO
will NOT stay in tissue during lengthy dehydration. If it is very quick
(as for dehydrating sections which are already fixed to slides) then TBO can
go through ethanol (my own experience). I don't know if it will stay in
place in xylene or TBA.
Here are two references with extensive descriptions of in situ methods.
Please send me your address again, and I'll send you some protocols by mail
with comments about my results.
Univ. of Toronto
Tautz and Pfeifle (1989) Chromosoma 98:81-85
Hemmati-Brivanlou et al. (1990) Development 325-330
Good on DIG methods, but for sections
Tanimoto and Rost (1993) in Methods in Plant Biochemistry v 10, pp 141-158
Academic Press (I don't know the editors, sorry)
Dear Yi-Fang Tsay,
The most useful methods for you will be some written up by Cindy Lincoln,
another post doc in the lab, who has used bits of mine and Vivian Irish'
protocols and has great results with dig in situs in arabidopsis.
One thing I recommend is that you try to get a good positive control-
something like tubulin or a ribosomal probe which is highly expressed and
will give you intense staining. That way you know that if your other
probes dont work the problem may be one of sensitivity rather than just the
method itself not working. Whilst dig is great for in situs, it is still a
little less sensitive than radioactive methods.
Feel free to email me if you have other questions, Dave
There is a protocol in Plant Molecular Biology Report vol 8(4), 1990.
Hope you find it helpful.
The answers to your specific questions are in the protocol, but I thought
I would also answer them directly. The dehydration through ethanol to
xylene after detection can decrease the signal, however when the signal
is strong and the washes are only a few seconds then it serves nicely to
eliminate any background. Another option would be to use an aqueous
mounting media, but I have heard that this is good for the signal and not
the tissue morphology. As well, when using a xylene-based mounting medium,
don't use Permount, use another such as Entellen from BDH. Second: the signal
from DIG using the standard X-phosphate- NBT substrate will be in the form of a
blue precipitate -- the use of any counterstain will obscure the signal, perhaps
totally. As stated in our protocol, one option is to
stain with 0.1% Fluorescent Brightener 28 (Calcofluor White, CI 40622) --
another option, which is the one we use, is to use phase-contrast to see the
Botany Dept, UBC
Vancouver, BC V6T 1Z4
western at unixg.ubc.ca
Sorry for this late reply. We are currently using DIG labeled probes for
in situ hybridization. For Arabidopsis this has worked very well on
sections for a range of probes. We've had only problems with the quality
of the sections. Parafin results in a bad preservation of young growing
tisues. We therefore use methacrylate embedding. Methacrylate is a resin
that can be removed after sectioning using acetone. It works well for in
situ hybridization and immunocytochemistry. We have published a short
technical paper on this: Kronenberger et al. (1993) Cell Biol. Int. 17
(11) 1013-1021. I can send you a copy if you could give me your complete
As for the dehydration before mounting we haven't had any problems so
far. By the way, in case of weak signals you can leave the substrate on
the sections for long periods (up to three days). Of course you need to
combine this with the right controls (and sense controls are not always
the best ones, since they can give aspecific labeling).
INRA, laboratoire de Biologie Cellulaire
Route de saint Cyr
78280 Versailles cedex
Both Dr. Tamara Western and Dr. Neil Butt have sent me a complete protocols. If
anyone is interested in it, I can e-mail you a copy.