Message: 9
Date: Fri, 12 Jan 2007 15:01:25 -0500
From: Melissa Mazan 
Subject: [Histonet] immunocytochemistry
To: histonet@lists.utsouthwestern.edu
Message-ID: <20070112150125.bxbafrwguo8ogck4@webmail.tufts.edu>
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Hi all, I haven't done immunofluorescence on cultured cells before, and 
one of my colleagues would like me to do some - he's grown up the cells 
on cover slips, then fixed with methanol/acetic acid.  They've been 
stored at room temp for the past week. Prior to staining, do I have to 
rehydrate them with graded alcohols? or just rinse with PBS or 
PBS/triton-x?
Thanks! Melissa



Hi Melissa

You can stain 24 well plates instead of cover glasses if you want. The
coverslips can go directly into PBS- Hopefully they were rinsed after
fixing and not air dried with the acetic acid left on. The other problem
I have had is fixing cells without rinsing the media with FBS or other
kind of serum.  That tends to leave a bad haze on my cells. I do not let
my cells dry out at anytime. I personally will avoid cover glass
staining if possible. I stain the plastic plates or use chamber slides.
The same reagents could be used for cover glasses.

For the antibodies I stain I have not found methanol to be a good fix,
especially the cell membrane markers so I do not use it for anything.
In the future I would fix your cells, rinse them and hold them in buffer
for up to about a week before staining.  Many cells will start to lift
off the plate or slide after a week.  My preferred antibodies are
PharMingen and I also use a lot of Lab Vision.
Here is the "cheat sheet" I give everyone here- it covers all the IF we
do.  We stain many plates (1-14 at a time with 1-6 different antibodies
per plate) with excellent results. I do cover them when in secondary and
store them in the fridge, usually in the cardboard Revco boxes that fit
2 or 3 plates. The formatting got lost so if you want an easier copy to
read, let me know and I will attach one.

Good luck!


Donna

Donna Harclerode, HT, (ASCP), HTL, QIHC
Scientist / Immunohistochemistry
Cytori Therapeutics
3020 Callan Rd.
San Diego, CA 92121
858-458-0900 ext 5416
fax 858 200-0945
dharclerode@cytoritx.com

Basic Fluorescent Immunocytochemistry for Plates or Chamber Slides
DAB on final page

Remove media from live cells
1.	Rinse live cells in media containing no serum or in PBS 1 times
1-5 minutes.
2.	Fix cells in 4% PFA made from concentrate (32% Electron
Microscopy Science) using PBS as a diluent for 15 minutes and remove
fixative
3.	Rinse in PBS 3 time 5 minutes each
4.	Make up correct dilution for primary antibody in Dako Antibody
diluent (I do not use serum usually in any antibody solution but you can
add 5% Normal Donkey serum to the diluent used for the primary antibody-
all my secondaries are made in donkey) Just prior to adding antibody
gently blot the plates on paper towel to limit dilution of antibodies.
	and add 100 ul to each chamber on a 24 well plate to be stained.
Rotate the 	plate to be sure the entire well is covered with
reagent.
FOR OVERNIGHT CHAMBER SLIDES- USE A HUMID CHAMBER AND 
100ul per well on a 4 well chamber slide or a 24 well plate 
or 200ul on a 2 chamber slide
TC plates also benefit from wet paper towels being placed on top of each
plate
5.	Place in 4oC Fridge overnight.  Alternately you can use a rocker
at room temperature for 2 hours.  
6.	Remove the primary antibody and rinse 3 times in PBS each for 5
minutes (10 minutes for each rinse may be preferred)
7.	Incubate with secondary fluorophore made against the species
that the primary antibody is made in and incubate on rocker for 30
minutes to 1 hour. All secondaries are from Jackson ImmuoResearch
diluted at 1:100 with Dako Antibody diluent (NO serum) (up to 1:200
dilution of secondary can be used).

If desired add 1ul of DAPI solution (frozen aliquots) for each ml of
secondary for nuclear detection

Unless otherwise requested for specific experiments use whole IgG
secondaries.  Fab' fragment secondaries have sometimes shown not to be
as bright as whole IgG.  If there is possible Fc receptor interaction a
small comparison study could be used to see if the Fab' secondary
decrease background.  
Whole molecule secondaries are much less expensive than Fab' fragment
8.	Rinse plates 3 times with PBS each for 5 minutes
9.	Plates can be viewed without coverslipping if desired or a
mounting media made for fluorescent dyes may be used.  
10.	For chamber slides remove the collar rinse again in PBS and
coverslip with Aqua Mount or other mountant for fluorescent preparations

11.	Slides may be sealed with nail polish, but this will prevent
removal of the coverslip and remounting if there are problems with the
slide.  I highly recommend not ringing the aqueous sections with nail
polish for this reason.  If they are coverslipped correctly and kept
flat in the folders, the covers slips stay where they belong





Notes:

Do not allow plates to dry out at any time.  Adding different antibodies
to 24 well plates requires planning and concentration.

A rocking plate is used for all rinses and room temperature incubations.
An orbital rocker should not be used, as the center of each chamber is
likely to have less contact with the antibody

>From the secondary antibody (fluorophore) on, keep the slides or plates
from direct light exposure.  It is not necessary to seal them in foil,
but blocking the overhead light with a box or towel is good.

Antibody concentrations used for fluorescent tissue sections IHC
staining will be close if not exactly the correct concentration for
chamber slides
 
Dako diluent contains both a non serum block for background and a
detergent for permeablization of the cells. The non serum block will
prevent artifact staining of Fc receptors and secondary cross reactivity
with the primary.

Add antibody and PBS gently on the side of the well so the cells will
remain adherent to plate or slide.  To remove reagents use an aspirator
or just gently turn the plates onto clean papers towels and blot gently.


Results tend to be crisper when primary antibody is incubated overnight
at 4oC than 2 hours at room temp

Do not use any permanent mounting media containing alcohol or other
solvent on fluorescent preparations.

Chamber slides can have the collars removed before fixation if the
antibodies require acetone or any other solvent based fixative.  They
must then be stained in humid chamber flat and cannot be used with the
Sequenza system used for standard slides because of the dividers on the
slides. The unfixed chamber slides are very fragile, they must be
treated very gently or cells will be knocked off the slide.

Any or all PBS rinses can be increased to10 minutes or more if
convenient or if any type of matrix is used with the cells.

Slides or plates may be stored at 4oC in the fridge for up to 2 months.
Some signals may last longer, but these will be on a case by case basis

Alternate fixatives to 4% PFA can be used.  Acetone -Ethanol 50% of each
can be used in plates.  We did not find a significant improvement to
date with any abs we tested.  Methanol is also recommended by some
papers.  I have found serious problems with methanol fixation and would
try it only as a last resort. 

Ideally plates will be stained immediately after fixation. Plates can be
fixed and then stored in PBS after 3 washers at 4 oC for up to one week.

Prepared antibodies are stable if made in Dako diluent and stored at 4oC
for at least 1 month.  Secondary preparations are stable for at least 2
weeks if kept at 4oC and light protected.

Step 1-6 
Rinse cells, fix plates, wash with PBS 3 times 5 minutes

7. Add 2-3 drops per plate of HRP block (Dako Endogenous Peroxidase
block) if there is any chance that there will be RBCs or other HRP cells
(granulocytes) for 10 minutes

8.  Rinse 3x with PBS (if you have cells with endogenous biotin, you
will also need to block for that)

9 Add 100ul properly diluted primary ab to a 24 well plate and incubate
on rocker for 1-2 hours

7. Rinse Plate 3 times each for 5 minutes with PBS

8.  Add 100ul of  secondary antibody to each well (unless using a
biotinolated primary) made in Dako diluent at 1:200 biotin anti mouse or
rabbit or whatever and incubate for 30 minutes on the rocker

9 Rinse the plates 3 times for 5 minutes each 

10.  Add 2-3 drops LSAB (labeled strep avidin biotin) to each well and
incubate 30 minutes on a rocker

11. Rinse 3 times with PBS each for 5 minutes

12 Always wear gloves and perform the DAB step on paper that will be
disposed of after use.  It will dye your clothes and is a mild
carcinogen.
Make up DAB solution 1 ml diluent 1 drop reagent.  4-5 ml is a good
amount for a 24 well plate

13 Add 3-5 drops of DAB per well to stain the reactive cells.  If all
wells are not done at the same time, keep buffer in the well and do not
allow to dry out.  Stain from 30 seconds to 5 minutes (usual is 1-2
minutes.

Rinse plates well in DI water

Counterstain with Gill 1 hematoxylin if desired 5-10 seconds

Rinse with DI water

Blue with Bluing reagent (or sodium bicarbonate of other mild base)

Rinse well with DI water and allow drying.

DI water can be added to plates if desired for viewing or visualized
dry.  Do not store wet plates long term as they will eventually grow
mold.



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