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Salmon Lab Protocols: Immunofluorescence

Making Boiled Donkey (or whatever) Serum (BDS) for Blocking
You want a final working concentration of 5% Serum.  Order the serum from the same company and organism from which you buy secondary antibodies, Jackson Immunolabs for the Salmon Lab. The reason we use this for all our incubations is because of our secondary antibodies.  We use donkey secondary antibodies and thus any source of non-specific staining or background will be best blocked with donkey serum.  That way anything your antibodies will bind in serum will bind before you put on the secondary.
1) Boil normal donkey serum in the portion of ddH2O required to make a 5% serum solution in PHEM (which you'll add from your 2x stock after boiling) for 10 minutes.
Note: use caution when boiling as this tends to easily boil over.  Watch the solution boil.
2) Allow to cool.
3) Add appropriate amount of 2x PHEM and sodium azide to 0.05%.
4) Spin at 18K in Sorvall for 1 hour.
5) Filter through 0.22 um filter and store in the fridge.

 

Making Heat-Inactivated Serum for Blocking
You want a final working concentration of 20% Serum.  Order the serum from the same company and organism from which you buy secondary antibodies, Jackson Immunolabs for the Salmon Lab. The reason we use this for all our incubations is because of our secondary antibodies.  We use donkey secondary antibodies and thus any source of non-specific staining or background will be best blocked with donkey serum.  That way anything your antibodies will bind in serum will bind before you put on the secondary.
1) Heat normal donkey serum to 60-70oC in the portion of ddH2O required to make a 20% serum solution in PHEM (which you'll add from your 2x stock after heating) for 1 hour.
2) Allow to cool.
3) Add appropriate amount of 2x PHEM and sodium azide to 0.05%.
4) Filter through 0.22 um filter and store in the fridge.

5) You will have to dilute this for blocking, do a test dilution with your favorite antibody, likely this will best work at a 5-10% serum solution. The last batch, for example, worked at 6%, but the next batch worked at 3%.

 


Fresh 4% Formaldehyde
DO ALL STEPS IN THE HOOD
In 10mls ddH2O, add 0.8 g paraformaldehyde
Bring to 80oC in 50 ml conical tube in a beaker of distilled water or use the same flask each time*
NOTE: if you're using conical tubes, it is crucial to remember the thing is boiling or you'll make a huge mess and be very embarrassed!
When solution begins to clear, remove from heat
Add drops of 1N NaOH until completely clear
Add 10 mls 2x PHEM
Filter through 0.2 um filter to eliminate any polymer formed during heating.
Allow to cool before adding glutaraldehyde (if applicable)
Allow to cool before fixing cells

 


Mounting Media
20mM Tris pH 8.0
0.5% N-propyl gallate
90% Glycerol
Store at 4oC

 


PHEM (500 mls) 2x
18.14 g Pipes
6.5 g Hepes
3.8 g EGTA
0.99 g MgSO4
pH 7.0 w/ KOH

 


PBS (5x in 500 mls)
20.45 g NaCl
0.465 g KCl
10.142 g Na2HPO4*7 H2O
0.545 g KH2PO4
pH 7.2

 


F-Actin/MT Fixation Protocol (5/19/98)
updated 4/2/01
Note: My cells haven't been sticking so I now use a 20 second quick fix to just affix  the outside of the cells to the cover slip prior to permeablizing.
1) Permeablize cells in 0.5 % TritonX-100 (freshly made, sonicated, and preheated to 37oC) in PHEM for 5 minutes alone
2) Slowly dribble in about 6 drops of fix (freshly made 4 % Formaldehyde/0.5 % Glutaraldehyde in PHEM) into Permeablization buffer.  Let sit for another 2 minutes.
3) Fix in Fresh PHEM Fix for 20 minutes.
4) Rinse 3 times for 5 minutes each in PHEM.
5) Quench with Sodium Borohydride (a small pinch) in PHEM for 5 minutes.  Be very careful not to let big bubbles form, which will damage fine actin structures, by lifting out every 10-20 seconds or so.  The time will vary, so keep a close eye on it.
6) Quench 2 times in Sodium Borohydride (a small pinch) in PHEM (no Triton yet) for 4 minutes each following the same extreme caution as above.
7) Rinse 3 times for 5 minutes each in PHEM-T (PHEM + 0.1 % TritonX-100 made fresh and heated to 37oC (sometimes I sonicate and sometimes I forget, but it's probably better to be safe and sonicate to be sure the detergent is evenly distributed in the buffer).
8) ALL BLOCKS AND INCUBATIONS SHOULD BE DONE WITH THE COVERSLIPS FACING UP ON PARAFILM SO THE FINE STRUCTURES DO NOT GET DAMAGED.
9) Block at 37oC in Boiled Donkey Serum for at least 45 minutes,  ideally 60 minutes.


Note: you can leave in either block or in primary Antibody overnight at 4oC and it still looks beautiful.


10) Put on primary DM1A (mouse) at 1:300 in block (this concentration works great for my repeated frozen/thawed aliquot, but maybe if it's a  fresh aliquot use it at 1:400) for 30 minutes at 37oC.
11) Rinse 3 times for 5 minutes each rinse in PHEM-T.
12) Put on secondary (I use Donkey anti-mouse-Rhodamine Red X) at 1:100 in block for 30 minutes at 37oC.
13) Rinse 3 times for 5 minutes each rinse in PHEM-T.
14) For Alexa 488-Phallodin, I use 3 uls/coverslip and I use about 150uls of block/coverslip to apply it.  This timing is sort of crucial, only leave it in for 20 minutes at room temperature.
15) Rinse 1-2 times in PHEM-T
16) I do DAPI here... about 1 ul/10 mLs of PHEM-T for 60-120 seconds
17) Rinse 3 times in PHEM-T for 5 minutes each.
18) Rinse 1 time in PHEM for 5 minutes to get the bubbles off, etc (I don't really know why I do this step, but it feels right to remove the detergent for mounting).
19) Mount in 6-7 uls (I have a hard time avoiding bubbles so it helps to use this larger volume for me) of 90% Glycerol + N-Propyl  Gallate.

 

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