The Mitchison Lab has an excellent guide on staining and fixing cells for Actin and Microtubules which is worth reading [Link]
Most coverslips come with a fine film coating to stop them sticking to each other. This can reduce the ability of coating agents such as poly-L lysine from working properly, and can thus reduce the ability of cells to properly adhear to the glass. As most mitotic cells ’round’ up and have a much weaker attachment, a poorly coated coverslip can dramatically reduce the numbers of cells you finally have to look at down the microscope. Thus it is always important to first clean the coverslips and then coat them with either Histogrip, Fibronectin, or Poly-L-lysine.
1) Boil coverslips in dH2O in a large beaker for several minutes in a microwave
2) Add HCl to a final concentration of about 1M to the hot water. Careful of fumes do in a fume hood if possible.
3) Cover the beaker with some parafilm, and gently stir/rock the coverslips on for 4-16h or until cool.
4) Rinse the coverslips several times in dH2O.
5) Then rinse 3-5x with 100% Ethanol, leave coverslips in EtOH and go to TC hood
6) In TC hood, separate individual coverslips out onto large piece of clean Kimwipe or similar blotting paper and allow to air dry.
7) They can now be stored (for unto a year) in a 10cm plate or 50ml falcon until coating. Some people like to autoclave them but it is not necessary.
1) In a TC hood, make a 1/10 dilution of the histogrip into 100% Acetone in a 50ml Falcon tube. Normally 10-15ml final volume is plenty.
NB: most TC plastic plates will be dissolved by the acetone, but most 50ml Falcons should be ok, but check first.
2) Have a second empty 50ml falcon ready.
3) Drop about 10-20 individual coverslips one by one into the 50ml falcon with the Histogrip solution. Re-cap and invert tube gently several times.
4) Decant the Histogrip solution into the empty 50ml falcon.
5) Place coated coverslips into a 3rd Falcon full of TC clean H2O
6) Repeat steps 3-5 until you have coated enough coverslips
7) Remove H2O and wash coated coverslips 3x with H2O
8) Separate individual coverslips out onto large piece of clean Kimwipe or similar blotting paper and allow to air dry.
9) They can now be stored (for unto a year) in a 10cm plate or 50ml falcon.
25 mM HEPES
1 mM EGTA
60 mM PIPES
2 mM MgCl2
pH = 6.9
(Add in this order.)
Antibody Blocking Solution (ABS)
3% BSA (or 5% Fetal Calf Serum)
Mix well and filter, aliquot and store at -20°C
Formaldehyde Fixation in PHEM buffer
(Good general use fixation, good for kinetochore proteins, ok for microtubules)
1. Wash coverslips 2x with 1X PBS.
2 . If staining a cytoplasmic protein or if you have high background then try a short pre-permeabilize of cells using 0.1-0.5% Triton in PHEM buffer 30 sec -1 min at room temperature (RT).
3. Carefully fix cells with 3.7% Formaldehyde (fresh is best) diluted in PHEM + 0.5% TritonX-100 for 10 min at RT.
4. Wash 4x with 1X PBS at RT. Can be stored at 4°C for 2-3 days at this stage.
5. Block coverslips for 15-30 min in ABS, then proceed to Antibody Staining
Fixation with -20°C Methanol
(Good for microtubules and most proteins)
1. Wash coverslips 2x with 1xPBS
2. Remove all PBS (but do not allow the cells to dry), and immediately add enough -20°C methanol to cover the coverslips. About 2-3ml if using a 6 well plate.
3. Put the plate in a -20°C freezer for 5 min (NB: can be stored for weeks as long as you keep the coverslips covered in MeOH).
4. Remove coverslips from MeOH, and rehydrate them in 1xPBS with 0.1-0.5% Triton-X-100 for 10-20 min
5. Block coverslips for 15-30 min in ABS, then proceed to Antibody Staining.
1. Incubate with ABS for 15-30 min at RT.
2. Incubate with 1°Ab diluted in Cell Blocking solution in moist chamber.
3. Wash 3x 1X PBS-Tween for 5 min at RT.
4. Incubate in 2°Ab +DAPI in PBS-T in moist chamber.
5. Wash 3x 1X PBS-T 5 min at RT.
6 . Mount with Prolong Gold or Mowiol mounting medium on clean glass slide.
Mowiol 4-88 Mounting Medium
Mowiol 4-88 is a high-quality mounting medium with good anti-fade characteristics. It hardens and matches the refractive index of immersion oil, and thus is particularly suited for this form of microscopy. Additional anti-fade (DABCO) is added to further retard photobleaching.
Mowiol 4-88 (Calbiochem; 475904), DABCO (Sigma; D-2522)
1. Add 2.4g Mowiol to 6g glycerol and stir briefly with a pipette.
2. Add 12ml dH2O and stir at room temp for several hours or overnight.
3. Add 12ml 0.2M Tris (pH 8.5) and heat to 50oC for 1-2 hrs while stirring.
4. When the Mowiol has dissolved, clarify by centrifugation @ 500 x g for 15mins.
5. Add DABCO to 2.5% (0.72g), aliquot and store at -20oC. Bubbles can be removed by centrifugation. Aliquots can be stored for up to 2 weeks at 4°C or frozen to -20°C for months