Staining
Fixing the Cells
Formaldehyde Fixation for Non-Adherent Cells
1. Coat coverslips with an excess of poly-L-lysine (0.01% solution, Sigma catalog #P-4707) for 10 minutes at room temperature.
2. Aspirate the poly-L-lysine solution and allow coverslips to dry completely.
3. Transfer cells in medium to 50 ml tubes.
4. Centrifuge (400 x g, 15 degrees C) for 5 minutes.
5. Aspirate the medium and resuspend the cells in 30 ml PBS.
6. Cover the dried, treated coverslips with the cell suspension.
7. Incubate at room temperature for 10-15 minutes.
8. Aspirate excess cell suspension.
9. Continue with cell fixation.
10. Grow cells on glass coverslips.
11. Rinse briefly in phosphate-buffered saline (PBS).
12. Immerse for 10 minutes in 3.7% formaldehyde solution (for 100 ml: 10 ml 10X PBS, 33.4 ml of 11.1% formaldehyde, 0.6 ml of 30% Triton-X, 56 ml distilled water). The solution is stable for 1 week at 4 degrees C. Formaldehyde should be freshly prepared from its para-polymer. It is toxic and should be handled appropriately in a fume hood.
Formaldehyde Fixation for Adherent Cells
1. Grow cells on glass coverslips.
2. Rinse briefly in phosphate buffered saline (PBS).
3. Immerse for 10 minutes in 3.7% formaldehyde solution (for 100 ml: 10 ml 10X PBS, 33.4 ml of 11.1% formaldehyde, 0.6 ml of 30% Triton-X, 56 ml distilled water). The solution is stable for 1 week at 4 degrees C. Formaldehyde should be freshly prepared from its para-polymer. It is toxic and should be handled appropriately in a fume hood.
Methanol/Acetone Fixation for Non-Adherent Cells
1. Coat coverslips with an excess of poly-L-lysine (0.01% solution, Sigma catalog #P-4707) for 10 minutes at room temperature.
2. Aspirate the poly-L-lysine solution and allow coverslips to dry completely.
3. Transfer cells in medium to 50 ml tubes.
4. Centrifuge (400 x g, 15 degrees C) for 5 minutes.
5. Aspirate the medium and resuspend the cells in 30 ml PBS.
6. Cover the dried, treated coverslips with the cell suspension.
7. Incubate at room temperature for 10-15 minutes.
8. Aspirate excess cell suspension.
9. Continue with cell fixation.
10. Grow cells on glass coverslips.
11. Immerse coverslips in ice-cold methanol:acetone (1:1), incubate at -20 degrees C for 10 minutes.
12. Air dry coverslips.
Methanol/Acetone Fixation for Adherent Cells
1. Grow cells on glass coverslips.
2. Immerse coverslips in ice-cold methanol:acetone (1:1), incubate at -20 degrees C for 10 minutes.
2. Air dry coverslips.
Blocking
Place the coverslips cell-side-up on a piece of filter paper inside a petri dish. Cover the cells with blocking buffer (1% BSA in PBS) for 10 minutes to minimize non-specific absorption of the antibodies to the coverslip (25-50 ul is usually sufficient).
Incubation with Primary Antibody
1. Remove the blocking buffer by holding each coverslip on its edge with forceps and draining it onto a sheet of fiber-free paper.
2. Dilute primary antibody to 1.0-10 ug/ml in blocking buffer. (The concentration of antibody to be used will depend on several variables, such as the affinity of the antibody and the abundance of the antigen.)
3. Distribute 4-40 ul of the primary antibody on each coverslip and incubate for 45 minutes at room temperature.
Incubation with more than one Primary Antibody
Very often, it is desirable to examine the co-distribution of two different antigens in the same cell. For this reason, a double immuno-fluorescence procedure may be used in which cells are incubated simultaneously with two separate primary antibodies. For this procedure, the antibodies must be raised in different species and must be monospecific.
1. Incubate coverslips with a mixture of the diluted primary antibodies for 45 minutes at room temperature.
2. Decant the primary antibodies solution or remove it by aspiration.
3. Wash coverslips three times in PBS, 5 minutes each wash.
Incubation with Secondary Antibodies
Note: If the primary antibodies are already conjugated to a flurochrome, incubation with secondary antibody is unnecessary. The coverslips are now incubated with secondary antibodies conjugated to a fluorochrome (e.g. anti-mouse IgG:FITCor, anti-rabbit IgG:TRITC, depending on the donor species of the primary antibody. We recommend the use of cross-absorbed and affinity-purified secondary antibodies to minimize background and non-specific reactivity from the secondary antibodies. High-quality conjugated antibodies are essential for the avoidance of cross reactivity between two different antibodies in double immunofluorescence protocols).
1. Place coverslips cells-side up on a piece of filter paper inside a petri dish.
2. Dilute the secondary antibodies to the appropriate concentration in blocking buffer. Add enough secondary antibody solution to cover the surface of each coverslip (usually 10-25 ul).
3. Incubate for 30 minutes at room temperature.
4. Remove the secondary antibody by blotting the edge of each coverslip on fiber-free paper.
5. Wash coverslips three times in PBS, 5 minutes each wash.
Preparation for Microscopy
1. Invert each coverslip onto a slide containing 10 ul of mounting media (PBS pH 8.0. 50% glycerol, 0.1% p-phenylenediamine dihydrochloride).
2. Remove the excess mounting media with fiber-free paper without disturbing the coverslip. Seal the edges of each coverslip with regular transparent nail polish and allow to dry for 3 minutes. This will provide semi-permanent preparations. The cells are now ready for microscopic viewing.
In-situ Staining for beta-galactosidase Activity
1. Wash cells 2 times with 1-2 ml PBS.
2. Fix cells with 4% paraformaldehyde for 15 minutes at room temperature.
3. Rinse 3 times with 1-2 ml PBS.
4. Add 1 ml X-gal solution (0.2% x-gal, x-gal stock is made 2% in DMF; 2 mM MgCl2; 5 mM K4Fe(CN)6.3H2O; 5 mM K3Fe(CN)6; solution is in 1x PBS) and incubate at 37C.
5. Observe under light microscope after 4-6 hours. Shorter incubator is okay if transfection efficiency is good. Overnight if efficiency is very low.
Staining Methods for Cell Death
I. The simplest way: trypan blue.
Dead cells stain blue.
II. Non-fixed cells: FDA (fluorescein diacetate) - green, alive cells; P.I. (propidium iodide) - red, dead cells
For 35 mm plates:
1. To 2 ml medium, add 1-2 ul mg/ml P.I. and 6 ul 5 mg/ml FDA.
2. Incubate at room temperature for 3-10 minutes.
3. Examine cells under the scope.
Note:
1. Can add 0.5-1 ug/ml P.I to the culture medium at the time of stimulation. P.I. is not toxic at this concentration to cultured neurons.
2. If the P.I. staining is not strong enough to be picked up easily under your scope, use 2 X p.I.
3. After staining, need to examine the staining right away. Otherwise the green staining get diffused. You can leave cells at 4 degrees C for a few hours or overnight to slow down the diffusion (I have tried 3T3, do not know if it works for neurons - probably not for DIV 7 cortical neurons. PBS lysed neurons).
4. After fixation, the PI staining is lost gradually, even after mounting.
5. This method stains for non-fixed cells.
6. P.I.: Sigma, dissolve in PBS; FDA: Sigma, dissolve in acetone
III. P.I. Staining for Fixed Cells
1. Fixation:
a. EtOH fixation - gives brighter P.I. staining: gently overlay over media 4X media vol of EtOH precooled to -20 degrees C, room temperature for 3 minutes, gently mix media and EtOH with pipet, room temperature 5 minutes
b. Paraformaldehyde fixation (4% paraformaldehyde/4% sucrose in PBS, pH 7.2-7.6). Gently overlay over media 2x media vol. of 8% formaldehyde/8% sucrose in PBS, gently tilt the plates to mix; or remove culture medium, fix cells with 4% paraformaldehyde/4% sucrose in room temperature for 15 minutes.
2. Aspirate off media
3. Staining: 4 ug/ul P.I./0.1% triton X-100/0.5 mg/ml RNaseA in PBS, room temperature for 5 minutes, then examine under the scope or mount with coverslips.
Note:
1. P.I. will stain for both DNA and RNA. It is critical to include RNaseA to eliminate the cytosolic RNA staining background. If use EtOH fixation, it is less critical to include RNaseA in staining solution.
2. This will stain both alive and dead cells. Alive cells should have evenly stained nuclei. Nuclei from apoptotic cells show condensed, or fragmented morphology. Can not distinguish necrosis.
IV. Hoescht Staining
1. Fix cells:
a. Remove media, fix with 4% paraformaldehyde/4% sucrose in PBS, neutral pH, room temperature for 15-45 minutes.
b. If cells are not adhering well to the plates: gently overlay over media 2X media vol. of 8% paraformaldehyde/4% sucrose in PBS, pH 7.2-7.6, gently tilt the plates to mix; room temperature for 15 minutes.
2. Stain cells with 2.5 ug/ml Hoeschst 33258 in PBS/0.1% triton X-100., room temperature for 5 minutes.
3. Mount with coverslips. Examine cells under fluorescence scope using DAPI filter.
Note:
1. Alive cells should have evenly stained nuclei. Nuclei from apoptotic cells show condensed, or fragmented morphology.
2. Hoeschst 33258, sigma B-2883 (bis-Benzimide), 5 mg/ml in H2O stock. Light sensitive. Non-fixed cells.
3. Hoeschst 33258 stains permeablized cells. Hoeschst 33342 (bis-Benzimide, Sigma B2261) is permable, can stain both fixed and non-fixed cells.
DAB
1. PBS/NAF (NAF should be protected from light).
2. NaBH4 1% (5 minutes or 10 minutes).
3. PBS
4. Block, 10% EtOH 1% H2O in PBS, 2 times for 15 minutes each.
5. PBS rinse.
6. PBS, 0.2% Triton, 0.2 Gelatin dilution buffer.
7. Blocking, PBSTG/10% serum (secondary antibody goat, 2% BSA, 0.1 M glycine) for 1-2 hours at room temperature
8. Primary in blocking
9. Incubate for 24-48 hours in 4 degrees C
10. Rinse four times for 15 minutes each.
11. Incubate secondary antibody (1:250) in blocking solution for several hours at room temperature.
12. Rinse four times.
13. Dilute A, B solution in blocking (1:250). (Preincubate in dark for 30 minutes.) Incubate for 1-1.5 hours.
14. Rinse in PBSTG.
15. Rinse in PBS twice for 10 minutes each.
16. 0.1 M Tris (pH 7.5) twice for 5 minutes each.
17. DAB development: 0.05% DAB in 0.1 M Tris
18. Optional: Add 0.0125 NiSO4 (4% in H2O stock) for 5 minutes.
19. Stop by adding ice-cold 0.1 Tris (pH 7.5)
20. Rinse with Tris.
21. Mount.
22. Dry overnight.
23. Dehydrate in EtOH, from 70% to 95% to 100% and 100% again. 5 minutes each step.
24. Put 2x xylene.
25. Mount in DPX, coverslip.
Keywords.
Adenylyl cyclase (AC), amyloid precursor protein (APP), Alzheimer’s disease (AD), brain derived neurotrophic factor (BDNF), cAMP, cAMP-responsive element (CRE), CREB, extra-cellular regulated protein kinase (ERK), FE65, long-term potentiation (LTP), long-term depression (LTD), memory formation, neuroplasticity, local protein synthesis.