Staining

Actin Staining

    1. Fix cells in 70% acetone/30% methanol for 5 minutes (~1ml per well)*

    1. Aspirate fixant, then permeabilize/block with (~1ml per well) 15 minutes:

      1. 3% BSA

      2. 0.1% Tween-20

    1. Rinse 3X with PBS for 5 minutes each

    1. Incubate coverslip upside down onto 1 mg/ml fluorescently labeled phalloidin for 30 min (use about 50mL of antibody solution per coverslip)

    1. Rinse 3X with PBS for 5min. each (~1mL per well)

    1. Add 1mg/mL DAPI (in methanol) for 5 min. (~1mL per well)

    1. Rinse briefly with ddH2O

    1. Dry in coldroom for ~15 minutes to get rid of excess moisture

    1. Coverslips: mount on slides (upside-down) onto Gel-mount w/DABCO

Hydrogels: store in water in fridge (cover and wrap in parafilm)

    1. visualize

* Leave coverslips right-side up in well plates except in the antibody incubations

Antibody Staining (General)

    1. fix in 70% acetone/30% methanol for 5 minutes (~1ml per well)*

    1. aspirate fixant, then permeabilize/block with (~1ml per well) 15 minutes:

      1. 3% BSA

      2. 0.1% Tween-20

    1. rinse 3X with PBS for 5 minutes each

    1. incubate coverslip upside down onto 1° antibody for 1 hour in humidity chamber (use about 50mL of antibody solution per coverslip)

    1. rinse 3X with PBS for 5min. each (~1mL per well)

    1. incubate coverslip as in #4 with 2° antibody (1 hour)

    1. rinse 3X with PBS (~1mL) for 5 min. each

    1. add 1mg/mL DAPI (in methanol) for 5 min. (~1mL per well)

    1. rinse briefly with ddH2O

    1. dry in coldroom for ~15 minutes to get rid of excess moisture

    1. Coverslips: mount on slides (upside-down) onto Gel-mount w/DABCO

Hydrogels: store in water in fridge (cover and wrap in parafilm)

    1. visualize

* Leave coverslips right-side up in well plates except in the antibody incubations

LIVE/DEAD Assay (Molecular Probes)

1) Prepare Dye Solution

a. For 1 ml in PBS (or any other phenol red free cell compatible solution) add:

i. 0.5 μl Calcein

ii. 2.0 μl Ethidium Homodimer

2) Add enough dye solution to completely immerse construct or cells

3) Incubate (37°C) for 10 minutes

4) Rinse sample with PBS to remove excess dye

5) Mount cells with Fluromount mounting solution

6) View sample with fluorescence – View live cells under blue excitation (green emission), dead cells under green excitation (red emission)

CFSE Staining

1. Pellet Cells

2. Using 10x CFSE solution, add 1mL 1X CFSE (0.9 mL PBS and 0.1 mL 10X CFSE)

3. Incubate at 37 deg C for 15 min

4. Add a few mLs of PBS

5. Pellet

6. Resuspend in desired volume of media

MSB (Martius Scarlet Blue)(after Lendrum17)

http://www.histosearch.com/histonet/Jun03/MartiusscarletbluestainfoA.html

Three dyes of different molecular sizes are used in this method to selectively stain connective tissues. A yellow dye, of small molecular size, in the presence of alcoholic phosphotungstic acid, selectively stains red blood cells and sometimes early deposits of fibrin. A red, intermediate sized dye molecule is then used to selectively stain muscle and mature fibrin. Staining of collagen is prevented by treatment of the section with aqueous phosphotungstic acid which removes red staining from collagen after staining in the red dye. A large molecular size blue dye is then used to stain collagen and old fibrin.

SECTION PREPARATION:

Paraffin sections are cut at 3 to 5 mm from tissue fixed in neutral buffered formalin. A control section from foetal lung in hyaline membrane disease, should be included.

REAGENTS REQUIRED:

Martius yellow solution

Martius yellow (CI 10315) 0.5 g

Absolute ethyl alcohol 95 ml

Phosphotungstic acid 2 g

Distilled water 5 ml

Dissolve the dye in absolute ethanol and the phosphotungstic acid in distilled water. Combine the two solutions.

Crystal ponceau solution

Crystal ponceau 6R (CI 16250) 1 g

Distilled water 97.5 ml

Glacial acetic acid 2.5 ml

Aniline blue solution

Aniline blue (CI 42755) 0.5 g

Distilled water 99 ml

Glacial acetic acid 1 ml

1% acetic acid

Glacial acetic acid 1 ml

Distilled water 99 ml

1% aqueous phosphotungstic acid

Phosphotungstic acid 1 g

Distilled water 100 ml

Weigert’s haematoxylin

METHOD:

    1. Dewax and rehydrate sections.

    2. Stain with Weigert's haematoxylin for 5 minutes.

    3. Wash in tap water.

    4. Differentiate, if necessary, in acid alcohol then blue in running tap water (or suitable alternative) for 5 minutes.

    5. Rinse in 95% ethanol.

    6. Stain with martius yellow for 3 minutes.

    7. Rinse in distilled water.

    8. Stain in crystal ponceau 2R solution for 10 minutes.

    9. Drain the stain from the slide.

    10. Mordant and differentiate with 1% phosphotungstic acid for 5 minutes.

    11. Rinse in distilled water.

    12. Stain in aniline blue for 1 minute.

    13. Rinse briefly in 1% acetic acid.

    14. Dehydrate, clear and mount with a neutral mounting medium.

General Antibody staining using an HRP-conjugated secondary

Materials

    • 10% formalin

    • PBS

    • FBS or BSA

    • 3% H2O2

    • 0.1% Triton-X in PBS

    • primary antibody

    • secondary antibody (HRP-conjugated)

    • AEC (or other suitable HRP chromogen)

    • TBS (0.05 M Tris-HCl, 0.15 mM NaCl, pH 7.6)

    • 37 mM NH4OH

Methods

1. Remove media and rinse cells with PBS

2. Incubate cells with 10% formalin at room temp for 10 minutes

3. Rinse 2x with PBS

4. Permeabilize fixed cells by incubating in Triton-X solution for 2 minutes at room temp

5. Rinse 4x with PBS over 5 minutes

6. Remove PBS and incubate cells in H2O2 for 5 minutes (this quenches endogenous peroxidase activity of the cells)

7. Rinse with diH2O, then incubate in TBS for 5 minutes

8. Add 1° antibody (i.e. anti-PCNA), leaving 2 wells untreated. Incubate for 60 minutes at room temp in a humidified chamber

9. Rinse 3x with PBS over 5 minutes

10. Add 2° antibody (i.e. anti-mouse IgG HRP), leaving 1 previously untreated well untreated. Incubate for 40 minutes at room temp in a humidified chamber

11. Rinse 3x with PBS over 5 minutes

12. Incubate all wells with AEC solution for 5-15 minutes

13. Rinse with diH2O

14. Counterstain with Mayer’s hematoxylin

a. Cover cells with hematoxylin and incubate for 5 minutes

b. Rinse with diH2O

c. Add NH4OH sol’n and incubate for 1 minute. Remove and repeat until stain turns blue.

d. Rinse with diH2O

15. Mount with aqueous mounting medium

Notes:

· Make all antibody solutions in 3% BSA or FBS in PBS (blocking buffer).

· This protocol allows for two staining controls: a) one well that receives no 1°, but has 2° and AEC, and b) one well that receives neither 1° nor 2°, but does have AEC.

· Staining control wells should be kept in blocking buffer when other wells contain antibody solutions.

· Antibody dilutions are not given here, as they will vary with the antibody source.

· Aqueous mounting media cannot be used with some chromogen solutions.

Staining ES cells for SSEA-1 – Method 1: 6-Well Plate

    1. Coat wells in 6-well dish with gelatin.

    2. Plate cells onto each well. Allow cells to adhere overnight.

    3. Once the cells have adhered, suck off the media, being careful not to disturb the adhered cells.

    4. Add a few drops of 4% paraformaldehyde to fix the cells.

    5. Incubate in a humidified chamber 30-60 minutes (ie wet paper towels).

    6. Prepare 1X PBS. (Add 200 ml 10X PBS to 1800 ml distilled water).

    7. Suck off remaining paraformaldehyde.

    8. Wash wells with 1X PBS twice.

    9. Dilute 1˚ antibody (SSEA-1, or MC-480) 1:10 in DAKO antibody diluent or in 1% BSA in PBS.

    10. Add enough 1˚ antibody to coat surface.

    11. Incubate in a humidified chamber 30-60 minutes.

    12. Suck off remaining 1˚ antibody.

    13. Wash in 1X PBS twice.

    14. Dilute 2˚ antibody (anti-mouse IgM) 2:1000 in DAKO antibody diluent or in 1% BSA in PBS.

    15. Add a few stops of 2˚ antibody to coat slide surface.

    16. Incubate in a humidified chamber 30-60 minutes.

    17. Suck off remaining 2˚ antibody.

    18. Wash in 1X PBS.

    19. Wash in distilled water.

    20. Use Fluoromount-G to mount coverslip.

    21. Visualize under fluorescence microscope

Staining ES cells for SSEA-1 – Method 2: Glass Slides

    1. Coat center of slides with gelatin.

    2. Plate cells onto each slide. Allow cells to adhere overnight.

    3. Once the cells have adhered, carefully remove excess media with Kim wipe.

    4. Add a few drops of 4% paraformaldehyde to fix the cells.

    5. Incubate slides in a humidified chamber 30-60 minutes (ie wet paper towels).

    6. Prepare 1X PBS. (Add 200 ml 10X PBS to 1800 ml distilled water).

    7. Remove remaining paraformaldehyde.

    8. Wash slides with 1X PBS twice.

    9. Dilute 1˚ antibody (SSEA-1, or MC-480) 1:10 in DAKO antibody diluent or in 1% BSA in PBS.

    10. Add a few stops of 1˚ antibody to coat slide surface.

    11. Incubate slides in a humidified chamber 30-60 minutes.

    12. Remove remaining 1˚ antibody.

    13. Wash in 1X PBS twice.

    14. Dilute 2˚ antibody (anti-mouse IgM) 2:1000 in DAKO antibody diluent or in 1% BSA in PBS.

    15. Add a few stops of 2˚ antibody to coat slide surface.

    16. Incubate slides in a humidified chamber 30-60 minutes.

    17. Remove remaining 2˚ antibody.

    18. Wash in 1X PBS.

    19. Dehydrate slides in 70% ethanol (3 min), 95% ethanol (3 min), 100% ethanol (3 min), xylene (at least 5 min).

    20. Mount slides by placing several drops of cytoseal onto cells. Apply coverslips and remove bubbles by pressing on coverslips.

    21. Visualize under fluorescence microscope

Immunofluorescent Staining (and DAPI)

Materials

    • 4% formaldehyde

    • PBS

    • Blocking buffer (0.1% Tween-20 in PBS)

    • 0.1% Triton-X in PBS

    • primary antibody

    • secondary antibody (fluorescent conjugate)

    • DAPI: diluted 1:100 or 1 ug/ml in PBS

    • Fluromount mounting solution

Methods

    1. (Optional) Tether cells to coverslip

    2. Remove media and rinse cells with PBS

    3. Incubate cells with 4% formaldehyde at room temp for 10 minutes

    4. Rinse 2x with PBS

    5. Permeabilize fixed cells by incubating in 0.1% Triton-X solution for 2 minutes at room temp

    6. Rinse 4x with PBS over 5 minutes

    7. Add 1° antibody to wells or invert coverslip (cell side down) over large drop of 1° antibody on Parafilm..

    8. Incubate for 60 minutes at room temp in large slide box with moist Kimwipe.

    9. Rinse 3x with PBS over 5 minutes

    10. Add 2° antibody to all wells or invert coverslip (cell side down) over large drop of 2° antibody on Parafilm.

    11. Incubate for 30 minutes at room temp inlarge slide box with moist Kimwipe

    12. Rinse 3x with PBS over 5 minutes

    13. Counterstain with DAPI diluted 1:100 (or 1µg/mL in PBS)

    14. Incubate for 1-5 minutes

    15. Rinse several times with PBS

    16. Rinse 1x with DI H2O

    17. Mount with Fluromount mounting solution

Notes:

· Make all antibody solutions in blocking buffer

· Staining control wells should be kept in blocking buffer when other wells contain antibody solutions.

· Antibody dilutions are not given here, as they will vary with the antibody source.

Suggested Antibody and Stain Dilutions

Stains

DAPI (from working stock) 1:100

Fluorescent Phalloidin 1:100

Antibodies

1˚ SSEA 1:20

2˚ IgM-PE 1:500

1˚ Troma-1 1:10

2˚ Texas red IgG donkey-a-rat 1:100

IgG-PE goat-a-rat 1:100

IgG2a-PE goat-a-mouse 1:100

Primary Antibodies

Different combinations:

1) DAPI / CK / Insulin

2) DAPI / CK / AFP

3) DAPI / CK / Abumin

4) DAPI / Insulin / Glucagon

5) DAPI / Insulin/ PP

6) DAPI / Insulin / Somatostatin

7) DAPI / Nestin / Insulin

8) DAPI / Nestin / Albumin

9) DAPI / Nestin / Glucagon

10) DAPI / Nestin / AFP

11) DAPI / Insulin / PDX

12) DAPI / Insulin / Shh or FoxA2 or Pax6

· Crossed out antibodies are not required for the experiments

· ** indicates must be bought * indicates may be needed for secondary experiments

· # obtain from Melton