Carmine Staining

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Carmine stain preparation

2.5g Alum potassium sulfate

1.0g Carmine (C-6152 Sigma)

to 500mLs dH2O

Boil for at least 40 minutes and keep hot.  Filter through Whatman #1 paper and adjust to final volume of 500mLs.  There will be a lot of stain that doesn’t dissolve.  This can be minimized by keeping the solution at boiling while filtering a small amount at a time.

I store this in the refrigerator.  The stain can be re-used several times, when the stain gets depleted it will take longer to get good staining.  I re-filter after each use—but again only through Whatman #1

Whole mount preparation

Collect the fat pads and spread on a clean glass slide.  Allow to air dry a minute or two and then immerse in Formalin.

Fixing and Staining

  1. Fix glands in formalin overnight.  Wash in dH2O for 5 minutes.
  2. Stain in Carmine for 1-3 days.  You can see when the stain has penetrated through by checking the back of the fat pads through the slide.  Fatter glands may take longer than 3 days.
  3. Dehydrate in 70% EtOH overnight.  Wash in 95% EtOH 60 minutes.  Wash in 100% EtOH 60 minutes.
  4. Transfer to Xylenes overnight to clear the gland.  Fatter glands may take a few days to clear.
  5. Dip into 100% EtOH to remove xylenes
  6. Transfer to glycerol.  Store the glands in glycerol in open coplin jars in the hood until the xylenes are truly gone (a couple of days).

Steps that can be more than one day (i.e. over the weekend):

  1. Carmine
  2. 70% EtOH dehydration
  3. Xylenes

Carmine Stain Protocol (adapted from rat procedure by Amy Moser by Lisa Arendt)

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Lisa’s Carmine Stain Protocol

Carmine stain preparation

  • 2.5 g Alum potassium sulfate
  • 1.0 g Carmine (Sigma C-6152)
  • to 500 mls dH20
  • Boil for at least 40 minutes and keep hot. Filter through Whatman #1 paper and adjust to final volume of 500 mls. There will be a lot of stain that doesn’t dissolve.  This can be minimized by keeping the solution boiling while filtering a small amount at a time.

Fixing and staining

  1. Spread glands on a glass slide and allow to sit for ~10 min or so until they become ‘stuck’ to the slide.  All fixing and staining procedures can then be done in a coplin jar. Alternatively, flatten glands between glass slides during formalin fixation (do not allow to stick to glass) and then do all staining procedures with glands in tissue cassettes.
  2. Fix glands in 1:3 glacial acetic acid: 100% EtOH for 1 hour or formalin fixation overnight is fine.
  3. Transfer to 70% EtOH for 15 minutes followed by a short rinse in 50% EtOH (5 min) then dH20. (If fixing in formalin, a wash in dH20 should suffice). Following formalin fixing, glands can stay in 70% EtOH until you are ready to stain them.
  4. Stain in carmine for 1-3 days.  Fatter glands may take more than 3 days.
  5. Dehydrate in 70% EtOH overnight, then 90% and 100% EtOH with 60 minute washes.
  6. Transfer to xylenes overnight to clear the fat from the gland. Fatter glands may take a few days to clear.
  7. Wash with 100% EtOH to remove the xylenes.
  8. Transfer to glycerol (or mineral oil or methylsalicylate) in a clean coplin jar. Store the glands in glycerol in open coplin jars in the hood for a few days until the xylenes are truly gone (a couple of days).
  9. If the carmine is too dark, the glands can be rehydrated through graded alcohols to 70% EtOH and remain until some of the stain precipitates out of the glands.
  10. Visualize glands with coverslips over the tissue, use additional glycerol to fill the spaces around the glands (note, this will be a bit messy…make sure to clean up the microscope after use!).  If wanted, glands can be mounted with permount for long term storage.
  11. To submit for sectioning, scrape gently off of the slides and transfer to tissue cassettes for processing.

Hematoxylin Mammary Gland Whole Mount Protocol

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Mammary Gland Whole Mount Protocol

Inject animals with BrdU 2 hours before harvesting tissue (0.1cc per 10 g animal weight)
• Fix tissue for 2 hours in fresh, cold 4% paraformaldehyde (PFA) in ice.
• Rinse tissue in 70% ethanol until ready to stain (up to 2 weeks).
• All the following steps are conducted by submerging tissue-cassettes containing glands into the solution. Thus, many glands can be processed in one jar at once.

• Defat:
Acetone 30 min
Acetone 30 min
Acetone 30 min

Note: This step prohibits later immunohistochemical analysis by creating high background. If you want to do immunohistochemistry on these glands, skip this step.

• Rehydrate:
100% EtOH 30 min
95% EtOH 30 min

• Stain:

Hematoxylin stain:
0.13 g FeCl3·6H20
13.5 ml distilled water
Dissolve, and add 1.74 mL stock (10%) Harris hematoxylin *
Add 200 ml 95% ethanol
Adjust pH to 1.25 with concentrated (12N) HCl (pH is critical)
Make stain fresh each time, and check that stain turns blue by putting a tiny amount in a weigh boat and running under crude tap water. Stain should turn bright light blue. If not, throw out and try again.
Place glands in hematoxylin stain for 1 1/2 hours until whole gland looks purple.
If you did not defat in acetone (for future immunohistochemistry), stain O/N to allow hemotoxylin to permeate through fat. Monitor stain by holding gland up to the light.

• Rinse:
Crude tap H20 rinse (whole gland will turn blue)
Crude tap H20 overnight
Distilled tap H20 rinse

• Destain:
200 ml 50% EtOH + 416 µl 12N HCl for 30 min
Monitor destaining by holding gland up to the light. You should be able to clearly see blue epithelium with a fairly clear background.

• Dehydrate:
70% EtOH 1 hour
95% EtOH 1 hour
100% EtOH overnight
Complete dehydration of gland is critical for complete clearing. Incomplete dehydration will result in brown background in the fat. Use fresh 95% and 100% EtOH and don’t take shortcuts.

• Clear:
BABB 1-2 hours
BA = benzylalcohol
BB = benzylbenzoate
Mix these chemicals 1:2 (BA:BB)

Note: leaving glands in BABB for extended time results in the lightening of the hematoxylin stain.

Note: Xylenes or Histoclear can also be used for clearing glands. We have better luck with BABB, however.

For imaging, remove gland from tissue-cassette and press between two glass slides to flatten.

For long-term storage, transfer to methyl salicylate in glass scintiallation vials and KEEP DARK.
(If exposed to light, the blue will fade to brown)

* STOCK HEMATOXYLIN (10%): Harris hematoxylin (Fisher, powder)
Add 10g of powder to 100 ml of 95% ethanol, leave overnight stirring and will go into solution. Keep covered in foil. Let sit at least 3 weeks to 1 month for best stain.

IF Cytospun Slides (PJ Keller)

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IF Cytospun Slides

A.  Cytospin

-Spin cells at 20,000 to 50,000 cells per spot in 100-150 ml of media (preferably + at least 10% serum)

-Load 150-250 ml of sample into the chamber, spin at 500 rpm for 1’  with medium acceleration

-Fix spots in 100% ice cold methanol at –20 for 10’

-Air dry spots and freeze at –80 until use (better if used within 1-2 months, though I have stained years old slides as well with success)

B.  Immunofluorescence

  1. Allow slides to thaw at RT, dry any condensation with a kimwipe (avoiding spots, of course).  Circle spots with a pap pen.
  2. Permeabilize cells with 0.1% Triton-X 100 in PBS
    1. Add 150 ml per spot
    2. Incubate at RT 10’
    3. Wash 3X5’ in a coplin jar filled with PBS
  3. Block cells 1 hour in PBS + 1% BSA + 2% goat serum (for Alexa-fluor secondary antibodies)
    1. Add 150 ml per spot
    2. Incubate at RT 1 hour in a humidified chamber
  4. Add primary antibodies overnight at 4C or 1 hour at RT
    1. For Rabbit aCK14 use at 1:500 in PBS + 1% BSA
    2. For Mouse aCK18 use at 1:500 in PBS + 1% BSA
    3. Prepare a solution of both antibodies at 1:500 to double stain
    4. Add 150 ml per spot and incubate in a humidified chamber
  5. Wash slides 3×5’ in PBS to remove residual primary antibody
  6. Add secondary antibodies for 1 hour at RT
    1. Use goat anti-Rabbit or goat anti-Mouse Alexa fluor conjugated antibodies (Alexa 488 is green, Alexa 555 or 546 are red-orange)
    2. Prepare secondaries at 1:500 in PBS + 1% BSA (prepare a solution of both mouse and rabbit secondary antibodies to double stain)
    3. Add 150 ml per spot and incubate in a humidified chamber (in the dark!)
    4. Wash 3×5’ with PBS in the dark
  7. DAPI counter stain the nuclei
    1. Dilute 25 mg/ml stock in 1:100 PBS (i.e. 1 ml PBS + 10 ml DAPI)
    2. Add 150 ml per spot
    3. Incubate 5’ at RT in the dark, rinse with PBS 1X
  8. Mount slides with SlowFade kit and coverslip
    1. Blot excess PBS, add 1 drop equiibration buffer per spot
    2. Incubate at RT 5’, blot excess from the edge of the slide
    3. Add 1 drop slow fade solution per spot/well
    4. Cover with coverslips and seal with nail polish, store at 4C or –20 for longer term
    5. Clean backs of slides with windex before viewing

IF on Chamber Slides (PJ Keller)

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IF on Chamber Slides

A.  Culture

-Plate 5000-10,000 cells per well (8 or 4-well chamber slides, respectively) in growth media

-Allow cells to grow until 50-70% confluent

B.  Immunofluorescence (note: smaller volumes are recommended for 8-well chamber slides, larger for 4-well; regardless of the exact amount put on the cells, make sure that there is enough liquid to cover the cells completely as drying out can lead to high background staining)

  1. Fix cells in Methanol
    1. Aspirate media from cells
    2. Wash wells with PBS 1X
    3. Add 250-500 µl cold methanol
    4. Incubate at –20 C for 10 min
    5. Wash wells with PBS 1X
  2. Permeabilize cells with 0.1% Triton-X 100 in PBS
    1. Add 150 to 250 ml per well
    2. Incubate at RT 10’
    3. Wash 3X5’ with PBS
  3. Block cells 1 hour in PBS + 1% BSA + 2% goat serum (for Alexa-fluor secondary antibodies)
    1. Add 150 to 250 ml per spot
    2. Incubate at RT 1 hour in a humidified chamber
  4. Add primary antibodies overnight at 4C or 1 hour at RT
    1. For Rabbit aCK14 use at 1:500 in PBS + 1% BSA
    2. For Mouse aCK18 use at 1:500 in PBS + 1% BSA
    3. Prepare a solution of both antibodies at 1:500 to double stain
    4. Add 150 to 250 ml per well and incubate in a humidified chamber
  5. Wash slides 3×5’ in PBS to remove residual primary antibody
  6. Add secondary antibodies for 1 hour at RT
    1. Use goat anti-Rabbit or goat anti-Mouse Alexa fluor conjugated antibodies (Alexa 488 is green, Alexa 555 or 546 are red-orange)
    2. Prepare secondaries at 1:500 in PBS + 1% BSA (prepare a solution of both mouse and rabbit secondary antibodies to double stain)
    3. Add 150 to 250 ml per spot and incubate in a humidified chamber (in the dark!)
    4. Wash 3×5’ with PBS in the dark
  7. DAPI counter stain the nuclei
    1. Dilute 25 mg/ml stock in 1:100 PBS (i.e. 1 ml PBS + 10 ml DAPI)
    2. Add 150 to 250 ml per spot
    3. Incubate 5’ at RT in the dark, rinse with PBS 1X
  8. Mount slides with SlowFade kit and coverslip
    1. Remove the chambers with the provided tools
    2. Blot excess PBS, add 1 drop equilibration buffer per well
    3. Incubate at RT 5’, blot excess from the edge of the slide
    4. Add 1 drop slow fade solution per well
    5. Cover with coverslips and seal with nail polish, store at 4C or –20 for longer term