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In Situ Hybridization on Mammary Sections Using DIG Probes

From Jimmie Fata 2/18/03


In Situ Hybridization Method on Mammary Sections Using DIG Probes


Day 1


1.                  Dewax and Rehydrate paraffin embedded sections[1]


a.     2 X 2 minutes toluene[2]

b.     2 min 100% ETOH

c.      2 min in 95% ETOH

d.     2 min in 80% ETOH

e.     2 min in 70% ETOH

f.        2 X 2 min in 2X SSC buffer


2.                  Proteinase K Digestion[3]


a.     Pro K treatment @ RT for 10 min

b.     Wash Slides 2 X 5 min in 2X SSC buffer


3.                  Acetic Anhydride Treatment[4]


a.     Acetic Anhydride treatment @ RT for 10 min (add A.A. just before adding slides to the wheaton jar and put stir bar at bottom)

b.     Wash Slides 2 X 5 min in 2X SSC buffer


4.                  Prehybridization[5]


a.     Use a wax pen to delineate mammary gland on section

b.     Prehybridize in prewarmed Pre-Hyb. Buffer (100ml) for 2-4 hours @ 50 degrees C in a humidified chamber that has paper towels soaked in 50% formamide in 5X SSPE Buffer.


5.                  Hybridization


a.     Remove excess prehyb. Buffer w/kimwipe, avoid contacting tissue.

b.     Add probe (100 ng) directly to Pre-hyb. Buffer and (400 ng) tRNA[6].

c.      Hybridize laying flat in a humidified chamber[7]

d.     Incubate @ 50 degrees 12-36 Hours


Day 2


6.                  Washes


a.     Wash sections 4X in 4X SSC 10 minutes each


7.                  DIG Primary Antibody


a.     After last wash, rinse 5 min in Buffer 1

b.     Transfer to blocking buffer for 1hr @ RT

c.      Wipe slide with kimwipe, use wax pen to outline tissue then add 100ml antibody in blocking buffer, incubate for 4 hours @ RT in a light-sealed chamber with paper towels moistened in buffer 1


8.   Washes


a.      Wash 3 X 10 min in buffer 1

b.      Wash 5 min in buffer 2 for equilibriating


     9.    DIG Probe Color Detection


a.      After last wash, wipe slide with kimwipe, add 450 ml color    development solution.

b.      Incubate slide with C.D.S. for X hours[8] @ RT in a sealed container lined with paper towels moistened in buffer 2.

c. Stop Reaction by transferring slides to buffer 3 (To check the progress of the staining you can temporarily stop the reaction in buffer 2 then add more CDS).


        10.    Staining Nuclei[9]and Photography


a.   Dip 4X in dH20.

b.   Stain in nuclear fast red (.1mg/ml in water) 2 min

c.   Destain briefly in dH20.    

d.   Pass through graded alcohols 70% to 100% (2 min).

e.   Dip slides in Xylene

f.     Coverslip with permount.





Proteinase K treatment: (prepare fresh-20mg/ml pro K in 20mM Tris-Cl pH 7.5, 2mMCaCl)


20X SSC (1 Liter): (175.3g NaCl, 88.2g Na Citrate in DEPC treated H20), prepare 2X about 2 liters, and 4X about 


20X SSPE (1 Liter):  (175.3g NaCl, 27.6g NaH2P04.H20, 7.4g EDTA in 800mls, adjust pH to 7.4 with NaOH ( about 6.5 mls 10N NaOH), autoclave.


Acetic Anhydride Treatment:  (prepare fresh, 250 mls of .1M triethanolamine+1.25 mls acetic anhydride mixed on stir plate in hood, mix 30-60 seconds until all emulsion beads are gone)


Prehybridization Buffer:  (50% formamide, 5X SSPE, 1X Denhardts Solution)


Hybridization Probe:  (20ml prehyb, 100 ng DIG labeled riboprobe, and 400 ng tRNA)


Buffer 1 (1.5 Liters):  (100 mM Tris-Cl pH 7.5, 150 mM NaCl)


Blocking Buffer (30 mls): Buffer 1 + .3% triton X-100, 2% normal sheep’s serum)


Antibody Solution: (1:500 dilution goat anti-DIG antibody in blocking buffer)


Buffer 2 (50 mls): (100 mMTris-Cl pH 9.5, 100 mM NaCl, 50 mM MgCl2)-for equilibriating


Color Development Solution: (10 mls buffer 2, 35 ml 100 mg/ml NBT, 35 ml 50 mg/ml BCIP, 25 ml 1 M livamisol- to block endogenous peroxidase activity-60mg in 250 ml water = 1M)


Buffer 3 (30 mls):  (20 mM Tris-Cl, 10mM EDTA)


Nuclear Fast Red:  (.1 mg/ml in water)


DIG Stuff


DIG labeling mix (Boeheringer Mannheim)

5-bromo-4-chloro-3-indol-phospate (BCIP 150 mgs in 3 mls 70% DMF)


4-nitro blue tetrazolium chloride (NBT 300 mgs in 3 mls 70% DMF)


normal sheep’s serum            (Gibco)




















[1] All glassware should be washed with Absolve glassware cleaner (NEN-Dupont) prior to the protocol to remove RNASE.

[2] For frozen sections start here.

[3] Opens up tissue to allow probe access,  a timecourse experiment was performed on paraffin embedded embryos and little signal intensity was seen  between 5-10 minutes of digestion. After 20 mins the tissue began to fall off slide, no signal.

[4] Neutralizes positive charges on protein in the tissue, prevents charge interactions with probe.


[5] Blocks non-specific binding-Upright slide (Coplin Jars) work well 8-12 slides per 30 mls prehyb.

[6] TRNA is acts as a carrier for RNA.

[7] Prehyb and Hyb can be performed without coverslips. Dry slide w/kimwipe, circle tissue with was pen, add 100 ml prehyb and place in a sealed box humidified with paper towels soaked in 50% formamide/5X SSPE. After 2-4 hours at 50 degrees Celsius, add probe directly to prehyb and incubate at 50 degrees overnight.

[8] X hours can equal 30 minutes for very abundant mRNAs like U2 or up to 24 hours for less abundant ones like MMP-9. Most reasonably abundant mRNAs can be detected overnight (12-18 hours). Can stop reaction in buffer 2 to look at reaction, then just add more CDS. Stop reaction in buffer 3. For longer incubation times re-adding CDS is recommended. Background will increase with time therefore one must optimize incubation times for best signal to noise ratio.

[9] If background staining is light you can stain nuclei using this stain.