IN SITU HYBRIDIZATION (35S)
FOR PARAFFIN SECTIONS WITHOUT SPEEDVAC
LABELING RIBOPROBES
Before starting:
o Dry down a 10 µl aliquot of 35S-UTP [Perkin Elmer #NEG739H001MC] for 20 min in
speed vac
o Clean surface with RNase Away [Molecular Bioproducts #7002]
o Make sure to use riboprobe tips and Rnase-free tubes!
o Check water bath to make sure it is 37C
Mix Together (boxes 37, 46 and 47):
Reagent
1. 5X transcription buffer [Promega
#P118B]
2. 100 mM DTT [Promega #P117B]
3. rATP [Promega #P113B]
4. rCTP [Promega #P114B]
5. rGTP [Promega #P115B]
6. 35S-UTP
7. RNase inhibitor [Promega #N211A]
8. Linearized Plasmid
9. RNA polymerase
T3 [Fisher #BP3206-1]
T7 [Promega #P207B]
Sp6 [Promega #P108B]
Amount/reaction
4.0 l
2.0 l
1 l
1 l
1 l
6 l
1.5 l
1.5 l (~ 1.0 g)
2.0 l
Total volume: 20 l
1. Once tubes are prepared, incubate for 1 hour at 37C
2. Add further 1.0 µl polymerase, incubate another 1 hour at 37C (Can shorten to approx.
30 min.)
3. Add
 1 µl Rnasin
 1 µl DNAse [Promega M610A]
4. Incubate for 15 min. at 37 C
Updated 6/03/15 by LM
1
5. Add (can make a master mix, then distribute 59 µl to each probe)
 7 µl 0.5M EDTA
 14 µl 5X NTE (box 66)
 38 µl DEPC water
Purify probes using mini Probequant G-50 columns [GE #28-9034-08]:
o
o
o
o
o
o
o
o
Vortex column well, so that Sephadex is fully resuspended
Remove bottom from column and loosen cap a quarter turn
Place in sterile 2 ml collection tube provided by kit
Spin for 1 min. at 3000 rpm
Discard the collection tube with the eluted buffer
Place the columns in fresh, sterile, labeled microfuge tubes
Apply probe (71µl) to the middle of the column bed
Spin for 2 min. at 3000 rpm
Quantify probe activity:
o Fill 20 ml scintillation vials with 5 mL scintillation fluid. (Fisher #SX16-4) (One for
each probe and a wipe test.)
o Place 1 l finished probe in each vial and label
o Perform wipe test with kimwipe; put kimwipe into negative control vial.
o Count for 1 min.
o Counts between 500,000 CPM and 2 million CPM are desired
o If count is greater than 2 million CPM, repurify probe to remove un-incorporated 35SUTP
Updated 6/03/15 by LM
2
***subsequent protocol is for 24 or less slides in small glass Wheaton dishes
PRE-HYBRIDIZATION
Before starting:
o Clean surface and all containers with RNase Away; then rinse 3x with dH2O
o prepare 2 liters of 1X PBS (rinse bottle or beaker with RNase Away).
o 200 ml 10X PBS
o 1800 ml dH2O
1. Dry sections at 60C on slide warmer for at least one hour. This may be done in advance.
Drying time may also be increased as desired (ex. 3-6 hrs, overnight, etc).
2. On morning of in situ, prepare fresh 4% paraformaldehyde (PFA/PBS). [PFA, Acros
#AC41678]
 Add 225 mL dH2O to glass beaker (cleaned with RNase Away), pre-warm in
microwave (~1.5 min) to 60C, then place on heating block in hood set at “60C”.
 Add 250 µl 10N NaOH
 Add 10g paraformaldehyde powder (weigh out in fume hood)
 When powder goes into solution, turn off heat (about 10 minutes) and add 25 ml
10X PBS.
 Add 250 µl diethylpyrocarbonate [Sigma #D5758]. Let stir for another 10
minutes.
 Check that pH is between 7.0 and 7.4 using pH strip. (Add 10N NaOH or 12N
HCl if necessary)
 Proceed to step 3 once solution has cooled down to room temperature.
3. Deparaffinize, clear and rehydrate the tissue sections. Change xylenes and ethanols before
use!
 xylenes, 3x 5 min.
 100% ETOH, 3x 1 min.
 After ethanol, immerse in 1X PBS for 5 min.
4. 4% PFA/PBS for 15 min. (save this solution for step 7)
5. 1X PBS, 5 min.
6. 10 g/mL Proteinase K/PBS [Sigma #P2308], 15 min (box 63).
 250 mL 1X PBS
 250 µl 10 mg/ml proteinase K [Sigma #P2308] stock (add just before use!)
o To make stock solution, in a 15 ml sterile tube, bring 100 mg proteinase K
to 10 ml with DEPC H2O, aliquot into 600 µl lots, and store at -20°C
7. 4% PFA/PBS, 10 min. (re-use from step 4)
Updated 6/03/15 by LM
3
8. 1X PBS, 5 min.
9. 0.2N HCl, 10 min.
 300 mL dH2O
 5 mL of 12N HCl
10. 1X PBS, 5 min.
11. 0.1M triethanolamine [Sigma #T1377]/acetic anhydride [Sigma #A6404], pH 7.5
(TEA/AA), 10 min. (make just before use**)
 300 mL dH2O
 While slides are in step 10 (1X PBS):
o Remove ~5 ml water from dish, put into 50 ml Falcon tube, add 3.9 ml TEA,
mix well.
o Remove ~ 2.5 ml water from dish, put into another 50 ml Falcon tube, add
900 L AA, mix well.
 Pour mixed TEA solution into dish
 Transfer slides to dish, then:
o Pour AA solution over slides, shake slide tray gently, and start timer.
12. 1X PBS, 5 min.
13. 70% ETOH, 5 min.
 90mL dH2O
 210mL ETOH
14. 95% ETOH, 5 min.
 15mL dH2O
 285mL ETOH
15. Air dry ~ 1 hr.
o check thermometer in incubator to be sure it is 55C
Updated 6/03/15 by LM
4
HYBRIDIZATION
Before starting:
o Preheat water bath to 85C
o Check thermometer in incubator to be sure it is 55C
1. Make 50 ml hyb. juice:
Reagent
UltraPure Formamide [Sigma #47671]
1M Tris/HCl, pH 7.6
tRNA, 10mg/ml [Sigma #R8508]
50X Denhardt’s
50% dextran sulfate [Millipore #S4031]
5 M NaCl stock
10% SDS
ddH2O
Amount
25ml
500l
1ml
1ml
10ml
6ml
1.25ml
5.25ml
_____
Total volume: 50ml

Final Concentration
50%
10mM
200g/ml
1X
10%
600mM
0.25%
split into aliquots (will need 4-5 mL for 50 slide experiment), store @ -20C
2. Set up slides in humidified chambers (50% formamide/ 50% dH2O in bottom of chamber)
3. Warm aliquot of hyb. juice to bring SDS into solution
4. Defrost aliquot of 1 M DTT (stored @ -20C) and vortex to remix (boxes 60, 61 and 62)
o To make stock solution, bring 3.09 g DTT [Promega # V3151] to 20 ml with 10mM
sodium acetate, pH 5.2.
o Aliquot into 200 µl lots & store at -20°C.
5. Set up hyb. solution/probe in the following proportions (for 10 slides):
900 l hyb. juice
50 l 1M DTT
50 l probe (@~1 x106 cpm/l)
-vortex well and touch spin
o if more probe is needed, can use the following dilution (for 24 slides):
2160 l hyb. Juice
120 l 1M DTT
120 l probe (@~1 x106 cpm/l)
6. Denature hyb. solution/probe @ 85C for 3 min.
Updated 6/03/15 by LM
5
7. Add denatured probe solution to cover tissue section (80-100 l)
8. Place a small piece of parafilm over each section, rolling to prevent bubbles.
9. Hybridize overnight @ 55C in humidified chamber.
Updated 6/03/15 by LM
6
POST-HYBRIDIZATION WASHING
Before Starting:
o pre-warm water baths to 52C and 37C.
o Rinse all containers with dH2O 3 times; do not use Rnase Away!
o Make and pre-heat all incubation solutions and allow to equilibrate for 30 min. in water
bath before starting:
 For steps 2, 3, and 9 (2xSSC, 50% formamide, 52C):
 Prepare 450 ml of 4x SSC:
- 90 ml 20x SSC
- 360 ml dH2O
 distribute 150 ml of 4x SSC into each of the containers for steps 2, 3, and 9.
 Warm one container in microwave for 2.5 minutes to ~85C. (caution –
HOT!)
 Add 150 ml formamide in fume hood.
 Put container, with lid, into 52C water bath.
 Repeat for other two containers.
 For steps 4, 5, 6, 7, and 8:
 Prepare 1.5 liters of 2x SSC:
- 150 ml 20x SSC
- 1350 ml dH2O
 distribute 300 ml of 2x SSC into each of the containers for steps 4-8.
 Warm the container for step 6 (2x SSC, Rnase A) to 37C (about 45 seconds).
 Put container, with lid, into 37C water bath. Do not add Rnase A until at step
6!
*** Do not remove slides from incubator & place in staining rack until all solutions are
equilibrated!
1. 1x SSC @ RT, until all parafilm floats off. DO NOT PEEL OFF.
 15 ml 20x SSC
 285 ml dH2O
2. Incubate in 2x SSC/50% formamide, 52C, 5 min.
3. Incubate in 2x SSC/50% formamide, 52C, 5 min.
4. 2x SSC, RT, 1 min.
5. 2x SSC, RT, 1 min.
6. Incubate in 2x SSC/30 µg/ml RNase A @ 37C, 30 min (box 39).
 add 450l of 20 mg/ml RNase A [USB, #21195]/Tris-NaCl 20mg/ml just before use
Updated 6/03/15 by LM
7
o To make stock solution, in a 15 ml sterile tube, bring 200 mg RNase A to 10
ml with 10 mM Tris /15 mM NaCl buffer, pH 7.6. Mix by inversion. - Aliquot
into 900 µl lots and store @ -20°C.
7. 2x SSC, RT, 1 min.
8. 2x SSC, RT, 1 min.
9. Incubate in 2x SSC/50% formamide, 52C, 5 min.
10. Incubate 70% EtOH/0.1x SSC, RT, 3min.
 73,75ml dH2O
 175ml 100% ETOH
 1.25ml 20x SSC
11. Incubate 80% EtOH/0.1x SSC, RT, 3min.
 48.75 ml dH2O
 200 ml 100% ETOH
 1.25ml 20x SSC
12. Incubate 95% EtOH/0.1x SSC, RT, 3min.
 11.25ml dH2O
 237.5ml 100% ETOH
 1.25ml 20x SSC
13. dH2O @ RT, 10 seconds
14. 70% ETOH, RT, 1 minute
 75ml dH2O
 175ml 100% ETOH
15. Air dry ~ 1 hr.
13. Expose slides to Kodak Biomax MR single emulsion film [#870 1302] in an autoradiography
cassette, 24 hr. Place film so that notched corner is at bottom right in the cassette!!!
14. Develop film & expose, again for 72 hr.
* Exposures may be compared to determine time of exposure for slides to emulsion. (Emulsion
exposure is usually 3x that of the film.)
Updated 6/03/15 by LM
8
EMULSION
Before starting:
o Use Kodak Autoradiography Emulsion NTB 2, available from VWR Scientific
[#IB8895666].
o Aliquoting Emulsion (in darkroom!):
o When emulsion arrives, divide into 5 equal parts and put into small, opaque
bottles.
o Wrap each bottle with 3 layers of aluminum foil and label with content and
expiration date.
o Reserve dark-room 24 hrs. in advance, so fellow users doing timed exposures will know.
1. Pre-heat water-bath in darkroom to 45C. (Correct temperature is critical.)
o Prewarm some water in microwave; fill bath halfway. Place a 1 L beaker filled with
500 ml of 45C water into the water bath. Leave the lid open. Make sure the
temperature is 45C both inside the beaker and in the water bath.
2. Prepare 20 ml of 2% glycerol:
o Add 1 ml of 40% glycerol (stock) to 19 ml dH2O in a sterile 50ml Falcon tube
3. Pre-warm everything for 45 minutes.
o Place the dipping chamber (make sure no water in chamber) and metal apparatus into
the beaker in the water bath [dipping chamber + stand, Electron Microscopy sciences,
#70510]
o Place 1 aliquot of emulsion (do not unwrap foil!), and the 2% glycerol outside the
beaker in the water bath
4. Prepare slide boxes for each probe with desiccant packet.
o Activate dessicant in microwave (30 seconds). Place a blank slide, covered with tape,
in the box between the desiccant and the slide area.
o Remove slides from autoradiography cassettes
o Remember to keep orientation the same for each slide (example: frosted side on left.)
o Write out a label tape with probe, number of slides, date dipped, and date to develop;
place on appropriate box.
5. Take under-pads, slide draining rack(s) and slides to dark room. Also bring 3 pieces of
foil/box, and paper towels.
6. Turn on red light and turn off overhead light.
7. Add the emulsion aliquot to the 2% glycerol.
o Pour slowly and gently down the edge of the tube.
o Replace the lid and mix gently by inversion to prevent bubble formation.
o Do not leave the dark room for any reason after this point. Someone may walk in,
turn on the overhead light and expose everything.
Updated 6/03/15 by LM
9
8. Slowly and carefully pour the emulsion mixture down the side of the dipping beaker and
replace the beaker in the dipping apparatus.
9. Remove label tape from box, put on plastic draining rack, leaving space for slides above the
tape. Remove slides from the box one at a time, and carefully dip up to the frosted label into
the beaker for 1 sec. At the end of 1 sec., lift the slide partially out of the beaker and re-dip
up to the frosting 2 more times.
10. The slide must now be blotted on the paper towels to remove excess emulsion before placing
the slide frosted end DOWN in draining rack.
11. Repeat steps 9 and 10 until all slides are coated with emulsion, separating the slides from
each box with the label from that box on the draining rack.
12. When finished dipping, (or after 15 min., if a small number of slides is dipped) replace slides
in boxes, keeping all frosted ends on same side.
13. Replace lid and turn box on its side, oriented with frosted ends of slides toward BOTTOM.
Wrap box with 3 layers of aluminum foil, being careful to keep the orientation of the slides in
the box as you wrap. Remove the label for that box from the draining tray and place on the
top of the box.
14. Repeat steps 12&13 until all boxes are completed.
15. Store the slides at room temp. overnight, away from radiation and with label side ‘up’.
16. Transfer slide boxes to 4C with label ‘up’, away from radiation and expose appropriate
length of time.
Updated 6/03/15 by LM
10
DEVELOPING AND COUNTERSTAINING
1. Remove slide box(es) from refrigerator and warm to RT for at least 30 min.
2. Make Kodak Developer [#146 4726] and Kodak Fixer [#197 1746]:
o Dissolve 17g developer in a final volume of 250 ml dH2O. Pour into container
labeled ‘developer’. Solution should be slightly yellow.
o Dissolve 45g Kodak Fixer in a final volume of 250 ml dH2O. Pour into container
labeled ‘fixer’. Solution should be slightly cloudy white.
o Allow both solutions to equilibrate at RT for ~15 min.
3. Take solutions, slides, racks to hold total number of slides to be developed, and enough glass
staining dishes to hold all racks.
4. Set up dishes & process under red light, as follows:
o Developer (2 minutes)
o Fixer (4 minutes)
o dH2O (can leave in water until ready to counterstain)
* if doing more than one rack of slides, perform a rinse in dH2O in between developer and
fixer step, in order to keep fixer clean.
5. Counter-stain, as indicated by routine staining procedure: (note – make sure to use clean
ethanols and xylenes!)
a.
dH2O, 1 min
b.
Harris’s Hematoxylin, 1 min and 15 seconds.
c.
Cold tap H2O (carefully rinse until slides are blue, about 2 minutes)
d.
dH2O, 1 min
e.
dH2O, 1 min
f.
95% ETOH, 2 min
g.
EosinY/Phloxine,15 sec.
h.
95% EtOH, 2x 30 seconds
i.
100% ETOH, 5x 1 minute
j.
Xylene, 3x 2 min each
6. Coverslip.
7. Dry slides overnight
8. Next day, remove emulsion using Kimwipe and Windex.
Updated 6/03/15 by LM
11