View source for ISH on axolotl tissue sections

== Materials ==

* 20x SSC	175.3g NaCl
* buy 50x Denhardts from Sigma 
* 88.2g Na3citrate-2H2O (difficult to dissolve)
* add 800mls H2O
* 1M citric acid to pH6.0
* adjust volume to 1l


1) per Coplin chamber: total 40 mls:   40mls triethanolamine buffer (100 mM)
                                       0.1ml acetic anhydride
                                       1M triethanolamine
                                       100% acetic anhydride are in the “Gefahrenschrank”
2) for 100 mls Alkaline Phosphate buffer (use 1M stock solutions):
				75mls water
				10mls Tris
				 5mls MgCl2
				10mls NaCl
			          100µl Tween
3) for 35mls ISH denaturation mix:
				0.54g DTT (fridge 3)
				28mls 1xPBS
				7mls 10% SDS



''Hybridisation buffer A:'' 

					500 ml			50 ml
Dextran sulfate			50g			5g
Formamide				250 ml			25 ml
20x SSC				125 ml			12.5 ml
yeast RNA (50µg/µl)			10 ml			1 ml
5 mg/ml Heparin			10 ml			1 ml
50x Denhardt’s			10 ml			1 ml
10% Tween				5 ml			0.5 ml
10% CHAPS				5 ml			0.5 ml
0.5 M EDTA				10 ml			1 ml


''Hybridisation buffer B:''

50% formamide, 
5x SSC, 
0.1% tween

== Procedure ==

=== Probe preparation ===
# Mix the following at RT:
   10x transcription buffer		 2 µl
   dig Nucleotide mix			 2 µl
   Linearized plasmid			 x µl (1µg)
   RNase inhibitor			0.5 µl
   RNA polymerase (Sp6, T7)		 1 µl
   H2O 					 y µl
   ''Total		                         20 µl''
# Incubate 3-4h at 37C
# Check transcription by running 1µl on a gel. A diffuse band of greated intensity than the plasmid band should be visible (often also two bands).
# Add 90µl TE, 10µl 4M LiCl, 300µl EtOH 100%, incubate –20C for 1h – over night.
# Spin 10min, wash with 75% EtOH, air dry.
# Redissolve in 100µl RNase-free H2O. If probe doesn’t dissolve well heat to 65C for 5-10min. Store at –20C for few months (can also store probe in hybridization buffer). Run 1µl on agarose gel together with DNA ladder. RNA concentration is approx. 3x DNA concentration.
Use than 1µg/ml


=== ISH protocol ===

1. Prepare 16µm thick cryosections and mount them on SUPERFROST glass slides. Let them dry for several hours at room temperature. Make sure the sections stick evenly on the glass slide otherwise they will fall off during the hybridization procedure. (I usually cut the sections in the morning and start the in situ after lunch. Sections can also be stored at –20C for few months.) 

2. Draw a thick line with the wax pen around the sections on the slide and place them in a humidified chamber. Add PBS only on top of the slides for 5min. 

3. Replace PBS with ISH denaturation mix (work in a Coplin chamber)(2%SDS, 100mM DTT, 1xPBS - prepare everytime fresh!3) for 20min RT. Wash the slides 3x 5min in PBS/0.1%tween (in plastic containers).
.

4. Heat up 40mls PBS-Tween (0.1%) to 37ºC in a Coplin chamber (water bath) and add 40µl of Proteinase K (40µl aliquots at –80ºC, Tower 2, Box K, concentration: 1mg/ml → using concentration: 1µg/ ml))
-> Leave the Coplin chamber in the water bath(!)

-> Place the slides in the Coplin chamber and incubate them for exact 10min
Postfix immediately after (no washing step) with 4%PFA (can be frozen) for 10min in a humidified chamber

5. Wash the slides 3x 5min in PBS/tween. Treat slides for 15min with 100mM triethanolamine buffer and acetic anhydride (0.25% final conc) in a Coplin chamber. Wash 3x 5min in PBS/tween..

6. Prehybridise slides 15min to 1h at 70C with hybridization solution (heat up to 70ºC before using!!! 50% formamide, 5X SSC, 5X Denhardts, 500µg/ml hering sperm DNA and 250µg/ml yeast RNA (or 750µg/ml yeast RNA)-> store at –20ºC) in chamber with whatman paper wetted with post-hybridization solution.

7. Dilute probe in hybridization buffer A – recipe at the end - (1µl/ 1 ml hybridization buffer – work in a plastic chamber!!!) and denature at 70C for 30min. add the slides into the chamber an incubate for around 24 hours at 70 degree

7A) Save the Hybridisation buffer with the probe (is re usable) and wash the slides in buffer B o/n @ 70 degree use now the choplin chambers and wash the slides with different probes seperatly

7B) Wash the slides again 2 times for 30 min in buffer B @ 70 degree

8. Slides are washed 3 times for 30min at 70C in post-hybridization solution (50% formamide, 2x SSC, 0.1% tween- heat up to 70ºC before using!!!) 

9. Slides are washed 2x 5min and 1x 20min at RT in MAB/tween (100mM Maleic acid pH7.5, 150mM NaCl, 0.1%tween) on rocker. Then block in MAB/tween + 10% goat serum or FCS (store at 4ºC, sterile filtered) for 30min to 1h at RT in humidified chamber. 

10. Anti-DIG antibody is diluted 1:5000 in MAB/tween+10%serum and incubated over night at 4C in same chamber. -> in the fridge

11. Wash slides 2x 10min with MAB/tween and then 1x 5min and 1x 30min with Alkaline Phosphatase buffer (prepare each time fresh from 1M stock solutions2: 100mM Tris pH9.5, 50mM MgCl2, 100mM NaCl, 0.1% tween final).

12. Overlay slides with BMP purple (filter ready to use solution) and develop 30min-2 days at RT or 4C in humidified chamber. The reaction is stopped with PBS/tween , wash 3x5min. If reaction is done wash the slides briefly in water and let them dry. Then mount in aqueous mounting medium.