[Histonet] Need assistance with in situ protocol!!! (LONG)

[- -]

Dear histonetters,

 

I am in dire need of having my in situ protocol work!!  I cannot find the problem and feel there may be too many to fix the situation on my own.  Please if you are up for the challenge, look over my protocol and problems I am facing and offer up any suggestions or changes I should make.  Also, does anyone know of a good DIG-labeled RNA probe that a particular manufacturer makes and that can be used for targets found throughout different mouse tissues -- just trying also to ensure protocol is wrong and not probe??  

 

Thank you.

 

Note: Letter ‘u’ used to represent micro. Temps given in celcius.

Sorry – can’t tell you what probe I was using.  Only that it is a DIG-labeled RNA probe.

 

Problems:

Antisense and sense appear to both stain.

 

Large amount of background (endogenous peroxidase?)

 

At the end of my washes (after hybridization) tissue begins to disintegrate and cells begin falling away from the sections in large clumps – only tissue I am using is mouse kidney.  The kidney is perfused (saline then 4% PF/PBS – RNase free) and allowed to sit overnight in 4% PF/PBS.  Afer switching the 70% alcohol for ~3-5 hours, the tissues are routinely processed and embedded.

 

Sections are sectioned – 5 um, and placed onto positively charged barrier slides (Biogenex).  They incubate on a plate (covered) at 37 degrees and then before deparaffinizing are baked at 55 degrees for 1 hour.

 

At all times RNase free glassware/solutions are used during the entire process.  Bench tops and surrounding area wiped with RNase Away.

 

PROTOCOL:

 

Slides deparaffinized and brought down to distilled water, then washed 2x (5mins each) in 1x PBS.

 

Proteinase K digestion:

 

Slides incubated in 0.1M Tris and 50mM EDTA (pH 8)  prewarmed at 37 degrees containing 5 ug/ml Prot. K.  Incubation time 17mins (determined to be optimal).

 

After incubation, slides immersed in 0.1M glycine/PBS for 5 mins.

 

The slides were then post-fixed in 4% paraformaldehyde/PBS for 3 mins. Rm Temp.

 

Rinse with 1x PBS 2x (5 mins)

 
Acetylation
Slides placed in a glass staining jar containing 0.1M triethanolamine and while stirring, acetic anhydride was added to a final concentration of 0.25% acetic anhydride.

 

Slides rinse in distilled water and then dried on hot plate (37-40 degrees) for ~10-15 mins.

 

Hybridization

 

Buffer: Frozen at –20 degrees.  Placed in 50 degree waterbath before use.

 

Reagents                                  Final Concentration

Deionized formamide                    50%

20x SSC                                  5x

Dextran sulfate              10%

100x denhardt’s solution 5x

10% SDS                                 2%

10mg/ml denatured                     100ug/ml

sheared salmon DNA                            

Sterile ddH2O                           

Aliquots of 900ul made up and frozen.

Dilute riboprobe in buffer to final concentration of 2.5ng/ul ( DIG-label RNA probe)

To each section between 40-60 ul of probe was added to cover section – more for larger sections.

 

Sections placed in hybridization chamber (Boekel) at 42 degrees, covered each section with Parafilm to avoid evaporation.  

 

Slides incubated at above temp. 20 hours overnight.  Chamber kept humid with 5x SSC in the provided wells. 

 

Checked slides next day to ensure moisture still under parafilm and that there was no evaporation – none seen.

 

Slides placed in 2x SSC/0.1%SDS for 2mins.  Parafilm begins to float off to liquid surface.

 

Slides washed in 2x SSC/0.1% SDS at room temp shaking gently 2x (5 mins each)

 

Wash slides 0.1x SSC/0.1%SDS at the same temperature used for hybridization (prewarmed at 42 degrees) --  2x 10mins each

 

Rinsed briefly in 2x SSC twice (Room Temp) to remove traces of SDS

 

Slides incubated with 10ug/ml RNase solution in 2x SSC at 37 degrees for 15 mins

to selectively degrade single stranded RNA and reduce non-specific signal.

 

Wash in 2x SSC/ 50%formamide for ½ hr. at 52 degrees gently shaking

 

Rinse briefly in 2x SSC Rm. temp.

 

Placed into Buffer 1:

80ml 1 M Tris-HCl pH 7.5

40ml 3M NaCl

Add autoclaved deionized water to 800ml

 

Rinsed briefly

 

Place into fresh Buffer 1 for 5 mins.

 

Place into Buffer 2 for 30 mins. shaking Rm. Temp.

Buffer 2

1 M Tris HCl pH 7.5

3M NaCl

Triton X-100 (0.5%)

10% Blocking Reagent (1% final concentration)

 

Blocking reagent bought from Roche (final concentration is 1% or 1x)

Blocking Reagent Cat. No. 1 096 176 – made up as instructed by manufacturer.

 

Sheep Anti-DIG-AP antibody used (Roche) –0- Cat. No. 1 093 274

Buffer 2 tapped off and 100-150ul of antibody, diluted 1:50 in 6ml 2% normal sheep serum/0.5% Triton X-100 in PBS was added.

 

Slides incubated with antibody at 37 degrees for 2 hours.

 

Slides washed 3x in PBS (5 mins. each)

 

Rinsed in 100mM Tris-HCl (pH 9.5)

 

In darkness make up BCIP/NBT solution while rinsing in above Tris.

BCIP/NBT provided by DAKO – new kit, made up as specified.  Difficult to make mistake here.

 

Added BCIP/NBT solution 3-4 drops to cover section.  After 20-30 mins. Slides checked for signal.

 

Little seen except for tissue coming off slides (clumps of cells) and floating off.

 

Allowed to sit overnight at 4 degrees – next day there was signal – very strong in some slides – however, there was background, and sense and antisense slides had what appeared to be signal.

 

???  Any suggestions???

 

THANK YOU FOR YOUR ASSISTANCE

 


		
---------------------------------
Do you Yahoo!?
 Yahoo! Mail - Easier than ever with enhanced search. Learn more.