I'm completely desperate and hope someone from this forum can help me out! I'm working with brain slices (20um) and FISH and I'm having problems...
The protocol I'm using has worked several times with excellent results. But most of the times it simply does not work with no aparent reason. It can easily happen that one day works and the very same day I start a new in situ with the same reagents and doing the same things (obviously not) and it is a complete failure. I've been checking all sorts of combinations and changing all reagents and I haven't been successful. The last parameter I tested was, after reading genedetect protocol, the time I leave the slides out of the -70 freezer before starting the in situ. After section the brain I freeze it immediately with no fixation until the day of the in situ. In the genedetect protocol it clearly says DO NOT let them defrost before the PFA step (I was letting them defrost) therefore I tested 0 , 5, 10, 20 min out of the freezer but again I have had a new failure. Failure means no signal, greenish background. I copied the protocol I'm using below. I know this protocol works but I'm overlooking something CRUCIAL that I do differently, most of the times wrong and sometimes right.... Can anyone help me?
Thanks in advance!!!
DAY 1 Sections are fixated in PFA for 30 min at 4 degrees///1x PBS 1 min///0.2 M HCl 10 min///1xPBSwith triton 1xPBS 60 sec///50% Formamid/5xSSC 15 min/// Denaturation of labelled probes in hybridization solution, at 80 degrees for 5 min and then chilled in ice for 5 min. Per slide (100 ul hybridization solution +1 ul probe(biotin labelled probe)///18-24 hours at 62 degrees.
0.1 x SSC, for 2x30 min at 60 C. ///For equilibration blocking solution is used for 30 min: 600 ul Triton X 100 130 ml Tris buffer (Tris HCl 100 mM, NaCl 150 mM, pH 7.5) 20 ml 5% Block solution (0.75 g in 15 ml maleic acid: 100mM Maleic acid+150 mM NaCl, pH 7.5).///Apply 100 ul of diluted Streptavidin to the slide, cover with coverslip. Incubate for 1 h at 37 degrees in a moist chamber (1:500 in blocking solution).Wash the slides three times with Tris+Tween 20 (500ul/l) for 5 min. Staining in dark: apply 100 ul of the fluorophore tyramide to the slides, cover with a coverslip, and incubate for 10 min at 15-25 C. washig in dark: 3x5 min RT tris+tween20 (500 ul/liter).Water during 15 min. Mounting: Apply 20 ul antifading solution per coverslip.
Sat Oct 21, 2006 5:10 pm
Joined: Nov 18, 2005
Hi Laura, I perform FISH in my lab, but for formalin fixed, paraffin embeded tissue, so I can offer some advice you might want to try, but I may be way off when it comes to your frozens.
Anyway, usually you need to pretreat with a pepsin or protease. Pepsin from sigma, 50ug/ml, added to your HCL might help. You'll have to play around with times. Only pepsin works at those low pH's, though. If your going to add a proteinase K, you'll need a pH of about 7.6 in some Tris-HCl and you would need to skip the HCl step entirely. I find my number one FISH problems are pretreatment related.
You seem to be denaturing your probe, but not your tissue. I would denature your tissue and probe at a higher temperature, 73C for 6-10 min. Then hybridize at 37C for the 18-24 hours. This is only if your looking at DNA and not RNA (no denaturing necessary).
Your stingency wash on day two could be washing away any signal you might have had. 0.1X SSC is very stingent. Maybe try a 2XSSC at room temperature just to wash off the hybridization buffer, then go to a 72C 0.4XSSC at varying times and see if this helps.
Also, cutting your sections at 20um seems really thick for trying to count signals in individual nuclei. Maybe try cutting sections as thin as possible, like 7-10 um. I actually cut my brain tissue at 2um, but its ffpe.
I know what it feels like to have something work one week then fall apart the next. I used to do immunohistochemistry with never a problem. I took on FISH as a new challenge and that is all it has been.
Hope what I said helps. Tina.
Tue Oct 24, 2006 8:26 pm
Joined: Oct 21, 2006
Thanks a lot Tina!
I'll try out the ideas you've provided and let you know what turns out!
Wed Oct 25, 2006 6:26 pm
Joined: Oct 21, 2006
I've tried the different combinations proposed by Tina and the result is always the same: it fails. But now I've realized that on the second day, after the blocking reagent step I can already predict that it will fail because of the way the brain slices look like.
The difference between success/failure without looking under the microscope is that the succesful slides are let's say "more transparent" than the ones that fail. Even before the TSA step I already see that the slices have sort of lost the transparency that they had at the beginning of the procedure. Does this ring a bell to anyone?
Could it be wrong fixation i.e. I breakdown the PFA when I prepare it? Could it be the Blocking reagent? Or this could be simply anything and it is difficult to say?
Fri Nov 03, 2006 10:53 am
Joined: Oct 21, 2006
Could it be that with 1 min PBS after 30 min PFA is not enough to wash off excess of PFA?
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