In my lab we're doing in situ hybridization on brain sections of mice & rats. However, we don't have the facilities to generate DIG-labeled probes using the usual procedure (generating a plasmid containing the target sequence, transfect into bacteria, isolate plasmid, cut out probe). So, I'm thinking of a way to do it without bacteria. Here's what I've come up with so far:
1. Isolate RNA from brain tissue containing the mRNA of interest.
2. Generate cDNA using random primers or specific primers.
3. RNAse A treatment.
4. PCR with primers designed to generate a product that is (almost) as large as the original mRNA. The 5' primer will have a T7 promoter sequence added. Normal 3' primer.
5. Purify PCR product.
6. In vitro transcription using T7 RNA polymerase and DIG-UTP.
7. DNAse treatment.
If I'm correct I will end up with a DIG-labelled antisense RNA probe.
I would like to ask you for comments on my protocol. Since I have no experience with generating ISH probes, only with doing ISH, it might be that I made mistakes in my reasoning.
I also think your strategy is sound. If your transcript is more than 500 bases in length, you may want to consider doing a carbonate hydrolysis step to fragment your probe to a final length of 200-500 bases. This will allow better penetration of the probe during the hybridization step. It will also prevent intramolecular base pairing which may reduce the hydridization efficiency of your probe.
Thank you both for your comments! I will put an SP6 promoter sequence to my 3' primer to make a sense probe.
My mRNA of interest is ~600 bp and the primers I have designed will generate a product of just under 500 bp.
Try out of the protocol has been pushed back a bit due to other experiments with higher priority. I will report the results when I have them.
Bram,
Your approach looks quite interesting. As this post is fromabout a year ago, I'm curious about the outcome. Did you manage to get the probes as you expected ?
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