Q: Why don't you stop the Proteinase K reaction with glycine?
A: I think glycine inhibits the enzyme and stops the digestion immediately. I'm not sure when or why that step was removed from the protocol I'm using, but the repeated washes in PBT accomplishes the same thing more gradually. In the various protocols I've seen, people use a very wide range of ProtK final concentrations (from 4-50 µg/ml) and incubation times (1-10 minutes). In general, it seems like higher concentration goes with shorter incubation time and the use of a glycine buffer to stop the reaction: if the reaction time is short, the margin of error is smaller and you will have more control over stopping the digestion at the precise time by using the glycine. With lower concentration and longer incubation time, the timing of stopping the digestion is not so critical and the gradual removal of the enzyme with repeated washes is OK.
Q: About the Proteinase K: different manufacturers produce ProtK that differ in the amount of units of activity/mg. In the past, I used units/ml, rather than mg/ml. Did you ever pay attention to that, or do you think it is not reliable?
A: It doesn't matter what you use, units or mg per ml, as long as you know what the activity is for this application. Their 'units' don't measure digestion activity on fixed Drosophila embryos and you must discover the optimum performance conditions of your own stock of ProtK through testing. On the other hand, those units will give you a good estimate on where to start in the titration of your own stock if you are moving between brands or making up a fresh batch. One last comment about using ProtK: it definitely increases the absolute signal level of your stains, as well as reduces the background. It's worth the extra trouble.
Q: Why don't you use PCR to generate your probe templates? That way you don't need to grow up a lot of plasmid DNA.
A: Because I'm hopelessly old-fashioned and don't really know how to do PCR. Roche offers a PCR protocol for making labeled probes, which many people use, and the BDGP gene expression project uses a PCR protocol for its high-throughput embryo staining. I'm doing Qiagen midi-preps to purify good amounts of clean cDNA plasmids. However you like to make probe, do some quality control, such as running the reaction product on a gel, to make sure that the reaction went well, or detecting a spot of it on a membrane, to check the degree of labeling.
Q: Why don't you use DEPC-treated ddH2O? It's cheap and easy to make up.
A: I don't use it for any part of the embryo staining because I have no problem getting a lot of signal and therefore assume I've got no significant degradation of the probes after they are hybridized to the embryos. If you have an RNase contamination problem in your ddH2O water supply, then, by all means, you must fight back. Also, I prefer the commonly available non-DEPC treated, filtered, certified nuclease-free ddH2O in the probe synthesis reactions because I've heard even the smallest amount of residual DEPC can have a negative impact on the polymerase. Superstition? Probably, but 1 L of fancy RNase-free water, enough to synthesize about 50,000 probes, costs the same as 5 ml of DEPC and also you don't have to treat the ddH2O yourself.
Q: Why do you fragment your probes? I've heard you don't have to do this, and actually makes probes worse in some cases.
A: I don't claim any authority on this point, but I know what I've tried and the results I got. I am trying some experiments in which full-length, non-hydrolyzed probes are required, and find that they give terrible background problems. The exact same probes hydrolyzed give no background. I'm starting to get the impression that somewhere between 0.5-1 kb probe size, fragmentation becomes necessary. So some of the differing opinions on this point, to fragment or not to fragment, could be due to experiences with different probe sizes. I've heard also that there is an optimum average probe size to detect EACH gene's transcript. Why not?
Q: Sometimes after the hybridization steps I get big clumps of embryos that never break up. Why?
A: I'm pretty sure it's due to excessive Proteinase K treatment. Someone else once asked me about this problem, and apparently after modifying his ProtK digestion conditions, the problem was gone. I have no idea why this clumping occurs, and it is truly horrifying to see it happen to precious embryos.
Q: That is a very long hybridization step, I normally do only 12-18 hours. Are you sure the quality of the in situ increases so much with those extra hours?
A: In 50% formamide it takes probes a very long time to come to equilibrium with their proper targets, while being prevented from forming stable hybrids with their mismatch targets. Several people I have talked to about this have confirmed that longer hybridization times have dramatically improved the performance of probes that before gave weak stains. In these cases, the hybridization time was extended from one overnight (~12-16 hours) to two (~36-40 hours). I saw one reference where they went up to three overnights trying to detect very unabundant transcripts! I have not tested a time series myself. Currently, my embryos are coming out with very good signal at 20-24 hours, and also with very good morphology. I'm not sure, but it seems like the two overnight hybridization makes them softer and more prone to become warped on the slide. After all, we're baking a proteinaceous tissue structure for a long time in a strong denaturant.
Q: What about your 55° C. hybridization temperature? Why so low? I'm doing in situs at 65° C.
A: The original whole mount non-radioactive in situ hybridization technique, described in 1989 by Tautz and Pfeifle, employed labeled DNA probes and hybridization conditions that very closely resemble those used in the 1980's for DNA:DNA filter hybridizations: 50% formamide and 45° C. In the embryo, DNA:RNA hybrids were formed and those conditions worked. Shortly thereafter, people started using RNA probes, and the rule of thumb, that for a given sequence, the melting temperature of the RNA:RNA hybrid was 10° C. higher than the corresponding DNA:DNA hybrid, was probably used to establish the 55° C. temperature for RNA probes. I don't know exactly why or when people started using higher temperatures (60° C. and higher), but my suspicion is that it was in response to having problems with unacceptably high background staining caused by non-specific sticking of probes to the embryos. I've been hybridizing embryos at 55° C. for a long time without a problem, and the background problems I've encountered attempting these fluorescent methods have been due solely to the primary detection reagents, not the probes, according to my controls.