RNA Dotblot issues - Alkaline or formamide/formaldehyde desnaturation? Control (Jan/09/2007 )

Hi.
I work with Azospirillum amazonese, a nitrogen fixing bacteria, and I wanna know if a specific gene is expressed depending of the nitrogen disponibility.
I did a dotblot with these parameters: 4 ug of total RNA in each well (measured by absorption spectrometry), extracted after 1, 2 and 3 hours of cultivation with or without nitrogen, and these RNAs were desnaturated by alkalinity (NaOH). After the hybridization with a radioactive probe, I only got a weak signal, after almost one month!
Sambrook does not recomend to fill the well with more than 5 ug of total RNA. Did anybody try to use more than this quantity?
In Molecular Cloning, I saw a protocol where the RNA desnaturation is made by fomamide/fomaldehyde/MOPS buffer treatment. But it seems quite laborious to prepare MOPS buffer to me. Is it really necessary to make MOPS with DEPC water, sodium acetate DEPC-treated, EDTA DEPC-treated, adjust the pH to 7 (How can I do this, if the phmeter bulb is not RNAse-free?) and filtrate it? I usually migrate my RNAs in TEB buffer just autoclaved and they seems well to me. Can I do the same with MOPs buffer, just filtrating it? Is this kind of desnaturation more effective than alkaline desnaturation? I know that RNA is sensitive to alkalinity, but the transfer to membrane is so fast that I think there's no time to degrade them, LOL.
Another issue: Since I load the same amount of RNA in each well, do I need to normalize my results with a constitutive gene? Ribosomal proteins expression is affected by nitrogen starvation, for example. Which gene could I use as a normalizer?
Does anybody have a good protocol to label the probe by PCR?


Thanks,
Fernando

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Does that mean you had the membrane in the phospho cassette for a month, or did you repeat the experiment several times in a month?
There are too many things you could change in order to get a better result. You just have to look at your protocol more carefully, and try to trouble shoot.

I use only 1ug per well for a dot blot.



well, we are the worker bees, and it is the protocol. I made it with adjusted the pH, then DEPC treated the water, and then filter sterilised it. It's not that hard to do.

ABSOLUTELY!!!! I used GAPDH.


sorry, can't help you out there, but there are quite a few protocols on the net and here that could help you out.

V

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First of all, thanks for answering my questions.

The RNA was trasfered to membrane by mixing it with an ice-cold 10 mM NaOH/1mM EDTA solution.
I used 50 ng of probe and I labeled it with P32 by random priming. My membrane was blocked by SDS 7%/EDTA 1mM/0,5 M Na2HPO4 pH 7.2 and I washed it two times, one with SSC 1X at ambient temperature and one with SSC 0,5X at 60°C, both washings were taken in 10 minutes. I had the membrane in the phospho cassette for almost one month.


What kind of labeling do you do? Do you work with bacteria?

I don´t understand if you added DEPC-treated water or added DEPC to the solution. If you add DEPC to the entire solution (MOPS, EDTA, NaAc), how do you get rid of it, since MOPS is sensitive by autoclaving? If you add DEPC-treated water, how do you get rid of the RNAses that could come from the pHmeter?


Can I use 16S as a normalizer?

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well, assuming my benches are cleaned by NaOH treatment before RNAse assay, and assuming electroporation cuvettes are treated by100mM NaOH, seems that this is a strong way to denature fragile RNAs.
I loaded succesfully 10µg of RNA in dot blots (using a vacum device to ensure the solution is not dispersed). The RNAs were in 100%formamide prepred, and denatured by 56° 10' before applying on the membrane.
Normalization is essential for your experiments, as measurement by UV is not enough accurate.
EDTA can't be DEPC treated as DEPC reacts with amines. So Tris can't be treated as well. You should use clean powder, and DEPCtreated water (after autoclave)
Labelling the probe by PCR? you should have a look at klenow and random priming issues.

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the most i've had a membrane in the phospho cassette was for 2 days!! i can generally pick up a signal in a few hours.
i use a lot more probe than that. i don't do bacteria, but human cells, which were grown to almost 90% in a T75...
my washing with SSC were for 30 minutes each... but that isn't your problem. i'd say you're not using enough probe.
all the water you use has to be DEPC treated. then go on and make the other things.
i was using 32P to label transcripion from nuclear extracts.

V

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I did a Southern yesterday, and the signal was weak too. I agree with you, I think the problem is with the probe. Look at my protocol:

1.In a epp, add 25 ng of DNA with 125 ng of random primers in 43 uL of water. Close the tube and place in a boiling water for 2 minutes.
2.Place the epp on ice for 1minute. Spin. Place the tube on ice again.
3.Add to the tube: 1 uL 5mM dNTP;
5 uL 10X Klenow reaction buffer;
1uL dNTP 10uCi/uL 32P;
4.Add 5 units of the Klenow. Spin. Incubate the reaction at 37°C for 1 hour.

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Ok, maybe just tweak the protocol slightly.
try combining 25ng DNA in 5-20uL h20
Add dATP, dGTP, and dTTP and youre random primer buffer mixture.
then add the 32P dCTP (~50uCi). then add dilstilled water to 49uL. mix briefly, and pulse in a micocentrifuge.
Then denature the DNA by boiling for a few minutes, place immediatly on ice.
Add 1uL klenow , mix gently, and centrifge briefly.
incubate at RT for 1 hour.

or perhaps you could try the GibcoBRL Random Primer DNA labeling system.

V

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Thanks.
But I solved the problem. My colleagues label the probe with PCR, and to this type of labelling, they use 0,5 mM DNTP. Unfortunatelly, I did my labelling with this DNTP, but I should use 5mM DNTP. Doh! Certainly, it diminished the yield of the labeled probe.
PS: The DOTBLOT worked well.

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