CTAB of Fungi - (Jul/11/2012 )

oh not only taq - too many things can happen while extracting DNA, and too many secondary metabolites are present in fungi to omitt an internal positive control......I generally use three different primer pairs to test precious DNA extracts - because all the "universal" primers have one or the other bias.....

Thanks, I'm going to do another extraction - since we need more. I'll see if we can do the ITS region too as a test, beyond that we were going to have a series or tubes with different concentrations of the different reagents, including template, needed for PCR to find the optimal, and try gradient PCR too for examining temperature.

To update this, I've managed to achieve DNA yield with the following protocol, modified from the one above:

1). Add 100mg of flash-frozen ground mycelium to 500 micro-litres of CTAB buffer, add 10-20 acid-washed glass beads & vortex for 45 seconds.

2). Incubate in a water bath at 65 degrees for 40 minutes, invert several times during.

3). Let the tubes cool to room temp before adding 2 micro-litres of RNaseA and incubating at 37 degrees for thirty minutes.

4). Add 500 micro-litres of Phenol:Chloroform:Isoamyl Alcohol (25:24:1) and vortex until the phases are not distinct.

5). Centrifuge for 10 minutes at 4000rpm.

6). Carefully remove the aqueous supernatant to a fresh tube. Add 700 micro-litres of isopropanol. Invert once to mix.

7). Put tubes at -20 degrees for 2 hours.

8). Centrifuge at 13,000rpm for 20-30 minutes.

9). Remove supernatant & wash with 70% ethanol twice.

10). Make sure all ethanol is removed and re-suspend pellet in 50 micro-litres of TE buffer.

I have reasonable success with this and the yield is fairly good, however the isopropanol precipitation was at -20 degrees overnight because I was caught up in another part of my work. I'm concerned a few less desirable purity values may be caused by the fact that isopropanol at -20 that long will cause other things to fall out of the solution. If I wanted to solve this I see two possibilities, first keep the isopropanol precipitation at room temp for two hours instead, eliminating the need for -20 degrees. Second I can try ethanol precipitation although I need to know whether 2 volumes of 70% ethanol is adequate. I've seen protocols that use abs. ethanol and some also add a certain amount of ammonium acetate e.g. 50ul in one protocol or 1:5v/v of 7.5M to give a 1.5M conc. in the tubes in another. I'd also like to know how people manage a pellet of DNA which is being stubborn and not dissolving into TE at the end? I did my steps on friday and quantified DNA then with nano-drop but I noted my tips would sometimes get a bit clogged because of the pellet. It may have dissolved more over the weekend. I've seen a suggestion to put tubes at 60 degrees and it speeds things up, but they didn't say how long for or if it was possible to do it too long.

Thanks,
Ben W.

I doubt the longer freezing of the isopropanol solution would precipitate substantially more impurity. For ethanol, use 2.5 volumes of pure ethanol (not 70%). It can take days to dissolve a genomic DNA pellet that is dry. Vortexing periodically, heating, and waiting will eventually work.

I've read a paper in which during ethanol precipitation 50 micro-litres of 7.5M ammonium acetate was added and then two volumes of ice cold ethanol. After centrifugation, the pellet was resuspended in 300 micro-litres of 0.2M ammonium acetate and then re-precipitated with 2 volumes of ethanol, before drying with vacuum and re-suspension in 50 micro-litres of TE. They don't describe washing. I wondered what the purpose of ammonium acetate is here and whether the resuspension in it and second precipitation replaces washing?

The NH4oAc is the salt component of the precipitation, similar to adding NaCl or NaoAc. You need a certain amount of salt to get efficiently precipitate nucleic acids, these are often provided by the lysis buffer(s) but can also be added at the end. The amount of salt depends on the compound for some reason that escapes me at the moment.

Just tried a PCR amplification, no luck and no amplicons, when I consider PCR it's probably worth a new thread, but my purity values for my 5 samples I'll list here, if they're not pure enough I'll need to do them again. I'm not that experienced so I don't know what level of purity I can get away with. I know 1.8 is good A260/A280 and 2.0 is good A260/A230 however.

A260/A280 A260/A230 DNA Conc.
Sample 1 = 1.760 1.477 147 ug/ml
Sample 2 = 1.848 1.410 974 ug/ml
Sample 3 = 1.362 1.219 64 ug/ml
Sample 4 = 1.594 1.320 80.5 ug/ml
Sample 5 = 1.688 1.093 153 ug/ml

I've also done the protocol from post #13 but with the 50ul of 7.5M ammonium acetate and 2 vol. of ethanol. I noted a greater increase in DNA yield according to spec. Which is interesting given sources that say DNA is less soluble in Isopropanol so precipitation of larger species and smaller concentrations (although the same source also states ethanol is better for small volumes and small DNA concentrations). Does anyone have any advice as to the concentration of Ammonium Acetate for Ethanol precipitation? I've seen 0.1M or 2M or 50ul of 7.5M, and even 1.5M and then ethanol precipitation, followed by 200mM and another ethanol precipitation (no wash afterwards). Or is Isopropanol really best? - I seem to see conflicting opinions in the literature or on sites.


I think I need to consider ways to improve on those purity values, but I already have a phenol component to get rid of protein and I thought CTAB handled polysaccharides fairly well. Tomorrow I'm using 50mg of material and not 100mg, and I think maybe a smaller pipette to prevent possible phenol contamination, I noticed even if I'm careful with a p200 the boundary get's a little disturbed, I could take a conservative 200ul of the supernatant rather than my 400ul I manage with the p200, to ensure boundary layer is not disturbed. Use of a step where the supernatant is incubated at 4 degrees for 15 mins in 2M NaCl and 4% PEG is used to increase yield and 2M of NaCl is also said to help with polysaccharides, I don't know if anyone else knows of why this is or has even done it. The same paper also uses a wash solution of 15mM Ammonium Acetate in 75% Ethanol.

Is there anything I can do already to my DNA so I don't have to go through everything all over again. I'm thinking of also testing for DNAses by incubating at 37 degrees overnight.

As a final consideration - 1ul of my DNA samples are added to a working volume of 25ul for my PCR mix I'm wondering if, impurities considered and the concentrations of my DNA, if it's just too much.

As always your hints and advice are greatly appreciated, I've been reading a lot but it doesn't make up for being a 1st molecular project newbie.

Just a little piece of thought. Did your template work in other primers set? This is because I think there is a very high possibility that your primers are not working rather than you have a bad template. Based on my memory, I used to have such quality template to do my PCR and I still got my amplified amplicons.

EDIT: spellings

The ethanol will not precipitate DNA without the presence of salt. The ammonium acetate is the salt in this situation. Certainly the resuspension and reprecipitation will remove contaminants from the first preciptation, but it will not remove the salt (ammonium acetate), which is the purpose of the 70% ethanol wash in the standard protocol. The vacuum may remove the ammonium acetate, given enough time, since its decomposition products are volatile. The DNA will be very hard to resuspend after this, since it will be so dry.

I've tried another PCR reaction and it has worked this time, albeit I need to re-run it because I got some smear in my negative control. However the expected RAMS bands predicted to appear based on a previous student's experience did appear. I'm not entirely sure why the first reaction did not work, perhaps DNase, but if it were in my reagents then I would have expected the run that worked to fail as well. I have noticed I have a tendency to over-dry my DNA, but after a day or so of heating for a few minutes and then mixing sorts this out. My next goal is to try and vary the component amounts to reduce some of the noise and get an optimal result.