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CTAB of Fungi - (Jul/11/2012 )

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Hi all,

I'm trying to extract genomic DNA from Ganoderma boninense of 5 different strains, so as to use it for PCR with redundant primers to amplify micro-satellites, for purposes of observing diversity. Once optimised this will be used in the field. What I'm doing is more of an optimization.

Unfortunately this is my first independent stab at molecular techniques - my first taught or guided stab being in a module where we were supposed to do a RT-PCR, it failed from the offset ad the entire class spent the rest of the time filling out the coursework book "as if" it had worked - so I'm not 100% happy with it all.

I've done a CTAB as follows:

1). 100mg of mycelia added to 1.5ml tube (prepared by liquid nitrogen freezing and grinding in pre cooled mortar and pestle & stored at -80 degrees C).
2). 500 micro-litres of CTAB buffer added, incubated at 65 degrees C for 30 minutes.
3). 500 micro-litres of Chloroform :Isoamyl alcohol added. Mixed until homogeneous and then centrifuged in a micro-centrifuge (13,000 rpm) for 5 minutes.
4). Top layer extracted to a new tube - avoiding the lower layers. Carried out the Chloroform step again.
5). Transferred top layer again, added 100 micro-litres/ml of RNase A, incubated at 37 degrees C for 30 minutes.
6). Added 0.54 volumes of chilled Isopropanol, left at -20 degrees C overnight to precipitate.
7). Centrifuged at 13,000 rpm for 40 minutes afterward I could see, some smear on the bottom I took to be the DNA.
8). Removed supernatant, avoiding disturbing the bottom pellets/smears.
8). Added 100 micro-litres of chilled 70% ethanol, flicked & inverted to mix, spun again at 13,000 rpm for 5 minutes.
9). Removed ethanol avoiding pellet, added 100 micro-litres of TE buffer, flicked and inverted to mix.

I'm sure I saw precipitate as smears/pellets at the bottom of the tubes, but when I ran 5 micro-litres of the stuff with 2 micro-litres loading dye on a 1% agarose gel at 50volts I didn't get any visible bands, the ladder I loaded did work but wasn't all that vibrant either.

Does anyone have any ideas or advice? am I doing something wrong? I'm using autoclaved pipettes and doing most of this in a fume hood because of the chloroform:isoamyl alcohol. I'm going to try again with fresher samples and a new CTAB buffer, but other than that I can't imagine why it's not working, I know some people heat the CTAB but many other papers out there don't bother, or at least don't state it. I'm wondering if centrifuging is too long at it's meaning the stuff at the bottom isn't re-suspending.

Sorry for the long post - I'm working to optimize this RAMS protocol from a past student's thesis, and they left some key details out. In fact in the study that was published from the part of the thesis I'm working from it says a commercial kit was used. In the thesis it states some of it was kit, some was CTAB.

Ben W.


That protocol should give you nice DNA. I suspect that you have added the final TE too soon after removing the ethanol - meaning that there is still some residual ethanol, which will prevent the DNA going back into solution. After you have removed the ethanol just leave the tubes (inverted on a tissue) on the bench to dry, if you can still smell the ethanol, then you should let them dry some more. Most protocols say to leave for 10 min, but I have found that 1-2 hours is usually fine.

Isopropanol pellets are usually clear or white (some protein carry over usually) and gel-like, and should be attached to the side of the tube near the bottom, instead of being directly in the bottom. If you place all the tubes in the microfuge with the hinge side to the top, the will always know where the pellet is/should be - directly down from the hinge, near the bottom of the tube.


agree with Bob1, your DNA should be fine, and you should take care when drying the pellet - I usually put the tubes open on a heating block @ 37°C - so the EtOH is usually evaporated after ca. 15 - 20 min.
Other ideas for trouble shooting:

first of all are you using a fungal culture (i.e. mycelium from petri dishes) or fruiting bodies? There often is an enourmous difference in the outcome of DNA extraction depending on the material used.

second can you tell us what is in your CTAB Buffer - there are a lot of slightly different recepies around - which might influence your results

I would think about including an Acetat (NH4 works best for me, but also Na, or K Ac do the job) step at some point - to remove Polysaccharides etc which often are a problem when extracting DNA from fungi (also aids the DNA preciptiation later on)

You are using a considerable amount of fungal material - so you will have quite some protein carry over - as already pointed out by Bob1, but you should get a clearly visible band on an agarose gel
- best probably is to start again and monitor every step critically, because when you are new in the lab sometimes things go wrong without you noticing it because there are too many things to care about at once - and when you do it a second time suddenly everything works fine :)

keep us informed about your progress


My CTAB buffer is made of:

20g of CTAB powder
280ml of 5M NaCl.
40ml of 0.5M EDTA pH8.
100ml of 1M Tris-HCL pH8.

Made up to 1 Litre with ddH20 (autoclaved). All mixed in a sterilized bottle and flea.

Then when I want to use it I transfer about 50ml to a plastic, pre-sterilized tube and add 1% 2-mercaptoethanol.

I'm using a mycelium that was growing in Potato Dextrose Broth for 2 weeks, since Ganoderma is a slow grower, then using sterile filter paper and funnels to remove the broth, then washing with autoclaved distilled water. Then I take the mycelium and grind in liquid nitrogen in a pre-cooled mortar & pestle. I made sure the material didn't thaw.

One method I've seen did use 1:5 v/v ammonium acetate (I think of 7.5M, but it wasn't specified) added to the aqueous supernatant after a second chloroform:isoamyl alcohol step, during precipitation with 2:1 v/v absolute ethanol and then 500 micro-litres of 0.2M ammonium acetate with a second 2:1 v/v absolute ethanol precipitation. I left this out though as the method I emulated this time did not feature it.

Do you have any recommendations as to the amount of material I should be using? Most methods I'm reading just say "material", I'm wondering if there's some kind of rule of thumb - so much material for such and such a working volume or something like that.

I'm currently harvesting fresh material now as the stuff I've been using is a little older than 2 weeks now. My lab partner found a protocol he thinks worked quite well and is going to give me the details shortly when he gets a break. I'll update when I get it.


I usually use material from agar plates, where the rule of thumb is to use approx 1 cm² of mycelium (without taking any agar!!!!!!). This works fine for RFLPs, sequencing, AFLPs, fingerprinting (RAPDs) etc.

How much fungal material are you able to harvest from each flask? Because if the 0.1g is all the material you harvest after two weeks, I would think about either using agar plates or using a longer incubation time to get more starting material - the wet weight of your culture is much higher than the amount of fungus.....

The buffer sounds alright, I usually omitt the ßMercaptoethanol - because it had adverse effects on the DNA extraction efficiacy when I used it with fungal cultures, especially basidiomycetes.


The freezing and grinding mycelia from liquid culture gives me enough frozen mycelium powder to fill about half a 50 ml sterile plastic centrifuge tube. These are stored in a freezer at -80 degrees C and are retrieved whenever I need them, I harvested a new batch today. The fungus is also very slow growing on agar too, but I have some mature plates still in the incubator room. I could try without the 2-Mercaptoethanol - the stock buffer I made does not contain it because I heard it decreases the shelf life of the buffer to about 3 days. It would also explain why a previous student did not include it in her buffer - according to the lab book.


I completed the protocol, ensuring this time no observable ethanol was remaining and left to air dry a little longer than usual. From my five samples I got concentrations of 143, 206, 27, 25 & 72 micrograms a ml. with A260/A280's of 1.653, 1.953, 2.101, 1.939 & 1.973. Higher values than 1.8 according to what I read are not a problem unless really to high. The A260/A230's were 0.905, 1.378, 0.464, 0.327, 0.847. Which are really quite low, possibly suggesting polysaccharides or protein. RNase was also added and there was no ammonium acetate step this time. MilliQ water was the reference for the nanodrop though and not TE buffer, which has different properties in pH and such. I'm thinking of measuring again with a TE reference, possibly putting the DNA I have through some procedures I've seen designed to remove polysaccharides e.g. .


The PCR amplification of micro-satellites was unsuccessful, no product, so now I'm in troubleshooting phases.


I would not trust nanodrop too much - and without the right reference (TE) the readings cannot be used to estimate DNA conc. When using RNAse the a230/a260 reading should be low.

Have you tried to run a gel to see how your DNA looks like?

Have you tried a reference PCR with your samples (i.e. ITS primers) - and if did this control reaction work? Such a PCR tells you if your DNA is ok or if you have a probem with your DNA extract.

Too much DNA is often a problem esp when using microsatellites (it was in my hands) - I would start my troubleshooting by trying to dilute the samples.


Agree with geb, see if your ITS primers works on the template. It could be your Taq polymerase is not working after all.

-Adrian K-
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