muscle tissue DNA extraction

Harry Witchel Harry.Witchel at
Thu Dec 4 14:20:39 EST 1997

Hello snail lovers!
 I have recently received an e-mail asking for an explanation of what a
CTAB DNA prep is, so I assume that there may be more people out there who
would like to know.  CTAB is formally known as "hexadecyl trimethyl
ammonium bromide" (Sigma H-6269), and it is a detergent with the property
that DNA will precipitate out of a CTAB buffer unless there is high salt
(typically NaCl > .7 M).  There are many different kinds of CTAB preps
(surprise! surprise!).  There is one in Current Protocols in Molecular
Biology (Ausubel -- The Red Book), although I do not see one in Maniatis
1989.  If you have a choice between Maniatis or the Red Book, buy the Red
Book because it has a much better index which makes it much more useful.
 The protocol I present below is adapted from Winnepenninckx (No I can't
pronounce it either), 1993, Trends in Genetics, vol. 9, p.407.  Essentially
the prep is
1)  Freeze and grind tissue
2)  Solubilize tissue in CTAB buffer
3)  Add Proteinase K and digest homogenate
4)  Chloroform extract proteins
5)  Alcohol precipitate DNA
 I tend to get full length DNA for Southerns, but I also get a bit of RNA
contamination, which I now deal with by doping the final product with a bit
of RNase and just letting the RNase do its stuff indefinitely.  Here it is:

CTAB Buffer
2% w/v CTAB
1.4 M NaCl
20 mM EDTA
100 mM Tris-HCl pH 8
.2% v/v B-mercaptoethanol (optional)

I tend to use rather a lot of tissue, such as 10 cerebral ganglia or 10
buccal glands from a common garden snail.  A buccal gland is 9 cubic mm.  I
pour liquid nitrogen in my mortar, put the tissue in to freeze it, and then
grind.  The mortar and pestle are sturdy ceramic unglazed ones which I got
from a kitchen store; they can be autoclaved, but you should probably use a
a different set for each species.  If you are doing many species, you may
be better off grinding them in disposable plactic.
I use 5-10 ml of preheated (55C) CTAB buffer in a 20 ml disposable plastic
universal tube.  I add the powder a little at a time and mix the homogenate
repeatedly while doing this by inverting the tube; this prevents
undigestible clumps of tissue from forming.  Remember that genomic DNA is
very easily sheared, so you should mix the solution by inverting a capped
tube (rather than by shaking), and you should not even dream of using a
dounce homogeniser.
Proteinase K stock (20 mg/ml in water) is as per Maniatis.  As usual, mix
by inversion.  The incubation takes as long as necessary for all the tissue
to solubilize.  At a minimum, you should leave it for 30 minutes.  I tend
to go for 1-3 hours, but if the tissue is recalcitrant and will not
solubilize, I might add another 75 ul of Proteinase K and let the reaction
run overnight at 55C; the DNA still seems full length.  If you have clumps
of tissue which will not disperse, or if your tissue contains a lot of
elastic fibers, it will never fully solubilize, in which case you should
decant the fluid and discard the solids.
This eliminates the Proteinase K as well as the tissue protein.  Chloroform
as usual contains 1:24 of isoamyl alcohol.  Most protocols with CTAB use
only chloroform and not phenol, and I assume there is a good reason for
that.  I do my chloroform extractions in corex tubes -- I prefer the 30 ml
tubes, but most people only have 15 ml tubes.  I gently pour my homogenate
into the corex tube.  I mix the chloroform with the homogenate by squirting
the chloroform (but not the DNA homogenate) through a 10 ml glass pipet;
using a pipet bulb I squirt the chloroform into the homogenate, then I suck
the unmixed chloroform from the bottom of the tube back into the pipet, and
I squirt that chloroform back into the homogenate.  I think squirting is
safer than covering the tube with two layers and parafilm and shaking
because chloroform has a habit of blowing holes through parafilm.  I then
centrifuge the mixture fairly hard: SS-34 rotor, 10-12,000 rpm, 10C, 20-30
minutes.  The first extraction will have a lot of junk at the interface,
and you may be forced to leave over between .5-1 ml of homogenate to avoid
getting the interface.  Sometimes the homogenate will be brown, but in nice
tissues like buccal gland, the homogenate ends up clear.  To remove the
upper homogenate layer to a new tube, I use a 1 ml pipetman and
commercially available "wide bore pipet tips".  You can make your own wide
bore pipet tips by sterilely cutting the end off some blue tips.  If you
use a glass centrifuge tube, it will be much easier to see the interface,
and you will know whether you have gunk at the interface.  If you have much
gunk at the interface after 2 chloroform extractions, try a third
At this point I have put my cleared homogenate into a 20 ml plastic
universal tube again, and I gently add the isopropanol and mix by
inversion.  I then leave the DNA to precipitate for a few minutes at room
temperature while I make a "spool".  A spool is just a glass rod with a
tiny bend at the end.  To make a spool, use a long pasteur pipet, and close
off the skinny end using a bunsen burner.  Then make a tiny bend (I use a
45 degree angle, but I don't think it matters) about 2-3 mm from the end of
the pipet.  Let the glass cool for a minute, then stick the thin end of the
spool into your DNA solution, and capture all the strands of DNA onto the
spool by moving the spool in circles around the edge of the the universal
tube.  The DNA ideally should be made of white threads, but it may be clear
(RNA) and it may be brown (oops! Don't give up, it may still be good).
Once all the DNA is on the spool, you can gently remove it from the
isopropyl solution and drip the alcohol off the DNA by touching the DNA to
the edge of the tube.  If you do not see any strands of DNA at this point,
there is probably too little to spool.  You will have to let the DNA
precipitate overnight at room temperature, then centrifuge the solution at
10,000 rpm.
70% ethanol will get the salts off the DNA and into the water, then 100%
ethanol will get the water off the DNA.  I wriggle the spool in a tube of
each ethanol for 30 seconds or so.  If you had to centrifuge the DNA, the
washes are done on the pellet in the centrifuge tube, as you would with a
plasmid prep.
Get as much alcohol off the DNA as possible, then scrape the DNA off the
glass spool using the edge of the eppendorf tube or a clean yellow tip.  If
there is any fluid at the bottom of the tube, pipet it out.  The reason you
dry the DNA is that ethanol interferes with gel electrophoresis of high MW
DNA, so if you are going to continue to purify the DNA using some extra
steps, it is unnecessary to dry it at this point.  I dry my DNA for 15
minutes under a low vacuum by covering my eppendorf tube with parafilm,
poking holes in the parafilm using a syringe needle, and putting it in the
aspirator.  If there is ethanol (from the side of an undried tube) in your
final DNA sample, it may not run correctly in the gel.
T.1E is 10 mM Tris pH 8/.1 mM EDTA, which is ultimately better for doing
reactions later.  If I have to centrifuge the DNA, I may try as little as
20 ul of T.1E, but if I have a huge wadge of DNA floating on the spool,
then I might use 100-200 ul.  At the very least the DNA should be left to
redissolve at 4C for an hour, but I like to give my DNA overnight to
digest.  Then I electrophorese the DNA to see if it is any good.

Problems with electrophoresis of high MW DNA include:
 the DNA may float out of the well (ethanol or organics present in sample),

 it may refuse to leave the wells (voltage too high, high salt, or DNA is
not redissolved),
 it may have a shooting star effect (too much DNA in a small lane), and
 there may be a streak running down the center of the lane (RNA or smaller
DNAs dragging through the high MW DNA).

To reduce electrophoresis problems, you may try any of the following: dry
DNA (no more than 15 minutes in a vacuum), run low percentage (.6%) agarose
gels, and run the gel at a lower voltage (especially during the first 15
minutes as the DNA attempts to enter the gel matrix).

Harry Witchel
Robert Meech's Laboratory
Dept. Physiology
Medical School

D. Knaebel, wrote:

> I have some students interested in extracting DNA from muscle tissue
> (actually, the buccal mass from a salt-water invertebrate. Does anyone
> have any good techniques for this, or know of modifications to standard
> ones that would work?
> Thanks,
> Dave Knaebel
> Biology Department
> Clarkson University

Harry J. Witchel, Ph.D.
Dept. Physiology
Medical School
Bristol  BS8 1TD

Harry.Witchel at

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