[NHCOLL-L:1457] RE: Formalin & DNA

[Simmons, John E]

Tonya,
I agree with most of what Ross says, with a few minor exceptions.

There has been very little published concerning extracting DNA from
formalin-fixed specimens, but there are many anecdotal stories if you talk
to people working in labs.  Formaldehyde works as a fixative by snipping
protein chains into short lengths to prevent autolysis.  Because of this, it
is extremely difficult to get "good" DNA from formaldehyde-fixed specimens.
There are some unpublished protocols around for getting some DNA, but none
seem to be very satisfactory.  One problem is the great number of variables
in formaldehyde fixation, particularly the time interval between death and
fixation, the buffering system used, and the resulting pH.

I agree with Ross that hauling a liquid nitrogen thermos around in the field
is extremely difficult and often impossible to do.  We, too, have gone over
to using 95% ETOH in the field.  There are several caveats with this,
however.  You must keep the tissue away from light as much as possible,
particularly away from UV exposure.  The vials have to be tightly sealed.
We use screw-cap vials, tighten the lids down, and then wrap the lid/vial
junction with masking tape so the lids are less inclined to come loose.

What I am hearing from the people doing DNA is that ETOH is not a good
long-term storage system.  What is being recommended now is that you don't
keep tissues in ETOH longer than 10 years.  Once back in the lab, the ETOH
should be decanted and the tissues frozen in an ultracold.  I don't do DNA
work myself (preferring to cling to what sanity I have left instead), so I
would be interested to hear what others think about this.  Keep in mind that
(as far as I know) there are no long-term controlled studies on DNA
degradation comparing ETOH storage to ultracolds.  The latest edition of
Hillis et al. recommends against the use of buffers for storage of tissues,
also.

You cannot "fix" specimens in ethanol.  By definition, fixation is the
formation of covalent bonds (crosslinks) to link the molecules composing the
tissues.  Aldehydes form these bonds; of the available aldehydes, only
formaldehyde has a penetration ability sufficient to fix whole animals.
Ethanol is termed a "pseudo-fixative," because it "fixes" the tissues by
altering patterns of hydrogen bonding by removing water, rather than by
forming crosslinks.

Until the late 1890s, almost all fluid-preserved specimens were preserved
directly in alcohol, because formaldehyde was not discovered to be an
effective fixative until 1893. It was not the case of "some European
collections" doing this, but rather everyone who was making fluid preserved
specimens.  The only exceptions to direct alcohol preservation were a few
individuals who used other chemicals along with the alcohol.  Thus any
specimen collected and fluid preserved prior to the 1890s was most likely
preserved directly in plain old ethyl alcohol.  Formaldehyde fixation became
common not because it produced better specimens, but simply because it was
easier and cheaper to carry around and use.  Many people feel that
formaldehyde fixed specimens are better for various subjective reasons
(e.g., they are "harder") but formaldehyde fixation has its own problems.
By using perfusion rather than just injection, you can prepare specimens in
the field in ethanol and get extremely well-prepared specimens.  However,
the problems of carrying ethyl alcohol in the field are even worse than the
problems with carrying liquid nitrogen.  I don't know of anyone willing to
go to all this trouble just to avoid using formaldehyde.

The sooner after the animal's death that you remove the tissue samples, the
better.  The sooner that you preserve the specimen after its death, the
better.  I am appending below a section from a forthcoming publication on
this topic, which provides a bit more detail and a list of references.  As
this manuscript is currently undergoing review, I would appreciate any
comments anyone would care to make about it.

Thanks,
John

Section from:
HERPETOLOGICAL COLLECTING AND COLLECTIONS MANAGEMENT
Revised edition 
Fall 2002

John E. Simmons
Collection Manager, Natural History Museum and Biodiversity Research Center
and
Coordinator, Historical Administration and Museum Studies Program
University of Kansas
1345 Jayhawk Boulevard
Lawrence, Kansas 66045-7561


Blood and Tissue Samples

Although some success has been achieved in extracting DNA from formaldehyde
fixed specimens preserved in ethanol (Chatigny 2000), it is preferable to
prepare fresh blood or tissue samples expressly for this purpose.
Techniques for vein and heart puncture and subsequent blood collecting may
be found in Bennett (1986), Branch (1973), Cooper and Jackson (1981),
Dessauer (1970), Duguy (1970), Esra (1975), Frye (1991), Haskell and Pokras
(1994), Kuch et al. (1999), Luck and Keeber (1929), Maxwell (1979), Powell
and Knesel (1992), Reinert and Bushar (1991), Sooter (1955), Stephens and
Creekmore (1983), and Wibbels et al. (1998).

Non-lethal techniques for obtaining tissue samples (including from
automotized tails and shed skins) may be found in Bricker et al. (1996),
Cameron et al. (1998), Clark (1998), Cordero et al. (1998), Gonser and
Collura (1996), Kuch et al. (1999), and Mockford et al. (1999).

The following instructions for taking and processing tissue samples are
based on the recommendations of Dessaur et al. (1996).    First, make sure
that your permits allow you to take tissue samples.  Although most
jurisdictions consider tissues to be the same as specimens, in some areas,
special permission is necessary to take tissues.

Once tissues are extracted and placed in containers, keep them out of the
light. Note in the field catalog which tissues were removed from each
specimen.

The tools used to collect samples for DNA analysis must be kept very clean.
Rinse the forceps, scissors, and scalpel blades immediately in clean water
(before the blood on them dries) and rinse them in 95% ethyl alcohol before
taking each sample.  The preferred tissues to sample in reptiles and
amphibians are a piece of liver and a piece of thigh muscle without skin.
Each sample should be about the size of a pencil eraser.  

Using clean tools, cut through the skin of the animal's left thigh and
remove a chunk of muscle about the size of a pencil eraser.  Do not cut the
femoral artery.  If the animal is legless, take the sample from the muscle
from along the vertebral column or the neck.  Place the muscle tissue in an
appropriate, labeled container, and clean the tools immediately.  Make a
small lateral incision in the left side of the abdominal cavity.  With small
forceps, grasp the liver and pull it through the opening.  Remove a piece
about the size of a pencil eraser, and place it in an appropriate, labeled
container.  Clean the tools immediately.  Label each sample immediately with
the specimen's field number written in pencil on a small piece of acid-free
100% rag paper or spunbonded polyethylene material placed in the tube, vial
or bag with the tissue sample.  If using plastic cryotubes, scratch the
field number on the tube with a needle, and label the tube with a
cryomarker. 

The preferred way to preserve DNA samples is to immediately freeze them in
liquid nitrogen.  This requires hauling around a thermos of the stuff, which
may be difficult to do in the field.  Tissues in liquid nitrogen should be
placed in (in order of preference) (1) plastic cryotubes; (2) polyethylene
bags (with air removed); (3) or wrapped tightly in extra heavy-duty aluminum
foil. 

If it is not possible to freeze tissues, they may be preserved in (1) 95%
ethyl alcohol for 1-2 hours and then placed in fresh 95% ethyl alcohol; (2)
75% ethyl alcohol for 1-2 hours and then placed in fresh 95% ethyl alcohol;
or (3) isopropyl alcohol instead of ethyl alcohol.  You may add 100 µmol of
EDTA (ethylenediamine tetra-acetic acid) per liter of alcohol for
stabilization.  Tissues in alcohol should be placed in glass vials with good
closures.  Wrap the closure/container junction with masking tape. 

The use of buffers such as DMSO (dimethyl sufoxide) is not recommended for
tissue collection or storage.  They do not provide long-term stability for
storage of tissues.

After being frozen in liquid nitrogen in the field, tissues may be
transported back to the museum in liquid nitrogen or on dry ice (solid
carbon dioxide).


Bennett, J. M.  1986.  A method for sampling blood from hatchling loggerhead
turtles.  Herpetological Review 17(2):43.  
Branch, B.  1973.  The collection of blood by cardiac puncture from
surgically anesthetized snakes.  Journal of the Herpetological Association
of Africa 11:5-6.  
Bricker, J., L. M. Bushar, H. K. Reinert and L. Gelbert.  1996.
Purification of high quality DNA from shed skin.  Herpetological Review
27(3):133-134.  
Cameron, K. D., S. B. Broyles and P. K. Ducey.  1998.  Non-lethal technique
for obtaining tissue for molecular studies of Ambystoma salamanders.
Herpetological Review 29(1):20-23.  
Chatigny, M. E.  2000.  The extraction of DNA from formalin-fixed,
ethanol-preserved reptile and amphibian tissues.  Herpetological Review
31(2):86-87.  
Clark, A.  1998.  Reptile sheds yield high quality DNA.  Herpetological
Review 29(1):17-18.  
Cooper, J. E. and O. F. Jackson.  1981.  Diseases of the Reptilia.
Academic Press,  New York.   
Cordero, P. J., A. Salvador and J. P. Veiga.  1998.  A method of DNA
sampling in lizards with tail autonomy.  Herpetological Review 29(1):23-25.

Dessauer, H. C.  1970.  Blood chemistry of reptiles; physiological and
evolutionary aspects.  Pp 1-72 in Gans, C. and T. H. Parsons (editors).
Biology of the Reptilia, Volume 3. Academic Press, New York 
Dessauer, H. C., C. J. Cole and M. S. Hafner.  1996. Collection and storage
of tissues.  Pp. 29-47 in Hillis, D. M. and C. Moritz (editors). Molecular
Systematics.  Second edition.  Sinauer, Sunderland.  xvi + 655 pp.  
Duguy, R.  1970.  Numbers of blood cells and their variation.  Pp. 93-109 in
Gans, C. and T. Parsons (editors).  Biology of the Reptilia. Academic Press,
New York.
Esra, G. N.  1975.  Blood collecting technique in lizards.  Journal of the
American Veterinary Medical Association 167:555-556.  
Frye, F. L.  1991.  Biomedical and Surgical Aspects of Captive Reptile
Husbandry.    Krieger Publishing Company, Melbourne, Florida.  325 pp.  
Gonser, P. A. and R. V. Collura.  1996.  Waste not, want not: toe-clips as a
source of DNA.  Journal of Herpetology 30(3):445-447.  
Haskell, A. and M. A. Pokras.  1994.  Nonlethal blood and muscle tissue
collection from redbelly turtles for genetic studies.  Herpetological Review
25(1):11-12.  
Kuch, U., M. Pfenninger and A. Bahl.  1999.  Laundry detergent effectively
preserves amphibian and reptile blood and tissue for DNA isolation.
Herpetological Review 30(2):80-82.  
Luck, J. M. and L. Keeler.  1929.  The blood chemistry of two species of
rattlesnakes, Crotalus atrox and Crotalus oregonus.  Journal of Biological
Chemistry 82:703-707.  
Maxwell, J. H.  1979.  Anesthesia and surgery.  Pp 127-152 in Harless, M.
and H. Morlock.  Turtles: Perspectives and Research. John Wiley & Sons, New
York.  xiv + 695 pp.
Mockford, S. W., J. M. Wright, M. Snyder and T. B. Herman.  1999.  A
non-destructive source of DNA from hatchling freshwater turtles for use in
PCR base assays.  Herpetological Review 30(3):148-149.  
Powell, S. C. and J. A. Knesel.  1992.  Blood collection from Macroclemys
temmincki (Troost).  Herpetological Review 23(1):19.  
Reinert, H. K. and L. M. Bushar.  1991.  A safe and simple method of blood
collection from rattlesnakes.  Herpetological Review 22(2):51-52.  
Sooter, C. A.  1955.  Technique for bleeding snakes by cardiac puncture.
Copeia 1955(3):254-255.  
Stephens, G. A. and J. S. Creekmore.  1983.  Blood collection by cardiac
puncture in conscious turtles.  Copeia 1983(2):522-523.  
Wibbels, T., J. Hanson, G. Balazs, Z. Hillis-Starr and B. Philips.  1998.
Blood sampling techniques for hatchling cheloniid sea turtles.
Herpetological Review 29(4):218-220.