Re: Long term tissue storage
Dear All,
Thank you for all your responses on the issue of long term tissue storage.
Interestingly, there does not seem to be a good consensus on the best
storage. By far most responses favor storage in ethanol in a cooled
environment (-80oC - 4oC) – though, on a larger scale, these methods
require spark proof cooling units which are not cheap and not always
feasible given local fire regulations. Preserving ethanol tissues at low
temperature may have the advantage over frozen tissue that a freezer
failure would not lead to the complete loss of the collection – for that
reason, expensive alarm systems would also not be required.
Some scientists use DMSO, which is easier to ship, but a nasty chemical.
Desiccating methods seem to be increasingly used, though there seems to be
little experience on conservation problems under realistic conditions.
For long-term storage, many people recommend storage in liquid nitrogen –
this may not always feasible in the long term, given potential loss of
collections because of failure to refill containers. However, a curating
service such as that offered by the Ambrose Monell Cryo Collection at the
American Museum of Natural History may be the best option for most of us.
Below are responses, organized by the methods they propose. I also include
some useful reviews at the end of this message.
Ethanol cooled
* 4 deg is much preferred unless (a) samples are in small containers,
e.g. 2ml, (which fin clips would be) with screw caps (including rubber
o-ring), have a low tissue to Etoh volume, and are not watery tissue (fin
clips ok I reckon; tissue lumps less so). Even so I would still seal any
box that they are in with tape around the lid seam and avoid warm rooms.
For big tissues (e.g. whole fish, etc.), I can't really see a viable way
around 4 deg storage, unless you have a robot who likes topping up and
perhaps completely replacing Etoh periodically. (David Weetman, Hull, UK)
* If possible, I'd invest in a -80°C freezer and store your tissues (in
ethanol) there. The ethanol won't freeze but it will be very cold and your
tissue definitely won't degrade. I've worked in a few labs (mostly birds,
but also marine mammals and other critters) and think the ethanol/-80°C
combination makes the most sense (especially if your tissues are in 2ml
cyrovials - or something similar - b/c you can get an awful lot of tissue
in a relatively small space). (Tammy Steeves, Canterbury, NZ)
* I too have found that freezing or refrigerating ethanol preserved
samples increases the longevity of the samples for DNA extraction. In the
past I have simply transferred the samples into small cryotubes and kept
them in a -80 freezer. (Kevin Roe, Curator of Mollusks, Delaware Museum of
Natural History, USA)
* There was a poster at the Conservation Genetics Meeting in 2001
comparing different storage methods. I can't recall whose it was though.
The best way to store DNA for a long time was lyophilized DNA at -80^C. We
keep samples in alcohol (changed after the first day) at -80^C. You might
also try -20^C, but I strongly discourage you from trusting 4^C for long
term DNA preservation. (Natalia Martinkova, York, UK)
* We are keeping our collection in -80 freezers, and that seems to work
out fine (specimen are preserved in 95-100% ethanol). (Katharina Dittmar de
la Cruz, BYU, USA)
* I've also been storing tissues, including lots of fish, at room temp,
and also in the fridge and sometimes in a -20. I would like to hear what
you find out. And, I'd also like to know more about the issue of
spark-proof cooling. (Sarah Cohen, SFSU, USA)
* A few years ago, I extracted DNA from invertebrates stored in 75%
EtOH since the early 1980s in room temperature and I was able to amplify
mtDNA without any problems, but I personally keep my EtOH-preserved samples
at -18C (freezer) and, if I can, -80C is better. (Sergios-Orestis
Kolokotronis, American Museum of Natural History, USA)
Ethanol at room temperature
* We had insect samples (mainly honeybees) stored for 5-7 years in 95%
Ethanol at room temperature - no problems. (Ute Kryger, Pretoria, SA)
* Our lab has not noticed any problem with ambient storage of salmon
tissues in ethanol (many 10s of 1000s of fin, muscle, and blood samples in
our archive). 10- or 15-year-old samples typically amplify just fine;
however, I can't say we've worked with alcohol-preserved fin clips much
older than that. Our greatest concern for the alcohol-preserved samples,
based on anecdotes from our museum curator colleagues, is that the
laser-printed accession numbers on the labels (inside the tubes) might
eventually fall off the paper. Fortunately, we've not yet seen this in any
samples going back 15 years or more. Some years ago, we thought we saw
problems with some types of denaturent use in commercial ethanol. Since
then, we've used only non-denatured ethanol (mind your field help!) or
ethanol denatured with methanol. The most critical issue in preserving
tissue is that there be sufficient alcohol-to-tissue-volume ratio to
immediately and fully dessicate the sample. We try for a ratio of at least
5:1. We typically fill 1.5 mL snap-cap tubes with 1.3 uL of 95% ethanol
and then tell the field crews that if the tube overflows, you've collected
too much tissue. Incidentally, we don't check gravimetric alcohol
concentrations and refresh evaporated samples, as is standard practice in
museums. A certain fraction of samples dry out and become "potato chips."
Nevertheless, these samples produce ample, high-quality DNA, essentially
indistinguishable from newly-preserved samples. In recent years, we've
begun collecting for some projects by simply sticking cut fins to Whatman
paper printed with 96-cell grids. Of course this option raises storage
issues wrt dermestid beetles (every curator's friend/foe). (Paul Moran,
NMFS, USA)
DMSO
* Don't you think that storage in 25% Dimethylsulfoxide (DMSO) in
saturated NaCl would help you in your goal? We have used it here with
several sample types, fish inclusive. (Charles Masembe, Makerere, Uganda)
Dehydration
* I have had very good luck recovering high quality DNA from desiccated
tissue. I mainly work with amphibians, but I expect the procedure would be
the same for fish. I just put the tissue (usually a toe clip) into a 1.5
ml tube containing indicting drierite. The tissue quickly mummifies and
can be stored at room temp indefinitely. (Eric Hoffman, Georgia Tech, USA)
* For both short- and long-term tissue storage, we prefer dried fin
clips. They are stable at room temp and easy to store, plus no problems
with evaporation, etc. I have used ethanol to store fin clips in the
short term (few months) with good results upon extraction and subsequent
PCR, but don't recommend ethanol for long term storage because our most
problematic samples are those kept in ethanol. (Libby Gilbert-Horvath,
National Marine Fisheries Service, USA)
Liquid nitrogen
* Currently, at my company we do DNA and RNA work and have choosen to
place tissues samples (whole organs) in 1.5ml tubes held under in a Liquid
Nitrogen bank because very little, if any, degradation of nucleic acids
occurs. It is a rather cheap, however not so safe alternative as you have
to be very cautious handling liquid nitrogen. (Kambiz Kamrani)
* Nothing beats liquid nitrogen. I have definitely had the best results
with tissue that was frozen quickly and maintained at that temperature. The
rule in tissue storage is "as cold as possible, as early as possible".
Certainly if you want it to last for decades, that's the safest way to go.
(J. Harshman)
Curation services
* Your best bet is to have your samples curated (at no cost) by the
Ambrose Monell Cryo Collection at the American Museum of Natural History.
Their web site is:
http://research.amnh.org/amcc/ (Marian
Litvaitis, New Hampshire, USA)
* I am the director of the Ambrose Monell Cryo Collection (AMCC) here
at AMNH. We essentially store our samples in cryogenic freezers for (very)
long term preservation. We have done quite a bit of work on preservation at
below zero temps (please see our website at
http://research.amnh.org/amcc/), and from
our findings, it is evident that DNA still degrades at -20 and, to
everyone's surprise, at -80. Having samples below -120 celsius prevents
crystallization, and virtually stops biological activity and time. The AMCC
offers researchers such as yourself the possibility to store their samples
in our freezers (free of charge, of course, since we are an academic
institution) and would be happy to help you do so if you desired to. Please
have a look at our website
(http://research.amnh.org/amcc/) and see if
this is something you would like to explore. (Angelique Corthals, AMNH, USA).
Reviews:
* Tom Pickerell, DEFRA, UK, sent some paragraphs from his thesis
reviewing DNA storage.
* Kilpatrick CW (2002) Noncryogenic Preservation of Mammalian Tissues
for DNA Extraction: An Assessment of Storage Methods. Biochemical Genetics
40, (1/2), 53-62.
* Dawson MN, Raskoff KA, Jacobs DK (1998) Field preservation of marine
invertebrate tissue for DNA analyses. Molecular Marine Biology and
Biotechnology (1998) 7(2), 145–152
* Srinivasan M, Sedmak, Jewell S (2002) Effect of Fixatives and Tissue
Processing on the Content and Integrity of Nucleic Acids. American Journal
of Pathology, Vol. 161, No. 6, 1961-1971
* Smith LM, Burgoyne LA (2004) Collecting, archiving and processing DNA
from wildlife samples using FTA® databasing paper, BMC Ecology 2004, 4:4
* Prendini, Robert Hanner and Rob DeSalle. Obtaining, Storing and
Archiving Specimens and Tissue Samples for Use in Molecular Studies
(http://research.amnh.org/amcc/storage_collecting.pdf)
Dr Lorenz Hauser
Assistant Professor
School of Aquatic and Fishery Sciences
University of Washington
1122 NE Boat Street, Box 355020
Seattle, Washington, 98195-5020
Tel: (206) 685 3270, Fax: (206) 685 6651
http://www.fish.washington.edu/people/hauser/
Lorenz Hauser