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