G'day folks My original post is followed by the various suggestions I got for alternative methods for preserving DNA. Thank you very much to all the people who sent me messages. I tried to incorporate most of the emails I received, but some were a little redundant. Cheers Peter G'day folks I was wondering if folks have any experience with preserving DNA in the field in liquids other than ethanol? My target organisms are fishes, but I'm sure different liquids work on most groups just fine. It some parts of the world ethanol can be very difficult and/or expensive to obtain. Are there any good viable alternatives that can be obtained in third world countries that should be ok that people have experience with? Isopropanol is often available it seems, but I've only seen one report that suggested it should be ok. Also, I guess the isopropanol one would buy in a pharmacy varies depending on who makes it and the country (wikipedia says that in the USA and the UK Isopropyl Rubbing Alcohol is made of ethanol, not isoproyl). One paper looked at preservation of spiders, and found RNAlater and propylene glycol worked well. Another paper on aphids found acetone was good, as was diethyl ether, and ethyl acetate. I'm not sure though how easy these are to obtain, or safely these can be transported though (RNAlater is too expensive to consider). I also searched the archives, but suprisingly this doesn't seem to be a topic that has come up before. Thanks Peter Unmack Details of the different things people suggested are given below. It seems like laundry detergent and salt may be the best options, while silica gel is good for small tissue samples. I'm rather leary of DMSO as that stuff has the potential to be dangerous (it makes your skin highly permeable to everything), and as noted by one respondant, some tubes tend to leak. I'd be really interested to hear anything further from folks who have used detergent or salt for preservation. Here are the responses. You can also consider dry storage of DNA, as implemented in FTA cards: http://www.whatman.com/FTAandFTAElute.aspx At our lab we work almost exclusively with fish tissue, from both marine and freshwater species, and our preferred method of preserving DNA in the samples prior to extraction is air-drying in blotter paper, with each sample in a separate coin envelope. I am not sure if this would work for you in a humid location, as the key to high quality DNA seems to be rapid drying of the tissue, but barring access to a dessicator, a sunny windowsill works just fine. This method also makes storage and transport of tissues a breeze. DMSO 20% DMSO saturated with NaCl for a variety of species, including fish - easier to transport on planes than alcohols, and preserves well at RT for extended periods (see Amos, W. & Hoelzel, A.R. 1991. Long-term preservation of cetacean tissue at ambient temperature for molecular genetic analysis. IWC Special Issue 13:99-104). DMSO is typically available as 'horse liniment' or veterinary rubbing compound in country stores in the US, or from local vets. Pour in NaCl or table salt to saturation, collect the supernatant fluid. DMSO tends to leak from regular eppendorf tubes, you'll want to use gasketed tubes or parafilm. NaCl I used 5M NaCl solution to preserve bat wing membrane in the field, prior to extraction. For memory, I think it also worked well for Quokka ear tissue. Saturated table salt (NaCl) in clean water works well for small arthropods. silica gel When ethanol is not available then the best alternative is dessication with silica gel. I know of a few studies that used acetone successfully, but in my experience isopropanol and rnalater are not reliable. Laundry detergent. Someone once told me that they used washing powder to preserve specimen when they go sampling in muslim countries where it is rather hard to get alcohol. There is actually a protocol for extracting DNA with laundry detergent - I attach the pdf! However, since it is tricky to do all the extraction steps "in the wild" preserving your samples in detergent is also an option. I have tried to preserve freshwater snails when I had no access to alcohol and it seems to have worked ok. I haven't "properly" tested the quality of the DNA but I got quite a lot of DNA out of those animals using commercial kits and a chelex extraction protocol. In the field I just put the snails as they were into the powder, hoping that their excess water would mix with the detergent powder which would then seep into the cavities preserving the tissues. However, I also prepared some snails out of their shells before I stored them in the powder and they gave much better results. Perhaps it would be even better to dissolve the washing powder first and then store the tissue in the liquid. I DID put the snails straight into the powder, but I think it would be better to disolve the washing powder first, so it can penetrate the tissues better. The snails we preserved were of different sizes - Potamopyrgus, which is about 3mm long, and Lymnea ovata, which is much bigger, something like 1.5cm. The potamos I stored whole, but with those ovatas I found it better to dissect the foot off and only preserve that. I kept the snails in the powder for about three weeks, then transferred them to EtOH. I rinsed the washing powder off the snails as much as possible before I pickled them. A. Bahl and M. Pfenninger. 1996. A rapid method of DNA isolation using laundry detergent. Nucleic Acids Research, 24, 1587-1588. Kuch U, Pfenninger M, Bahl A. 1999. Laundry detergent effectively preserves amphibian and reptile blood and tissues for DNA isolation. Herpetological review, 30, 80-82. RNAlater I used RNAlater with success for DNA and RNA. I always buy my RNAlater but I have a recipe for it (never try but it should work). You can check the web site where the recipe is described: http://www.patentstorm.us/patents/6204375/fulltext.html sarcosyl-urea One good option is sarcosyl-urea. I came across this stuff just recently when I moved jobs to South Carolina. It was developed by someone who works in the genomics section of the NOAA lab next door and has been used for more than 20 yrs. Apparently, sequencing of samples kept for 5-6 yrs (at room temp, I think) is no problem. 20 yr samples appear too fragmented for long reads, but are probably fine for short reads and microsats etc. The recipe for sarcosyl-urea (not available commercially) is: 1% sarcosyl 8M urea 20mM sodium phosphate 1mM EDTA pH 6.8 It takes a little while to make up and pH is fairly critical, apparently. It precipitates at 4 deg C (but goes back into solution if warmed). At the lab here, they use it routinely during field work for collecting small (2-3 mm) fin clips from fish and storing in about 1ml liquid. It actually does the digestion step for you, so your sample will not be visible after a short while. This means that it's probably a good idea to be fairly clean as you collect (e.g. rinse sample and clean forceps between samples). I've asked about that here (where the field collection method is quite 'dirty') - occasionally a little contamination can be a problem sometimes, but as long as your tissue sample is the main thing in there, it will outcompete during PCR amplification. See these for a couple of refs where they've used the stuff: 1) http://www.nmfs.noaa.gov/mb/sk/pdf/Report_11.pdf 2) Chapman et al, 2004. Diseases of Aquatic Organisms, 59: 179-185. I use a Sarcosyl-Urea solution that in addition to Sarcosyl and Urea, contains EDTA and maybe one other component (Anhydrous di-basic sodium phosphate maybe?). I find it to be a great storage buffer after an initial heating period (usually done by placing my samples on the dashboard of the vehicle I'm using to drive between sampling site - supposed to be something like 50-60 C overnight, although depending on the tissue and tissue type I find it might take a little bit longer), you can store it at room temperature. I don't have the recipe in front of me right now, but I could send it to you if you like. It's essentially non-toxic and have sent it through the mail and/or gotten it through customs before; you just might have to carry some paperwork with you. As for availability, these are basic molecular biology reagent and you should be able to get them through any University wherever you are working. Or, if you're in a pinch and don't want to do that, I always make up sampling kits ahead of time and just bring them with me. I am doing both microsatellite work and mtdna sequencing and the sarcosyl has worked great and seems to have a pretty decent shelf life although I can't vouch for that (however, over the course of my 6-year study it's been fine). other alcohol In a bind I use high percentage drinking alcohol to preserve coral DNA. Vodka and Rum work just fine. Longmire I use Longmire's buffer for collecting and storing field samples collected by other people. Longmire J, Maltbie M, Baker R (1997) Use of "Lysis Buffer" in DNA isolation and its implication for museum collections. In: Occasional paper, Number 163. Museum of Texas Tech University. The tubes of buffer often sit in a vehicle for sometime before they are used and samples may take a while to reach me. This buffer is also accepted on airline flights whilst ethanol is not. I have stored avian blood and tissue successfully in this buffer. If I remember correctly it has been tested for use in the field at ambient/high temperatures and found suitable for relatively long periods of time (years). As an aside - I once received avian tissue from a bird that I understand had been preserved in home brewed Mauritian rum for some years. The sample was also sent to me in the rum - I extracted DNA suitable for microsatellite analysis from this without any problems. I have been using EDTA for blood and TE (Tris-EDTA) for the mouth-swab samples from marsupials in rain forests in Brazil. It have been very good for us. You just have to centrifuge well before you start DNA extraction. One option might be to do your dna extracts in the field with 5% chelex. I did this in the field in remote areas of Australia with wombat hair. You'd have to check whether this is a suitable method with your type of sample. I've used Queen's lysis buffer for storage before. I'm not sure how easy the reagents are to get in third world countries, but it's another options. I've attached a pdf I found with a whole bunch of recipes including one for the buffer I am talking about. I had good luck with isopropanol for forest elephant dung storage in tropical conditions. Yes, EtOH can be really hard to find in some places. I haven't tried methanol (CH3OH), which is easier to find, but I'd avoid it. I've also used DMSO and generally speaking salt- saturated SDS solutions. See http://www.ajtmh.org/cgi/content/abstract/39/1/33 Also, keep in mind that you can't fly with such flammable liquids, like ethanol and isopropanol, and since rubbing alcohol varies in its constituents across countries (maybe even in the same supermarket) and that it can even be alcohol-free ketones, you may want to have DMSO or salt-soap-like media with you to avoid bad surprises. It's happened to me before. I did a literature search on this recently and found the references listed at the end of this message. I was more concerned with transportation - ethanol is readily available in most places I go. I noticed, as you did that several of the studies in the literature give conflicting recommendations. In the end I decided to continue with ethanol as usual, and remove it before transportation. I haven't used these specimens for DNA yet, and I haven't done any experimentation myself with the different fluids. I hope that helps, but maybe it just adds more confusion! Austin, A., Dillon, N., 1997. Extraction and PCR of DNA from parasitoid wasps that have been chemically dried. Australian Journal of Entomology. 36, 241-244. Dillon, N., et al., 1996. Comparison of preservation techniques for DNA extraction from hymenopterous insects. Insect Molecular Biology. 5, 21-24. Gurdebeke, S., Maelfait, J. P., 2002. Pitfall trapping in population genetics studies: Finding the right "solution". Journal of Arachnology. 30, 255-261. Mandrioli, M., et al., 2006. Factors affecting DNA preservation from museum-collected lepidopteran specimens. Entomologia Experimentalis et Applicata. 120, 239-244. Post, R. J., et al., 1993. Methods for the Preservation of Insects for DNA studies. Biochemical Systematics and Ecology. 21, 85-92. Quicke, D. L. J., et al., 1999. Preservation of hymenopteran specimens for subsequent molecular and morphological study. Zoologica Scripta. 28, 261-267. Vink, C. J., et al., 2005. The effects of preservatives and temperatures on arachnid DNA. Invertebrate Systematics. 19, 99-104. Williams, S. T., 2007. Safe and legal shipment of tissue samples: does it affect DNA quality? Journal of Molluscan Studies. 73, 416-418. peter.mail2@unmack.net