Dear list: My request for protocols for getting DNA out of bone and formalin-fixed generated lots of response. Thanks for your time and information! Below is a collection of all the responses so far. I can't comment on the efficacy of any one of them yet, but I'm working on it. Thanks, Steve ** If you are not having success I suspect your problems are one/some of the below a) There is absolutely no DNA in the samples - in particular if the material was in formalin for more than ca. 12-24 hours it will have been heavily crosslinked, and even fragmented if it was in unbuffered formalin b) There is DNA but the primers you are using are targeting too big a fragment. How old are the samples and how long were the fixed ones fixed, and how long ago where they fixed? In general, a simply and effective bone protocol is that by Rohland et al, attached. If that doesn't work, and you are using really short PCRs (ca 100bp) my guess is it won't work. For formalin, I once did a lot of work on it. Please see the extremely tedious paper that I wrote as a result. As you will gather from it, heat + alkali can help matters (the Shi 2004 paper I cite), but in general by increasing yield only. If there is no DNA...it won't do much. Happy to help with any specific questions. ** Hi Steve, Dealing with formalin-fixed material is not an easy task, as you might already know. We recently published a paper using a new method based on Tetramethylsilane (TMS)-Chelex to reduce costs compared with Fang et al. (2002). Most importantly, we observed that neither an indigestible matrix of cross-linked protein nor soluble PCR inhibitors impede PCR success when dealing with formalin-fixed material. Instead, amplification success from formalin-fixed tissue appears to depend on the presence of unmodified DNA in the extracted sample. We were not able to get DNA sequences out of every sample, but we provide a 'test' to check if DNA will be useful for PCR. You can obtain a pdf of our paper at http://www.icm.csic.es/scimar/index.php/secId/6/IdArt/3907/ I hope this helps. Cheers, ** Dear Steve, Take a look on this article ( http://www.pnas.org/content/105/40/15464.figures-only). The authors extracted DNA from museum specimens (mostly bones) of the giant tortoises of Galapagos. Take care ** Hi Steve, I think you're wasting your time with formalin-fixed tissues, as this issue has appeared on EvolDir about twice or thrice a year for fifteen years...but you should be able to get DNA from the carapace. For ideas, see Biotechniques. 1999 Jun;26(6):1086-8. Microsatellite locus amplification using deer antler DNA. O'Connell A, Denome RM. PMID: 10376147 [PubMed - indexed for MEDLINE] also http://oldwww.wii.gov.in/ars/2006/imran_khan.htm ** If you are interested in using a kit to do the extraction, there is an excellent protocol on the Qiagen website for extraction of DNA from bone. It is called "Purificant of total DNA form compact animal bone using the DNeasy Blood and Tissue Kit". I recently used it to extract over 40 toe bone specimens from a mouse species with a lot of success. If not I also have a standard ethanol precipitation method that I have used with success in bones as well. Let me know if you would like that. ** I would suggest you post your message on the ancient DNA forum at: https://www.jiscmail.ac.uk/cgi-bin/webadmin?A0=ancient-dna Several people worked on Museum specimen. I work on bones so let me know if you want a few protocols. Usually the one from Hofreiter & Rohland is the one most aDNA people use. ** Steve, I saw your message this morning on Evoldir. Coincidentally, I am working on a very similar problem with the goal of extracting DNA from turtle claws. We have had success with fresh tissue and we have been able to amplify a 630 bp fragment from the mtDNA control region of painted turtles. During the past month, we have been conducting experiments where we expose claws to buffered formalin for 48 hours followed by 4-5 days in 70% EtOH (modeling museum protocol), and then attempting extraction using a modified protocol from the Qiagen blood and tissue kit (basically the addition of DTT, additional proteinase K, and longer digestion times). So far the results are not too encouraging but we are actively trying some different things and have some claws from specimens of different collection date from the Field Museum that we will test once we're confident in a protocol. ** Dear Steve, There are many methods that can be used for extraction and almost all of them work but it is also a matter of how are you testing the extracts to determine their success. With degraded DNA you have a few problems that have to addressed; 1) Quantity 2) Quality 3) Inhibition If you are trying to quantify using real-time qPCR then you have to solve the other two problems first as these will both affect your ability to amplify the DNA thus quantify. If you are determining the concentration in other ways you may also get background or contaminating DNA that may not contaminate the PCR but are there as background DNA because the sample has sat in a museum for a long time. If you are determining success based on PCR amplification (success or failure) then you also have to consider 2 and 3 first. To overcome quantity you add as much material as you can to ensure quantity is abundant. Some people have used up to 5g. Once you have resolved all the other problems then you can optimize quantity because 5g is a lot of material and sometimes too much of a very valuable sample. I regularly use between 10-50mg of bone material. There are a number of problems with degraded DNA that affect their quality. 1) Fragmentation 2) modification 3) cross-links All degraded DNA will be fragmented with single stranded and double stranded DNA breaks. Most research agree that 200bp is the norm in nDNA. So if you are trying to determine success by PCR your primers have to amplify less than 200bp. 200bp is the amount of DNA that wraps around the histone complex and in a way protect the DNA where the breaks more frequently occur in the inter-linker regions between the histone complexes (Look at the Apoptotic 200bp ladder for proof). All degraded DNA is modified this is where you get abasic (AP) sites due to depurination and depyrimidation, modified bases where the amine is lost through deamination. There is also oxidative damage generating hydantoins amongst many other types of modified bases. Some of these will prevent your DNA from being amplified. Some of these will generate miscoding lesions where the sequence is not authentic due to an incorrect base pair generating an erroneous sequence. Some researcher have incorporated repair methods to overcome this step (as have I). The other form of damage is cross-links which can occur between the DNA and other molecules like metals and proteins (amongst others). There has been some research to develop methods of cleaving these cross-links off to allow the DNA to be analysed. All of these have to be considered for degraded DNA. Finally there is inhibition. If you do not resolve this problem first you will not be able to do anything. Inhibition is going to be unique to the site, the collection or even the sample. It must be determined if you have it (spiked PCR) or ensure you have removed it through various purification systems. So with these points in mind go back to the methods in the literature and you will find they will work. The most common methods in degraded DNA studies are proteinase K methods and Guanidinium thiocyanate but there are other chaotropic salt methods that can also work very well. I can give you dozens of methods of extracting degraded DNA that I have personally used but without considering these factors you will not succeed with any of them. Let me know if you need any more information on any of these points or reference papers on any of these points. ** Hi Steve, It is almost impossible to get anything out of a formalin fixed specimen (I tried most of the available protocols on a muscle sample that was only 5 years old and got nothing and I have heard the same thing from most people that I contacted). As for getting DNA out of bone or alcohol fixed specimens, your luck should be better (although if the samples have been stored in a hot place, DNA will degrade much faster). Have you tried any of the bone protocols yet? Your best bets are the two attached. Also you need to be sure that you are doing all of your DNA extraction and PCR set up in a lab far from any labs that work on turtles (in my ancient DNA lab, we do all our work in the morning or on days that we don't go to the modern lab unless we go to the gym to shower before returning to the ancient lab). Finally, you need to be sure that your PCR primers are sensitive to low amounts of DNA and that they amplify products less than 200 bp in length (BSA added to your PCR also helps yield). The ancient DNA literature might be of help to you in terms of suggestions and recommendations for avoiding contamination from modern DNA. Good luck! ** Hey Steve, for the extraction of bone samples from museum specimens I can strongly recommend the latest version of the silica based protocol from Nadin Rohland and Michael Hofreiter. I attached the pdf. ** Hi Steve, I've used the Shedlock extraction protocol with success on seahorses and use the guanidinium non-destructive method on parrotfish bones from archaeological sites (1300 AD jaw bones and teeth). Have you tried these? Also, I would suggest you cut down your PCR product to ~150 bp...you may be getting DNA but its too short and degraded to amplify in large pieces. I amplify 150-250 bp fragments and piece them together. Design short primers, test them on modern good quality DNA to make sure they work, then use a good quality hot start Taq, touchdown PCR (like 65 - 45 C) for 65 cycles, with short extension times (30s denature, 30s annealing, 30 extension - will prolong the life of the taq over the large number of cycles), and relatively high MgCl2 (2.5 mM), high primer concentration 0.5 mM, and high dntp concentration (.25 mM) and BSA at 1 mg/ml. That way if there's any DNA in your PCR, it will work...but be very careful of contamination :) Hope that helps and feel free to write back with questions, -- Steve Kimble PhD student, Department of Forestry and Natural Resources Purdue University skimble@purdue.edu sjkimble@gmail.com 205.337.4843 http://web.ics.purdue.edu/~rodw/Steve%20Kimble.htm steve kimble