Dear all, As some of you requested, here are the replies I received (even the ones written in portuguese). I did a copy-paste on the answers but in case you need any further information, feel free to contact me. Once again, thank you for all the information and hope it helps for those (also) in need! Best regards, Carla Pereira Hi, I just read your thread on Evoldir. I sometimes had problems amplifying any sort of marker from my DNA preps (nothing molluscan though), even when they contained ample amounts of DNA. I found that an RNA digest of the DNA extractions got rid of that problem. Somehow RNA fragments seemed to interfere with my PCRs. Not sure if you do an RNAse digest with your extractions anyway, but if not that might be worth a shot. Cheers, Basti Bastian Bentlage Dept. of Ecology and Evolutionary Biology The University of Kansas Lawrence, KS USA hello, you might've tried this already, but, i've extracted RNA from mussels and found that doing a lithium chloride re-precipitation (after you've extracted the DNA fully and resuspended do another precipitation). The protocol is in Molecular systematics (hillis, moritz, and mable) in the rna/dna extraction chapter. best of luck! Anne Dalziel University of British Columbia Vancouver, Canada Hi, We have been working on a few molluscan species in our lab - mostly squid and abalone. Generally we have fouund the Qiagen DNEasy kits to work pretty well for DNA extractions for these species. However, you did not mention how your tissue was preserved, or how old the specimens were. One thing we have found is that samples more than a couple of years old can be a bit hit and miss, especially if they've been stored in less than perfect condiditon. Basically, the fresher the sample, the better the chance of successful DNA extraction and PCR amplification. I think generally, from my experience, alcohol preservation works better than freezing for molluscs and other species that have high mucopolysaccharides. You also havent mentioned what primers/approach you were using for the PCR. Again we have found that most "universal" primers are unreliable for molluscs and that it is pretty important to develop species-specific primers. If you have managed to get at least some individuals to PCR and sequence, then check how well your primers match, it may be that there has been a small change in the primer-binding site that is causing your problem and it might be a good idea to try to design new primers. I hope this helps Regards Karen Miller Dr Karen Miller Institute of Antarctic and Southern Ocean Studies University of Tasmania AND Australian Antarctic Division Private Bag 77 Hobart, 7000 Tasmania Australia Hi, I have recently worked with a number of marine organisms, including molluscs, crustaceans, fish and ascidians, and I generally find that the molluscs and fish are among the easiest to work with (I worked with snails of the genera Siphonaria and Nassarius, as well as the mussel Perna perna). You lack of success may be due to a number of other factors, e.g. the tissue you are targeting is not ideal (I usually cut off a very small piece from the foot of the snails), you are using too much tissue, the samples were not preserved well (frequent freezing and thawing degrades DNA, have you tried fresh tissue?) or the primers you are using are not working for all your species. I have found that if it's mtDNA you're after, Folmer's COI primers usually work well (Folmer et al. 1994), and Palumbi's (1991) 16Sar and 16Sbr combination is also reliable. But I have just found a Siphonaria species for which the COI primers do not work, and have used a different design for that. If you're suspecting that the primers are the problem, I suggest you get both of these and if one works and the other doesn't, you know where the problem is. Hope this helps and best of luck, Peter Dr Peter R. Teske Postdoctoral Researcher Molecular Ecology Lab Dept. of Biological Sciences, E8C Macquarie University Sydney, NSW 2109 Australia Hi, Try adding BSA (Bovine serum albumin) to your PCR to around 0.8 µg/µL final concentration. It works well in removing inhibitors. Good luck Jonas Hello, I would much appreciate if you could forward any answers you might get to me (or the list server), since we are facing similar problems and are quite interested to hear possible solutions. Thanks a lot in advance, Stefan We always extract our Littorina samples with this protocoll: Mitochondrial cytochrome b sequencing. DNA was extracted and purified using a modified CTAB protocol (Winnepenninckx et al. 1993): a small piece (1-2 mm 3)* *of foot tissue was homogenized in 300 μl *[You can increase this up to 700]* CTAB buffer (100mM Tris-HCl pH 8.0, 1.4 M NaCl, 20mM EDTA, 2% hexadecyltrimethylammonium bromide [CTAB] and 0.2% β-mercaptoethanol) and 10μl of 10mg/ml proteinase K *[You can increase this to 20]* and kept a minimum of 3 h at 60 °C *[over night works fine as well]*. DNA from the homogenate was extracted using a standard chloroform-isoamyl alcohol (24:1) extraction [same amount as CTAB buffer] with a triple alcohol precipitation *[First time isoamylalcohol, same volume as CTAB; second and third times 70% EtOH, double volume compared to CTAB. Keep in -20 for one hour min. before you gently pour out the alcohol]*. Due to the mucoid nature of the tissue Phase Lock Gel Light (Eppendorf) was used during centrifugation to increase yields *[read the manual]*. This is from a paper I'm writing *[additional comments]*. Good luck! /Petri Petri Kemppainen, PhD student Tjärnö Marine Biological Laboratory, 45296, Strömstad, Sweden Hi, The problem you are facing is very common in the molluscan world. Usually the CTAB 2x extraction protocol works pretty well in most molluscan samples. I would try it for sure. Find attached the protocol. I wish you good luck. Aris Oi, Eu vou temptar falar portugues, mas na verdade nao posso falar muito :). Aqui as minhas suggestoes: Tenta usar protocoles de extracao a base de CTAB (buscar no internete por isso, eu annexei um protocole em baixo). A metode de CTAB e melhor por extracoes com muitos mucopolysaccharidos - esse tipo de probleme occore se com varias tecidos de planta - possivelmente e uma boa ideia de dar um olhada en ese literatura tambem. Hope this helps - please don't post my reply back to evoldir :) Tudo de bom, -Kai Hi, I have had some similar problems working with both bivalves and with bat wing tissue samples. In the case of bivalves, we have occasionally had to re precipitate/wash the samples with an additional LiCl precipitation after a salt-based DNA extraction to get rid of PCR inhibitors. In the case of the bat tissue, it appears as if too much pigment interfered with PCR. This was remedied to some extent by using a Chelex based method of DNA extraction (BioRad's Instagene matrix). This method can remove or decrease inhibitors sufficiently to allow amplification for some samples. In any situation where you suspect PCR inhibition, you might want to set up the following experiment to test the hypothesis of PCR inhibition. The test consists of performing PCR on three DNA samples: the first tube will contain a positive control (i.e., a template that consistently works), the second tube will contain your problematic sample that doesn't work, and the third tube will contain a mixture of the first two samples. IF PCR inhibitors really are the problem, then the third tube shouldn't work because the inhibitors in the second sample should "shut it down". If it does work, then maybe you have a different problem. Cheers, Donald Stewart Hi, have you tried the protocol from Winnepenninckx et al (1993) Trends in Genetics? It is a quite old one, but work well on freshwater molluscs. I have been using it mainly for Radix. A colleague used to do the chloroform step twice to get rid of more proteins... We are performing both microsatellites analysis and sequencing with these DNA extractions... Please note in the paper: it is 2 to 3 times the volume in isopropanol, and not two third of it. It is a bit confusing. I hope you get better results with that... Good luck Mathilde -- PhD student Mathilde Cordellier Abt. Ökologie & Evolution J.W.Goethe-Universität BioCampus Siesmayerstraße D 60054 Frankfurt am Main Germany Hi, I have worked with molluscs (Hydrobia and Potamopyrgus) and met similar problems. The best recepie for dNA extraction is to have the samples in liquid nitrogen, extract fresh live samles or samples that were straight away stored in -85. I found that samples used in alcohol or samples stored in higher temperatures than -85 centigrades were almost impossible to yield nice DNA from. The best ones were the ones where a bit of fresh tissue was used immediately I found that the QiaGen DNA extraction kit by using the CTAB protocol gave good DNA and enabled both nice sequences and microsatellites. If you look in the literature you can see that many researchers try to transport their molluscs alive to the laboratories. I hope you have the best of success Good Luck! Micaela Hellström PhD Student Åbo Akademi University/Stockholm Universiy Hi, I have received your email from someone working at the Utrecht Behavioural Ecology department. I myself am a student from Leiden University, and have been working with molluscan tissues too. In my study I've tried to collect mitochondrial DNA (COI, 16S) from several species found at the Azores, (Columbella adansoni, Nassarius incrassatus and Anachis avaroides) I've collected these gastropods in 2006-2007 and started lab work in march 2007. However no PCR would yield any amplified fragments or if something was visible after gel electrophoreses it was one big smear of DNA..... Just like you I've tried several extraction kits (Ezna Mollusc Kit and Qiagen Extractionm kit) to no avail. I also tried chloroform (part of the Ezna protocol) and also extra chloroform steps to clean the DNA. Dilutions 1:10 to 1:100000 didn't yield results either..... BUT! When after 5 months I wanted to give up and tried one last time some simple idea of mine I got good results... Now I do not know whether it will help you... but the protocol is very cheap and simple so what harm can it do? I will explain what I think the problem was and it will be followed by the solution that worked for me... At the Azores I stored my samples in alcohol which I derived from the supermarkets, the so called medical alcohol. We (my lab colleagues and I) think that this was the culprit but we have yet to prove it. We think in order to keep people (alcoholics) from drinking the medical alcohol, the producer puts ether in it (happens in other countries), which has a very bad taste. However ether is known to bind to DNA... so you see where this is going... No extraction kit contains chemicals to get rid of the ether and apparently chloroform cannot separate the ether-dna complex either. So what did I do? - I took my sample, - crushed the shell, or pulled out the tissue with a pincer - cut up all the tissue into very tiny fragments. - placed the tissue in clean laboratory alcohol - let it stay for a few days, maybe shake it a few times a day - refresh with good alcohol - again let it stay for a few days - repeat if nessecary (for me letting it stay between 1 and 2 weeks was enough as long as I kept the alcohol refreshed often) - use an extraction method with chloroform cleaning (without worked also, but chloroform went better) - I used 1:100 crude dna dilutions in the next steps (sometimes 1:10) - pcr, clean, sequence etc this simple thing got me proper results... I think the tissue and alcohol were saturated with ether (or maybe other inhibitors) and by rinsing it with fresh alcohol without ether it dissolved again from the tissue into alcohol and could thus be discarded. DNA yields were still low (nanodropping gave about 30-50 ng/microliter) but enough to get proper sequences. As mentioned before we are not certain if it was ether (since there is no contents on the alcohol bottles, but collegues have seen the same with other samples stored in this, hence our 'educated' guess) However it might have well been some other cause, which you might have encountered too. That's why I've sent you this rather long email, hoping it might help you... Good luck with your work, With best regards, Henk van Goor Hi, I've had a lot of problems extracting DNA from sea urchin, and even if I'm not so experienced in the field I'll try to give you an advice. I don't know if this is your case but with my sea urchins the DNA yield was good, and the DNA quality seemed to be fine, nevertheless we were not able to amplify. If you think your main problem is the lysis of tissues and cells or some inhibitor you can try to homogenize a small portion of tissue into a potter with a buffer (0.25M sucrose, 5mM HEPES, 1mM EDTA pH 7.2) keeping the potter in ice during the procedure because the friction releases a lot of heat! After that you can separate the mitochondria by simply spinning the lysate obtained at 500 x *g* for 10' at 4 °c and collecting the surnatant in fresh tubes. Than you can go on with the method from Palva & Palva (Palva & Palva, 1985, rapid isolation of animal mitochondrial dna by alkaline extraction.) for extraction of mtDNA. The extraction protocol that gave me the best results is a phenol/chloroform protocol involving a PEG (polyethylene glycol) 12% / NaCl 1.2 M solution for the DNA precipitation. Hope I've been helpful somehow, MB. PS: feel free to ask for further details if You are interested! *Michele Barbieri *Ph.D. student *Università di Pisa Dipartimento di Biologia Unità di Biologia Marina ed Ecologia *Via A. Volta 6, 56126, Pisa - Italy (I) 1. try a column based extraction (Qiagen DNeasy, Promega Wizard) after grinding under liquid nitrogen. 2. run it through the spec and dilute the template to <50ng/ul 3. try using a really good enzyme--high fidelity Taq etc. Otherwise I have had good luck (crustaceans, molluscs) with a few elaborate plant preps--they tend to be much more involved than the Winnepenicx et al. mollusc prep (which is based on a plant protocol anyway). Best of luck Paul Paul S. Schmidt Department of Biology University of Pennsylvania Hi, We've been working with Strombus gigas. We've had pretty good success using the Qiagen DNeasy kit, doing an overnight digestion with proteinase K. This has worked fine for mtDNA. Best regards Rich Richard M. Kliman Associate Professor Dept. of Biological Sciences Cedar Crest College 100 College Drive Allentown, PA 18104 Hi, I have done some previous work with mollusk tissue for DNA and RNA analysis. The first thing that I would look at would be the DNA quality (ran out on a gel and a A260/A280 ratio via spectrophotometer). Anything above 1.7 is generally considered to be good-quality. Lower readings indicate contaminants. Additionally, an A260/A230 ratio below 1.5 indicates increasing levels of salts and/or organic compounds may have co-purified with the DNA. A comparison of gel images and absorbance readings comparing the samples that have worked and those that have not may also be helpful. My preferred method of dealing with the polysaccharides associated with mollusks is to perform a CTAB extraction. Having used plant kits and various phenol:chloroform methods, I found the CTAB procedure to be the best method for the task. If your gel and absorbance readings indicate that the DNA you have already extracted is of high quality, it may be something in the PCR reaction that needs to be looked at. If it is determined that the quality of the DNA is less than optimal, you will have to determine the culprit and take measures to eliminate it. Good luck with your work and studies, Bernie Ball Department of Biology Box 90338 Durham, NC 27708 Hi, I just posted two protocols on my blog for you: http://shiftingthroughscience.blogspot.com/2008/03/mollusc-dna-extraction-ctab.html http://shiftingthroughscience.blogspot.com/2008/03/mollusc-dna-extraction-formalin.html Cheers, Andrew ------ Andrew G. McArthur, Ph.D. Email: amcarthur@mac.com, Web: http://mcarthurlab.blogspot.com Phone: 905.296.3252, Mobile: 905.745.2794, Fax: 647.439.0829 AIM: amcarthur@mac.com, Skype: agmcarthur * * Hello, I had a similar problem extracting DNA from a polychaete tubeworm I studied for my Masters. Seemed as though I would get a good amount of DNA that then would not perform well in downstream reactions. Here is a link to a primer notes paper I published that talks about the extraction procedure http://www.blackwell-synergy.com/doi/abs/10.1046/j.1471-8286.2003.00453.x Basically, everything you mentioned but we also used a CTAB step. The reference within the paper for: Winnenpenninckx B, Backeljau T, De Wachter R ( 1993) Extraction of high molecular weight DNA from mollusks. Trends in Genetics, 9, 407. might also be worth looking at. Good luck, James James B. Pettengill Behavior, Ecology, Evolution and Systematics 2174 Plant Sciences Building University of Maryland College Park, MD 20742 USA** Hi Carla two suggestions: use a CTAB method (no kits required!) to remove MPS often, MPS are cited as the source of problems, but another potential source is divergence of primer sites away from the conserved primer that you are using (eg a mutation of a single base at the 5' end would do it). Angus Hi, I saw your message posted on the EvolDir website. I completely understand what you are going through (very frusterating!) - my MS and PhD work focused on the mitochondrial genomes of oysters. The first thing I can suggest is to try the OMEGA Biotek E.Z.N.A. Mollusc extraction kit ( http://www.omegabiotek.com/genomicmolluscmini.asp) - if you have available funds. This kit is specifically designed to deal with mucopolysaccharides, etc. I've had very good luck with it. If your samples are preserved in ethanol, be sure to let the ethanol evaporate before you add it to any extraction buffer (ethanol with interfer with the proteinase K, etc). Also, if you are using a filter column, be sure that your tissue is fully disolved, otherwise your column will become clogged and adversely affect the extraction. Secondly, many mollusk species exhibit extremely high levels of polymorphism. Its possible that you are having difficulty amplifying due to polymorphism in the priming site. To determine if this is the situation, ask your advisor to help you find some universal primers (16S, COI, etc). If you get good amplification with universal primers, you can rule out any problems with DNA quality, your reagents, concentrations, etc, and then you can conclude that primers and polymorphism may be a problem. If you get good amplification with universal primers, you may want to redesign new primers to target highly conserved regions. You can also try running nested PCRs - this should help even out your amplification success. Good luck! Coren Milbury Hi Carla, I just talked with a lab tech who did his Masters project on mitochondial COI variation in a local gastropod. His prc didn't start working until he diluted down to 1:1000. He suggests stepwise dilutions until it starts to work. I use GeneReleaser to give me better amplification, but I don't work on gastropods. Allegra Allegra Briggs Graduate student, Kimmerer lab Romberg Tiburon Center, San Francisco State University * * * * Hi Carla, I have done some previous work with mollusk tissue for DNA and RNA analysis. The first thing that I would look at would be the DNA quality (ran out on a gel and a A260/A280 ratio via spectrophotometer). Anything above 1.7 is generally considered to be good-quality. Lower readings indicate contaminants. Additionally, an A260/A230 ratio below 1.5 indicates increasing levels of salts and/or organic compounds may have co-purified with the DNA. A comparison of gel images and absorbance readings comparing the samples that have worked and those that have not may also be helpful. My preferred method of dealing with the polysaccharides associated with mollusks is to perform a CTAB extraction. Having used plant kits and various phenol:chloroform methods, I found the CTAB procedure to be the best method for the task. If your gel and absorbance readings indicate that the DNA you have already extracted is of high quality, it may be something in the PCR reaction that needs to be looked at. If it is determined that the quality of the DNA is less than optimal, you will have to determine the culprit and take measures to eliminate it. Good luck with your work and studies, Bernie Ball Department of Biology Box 90338 Durham, NC 27708 * * Hi Carla, This is a well know problem with molluscs. I suggest first trying a DNA extraction kit for plants which should include a step to remove the polysaccharides. sincerely, Elizabeth Boulding. Carla, I had some of the same problems working with tissue from Acanthina sp. in the Gulf of California. I tried multiple extraction techniques and found the one that worked the best was the Qiagen DNeasy kit for animal tissue (blood). I used a small (grain of rice) piece of foot tissue and allowed it to lyse overnight. After the final step I also performed a cleaning step using EtOH. That worked well for me, yielding between 20 ng/ul-200 ng/ul of DNA. I used the final product for AFLPs. In order to optimize my reaction I did not dilute the PCR product. Hope this helps. Raena PS. I know that some people have used a CTAB extraction successfully. Hi Carla, When I was working on the gastropod Littorina using AFLP then I had difficulties with my extractions (particularly for restriction digests). The only satisfactory DNA extraction method that worked consistently is described in the paper which I have attached. Please note that the Wizard DNA clean up was critical. Craig** * * * * I would recommend using a chelex protocol, I am not in my lab now, I do not have any here but I am sure you can get it somewhere, (it worked for Solen marginatus!) Miguel Angel Varela Muinho Avoid using any gut tissue. There are many nasty enzymes in the gut. If you can snip of the head, for example, only use the head. We have used successfully 'DNAMITE for difficult cells' from Microzone Ltd. for 'difficult' barnacles. I always think that phenol ruins more than it makes good. Tried CTAB buffer? Good luck! Cheers, kirsten** * * * * Have you tried 2x CTAB in the extraction buffer. Seems to work well for mussels and Nucella. Look at Collins TC et al. 1996 Evolution for a reference. Best, Peter Hi Carla - > No doubt you have received numerous responses by now to your question, > but I work with all sorts of molluscs and have had excellent success > with the standard Qiagen DNeasy tissue kits. Although I have used > various approaches including CTAB, PCI, Plant kits, etc., Qiagen works > great and is very easy and non-toxic. Brenden Dear Carla, regarding your query on evoldir: Are you aware of Sokolov, E. P. 2000. An improved method for DNA isolation from mucopolysacharide-rich molluscan tissue. J. Moll. Stud. 66:573-575? I attached the pdf file. Colleagues of mine use it for molluscs and are happy. They modified it in such a way that they simply do it without (!) the phenol/chloroform steps. And it works fine. We also use it that way for insects and it yields good DNA for sequencing and AFLP. Anything you want to get rid of is simply salted out and precipitated with the potassium dodecyl sulfate. The rest is a simple ethanol precipitation. Good luck, Gregor Make sure you use a very small piece of tissue, maybe 5 to 10 mg wet weight. Your problem is not caused by having too little tissue, your problem is caused by having too much mucopolysaccharides. Using a CTAB procedure on a tiny piece of tissue should work well. Good luck! John H. McDonald Department of Biological Sciences University of Delaware Hi, I think you will have great success with CTAB, that was my last resort after trying everything else. Good luck and let us know how you are succeeding Kind Regards Micaela Ola Carla, Eu tive o mesmo problema que tu quando comecei o meu PhD. No meu caso eu estou a procura de genes sobre pressao evolucionaria em Cepaea *nemoralis*uma especie de caracol. Neste momento estou a acabar uma genomic library de microsatellites. E como ja reparaste e um ador de cabeça. Como tu tambem tentei liquid nitrogen, mas nao vale a pena. Pelo contrario, achei muito dificil de raspar o material para dentro do tube e aumentei a superficie em contacto com as enzimas. O que eu acho melhor e curtar um bocado do musculo 5x5mm e meter de imediato o tecido no estraction buffer. CTab requires a long incubation period, but it's worth it. Em termos de extraccao de DNA o melhor metodo em termos de yeld e extraccao com CTab. Mas se queres DNA de qualidade e nao te importas tanto com a quantidade Quiagen plant kit e muito bom e rapido. Tentei o kit para animais mas nao tive success. Mas lembra-te que eu uso genomic DNA. Boa sorte e espero que isto ajude. Bruno * * shortysquab@gmail.com