If you work on plants and you've extracted DNA from them in the last 20 years, chances are that you've used some version of the method described by Doyle and Doyle.1 According to Google Scholar, Doyle and Doyle has been cited more than 4000 times. The method is reasonably straightforward, but tedious: disrupt membranes with CTAB2, add a mixture of chloroform and isoamyl alchohol, precipitate DNA from the aqueous phase using cold isoamyl alcohol, wash the precipitate with 70% ethanol, and re-dissolve.In the most recent issue of the online-only Primer Notes and Protocols section of the American Journal of Botany there's a new paper by Dirk Bellstedt and colleagues describing a method for direct PCR amplification that may make life even simpler. You'll need to read the paper to get all of the details, but here's the basic procedure:
- Grind your plant material in a buffer.
- Centrifuge the mixture and mix the supernatant with a GES buffer.3
- Use the resulting mixture in PCR.
1Doyle, J. J., AND J. L. Doyle. 1987. A rapid DNA isolation procedure for small quantities of fresh leaf tissue. Phytochemical Bulletin 19:11-15.
2hexadecyltrimethylammonium bromide
3Recipe provided in the appendix.
Bellstedt, D., Pirie, M., Visser, J., de Villiers, M., & Gehrke, B. (2010). A rapid and inexpensive method for the direct PCR amplification of DNA from plants American Journal of Botany DOI: 10.3732/ajb.1000181
Plant DNA geek? Did I hear someone say my name?
I just read this via your link, and I will stay with CTAB. Out of the several groups of people they pitch this to, maybe some would go for it because it is fast, but I doubt it will hold much appeal for systematists. The instability of the samples would be a deal-breaker for me (storage for "several weeks" at -20 as opposed to years/decades for DNA preps), as well as the inability to extract from herbarium specimens.
The authors WAY overstate the time and agony involved with CTAB extractions in order to make their point. You can certainly have clean PCR-able (and stable) DNA samples from CTAB protocol on the day you start them. With a little mechanical tissue disruptor you could do 100 small preps in a day without a ton of effort.
This method probably has a place for certain applications, but I think the citation counts on Doyle and Doyle will keep climbing for now.
Robie
Cool. I don't do much wet lab work anymore, but I do remember long days spent on dna extractions from dozens of samples. Combining paint-shaker grinding with the ability to skip all the post-grinding steps could definitely speed up things up for large genotyping projects.
Thanks for the insight.
Have to look at the paper, but there's also the caveat that there are more copies of chloroplast DNA per cell than copies of nuclear DNA, so you can get away with less-than-precise methods. Also, any application I can think of is better done with DNA you can archive for the long term, for the sake of error checking, reproducibility, and the option to go back and do something totally different later.
I don't know enough about the chemistry of DNA extraction to comment on long-term stability. The authors say "Plant samples prepared in this way can be stored at –20°C for a period of several weeks...." As I read this, the meaning is ambiguous. It could mean (a) that the samples can only be stored and used for several weeks or (b) that they could be stored for at least several weeks and possibly longer. Is it clear for one reason or another from the extraction protocol that the DNA samples are likely to be stable for only a few weeks? If so, the method may not be as widely useful as I thought.