Cooking with soil

Beardyman knows how to combine the various beats in his kitchen to produce something delectable. The culinary analogy extends to analysing soil in the laboratory too and, while unlikely to grace the pages of the Sunday supplements, thinking about some complicated labwork in terms similar to baking a cake is a good way of making it less daunting – like baking, you need to be precise with your timings and temperatures, and not mix things that aren’t supposed to be mixed. Easy! Below is a recipe for a analysing the microbial community in a peaty soil (using a method called multiplex-Terminal Restriction Fragment Length Polymorphism, or m-TRFLP, but you didn’t really need to know that…). Don’t try this at home, obviously.

You will need:

  • hexadecyltriammonium bromide in salty water
  • polyethylene glycol in salty water
  • phenol
  • chloroform:isoamylalcohol
  • 70% ethanol
  • some very expensive primers
  • some restriction enzymes
  • a lysing machine
  • a centrifuge
  • a thermal cycler (PCR machine)
  • a PCR clean-up kit (I prefer Wizard)
  • a sequencer
  • plenty of ice
  • lots of gloves
  • DNA remover
  • a UV cross-linker for destroying DNA on tubes
  • lots of little tubes
  • some soil (finely chopped, frozen)
  • a couple of days
  • reserves of patience


  1. Measure out 0.5 g of soil, and add to a lysing tube with some phenol, hexadecyltriammonium bromide solution and chloroform:isoamylalcohol. Do this in a fume cabinet, otherwise things will go terribly wrong. Stir.
  2. Shake the mixture for about 30 seconds at 5000 meters per second in your lysing machine. Ensure that your lids are securely fastened to your tubes, otherwise your soily chloroform-phenol mixture will spray itself all over the inside of your lysing machine, and you’ll have to spend ages cleaning the machine while holding your breath.
  3. Put your mixture on ice and take it to the centrifuge. Once again, ensure that your lids are securely fastened. Whizz for ten minutes at 10,000 rpm.
  4. Carefully take your mixture out of the centrifuge and transfer the top, clear layer to a clean tube. Avoid hoovering up any of the gooey soily mix with your pipette. Add more chloroform:isoamylalcohol to the clear liquid and return to the centrifuge for five more minutes. Don’t forget to autoclave your hazardous waste!
  5. Transfer the top layer once more into a clear tube, and add some polyethylene glycol solution. Stir well, and leave on ice (not in the fridge, where someone might mistake your tubes for free samples of some quirky new energy shot product) for a couple of hours.
  6. Put your tubes back in the centrifuge at 10,000 rpm for ten minutes. You should now have an almost-invisible pellet of DNA at the bottom of the tube. A steady hand is required for the next step, so go easy on the coffee.
  7. Carefully remove the liquid from your tube, replace with ice-cold 70% ethanol, and centrifuge again for five minutes. Don’t be tempted to sample the ethanol.
  8. Remove the ethanol from your tube, leaving your DNA pellets in a warm, dry place for about half an hour. Once dry, add a drop of autoclaved water to your tube, and place in the fridge.
  9. At this point, it’s a good idea to check the concentration of DNA in your samples, to see whether your extraction has worked. Ask a grown-up to do this for you.
  10. Pre-program your thermal cycler. Add your choice of primers to your DNA, reading the instructions carefully. Stir well. Put your tubes in the thermal cycler for about three hours. If, at this stage, you’re exhausted, you can leave your tubes in the thermal cycler overnight, as long as you’re not planning a lie-in the following morning.
  11. Use a Wizard (kit) to clean-up your newly-amplified DNA. Pre-program your thermal cycler again, ready for the restriction digest.
  12. Add your restriction enzyme of choice (I like Hha1) to your cleaned-up DNA. Place in the thermal cycler for about one and a half hours.
  13. Check the final concentration of your DNA (ask a grown-up to help you with this). If you haven’t booked a slot on the sequencer, now is the time to go pleading to your lab manager.
  14. Place your tubes in the sequencer, overnight. In the morning, serve your data with some multivariate stats and a tentative garnish of interpretation. Delicious!

Weekly resource roundup: 20th May

I’ve just got back from a couple of days installing gas sampling kit, so this is a bit late, but better than never, here’s the selection for this week…

Surprising weather at Moor House

Surprising weather at Moor House

1.      Multiple to-do lists: Something I’ve been experimenting with recently is using three to-do lists to prioritise tasks. I use three notepads, for today, this week, and whenever, but there are other ways of organising it. I quite like it as a system for getting information out of my head, to make room for more!

2.      Gantt charts: Gantt charts are a good way of mapping out your project in the longer term. GanttProject is an easy-to-use program that runs on Windows, Mac or Linux and can be downloaded for free.

3.      May Methods Digest: online here

Weekly resource round-up: Monday 10th May

This week’s handy things:

  1. The latest issue of Methods in Ecology and Evolution is available online, and features some interesting stats articles (‘Do not log-transform count data’ for example).
  2. Interesting paper on free and open-source geospatial tools for landscape ecology, previously posted about here.
  3. This blog post features some useful tips for academic poster design.
  4. A few conference abstract deadlines are coming up:
    1. GFOE 40th Anniversary Meeting, Giessen, Germany: 25th May
    2. Organic Matter Stabilisation and Ecosystem Functions, Presqu’île de Giens, France: 15th May
    3. BES Annual Meeting, Leeds: 10th May (today!)

Weekly resource round-up: Monday 26th April

Methods in Ecology and Evolution journalI recently came up with the idea of composing a brief weekly email to send to students (I’m a student representative for PhD students at CEH Lancaster). I tend to hoover up items I think might be useful to myself and others during the week, compose the emails on Fridays, and set them free on Mondays (because who wants to get an inspiring list of things to look at last thing on a Friday?).

I’m archiving them on this blog; here’s last Monday’s:

  1. The Royal Geographical Society have produced a guide on publishing for new researchers. Although the title specifies the geographical discipline, I found the material to be general enough to be useful. It can be downloaded here.
  2. The new Methods in Ecology and Evolution journal is worth keeping an eye on if you’re into that kind of thing. The posts monthly digests of noteworthy methods papers published in other journals…
  3. …which brings me onto a really handy paper published in the first issue of MEE by Zuur et al. (2010): ‘A protocol for data exploration to avoid common statistical problems’. It’s already been downloaded over a thousand times, and the authors publish their R code and example data on their website. Here’s the paper:
  4. For anyone planning (but possibly too frightened) to get into using R for statistics, the following book reference is excellent: Zuur et al. (2010) A Beginner’s Guide to R. Springer: New York. Available online via SpringerLink – worth checking to see if your university has access.