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Troubleshooting 16S PCR from A/C filters (or other annoying substrates)

DNAExtraction copyOne of our goals at microBEnet is to increase communication and collaboration between people working in the disparate fields that make up the “microbiology of the built environment”.   Most of what we’ve posted about in the past relates to general information and events… but we’d also like to host more technical information that would help people doing the work on the ground.

Yesterday our lab was having some troubles with DNA extractions from A/C filters and I e-mailed some questions to folks at the BioBE Center and got back this extremely helpful response from Adam Altrichter (@atrickster).


Hey David — sounds like you’re having trouble with inhibitors! The good news is that if you’re extracting a dirty supernatant there’s a very good chance you should have DNA in that sample. Bad news is that the dirty-ness could be preventing you from getting any good, amplifiable DNA out of the sample. In general the power soil kits are pretty good at cleaning up samples, but not perfect.

For PCR troubleshooting, here’s what I usually do when I don’t get bands that I expect to see:
  1. Make sure you’re using a positive control (something you know will amplify) in your PCR so that you know the PCR should have actually worked (i.e. you didn’t forget a reagent or something).
  2. Try a dilution series on your DNA template (dilute in tris or water). 1:10, 1:100, 1:1000, even 1:10000 are not uncommon for soil samples.
  3. If you really suspect it’s inhibitor issues, which it sounds like it is, try spiking each of your samples with the positive control — if it doesn’t amplify it means something intrinsic to your sample is hindering amplification and not just low biomass. Try this set up:
  • Tube 1: sample DNA
  • Tube 2: Positive control DNA
  • Tube 3: Sample DNA + Positive control DNA
  • Tube 4: negative control

4. James mentioned BSA (bovine serum albumin) — try adding BSA (0.1-0.8ug/uL) which can help in samples with high impurities.
5. Try running an annealing temperature gradient, starting lower than you might expect.
6. Run more cycles…anywhere 25-40 cycles.
7. Do an additional phenol:chloroform extract of the template to remove impurities.
8. You can try to make all sorts of tweaks to your PCR — changing MgCl2 levels, adding more/less template or Taq, adding DMSO or betaine in combination with BSA has been shown to help.

Keep trying! There’s definitely some voodoo involved in this process. If you get desperate you might consider naming your thermocyclers and praying to the gods of PCR for mercy…
Now, about the extraction issues:
I might recommend doing the sample extraction slightly differently. This will add some time but should be a decent work around the absorbent filter issue:
– Place your filter in a 5 or 15 mL centrifuge tube — add 4-5 mL of sterile water or PBS
– Vortex for 5/10 min, spin it down quickly
– Then you can work with that liquid to extract instead of trying to extract straight from the bit of filter. If the solution is really dirty you should have plenty of DNA to work with. If it’s not, then you can try to concentrate it prior to adding to your extraction kit (you could try the Amicon ultra filter set up).
Alternatively, you could add more buffer to your bead beating tube (use a bigger bead tube, like the ones that come with the PowerWater Mobio kits) to prevent the filter from sucking it all up. However, I think just having the chunk of filter in the tube could be preventing efficient lysis of cells during the bead beating step — especially if you’re doing this step in the little 2mL tubes from the power soil kits. Make sure the ‘dirty’ side of the filter is facing inward in the bead tube too if you’re rolling it up to fit in the tube. You could also try to do a quick freeze-thaw in liquid nitrogen of the filter in the bead tube to break it up a little bit before bead beating (just make sure you reheat the tube at  ~65C for 10 min prior to bead beating to allow precipitants to go back into solution).

David Coil

David Coil is a Project Scientist in the lab of Jonathan Eisen at UC Davis. David works at the intersection between research, education, and outreach in the areas of the microbiology of the built environment, microbial ecology, and bacterial genomics. Twitter

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